Plant Techniques: Theory and Practice [1 ed.] 1032805137, 9781032805139

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Plant Techniques: Theory and Practice

Plant Techniques: Theory and Practice

S.M. Khasim Department of Botany and Microbiology Acharya Nagarjuna University, Guntur, India

K. Thammasiri Xishuangbanna Tropical Botanical Garden Chinese Academy of Sciences (P.R. China) S. Rama Rao Department of Life Science Sharda University, Greater Noida, Delhi M. Rahamtulla Department of Botany and Microbiology Acharya Nagarjuna University, Guntur, India

Capital Publishing Company NEW DELHI

KOLKATA

First published 2025 by CRC Press 4 Park Square, Milton Park, Abingdon, Oxon, OX14 4RN and by CRC Press 2385 NW Executive Center Drive, Suite 320, Boca Raton FL 33431 © 2025 Capital Publishing Company CRC Press is an imprint of Informa UK Limited The right of S.M. Khasim, K. Thammasiri, S. Rama Rao and M. Rahamtulla to be identified as author(s) of this work has been asserted in accordance with sections 77 and 78 of the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this book may be reprinted or reproduced or utilised in any form or by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying and recording, or in any information storage or retrieval system, without permission in writing from the publishers. For permission to photocopy or use material electronically from this work, access www.copyright. com or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. For works that are not available on CCC please contact [email protected] Trademark notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library Print edition not for sale in South Asia (India, Sri Lanka, Nepal, Bangladesh, Pakistan or Bhutan). ISBN13: 9781032805139 (hbk) ISBN13: 9781032822556 (pbk) ISBN13: 9781003503682 (ebk) DOI: 10.4324/9781003503682 Typeset in Times New Roman by Innovative Processors, New Delhi

Dedicated and presented to Our illustrious teacher Professor J.J. Shah, FNA (On the occasion of his 100th birthday celebrations on September 10, 2023)

Sardar Patel University Vallabh Vidyanagar, Gujarat, India

Preface The laboratory manual on Plant Techniques: Theory and Practice is the outcome of several decades of research and teaching in plant biology to undergraduate and postgraduate students of Plant Science, Horticulture, Microbiology, and Biotechnology. We the team of plant scientists always feel that there should be a comprehensive book that deals with principles and selected procedures together, so that the beginner, as well as the expert technician, could understand the basic concepts of Plant Science including botanical microtechnique and microtomy, staining techniques, molecular techniques, plant tissue culture, electron microscopy, and cryopreservation and germplasm storage, etc.; and it is very easy for them to pick-up and choose suitable procedure for studying plant material. To inculcate the basic knowledge among graduate and research students, the broad spectrum of plant science has been dealt with in this treatise. We strongly believe that this manual is incomplete and additional chapters may be incorporated in due course. We would appreciate hearing from fellow plant scientists and technicians for suggestions and modifications. One of the authors of the book, Professor S.M. Khasim is especially indebted to his illustrious teacher, Professor J.J. Shah, FNA (Sardar Patel University, India) to whom this book is dedicated with great respect. Prof. Khasim expresses his profound gratitude to his research supervisor, and mentor, Professor P.R. Mohana Rao (UGC: Emeritus Professor, Acharya Nagarjuna University; Sydney Sussex Fellow, UK), who is the source of inspiration throughout his academic career, for permitting to access his rich botanical library. Prof. Khasim is highly grateful to the Honorable Vice-chancellor of Acharya Nagarjuna University, Prof. Rajasekhar Patteti, for allowing him to work as an Honorary Professor in the Department of Botany and Microbiology. He further expresses his gratitude to his colleagues, Prof. Vishnu Vardhan, Prof. M. Vijaya Lakshmi, Prof. K. Ammani, Prof. G. Rosaiah, Dr. M. Raghuram (late), Prof. V. Uma Maheswara Rao, Prof. K. Mallikarjuna, Dr. A. Amrutha Valli (presently heading the Department of Botany and Microbiology), Dr. J. Madhavi, Dr. P.S. Raju, and Dr. Y.R.K.V. Tirupati Rao for their moral support and encouragement in his academic pursuit. Further, it is our pleasure to acknowledge all help received from all our colleagues and friends at Guntur

viii Preface

(India), Bangkok (Thailand), Xishuangbanna prefecture (P.R. China), and Shillong and Delhi (India). We appreciate the outstanding works of Dr. G. Stefano (Michigan State University, USA), Dr. F. Xu (Beijing Forestry University, Beijing), and Dr. M.S. Otegui (University of Wisconsin, Madison, USA) and we accessed some of their microphotographs and incorporated them in this book. We are especially thankful to doctoral students, Amrita Choudhury, Sohini Deb, and Banridor Kharbyngar from North-Eastern Hill University, Shillong, India for finalizing chapter 13 ‘Molecular Techniques’ under the supervision of Prof. S. Rama Rao. We are thankful to Prof. D. Ramachandran, Department of Chemistry, Acharya Nagarjuna University for drawing the chemical structure on ChemDraw software. Sincere thanks are also due to Mr. K. Omkar Murthy and Mr. K. Srinivasa Rao for word processing this original manuscript and providing the printouts very patiently and effectively. We are very much grateful to Mr. Raj D. Mirchandani and his team (Capital Publishing Company, New Delhi, India) and also to Taylor & Francis Group for their unstinted support and their effective execution of this project. S.M. Khasim K. Thammasiri S. Rama Rao M. Rahamtulla

Contents Preface About the Authors

vii xiii Part A: Theory and Basic Techniques

1. Introduction

3

2. Microscopy

7

2.1 Light Microscopy 2.2 Electron Microscopy 3. Botanical Microtechnique and Microtomy 3.1 Killing and Fixation 3.2 Dehydration and Infiltration 3.3 Embedding and Sectioning 4. Stains (Dyes) 4.1 Natural Dyes 4.2 Coal-Tar Dyes 4.3 Other Substances Acting as Stains 5. Staining Techniques 5.1 5.2 5.3 5.4 5.5

Types of Staining Differentiation Mordants Clearing (Bleaching) Staining Schedules

6. Centrifugation 6.1 Basic Principle of Sedimentation 6.2 Instrumentation

9 18 27 27 39 45 59 62 66 80 83 83 84 84 85 87 99 99 100

x

Contents

6.3 Mechanism of Centrifugation 6.4 Sedimentation Co-efficient 6.5 Classification of Centrifuges and their Uses 7. Colorimetry and Spectrophotometry 7.1 Colorimetry 7.2 Spectrophotometry 8. Common Adhesives and Mounting Media 8.1 Common Adhesives 8.2 Mounting Media 8.3 Substitute for Immersion Oil in Microscopy 9. Plant Collection and Herbarium Techniques 9.1 Plant Collection and Preservation 9.2 Herbarium 10. Biostatistics 10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10

Basic Statistics (Descriptive Statistics) Analysis of Variance (ANOVA; Components of Mean) Null Hypothesis T-test Coefficient of Variation Heritability Genetic Advance Correlation Path Analysis Residual Effect

102 104 104 107 108 115 121 121 122 126 127 127 133 137 138 161 163 163 164 167 167 167 168 169

Part B: Recent Advances in Plant Techniques 11. Histochemical Methods 11.1 11.2 11.3 11.4 11.5 11.6 11.7 11.8

Preparation of Sections for Histochemical Studies Carbohydrates Proteins Detection of Ions in Plant/Animal Tissues Nucleic Acids Lignin, Suberin and Cutin Cytokinins and Auxins Latex and Rubber

175 175 175 185 193 195 199 203 206

Contents xi

12. Electrophoresis 12.1 12.2 12.3 12.4

Modes of Electrophoresis Detection and Quantitative Assay Discontinuous (Disc) Gel Electrophoresis Protocols

13. Molecular Techniques 13.1 13.2 13.3 13.4 13.5 13.6 13.7 13.8 13.9

Genomic DNA Extraction Quantification of DNA Samples Isolated Quality of the DNA Samples Isolated Agarose Gel Electrophoresis Polymerase Chain Reaction Types of PCR Molecular Markers Other Approaches Protocols

14. Plant Tissue Culture 14.1 14.2 14.3 14.4 14.5 14.6 14.7 14.8 14.9 14.10 14.11 14.12 14.13 14.14

History Setup a Tissue Culture Laboratory Media Components and Preparation Explant Preparation Aseptic Technique Pathways of Culture Cells and Tissues Acclimatisation Commercial Production Culture Guide to Selected Plants Meristem Cultures and Production of Disease Free Plants Protoplast Fusion and Somatic Hybridisation Anther and Pollen Culture Micropropagation Experimental Protocols

15. Chromatography 15.1 15.2 15.3 15.4

Paper Chromatography Thin Layer (planar) Chromatography (TLC) Column Chromatography High Pressure Liquid Chromatography (or) High Performance Liquid Chromatography (HPLC) 15.5 Adsorption Chromatography

209 210 213 215 217 225 226 227 227 228 228 230 235 244 249 261 261 261 263 264 267 269 270 270 270 271 276 286 293 299 309 310 311 314 316 319

xii Contents

15.6 Gas Chromatography/Mass Spectrometry (GC/MS) 15.7 Protocols 16. Phytochemical Methods 16.1 16.2 16.3 16.4 16.5 16.6

Basic Equipment for Phytochemical Studies Steps Involved in Plant Collection Choice of Solvent Extraction Methods Preparation and Analysis of Plant Extracts Case Studies

17. Cytological and Vital Staining Techniques 17.1 Cytological Techniques 17.2 Vital Staining 18. Electron Microscopic Studies 18.1 Specimen Preparation for EM Studies 18.2 Electron Tomography in Plant Cell Studies 18.3 Scanning Electron Microscopic (SEM) Studies 19. Autoradiography 19.1 Protocols 19.2 Radiography of Botanical Material by Means of Low-energy X-rays (O’Brien and McCully, 1981) 20. Cryopreservation Development for Germplasm Storage 20.1 Principles of Cryopreservation 20.2 Steps in Plant Cryopreservation 20.3 Methods in Plant Cryopreservation 21. Microbiological Techniques 21.1 21.2 21.3 21.4

Isolation Methods Media Maintenance and Preservation of Cultures Preparation and Staining of Specimens

References Subject Index

319 320 329 329 330 331 333 336 339 341 341 366 373 373 383 385 395 397 401 403 403 411 413 427 427 429 430 431 453 477

About the Authors Professor Dr. S.M. Khasim is currently an Honorary Professor in the Department of Botany and Microbiology at Acharya Nagarjuna University, Guntur (India). Prof. Khasim was formerly the Head of the Department of Botany and Microbiology, as well as the Chairman of the Board of Studies in Biotechnology at the same university. He has been working for the last four decades teaching Plant Techniques, Plant Reproductive Biology, and Biotechnology at the post-graduate level, and he has made significant contributions to Orchid Biology and Conservation. He has collaboration with Mahidol University (Thailand) in the field of orchid biotechnology. The Orchid Society of India (TOSI), Chandigarh, has awarded him the Usha Vij Memorial Award 2018 for his outstanding contribution to orchid biology. He is the Fellow of the Indian Botanical Society (IBS), and member of the Indian Science Congress Association (Kolkata), and the International Society for Horticultural Science (ISHS) in Belgium. Professor Dr. K. Thammasiri is currently Professor at the Department of Gardening and Horticulture, Xishuangbanna Tropical Botanical Garden, Chinese Academy of Sciences in the Xishuangbanna prefecture (P.R. China). Prof. Thammasiri was formerly the Head of the Department of Plant Science, Faculty of Science, Mahidol University (Thailand). For the last three decades, he has been teaching and conducting research on Plant Tissue Culture, Orchidology, Plant Breeding, and Plant Ecology. He has written numerous orchid books, as well as the Plant Cryopreservation textbook. He was the convener for four International Symposia under the auspices of the International Society for Horticultural Science (ISHS), namely TSO 2016, Cryosymp 2018, BGL 2019 and Biotech 2021. Kasetsart University honoured him with a Distinguished Alumni award in research. He is a member of the Science Society of Thailand, the Society for Cryobiology, and the ISHS in Belgium. Professor Dr. S. Rama Rao is currently working as Professor and Head of the Department of Life Sciences at Sharda University in Delhi (India). Prof. Rao previously served as Head of the Department of Biotechnology and Bioinformatics at North Eastern Hill University in Shillong. He has been an active researcher in the field(s) of Plant Biotechnology and Conservation

xiv About the Authors

Biology, as well as a post-graduate teacher of Plant Cytogenetics and Plant Molecular Biology. He has actively collaborated with the National Bureau of Plant Genetic Resources in their exploration activities and has assisted in the collection of significant germplasm of many horticultural plants. For the first time, three new Vigna species and one Abelmoschus species have been collected from the Indian subcontinent. He is a member of the 7th Research Advisory Committee NRC Orchids, Pakyong, Sikkim, and the Fellow of the Association of Biotechnology and Pharmacy (India). Recently, he has been appointed as Ombudsperson of Central University of Andhra Pradesh at Anantapuram, India. Dr. M. Rahamtulla received his Ph.D. in Orchid Biology in 2022 from the Department of Botany and Microbiology at Acharya Nagarjuna University, Guntur (India). For the past seven years, he has been teaching Botany and Biotechnology at the undergraduate and postgraduate levels. He has published a good number of research articles in National and International journals. He is the Life Member of the Orchid Society of India (TOSI), Chandigarh.

Part A: Theory and Basic Techniques

1 Introduction The advancement of Plant Science has been correlated to the available technology. For studying the structure and function of plants and animals, it is necessary to learn and be trained in various techniques. There have been phenomenal achievements over the last 70 years in the field of instrumentation and methodology. In the recent past, there has been tremendous development in the field of molecular biology and biotechnology. In the course of time, new techniques have been coming up and old techniques have been improving in plant biology in general, and molecular biology in particular. Apart from that, various basic aspects such as fixation of the plant material, fixative mixtures, embedding of the plant material, embedding media, sectioning, and staining procedures have made it possible to study the structure and function of the plants. The invention of several pre-treatment chemicals for fixation has considerably aided the study of chromosome structure. The principle of pre-treatment is mostly to bring about physical changes in the cytoplasm and nucleus which would help in studying the details of chromosome morphology (Sharma and Sharma, 1972). In plant techniques, enormous new knowledge and techniques are being generated. This information is scattered mostly in research journals and some standard textbooks. Hence, the authors feel that there should be a comprehensive book needed containing an up-to-date account of plant techniques, their principles, and their biological significance. For the last four decades, authors have been working on angiosperms’ structure, function, conservation, and reproductive biology. Obviously, the book is confined to dealing with some techniques related to these aspects. Although it appears to be a broad spectrum of Botany, we tried to highlight some of the fundamental techniques to study plant structure and function, and other aspects of plant biology. In fact, this manual reviewed the above aspects of plant biology by considering the past literature. Every chapter has been written with a theory and principle of a particular aspect. The book is divided into two parts: Part A deals with the theory and principles of plant biology and some general techniques that have been carried out for undergraduate students; Part B with special techniques being employed to study the plant structure, development,

4

Plant Techniques: Theory and Practice

function, and conservation of germplasm. These chapters are briefly mentioned below: In Part A: Theory and Basic Techniques, Chapter 2 deals with Microscopy in which various types of light microscopes and electron microscopes and their working principles have been explained. In chapter 3, Botanical Microtechnique and Microtomy, various aspects related to this have been elaborately written; killing and fixative agents (coagulants and noncoagulants), dehydration and infiltration procedures, embedding and sectioning of plant materials are vividly explained. Chapter 4 is on Stains (Dyes), in which the chemistry of dyes and their mode of action on plant tissues have been discussed; various anionic and cationic dyes, and mordants being used to enhance staining are elaborately discussed in this chapter; some selected procedures are uniformly formatted, and biochemical explanation of staining reaction has been briefly explained. Chapter 5 is on Staining Techniques, in which different types of staining and some staining schedules are given; basic techniques such as Safranin (or Dalafield’s) hematoxylin technique (Johansen, 1940) and Tannic acid-ferric chloride-safranin triple staining, that is relevant to the budding scientists, have been given; an overview is provided related to staining techniques since the time of the Johansen (1940). Basic principles are explained with respect to Centrifugation and Colorimetry and Spectrophotometry in chapters 6 and 7 respectively. Chapter 8 gives a detailed account of Common Adhesives and Mounting Media. It is followed by the Plant Collection and Herbarium Techniques of chapter 9; preservation and pressing of the plant material and a list of world-class herbaria are provided in this chapter. Chapter 10 deals with Biostatistics; in which various statistical tools such as descriptive statistics, analysis of variance (ANOVA), T-test and coefficient of variation are given. In Part B: Recent Advances in Plant Techniques, experimental protocols, their biological significance, and results are elaborately given. As a part of this, chapter 11 deals with Histochemical Methods, in which detailed protocols have been provided to identify the various biological substances such as carbohydrates including callose, proteins, growth regulators (cytokinins, auxins), and detection of ions; besides that, histochemical identification of DNA and RNA has been done by Azure B method and Toluidine blue ‘O’ method respectively. The working principle of Electrophoresis has been given in chapter 12; the protocol for SDS-PAGE is elaborately provided here. Then, the latest account of Molecular Techniques has been provided in chapter 13; in this chapter, recent advances in molecular techniques such as FISH, blotting methods and flow cytometry, chromosome microdissection, and microcloning are precisely documented. Chapter 14 is on Plant Tissue Culture, in which very clear protocols on meristem cultures, protoplast isolation and somatic hybridisation, anther culture, and micropropagation of an epiphytic orchid, Dendrobium aphyllum (Rahamtulla and Khasim, 2022), Grammatophyllum speciosum (Sopalan et al., 2010) and Vanda coerulea (Wasiksiri et al., 2010)

Introduction 5

have been included. The basic principle of Chromatography is given in chapter 15; a brief account of HPLC and the latest protocol on GC-MS screening of solvent extract of Cymbidium aloifolium has been added (Venkatesh and Khasim, 2020). In chapter 16, Phytochemical Methods, extraction of alkaloids and flavonoids from orchid species is provided. Chapter 17 is on Cytological and Vital Staining Techniques; in addition to basic techniques, the recent protocol for staining plant cell vacuoles (Stefano, 2017) has been incorporated in this chapter. Chapter 18 deals with Electron Microscopic Studies, in which specimen preparation for EM and SEM studies is included; further a detailed protocol for orchid seed study with SEM (Ramudu et al., 2020) is incorporated. A brief account and general protocol of Autoradiography is given in Chapter 19. A succinct account of Cryopreservation Development for Germplasm Storage is provided in chapter 20; it also highlights the principles of cryopreservation and protocols for V-cryoplate and D-cryoplate methods. Finally, Microbiological Techniques have been provided in chapter 21; further, isolation methods of microbial strains and maintenance and preservation of microbial cultures are incorporated. This book covers the broad spectrum of plant techniques, in which basic principles and biological significance of various plant structures and natural products are dealt with; so that the beginner, as well as an expert plant technician, could understand the subject, and it is very easy for them to pick-up and choose suitable procedure for studying the plant material. All the experimental protocols given here have been uniformly formatted and the principle involved in the technique is briefly explained. However, the elaborate discussion has been omitted with brevity since it is outside this book’s scope. We hope both beginners and plant researchers could enjoy reading this book as much as we have enjoyed writing it in a tireless manner.

2 Microscopy Although the use of lenses for magnification has been known for centuries, modern microscopy began when Holland’s eyeglass maker, Z. Jannsen, lined up two lenses to multiply their magnifications effectively. Subsequently, Robert Hooke in 1665 and Leeuwenhoek in 1674 published the initial microscopic examinations of biological tissues (quoted in Prescot et al., 1999). A Microscope can be compared with a human eye since both have lens systems and, in both cases, images of the objects are formed. The most important principle involved in microscopy is to get a magnified image, in which structures may be resolved that otherwise cannot be resolved with the unaided eye. Resolving power: The ability of the lens system to distinguish two adjacent points as distinct and separate, in other words, the ability of the microscope to produce separate images of closely placed objects known as resolving power. This depends upon the wavelength of light and numerical aperture (NA). The Swiss physicist E. Abbe developed the theory of microscope resolution and expressed the Resolution Limit (RL) in the following formula: RL = 0.61 λ NA where λ = Wavelength of light [400-750 nm for visible light used in the compound microscope; blue light (400 nm) is better for resolution than redlight (700 nm)] NA = n sin θ (Numerical aperture) n = Refractive index of the medium present between the specimen and lens. Generally, immersion oil (n = 1.5) is better than air (n = 1.0) (Table 2.1) θ = Half of the angular width of the cone by the objective lens from the typical point of the specimen (Fig. 2.1) So, the numerical aperture is nothing but the light-collecting ability of the lens system. The higher the numerical aperture, the higher will be the degree of resolution, whereas, the lower the wavelength of the light, the higher will be the resolution.

8

Plant Techniques: Theory and Practice Table 2.1. Refractive indices of some mounting media Medium

Refractive index (n)

Distilled water Glycerol Cedarwood (immersion oil) Eucalyptus oil Carbon tetrachloride Olive oil *Euparal Sandalwood oil *Canada balsam *Polystyrene

1.33 1.47 1.51 1.46 1.46 1.47 1.48 1.51 1.54 1.59

* Permanent mounting media

Fig. 2.1. Determination of the angle.

Lowering RL can be achieved in three ways: (1) lowering the wavelength (A), (2) raising the refractive index, and (3) raising sin θ as lenses of shorter focal length are used, the distance between the object and the lens decreases (Table 2.2). *Table 2.2. Optical Properties of Objective Lenses Magnification

Focal length (mm)

Lens-specimen Distance (mm)

NA

10

16

5.5

0.2

40

4

0.6

0.65

95

2

0.1

1.32

* Adopted from Alberts et al. (1994)

If λ = 400 nm, n = l.5, sin θ = 0.99, and an optical lens of maximal NA, it gives a theoretical maximum resolution of the objective lens of about 0.2 µm which is 1000 times greater than that of the unaided human eye.

Microscopy

9

A wide variety of microscopes are currently available for studying plant structure, these are mainly categorised into two: (i) Light Microscope, and (ii) Electron Microscope. In the former one, magnification is achieved with the help of an optical lens system. Bright field, dark field, phase contrast, polarisation, and fluorescence microscopes come under this category. Whereas, in the latter category a beam of electrons is used in place of light to obtain higher magnification; transmission and scanning electron microscopes belong to this category. The resolution of various biological components is given below (adopted from Lewis and John, 1963): Structure

Scale Means of Resolution

Atoms, molecules 1A0

Macro Genes, Bacteria, Chromomolecules viruses cytoplasmic somes, cells organelles

10A0 100A0 1000A0 Electron microscope

1µ

Tissues

10µ 100µ Light Microscope

X-ray diffraction

2.1 Light Microscopy 2.1.1

The Stereo Microscope (Dissecting Microscope)

The stereo microscope, also known as a dissecting microscope, is an optical microscope, designed for low-magnification observation of a sample, typically using light reflected from the surface of an object rather than transmitted through it. It has two optical paths at slightly different angles, allowing the image to be viewed three-dimensionally. Stereo microscopes are the majority at low power, typically, between 10x and 200x, mostly below 100x. Stereo microscopes can be used to visualise surface features of objects, microsurgery, watchmaking, etc. It further allows students to observe photosynthesis in action as well as forensic engineering. It is also an essential tool in entomology. The stereo microscope should not be confused with a compound microscope equipped with double eyepieces and a binoviewer. In compound microscope, both eyes see the same image, with two eyepieces serving to provide greater viewing comfort. The image in such a microscope is no different from that obtained with a single monocular eyepiece. Stereo microscope has three key parts, these are: body/viewing head, focus block and stand. ‘Body/viewing head’ houses the optical parts in the upper position of the microscope; the ‘Focus block’ attracts the head of the microscope to the stand and focuses it; ‘Stand’ supports the microscope and consists of integrated illumination. Just like other microscopes, it has eyepiece lenses and objective lenses (Fig. 2.2).

10

Plant Techniques: Theory and Practice

Fig. 2.2. The Stereo Microscope (Dissecting Microscope).

Stereo Microscopes are of three basic types: (1) Stereo Fixed Microscope (2) Stereo Turset Microscope, and (3) Stereo Zoom Microscope. A stereo fixed microscope has fixed magnification using two objective lenses. The magnification has a fixed degree and is limited by the lens capability. To increase the magnification, changing the eyepiece is needed. Stereo tunet microscope comes with various mountings and one of which is the turset style. This type of mounting indicates an additional objective lens that you can rotate to your viewing position. The viewer can easily change the magnification by rotating the mounting of a turret. This type of microscope is cost-effective and more affordable than any other stereo microscope.

2.1.2

Compound microscope

2.1.2.1 Bright-Field Microscope

Bright-field microscope is indispensable to biologists. It provides magnification of the objects which are not visible to the naked eye. In this commonly used compound microscope, a dark image of the object appears against a bright background. This is accomplished by the condenser, a series of lenses, and diaphragms below the stage, on which the object is located.

Lens Systems A Compound microscope consists of three lens systems: (1) objective, (2) eyepiece or ocular, and (3) condenser (Figs 2.3, 2.4). Objective: Objective is the most important component of the light microscope since it affects the quality of image formation. Based on the degree of correction for chromatic aberration, objectives are of two types: (i) achromatic objectives that are corrected for two regions of the spectrum, and (ii) apochromatic objectives that are corrected for three regions of the spectrum (for details see Desai and Desai, 1980; O’Brien and McCully, 1981).

Microscopy

11

Fig. 2.3. The Compound Microscope.

Fig. 2.4. The path of light through Bright-field Microscope.

Light focused on the specimen is either differentially transmitted (absorbed) or, for the background, fully transmitted to the objective. This lens resolves the specimen to produce a magnified image and focus on the microscope tube. Generally, compound microscopes are fitted with three objectives with different magnifying powers. They are low power, high power, and oil immersion objectives. These are easily recognised by their length and NA value engraved on the barrel. Low power objective is the shortest and oil immersion is the longest. In a bright-field microscope, both direct (rays not hitting the specimen) and indirect rays (rays diffracted or scattered from the specimen) enter the objective, hence the dark image is seen against a bright background. Eyepiece (ocular): The main purpose of the ocular is to magnify the intermediate image and to correct certain aberrations produced by the objective. The ocular is composed of mainly two lenses; the upper component or eye lens is the magnifier, whereas the lower component is known as the field lens. Condenser: The quality and method of use of the condenser are important factors for good microscopy. The primary function of the condenser is to supply a sufficient cone of light to the objective to gain maximum resolution.

12

Plant Techniques: Theory and Practice

So, it should be properly positioned during the microscopy. The Abbe condenser, aplanatic condenser and achromatic condensers are available for general purposes. Condensers are also fitted with an iris diaphragm and filter holder to control the light intensity. All four optical components (lamp, condenser, objectives and eyepiece) are installed in the common axis in the Microscope. The following steps should be carefully taken for the effective use of a research microscope (see also Desai and Desai, 1980; O’Brien and Mccully, 1981): (i) Turn on the lamp; adjust it to a considerable intensity. (ii) Adjust the mirror in such a way that the light travels up the body tube of the microscope. (iii) Take out the eyepiece and examine the back focal plane of the objective. Close the condenser iris until about 3/4 of the field of vision is left clear. In some binocular microscopes, it is difficult to see the image of the condenser iris in the back focal plane of the objective. In this case, close the iris, then open it slowly and watch the ‘Circle of light’ as it increases. It comes to a certain position after which further increase in iris diameter does not increase the ‘Circle of light’. Close the iris a little from that position. (iv) Place the specimen on the stage. (v) Now keep the low power objective in position. Lower the body tube with the help of a coarse adjustment knob until the distance between the specimen and low power objective becomes just about 5-7 mm. (vi) Bring the specimen in focus with the help of a coarse adjustment knob by looking through the eyepiece. (vii) Sharpen the image with a fine adjustment knob. (viii) When higher magnification is needed, change the objective by shifting the nosepiece in the appropriate direction till it clicks into place. Care should be taken not to touch the objective lens without fingers. (ix) Focus the specimen again with the help of coarse and final adjustments. Adjustment of the iris is necessary again since different power objectives have different field areas. (x) To set up the oil-immersion objective, especially 90x or 100x, first rack the body tube up and place a drop of immersion oil (free of bubbles). (xi) By observing from the sides of the instrument, rack the objective down to touch the oil almost near the specimen. Care should be taken to see that the objective does not touch the specimen. (xii) By looking through the eyepiece, slowly focus upwards with a fine adjustment until a clear image appears. It comes quickly since the working distance of the oil-immersion objective is about 0.1-0.15 mm. (xiii) Remove and clean the oil from both objective lenses and slide with lens paper immediately after the use of the instrument.

Microscopy

13

2.1.2.2 Dark-Field Microscope

In a dark-field microscope, a bright image of the specimen is seen against the dark background. A specially devised condenser with a star diaphragm is used in this microscope. It blocks the light rays (direct rays) that would normally enter the objective lens. Instead, it supplies a hollow cone of light and illuminates the specimen. Only indirect rays scattered from the specimen would enter the objective. As a result, a bright image against the dark background is seen (Fig 2.5). In this microscope, special condensers of very high NA, such as cardoid and paraboloid, are employed with an oil immersion objective.

Fig. 2.5. Dark-field Microscope.

Since the contrast is quite vivid (just like stars against a dark sky), organelles such as mitochondria and lysosomes are easily detected by this microscope. It is also useful for the observation of plant structures and microbes in an unstained condition. It has diagnostic significance, especially in the case of syphilis (Desai and Desai, 1980). 2.1.2.3 Phase-Contrast Microscope

The phase-contrast microscope is based on phase-contrast principles, propounded by Fritz Zernike. He got the Nobel Prize in Physics in 1953 for his phase-contrast principles. According to him, light waves have a variable character for frequency and amplitude. The human eye cannot notice a phenomenon when two light rays have similar amplitude and frequency, but different phases (Fig. 2.6). This can be achieved by a phase contrast microscope (Fig. 2.7). When the light rays are passing through the vacuum, they travel at high speed. When they pass through the transparent cells, light rays become slow due to changes in velocity. When a beam of light changes speed, it is refracted at an angle depending on the magnitude of the velocity change. So, the light rays are refracted and alter their phase. This phase change may not be noticed by the human eye. In the phase-contrast microscope, both refracted rays (indirect rays) and direct rays are undergoing interference. As a result, a clear image of the specimen is seen.

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Plant Techniques: Theory and Practice

Depending on the type of phase-shifting element employed, the specimen appears darker against a light background (positive contrast) or lighter against a dark background (negative contrast) (see also O’Brien and McCully, 1981). The phase-contrast microscope is widely used to observe living cells and organelles.

Fig. 2.6. Properties of Light (a) Both rays are in phase (rays go up at the same time and down at the same time), (b) Constructive interference in which amplitude becomes doubled (2A°), brightness increased, (c) Rays are in & out of phase, and (d) Destructive interference, rays that are out of phase partially cancelled, brightness decreased and finally becomes dark.

Fig. 2.7. Phase-Contrast Microscope.

2.1.2.4 Polarisation Microscope

A polarisation microscope is used to visualise birefringents. Various biological structures like microtubules, microfibrils and crystals show birefringence properties because they refract polarised light in different ways depending on the direction from which the light strikes the specimen. Anisotropic materials are birefringents because they show double refractions. The principle of the polarising microscope is to deduce and measure the structural anisotropy of the specimen from the optical anisotropy. That it displays (see also Bennett, 1961; Bartels, 1966; James, 1976; O’Brien and McCully, 1981).

Microscopy

15

When a beam of polarised light passes through the birefringent object, the ray splits into two rays polarised in mutually perpendicular lines; the one which follows the ordinary laws of refraction is known as the ‘ordinary’ ray and the other, whose velocity through the object is different, is known as the ‘extraordinary’ ray. The difference in refractive indices (ne-no) is the value of birefringence (B). The two polarised rays, after emerging from the object, recombine but, because of different velocities through the object, one shows retardation when compared to the other. The value of ‘retardation’ (T) which is based on the birefringent property, is measured in the following manner: T = B×t Where T = Value of retardation B = birefringence, i.e., (ne-no) and t = thickness of the object Finally, T = (ne - no) t; it is expressed in terms of wavelength in Angstrom units (Sharma and Sharma, 1972). The polarisation microscope is fitted with two additional optical lenses: (1) a vertical polariser placed between the light source and substage condenser to produce polarised light, and (2) an analyser horizontally placed above the objective lens (Fig. 2.8). In this, a bright image of specimen against dark background appears. The polarising microscope is also useful for studying the optical properties of cell walls and starch grains.

Fig. 2.8. Polarisation Microscope.

2.1.2.5 Fluorescence Microscope

A fluorescence microscope is based on the principle ‘Fluorescence’. In this microscope, dyes are used that absorb light energy of one wavelength and emit light energy of longer wavelength; for example, the dye fluoresces in absorbs at 490 nm and emits at 520 nm appearing yellow-green and rhodamine has a characteristic absorption and emission spectrum (see also Pringsheim, 1963; Udenfriend, 1964, 1969). A fluorescence microscope is fitted with two filters, these are (1) a primary or exciter filter placed between the light source and specimen, which allows only the excitation wavelength, and (2) a secondary or barrier filter

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Plant Techniques: Theory and Practice

placed between the specimen and viewer, and it ideally transmits only the wavelengths emitted by the fluorescing compounds (Fig. 2.9). Fluorescence in histological specimens is induced either in compounds inherent in the tissue known as autofluorescence or in compounds (fluorochromes) that have been added to the tissue. Freehand sections, peels, and whole mounts are enough to study the autofluorescence of lignins and chlorophylls present in higher plants. Plant cuticles (van Gijzel, 1975) and suberised walls of cork (Mader, 1954) are also autofluorescent. The compounds of biological origin like berberine sulphate (from Berberis) and Primulin (Primula) and other compounds are still widely used as fluorochromes. Peirson and Dumbroff (1969) used an extract from Chelidonium roots to show effectively Casparian strips in roots. The other fluorochromes are acridine orange for DNA (Bertalanffy, 1963; Pearse, 1972; Kasten, 1967), ethidium bromide for DNA and lignified walls, and aniline blue for callose of sieve elements (Currier and Strugger, 1956; Eschrich and Currier, 1964; Smith and McCully, 1978; O’Brien and McCully, 1981).

Fig. 2.9. Fluorescence Microscope.

The major use of fluorescence microscopes in microbiology is in immunofluorescence studies. The antibody is made fluorescent by conjugating it with a fluorescent dye. With the help of a fluorescence microscope, it is possible to detect specific types of antigens using an antibody tagged with a fluorescent dye. 2.1.2.6 Confocal Microscope

A confocal microscope is often called a Confocal laser scanning microscope (CLSM) or Laser confocal scanning microscope (LCSM). Like fluorescence microscopes, fluorescence optics have been used in the confocal microscope. Instead of illuminating the whole sample at once, laser light is focused on a specific spot (defined spot) at a specific depth within the sample. This leads to the emission of fluorescent light at exactly this point. A pinhole inside the optical pathway cuts off signals, that are out of focus allowing only the fluorescence signals from the illuminated spot to enter the light detector (Fig. 2.10). The term ‘confocal’ denotes the presence of a diaphragm in the conjugated focal plane (confocal). This diaphragm is usually absent in multiphoton microscopes.

Microscopy

17

By scanning the specimen in a raster pattern, images of one single optical plane are created. 3D images can be visualised by scanning several optical planes and stacking them using a suitable microscope deconvolution software (Z-stack). It is also possible to analyse multicolour immunofluorescence staining using state-of-the-art confocal microscopes which contain lasers and emission/excitation filters.

Fig. 2.10. Confocal Microscope.

Some of the important features of various light microscopes are given in Table 2.3. Table 2.3. Characteristic differences between electron and light microscopes Electron microscope

Light microscope

1

Best resolution

0.0005 µm (0.5 nm)

0.02 µm

2

High vacuum

Necessary

Not necessary

3

Specimen

Dead; Ultrathin sections

Living and non-living material. Sections of 2-10 µm thickness

4

Radiation source

Electron beam

Visible light

5

Lenses

Electromagnetic lenses

Glass lenses

6

Source of contrast

Scattering of electrons

Differential light absorption

7

Staining

Heavy metals

Dyes

8

Focusing mechanism

Adjacent current to the magnetic lens

Adjacent lens position Mechanically

9

Specimen mount

Metal grid (copper)

Glass slide

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2.2 Electron Microscopy In an electron microscope, instead of light, a high-speed electron beam is used (Table 2.4) visualisation of minute structures like ribosomes, mitochondria and membranes are possible with this microscope. In 1924, F. de Broglie (quoted in Prescot et al., 1999) opined that electrons could behave as waves with the wavelength inversely proportional to the square root of the voltage used to generate the electrons: λ = 12.3/[(√V) (0.1)] nm

2.2.1

Transmission Electron Microscope (TEM)

In TEM, the commonly used voltage is 100,000, so the wavelength of the electrons is 0.004 nm. Aberrations inherent in the lenses of the instrument require its operation at very low numerical apertures (0.001). So, the resolution limit, according to Abbe’s formula, is approximately 0.2 nm. Compared to the unaided human eye (200 µmor 200,000 nm), this represents the magnification upto 1,00,000 X (see also Alberts et al., 1994). Although the source of illumination is an electron beam and not light, in general, the TEM is quite similar to the bright-field light microscope (Fig. 2.11). There are some differences between these two (Table 2.4). In TEM, high-speed electron beam generated from an electron gun is used. Electrons are focused by electromagnetic lenses since air molecules interfere with the movement of electrons, high vacuum is created within the microscope. Also due to the very poor penetrating capacity of electrons, sections should be ultrathin. These conditions do not allow the observation of living material through an electron microscope. There are three electromagnetic lenses in the electron microscope. The condenser lens, placed between the source of illumination and the specimen, collimates the electron beam on the specimen and an enlarged image is produced by two other lenses similar to the objective and ocular of a light microscope. Since the electrons are not visible to the human eye, the final image is projected on the fluorescent screen. Since most of the constituent elements in biological matter are of low mass, the contrast of these materials is very poor. This can be enhanced by staining with salts of heavy metals like uranyl acetate, lead citrate, etc. These metals may be either fixed on the specimen (positive staining) or used to increase the opacity of the surrounding area (negative staining). The latter is useful for the observation of particles and bacteria.

2.2.2

Scanning Electron Microscope (SEM)

The Scanning electron microscope (SEM) is mainly used to study the surface features of seeds, plant cells, tissues and microorganisms. The specimen

Microscopy

19

Table 2.4. Characteristic features of some important microscopes Type of microscope

Image formation

Magnification

Applications

Bright-field microscope

Dark image of the object against a bright background

1000-2000x

To study anatomy, embryology of plants and gross morphology of yeast, moulds, algae, protozoa, bacteria etc.

Dark-field microscope

Bright image of the object against a dark background

-do-

To study the gross morphology of the living specimen

Phase contrast microscope

Based on phase contrast principles; Darker against the light background (positive contrast or Lighter against the dark background (negative contrast)

-do-

To study the gross morphology of living specimens and cell organelles

Polarisation microscope

Bright image of the object against a dark background

-do-

To study the birefringents, viz., microtubules, microfibrils, crystals and to study the optical properties of cell walls.

Fluorescent microscope

Based on the fluorescent principle

-do-

To study the plant cuticles, suberised walls, lignins, chlorophylls, DNA, callose of sieve elements; immunofluorescence studies in microbiology.

Electron microscope

Viewed on a fluorescent screen

Up to 10,00,000x

To study the ultrastructure of cell organelles, microbial cells, viruses etc.

is surface coated with a thin layer of heavy metal and is then scanned by a narrow electron beam. The intensity of various radiations released from a portion of the specimen depends on the shape and chemical composition of the irradiated object. The image is obtained by scanning the specimen surface with a narrow electron beam, collecting and amplifying the generated signals and feeding them to the cathode ray tube for display. The final image can be

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viewed from the fluorescent screen. So, the topology of the object can be studied with SEM.

Fig. 2.11. Transmission Electron Microscope.

SEM consists of two main parts, viz., the probing system and the display system. In the probing system, the electron beam is generated by an electron gun (or electron probe) which consists of a V-shaped hairpin tungsten filament. When the filament is heated, electrons are emitted and accelerated through the anode. The diameter of the initially focused spot of electrons, called the spot size, is 50 µm. To increase resolution at higher magnifications, the electron source is demagnified (i.e., the spot size is decreased) to 2 mm by using electromagnetic condenser lenses. In the display system, when the electron beam strikes the specimen, many signals are generated, such as secondary electrons, backscattered electrons, and Auger electrons. These different signals provide different concentrations about the specimens; secondary electrons provide information on surface morphology, whereas X-rays are useful for elemental analysis and backscattered electrons for surface topology. Separate detectors have been used to collect each signal. 2.2.2.1 Surface Morphology

Secondary electrons are emitted by the sample and attracted by a grid (at positive potential) positioned in front of the detector. The detector collects all the electrons coming towards it. These electrons are accelerated towards

Microscopy

21

a scintillator by applying a strong positive voltage (10-12.5 Kv). When electrons hit the scintillator, they generate photons, which are guided by the light pipe to the photomultiplier where photoelectrons are amplified and fed to the cathode ray tube (display screen). The probe scans the specimen in a roster pattern synchronous to the electron beam of the cathode ray tube. A point-topoint correspondence between the specimen and the displayed image is thus achieved. Variations in secondary electron yield are then used to modulate the intensity of the electron beam in the cathode ray tube thus forming an image. The probe scans line-to-line on the specimen surface. The number of lines producing the image can be varied from 100-2000, thus giving different resolving power. To obtain a good picture, a slow scan rate is important so that the probe can stay at one spot for a longer time, thus increasing the number of secondary electrons. For viewing, a fast TV scan rate can be used. As the angle between the probe and the specimen surface varies, the signal strength and contrast of the image are engaged. The detector is placed on one side of the specimen; this enhances the specimen area and prevents signals from another area from reaching the detector.

Specimen Preparation for SEM The surface of an object to be studied by SEM must have the following characteristics: • It must be free from foreign particles or cell debris. • It must be stable when put under a high vacuum. • It should remain stable after exposure to the electron beam. It should develop as few surface charges as possible. • It must emit a sufficient number of secondary electrons. The biological specimen needs special attention because most of them are soft, full of extracellular fluids and have poor conductors. There are several preparative techniques depending on the nature of the tissue and the type of studies to be carried out. The preparation of specimens for SEM studies involves the following steps. Specimen Selection: Specimen size depends on (a) the stage capacity (x-y movement) of the SEM, (b) the type of fixation and, (c) the drying process used. It should be the smallest in size possible, to reduce the preparation artifacts. Tissue culture, single cells, or microorganisms can be processed in suspension on attached to a suitable base (coverslips) before going through preparative procedures. As extracellular products such as mucus, blood, other body fluids and tissue fragments may obscure the surface to be examined, they must be removed by gentle washing with cold isotonic buffer. Microorganisms and cell suspensions can be separated from their liquid environment by centrifugation.

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Plant Techniques: Theory and Practice

Sterilisation of specimen Apart from hard objects such as teeth, bone and wood, all other biological materials must be stabilised to prevent it from undergoing structural changes, which is done by chemical fixation or physical fixation (cryo-fixation). The fixative and the fixation protocol are the same as in the case of TEM specimen preparation. Washing and dehydration To remove the unreacted fixative, the specimen is washed in buffer (which was used for preparing the fixative), especially after secondary fixation with OSO4. The latter may be reduced by dehydrating agents, causing precipitates to form on the specimen thus obscuring the surface morphology. Chemically fixed objects must be dehydrated before drying. The purpose of dehydration is to remove the water before drying. The most commonly dehydrating is ethanol and acetone. To avoid initial osmotic damage to the specimen, starting solvent (acetone or ethanol) should be low (30% or lower). Then specimens are dehydrated using an increasing concentration of the dehydration agent up to the dry absolute acetone stage. Drying After dehydration, specimens are dried and there are different approaches to drying. Air drying Critical point drying Air drying It is the simplest method of drying, through it creates maximum distortion. It causes flattening of the specimen surface because of compressive forces of surface tension of the liquid-gas interface. Critical point drying. This is the commonly used technique to dry biological samples. This sample is transferred from an organic dehydration medium (acetone) to a drying medium (liquid CO2 or freon 13) in a chamber that is cooled and put under pressure. When the dehydrating agent (acetone) has been completely removed and impregnated with the coating can be done in two ways:

Thermal evaporation Many metals, when heated in a vacuum, evaporate readily into a mono-atomic state. The high temperature required to start the evaporation of the materials can be achieved by resistive heating: • The specimen is mounted on a rotating specimen stage to get an even coating. • Thin wire of metal to be evaporated is wrapped around a tungsten filament. • The chamber is evacuated.

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23

• The tungsten filament is heated by supplying voltage. • Metal will start evaporating and a thin layer will deposit on a rotating specimen. Under best conditions, the coating may be relatively uneven, and the changes of surfaces change accumulation or higher, hence most people prefer sputter coating. Sputter coating allows uniform coating on the specimen even on the parts that are not directly facing the metal to be evaporated. Hence, it is the most acceptable technique. Specimens are placed on a cooled base plate which functions as an anode. The cathode, which is the metal (e.g., gold) to be evaporated, is positioned above the specimen. The chamber, in which the cathode and anode are located, is evacuated. An inert gas such as argon is flooded into the chamber and is again evacuated. The negative high voltage is applied to the cathode. This result in glow discharge and argon gas molecules gets ionised. The positively charged ions move toward the cathode and strike it, thereby releasing atoms. The dislodged metal atom due to numerous collisions with the gas molecules gets deflected several times and hit the specimens at different angles. Thus, a uniform thin layer is deposited on the specimens. The thickness of the coating depends upon:

• Current applied • Anode to cathode distance • Sputtering time With Balzer SCD 020 sputter coater, using gold as the target (cathode), a coating of about 35 nm thickness can be obtained under the following conditions: Current Pressure Gas Cathode to anode distance Sputtering time

-

21.5 mA 0.05 mbar Argon 30 nm 1 minute

After the metal coating specimen are ready for observation in SEM. 2.2.2.2 Protocol for the specimen processing for SEM

Standard protocol for biological samples Primary fixation: 2.5% glutaraldehyde is Karnovsky’s fixative X 2-6 hours at 4º C

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Plant Techniques: Theory and Practice

Washing: 0.1 M phosphate buffer (3 changes, each of 15 minutes at 4º C) Postfixation (optional): 1% OsO4 for 2 hours at 4º C Washing: 0.1 M phosphate buffer (3 changes, each of 15 minutes at 4º C) Dehydration: All steps are to be carried out at 4º C 30% acetone 50% acetone 70% acetone 80% acetone 90% acetone 95% acetone 100% acetone (Dry acetone)

-

15 min 15 min 15 min 15 min 15 min 15 min 15 min 15 min × 2

Critical point drying Critical point drying with liquid CO2 at its critical point, i.e., 31.5º C at 1100 p.s.i. Mounting Mounting the samples onto the aluminium stubs. Sputter coating Using silver or gold, coat the samples with 20-30 nm thick film. Preparation of blood cells for SEM

• • • • •

Fix a drop of blood in 2% glutaraldehyde for 30 min Centrifuge for 5 min, at 1500 rpm Decent off the fixative and wash the blood cells with phosphate buffer Centrifuge and decant Resuspend in distilled water and repeat washing in distilled water (5 x times) • Decent and apply a thin film on a clean coverslip after resuspending in distilled water • Air dry • Coat with silver or gold Preparation of sperm sample for SEM

• • • • • • •

Fix 1 or 2 drops of sperm in 2% glutaraldehyde for 10 min Centrifuge for 5 min at 1500 rpm, decant the supernatant Wash the pallet with 0.1 M phosphate buffer and resuspend it Centrifuge and decant the supernatant Wash with distilled water, then centrifuge and decant Resuspend the pallet in distilled water Apply a thin film on a clean cover slip

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25

• Air dry • Sputter coat with silver or gold Image interpretation and magnification Details obtained from ultra-thin sections are hard to interpret because the section thickness acceptable for TEM is about 50 nm. i.e., above 1/10-wavelength of light. 1 mm size tissue can yield 10,000-20,000 ultra-thin sections and it is hard to translate the 2-dimensional image into 3-dimensional reality. Therefore, it is desirable to look at many sections before interpreting the results. So, a sound knowledge of tissue at the LM level is essential.

Magnifications on EM images • Organelles are measured in micrometers (µm) and nanometers (nm): 1 mm equals 1000 µm 1 µm equals 1000 nm 1 nm equals 10 Å • The true size of objects in micrographs can be calculated by using the simple and useful rule of thumb. • There are 1000 µm in one mm, therefore, a 1 µm object will appear to be 1 mm in size when the magnification is 1000x. • To put a scale mark representing one µm on a micrograph, simply draw a line as many mm long as there are thousands in the magnifications. Example: Magnification 50,000x i.e., 1 µm magnified by 50,000 times since 1 mm equals 1000 µm, one µm equals 50 mm or 5 cm i.e. 1 µm 5 cm Precise measurement of objects in micrographs: The original size of the organelle seen from the ultra-micrograph can be measured either in micrometers or nanometers by using the following calculation: Size of Micrograph (mm) × 1000 Magnification

= true size in µm

OR Size of Micrograph (mm) × 1,000,000 = true size in nm Magnification Calculating the total magnification of a picture from Bar Magnification 2µ Ex: 10 cm

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Plant Techniques: Theory and Practice

2 µm equals 10 cm 1 µm 0 = 5 cm 00000 = 50 mm Since 1 mm + 1000 µm 1 µm + 50,000 µm, i.e., 50,000 x

2.2.3

Scanning Transmission Electron Microscope (STEM)

In a conventional transmission electron microscope (CTEM), images are formed by the electrons passing through the ultrathin specimen whereas in a Scanning transmission electron microscope (STEM) the electron beam is focused on a fine spot (spot size 0.05-0.2 nm) which is then scanned over the sample in a roster illumination system constructed so that the sample is illuminated at each point with the beam parallel to the optical axis (Fig. 2.12). The rostering of the beam across the sample makes STEM suitable for analytical techniques such as Z-Contrast annular dark-field imaging and spectroscopic mapping by energy dispersive X-ray (EDX) spectroscopy or electron energy loss spectroscopy (EELS). These signals are obtained simultaneously, allowing direct correction of images and spectroscopic data.

Fig. 2.12. Scanning Transmission Electron Microscope Carl Zeiss Libra 200 MC (Courtesy: University of Waterloo).

A typical STEM is a conventional transmission electron microscope equipped with additional scanning coils, detectors and necessary circuitry. The first application of STEM for biological molecules was demonstrated in 1971 (Walls, 1971).

3 Botanical Microtechnique and Microtomy The branch of plant science dealing with the preparation of plant material to observe under the microscope is known as Botanical Microtechnique (Berlyn and Miksche, 1976; Khasim, 2002). The development of our knowledge about plant and animal tissues has been possible with the steady advancement in the field of microtechnique and it is a fact that even Robert Hooke could not discover the cell until he prepared thin sections of plant tissues. In this chapter, a thorough discussion has been done on various aspects, viz., Killing and Fixation, Dehydration and Infiltration, and Embedding and Sectioning.

3.1 Killing and Fixation The killing and fixation of the materials are the two important processes for permanent slide preparation. Both killing and fixing are achieved by means of a single fluid consisting of various chemical reagents. The term ‘killing’ means the sudden and permanent termination of the life processes. As soon as we collect the material, it should be put in a killing reagent, so that permanent retardation of life processes takes place. Any reagent used for killing should reach each cell of the organism or a piece of tissue and it works completely and effectively. Killing invariably proceeds fixation which is said to be the preservation and keeping intact of all cellular and structural elements as they are in living conditions. It is well-known that the fundamental structure of protoplasm is colloidal. Every protoplasmic structure, such as the nucleus, plastids and cytoplasm, is an intricate colloidal system. Boundary membranes like nuclear membrane and tonoplast around vacuole are still another colloidal system. All these colloidal systems belong to the hydrophilous colloid group. When fixative reagents encounter the protoplasm, the colloidal system undergoes an irreversible change of state. Some reagents used for killing purposes penetrate rapidly and others penetrate slowly. Those reagents which penetrate rapidly may damage the tissue if materials are kept in for a prolonged time. For example, fumes of

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Plant Techniques: Theory and Practice

osmium tetroxide kill the tissue within a few minutes; when the material is allowed to react in this reagent for more than 30 mins, tissue damage occurs.

3.1.1

Physical Fixation

In the physical fixation, the tissue is cut into very small (1-2 mm) pieces and their cellular details are well-preserved by freezing at a very low temperature. Tissue is wrapped in aluminium foil and immersed with great rapidity into the freezing solvent at -160°C (in a liquid nitrogen trough), for not less than a minute. The freezing solvents commonly used are Freon 22, isopentane, propane, nitrogen, and a mixture of methyl cyclohexane: isopentane or methyl cyclohexane: 2 methyl butane. The physical fixation process has been used extensively to study the histochemical localisation of many enzymes and lipids at the optical microscopic level.

3.1.2

Chemical Fixation

Generally, both killing and fixation are done in a single fluid which consists of various combinations of reagents. The general principle involved in combining substances to form killing and fixative fluid (fixative mixture) is to secure a balance between all the properties of the reagents involved. A substance that tends to shrink cytoplasm should be combined with a substance that swells the protoplasm. In other words, the disadvantages of one reagent are counterbalanced by another reagent directly opposite in nature. Two reagents having identical disadvantages should not be combined in the killing and fixative fluid. Also, reagents that oxidise easily should not be combined with the reagents which are powerful reducers. 3.1.2.1 Fixation Images

Fixation images are of two types. A. Acid fixation image: Chromatin, nucleoli and spindle fibres are wellpreserved; cytoplasm is fixed as a stringy network and, nucleoplasm and mitochondria are dissolved. B. Basic fixation image: Chromatin and spindle fibres are dissolved. Nucleoli, nucleoplasm and mitochondria are well-preserved. The cytoplasm is fixed as hyaloplasm, and vacuoles are also preserved. The correct procedure for describing the final image should include both the killing and fixative fluid, as well as dehydrating reagent and infiltrating medium. The final image, given by Bouin’s fluid with absolute ethyl alcohol and xylol as the dehydrating agents, is quite different from that of tertiary butyl alcohol used as dehydrating agent (Johansen, 1940). There are some overlapping images with certain reagents. For example, copper bichromate used in a solution at pH 4.8 preserves both chromatin and mitochondria (Zirkle, 1928).

Botanical Microtechnique and Microtomy 29

If the fixing fluid contains two or more chemical reagents, the resulting fixation image is determined primarily by the reagent which penetrates more rapidly (Zirkle, 1933). The various reagents penetrate at different rates and according to their respective concentrations. According to Baker (1958), all fixatives are classified into two groups: coagulants and non-coagulants depending on their reactions to solutions of albumen and albumen-gelatin gels. Coagulants denature albumen and form precipitation. In other words, protoplasm is transformed into a microscopical network. Non-coagulants do not precipitate the proteins, but they alter the properties of albumen and gelatin. Gelatin is rendered insoluble in hot water and albumen becomes non-coagulable. Clearly, non-coagulants cross-link proteins without causing extensive denaturation that leads to precipitation.

Coagulants Various coagulants reagents are given below: A. Ethanol (C2H5OH): Ethanol (absolute ethyl alcohol) is a fair killing and fixing reagent when immediate results are needed. Plant materials should not be kept in ethanol for more than one hour, after which the reagent is poured off. Then the dehydration and infiltration of the material are followed. Proteins and nucleic acids are precipitated by alcohol, but it destroys mitochondria. Fats and phospholipids are dissolved. Plant materials that are killed and fixed by standard fluids should be kept indefinitely in 70% alcohol for further studies. B. Mercuric chloride (HgCl2): Mercuric chloride, which is highly poisonous, is soluble in water, ethanol and benzene. It is a rapid fixer and a powerful precipitant of proteins and nucleic acids, and it tends to shrink tissues. Plant material becomes opaque as soon as fixed. After fixation, mercuric chloride is removed by iodine in an alcoholic solution (1 g iodine and 2 g potassium iodide in 100 cc of 70% alcohol). Iodine colours the tissues but it may be removed by soaking in 70% ethanol or in 0.2% sodium thiosulphate. C. Chromium Trioxide (CrO3): Chromium trioxide forms chromic acid (H2CrO4) in water. Chromic acid tends to shrink tissues. To counteract this, it is mixed with other reagents. It is a powerful coagulant of albumen and many other proteins including nucleoproteins. Fats are not affected; the nucleus and chromosomes are well-preserved. In paraffin sections, cell aggregates are well fixed. Nuclear sap is changed into a coagulum. Nucleus, nucleolus and chromosomes are well shown. Mitochondria and lipid bodies are not seen. The cytoplasm is well-preserved by chromium trioxide since the cytoplasm has a strong affinity for acid dyes.

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Non-Coagulants Non-coagulants fix protoplasm without forming a microscopic network. Formaldehyde, osmium tetroxide and glutaraldehyde are of outstanding importance in modern microtechnique. These are useful in histochemical studies and in electron microscopy. A. Formaldehyde (HCHO): Formaldehyde is soluble in water, and it penetrates rapidly into the tissues (Baker, 1969). It is one of the hardening agents. It gives a basic fixation image. It reacts with amino groups of proteins, but it will not form precipitation with proteins. Nucleus, mitochondria and lipid bodies are well-preserved in this fixative. The fumes of formaldehyde are extremely irritating to mucous membranes. The action of formaldehyde is chiefly on proteins. It reacts with amino groups of proteins, with the production of water, or simply attaches itself to the amino acid without the liberation of water (Fig. 3.1 A, B; see also Woodroffe, 1941; Baker, 1969). Levey (1933 quoted in Sharma and Sharma, 1972) suggested that two molecules of formaldehyde may react with one of the amino acids (Fig. 3.1C).

Fig. 3.1. A-C Formaldehyde Reaction with Amino Group.

Tissues fixed in the fixative containing formaldehyde often show wellscattered chromosomes, especially after sectioning from paraffin blocks. The cell volume increases considerably, resulting in the spreading of the chromosomes over a larger area. The constriction regions appear slightly exaggerated due to the contraction of the euchromatic segments. This effect is possibly due to the action of formaldehyde on chromosome proteins (Sharma and Sharma, 1972). Formaldehyde is widely used as a fixative in electron microscopic studies. Tissues fixed in formaldehyde are very transparent to electrons. It is necessary

Botanical Microtechnique and Microtomy 31

to stain the sections with dyes that can deflect electrons. Generally, electron stains like uranyl acetate and potassium permanganate are widely used in electron microscopic studies. Many marine algae including Cyanophycean members are successfully fixed in 6-10% formaldehyde dissolved in seawater. B. Osmium tetroxide (OsO4): Osmium tetroxide also known as osmic acid and highly expensive, is widely used for electron microscopic (EM) studies. It is soluble in water. It gives no coagulum with albumen solution, but it is believed to form cross-links in proteins by reacting with the double bonds of tryptophane and histidine and by complexing with sulphydryl groups (Baker, 1969; O’Brien and McCully, 1981). It does not coagulate nucleoproteins. OsO4 blackens unsaturated lipids but does not attack saturated ones. So, it reacts with double bonds (Baker, 1969). However, some lipid-rich structures are destroyed by OsO4 (Zee and O’Brien, 1970). For EM studies, now-a-days OsO4 is used as a post-fixative for tissues that are initially fixed in glutaraldehyde or acrolein, or formaldehyde. Under these conditions, it gives positive electron contrast to membranes and preserves unsaturated lipids. Osmium tetroxide penetrates very slowly. After glutaraldehyde fixation in root tips of 1-2 mm thick, OsO4 appears to penetrate more quickly and develops good membrane contrasts (Newcomb, 1967). The main advantage of OsO4 is that it does not cause much shrinkage, but on the other hand, there is a slight swelling. The texture of the osmiumfixed tissue, though not very soft, allows smooth cutting for observation under a light microscope. Osmium tetroxide does not allow ethanol to cause precipitation during dehydration. A serious limitation of OsO4 fixation is the blackening of the tissue. Bleaching with hydrogen peroxide is an essential step after fixation by OsO4, but it affects the chromosome stainability (Sharma and Sharma, 1972). C. Potassium dichromate (K2Cr2O7): Potassium dichromate is a superb hardening agent and is best for studying mitochondria (Johansen, 1940). It is a non-coagulant of proteins including nucleoproteins. Unacidified potassium dichromate dissolves DNA; cytoplasm is homogeneously fixed and mitochondria well fixed. With acidified potassium dichromate, the image is similar to chromic acid. In acidified conditions, chromosomes are well fixed; cytoplasm and chromatin are precipitated, and mitochondria are dissolved. It fixes lipids effectively. D. Acetic acid (CH3COOH): Since it is easily frozen, undiluted acetic acid is also known as glacial acetic acid. One of the primary advantages of using acetic acid in a fixative mixture is that it can be combined with any of the other fixatives so far studied. It can be mixed in all proportions with water and alcohol. It is commonly used in many fixative mixtures

32

Plant Techniques: Theory and Practice

since it tends to swell with cytoplasm which counteracts the shrinkage effects of other reagents in the fixative mixtures. However, it shrinks tissues (Zirkle, 1928b). Its importance lies in its fat-soluble nature, which penetrates rapidly and produces the acid fixation image. It has got importance in the pickling industry as a preservative rather than a fixative. It is a non-coagulant for proteins and precipitates DNA. Acetic acid is also a good solvent for aniline dyes. Due to this property, it is an important component of staining-cum-fixing mixtures, viz., acetic carmine, acetic-orcein, acetic-lacmoid, etc. E. Glutaraldehyde [CHO(CH2)3CHO]: Glutaraldehyde is a non-coagulant, and it probably gives the most faithful preservation of any known fixative (O’Brien and McCully, 1981). Glutaraldehyde penetrates slowly, but reacts rapidly, especially compared to formaldehyde (Flitney, 1966). It does not penetrate a thick cuticle (Mersey and McCully, 1978). It rapidly forms cross-links in proteins. It is widely used for EM studies as a prefixative of the specimen. The biological grade of glutaraldehyde supplied by Polysciences (8% glutaraldehyde) and J. B. EM services (25% glutaraldehyde) is the most reliable one for EM studies. F. Acrolein (H2C=CHCHO): Acrolein is a highly reactive aldehyde, and it gives excellent preservation of various structures for light microscopy. It penetrates a thick cuticle and reacts relatively rapidly with the cytoplasm (Mersey and McCully, 1978). Unfortunately, commercial samples of acrolein available in the market are usually impure and acidic. Distillation is needed to remove these impurities. If it is available in pure form, acrolein is an excellent additive to glutaraldehyde. Acrolein can be used to fix bulky objects intact and other specimens where rapid penetration is required. Mersey and McCully (1978) reported that a mixture of glutaraldehyde and acrolein produced good fixation of tomato hair cells and immobilise cytoplasm at a faster rate than all other fixatives studied (including glutaraldehyde and acrolein separately). Unbuffered acrolein can be used successfully for the fixation of large brown and red algae (McCully, 1966; Evans and Holligan, 1972). Ice-cold acrolein should be used. Since it is potent tear gas, acrolein should be used in a well-ventilated hood. 3.1.2.2 Fixative Mixtures

Baker (1969) divided the fixative mixtures into four groups. These are given below: Group A

Coagulant + acetic acid: These are useful for anatomical and histological studies, e.g., Clarke, Zenker, etc.

Botanical Microtechnique and Microtomy 33

Group B Group C Group D

Coagulant + non-coagulant + acetic acid: These are used for detailed histology and cytology, e.g., Flemming’s strong mixture, FAA, etc. Coagulant + non-coagulant: These are common fixatives for cytoplasmic inclusions, e.g., Helly. Non-coagulants only: These fixatives are used to study the cytoplasmic inclusions, e.g., Altmann, Regaud, etc.

Fixative Mixtures giving Acid Fixation Images A. Formalin-Acetic Acid-Alcohol: This fixative mixture is familiarly known as FAA (Table 3.1). It is a coagulant fixative widely used to study the anatomical and morphological aspects and is unsuitable for chromosome studies. Specimens are kept indefinitely in FAA without any damage. The minimum time required for fixation is 18 hrs. Later specimens are transferred to 70% ethyl alcohol for permanent preservation. 70% ethyl alcohol Glacial acetic acid Formalin

90 ml 5 ml 5 ml

Sass (1967) stated that the fixative need not be washed out prior to dehydration. However, woody specimens should be washed with running water and softened for 3-6 weeks in a 50% aqueous solution of hydrofluoric acid. Detailed analysis of the mode of action of the FAA is not known (O’Brien et al., 1973). It is likely that acetic acid kills the tissue first, later this image is modified by formalin and ethyl alcohol. FAA gives an acid fixation image. B. Formalin-Propionic-Alcohol (FPA): This is similar to FAA; instead of acetic acid, propionic acid is used in this fixative. It is good for bryophytes and pteridophytes. 50% or 70% ethyl alcohol Propionic acid Formalin

90 ml 5 ml 5 ml

C. Chromic Acid-Acetic Acid-Formalin (CRAF): CRAF is widely used as a coagulant fixative. It is not suitable for cytoplasmic studies. Fix the plant material in this mixture at room temperature for 12-24 hrs. Specimens can be stored indefinitely in this fixative and dehydrated without washing. Baker (1958) has discussed in detail the action of its individual reagents, but the action of this mixture is not known. It is likely that the tissue is killed by acetic acid and that the image is modified by formaldehyde (reducing agent) / chromic acid (oxidising agent) products (O’Brien et al., 1973; Mersey and McCully, 1978).

34

Plant Techniques: Theory and Practice Sass (1967) has given five recipes for increasing the strength of CRAF: Final fixatives in ml III IV 30 40

I 20

II 20

Acetic acid (1%)

75

--

--

--

--

Acetic acid (10%)

--

10

20

30

35

Formalin (37-40%)

5

5

10

10

15

Water

--

65

40

20

--

Chromic acid

V 50

(1% chromium trioxide)

Formalin is added just before use.

D. Carnoy’s Fluids: These are of two types. The penetrating capacity of these two fluids is very high: (1) Absolute ethyl alcohol 15 ml Glacial acetic acid 5 ml (2) Absolute ethyl alcohol 30 ml Glacial acetic acid 5 ml Chloroform 15 ml The first one is commonly used for cytological studies. The root tips are kept for 15 minutes and anthers for an hour in this fixative. Fixed material should be washed twice with 95% ethyl alcohol before proceeding to paraffin infiltration and embedding. Since it penetrates rapidly into the tissues, alcohol acetic fixative is more useful for large specimens and thick sections. E. Gilson’s Fluid: Gilson’s mixture consists of the following reagents: 60% ethyl alcohol 50 ml Distilled water 440 ml Glacial acetic acid 2 ml 80% nitric acid, 46° strength 7.5 ml Mercuric chloride 10 g This fixative is recommended for softer and gelatinous fungi. Specimens are kept in it for 18 to 20 hrs. Later specimens should be washed with 50% ethyl alcohol to remove mercuric deposits. F. Petrunkevitsch‘s Fluid: It is a good fixative for angiospermic megagametophytes which need deep penetration. Specimens are kept in this fixative, not more than 20 hrs. Wash specimens thoroughly with 50% ethyl alcohol for further processing: 40% ethyl alcohol 125 ml Glacial acetic acid 27.5 ml

Botanical Microtechnique and Microtomy 35

Conc. nitric acid Mercuric chloride

2.5 ml up to saturation

G. Chamberlain‘s Chrom-Osmo-Acetic: This fixative consists of the following reagents: Chromic acid 1g Glacial acetic acid 1 ml 1% aqueous osmic acid 1 ml Distilled water make up to 100 ml This mixture is used to fix freshwater algae and fungi. It is not suitable for root tips, stem tips or any other morphological material. H. Taylor’s Chrom-Osmo-Acetic for Smears: This fixative mixture is used to preserve chromosomes effectively. In this mixture, maltose is also added to preserve the identity of trabante (satellite) and to avoid the obliteration of constrictions: 10% aqueous chromic acid 0.20 ml 10% aqueous acetic acid 2 ml 2% osmic acid in 2% aqueous chromic acid 1.50 ml Distilled water 8.30 ml Maltose 0.15 g I. Sass Modified Bouins Fluid: This mixture gives good results in the case of flower buds and anthers of Liliaceae: 1% aqueous chromic acid 50 ml Saturated aqueous picric acid 35 ml Formalin 10 ml Glacial acetic acid 5 ml J. Fleming’s Chrom-Osmo-Acetic Mixture: This fixative is used for cytological studies: 1% chromic acid 50 ml 10% acetic acid 10 ml 2% osmic acid 10 ml Distilled water 30 ml The acetic acid in the mixture prevents excessive damage. Osmic acid preserves the chromosome structures and blackens the tissues. So, it is necessary to bleach the fixed material before staining. K. Weak Chrom-Acetic (Johansen, 1940): This fixative is used to preserve algae, fungi, bryophytes, prothalli of pteridophytes, and moss capsules which are easily penetrated: 10% aqueous chromic acid 2.5 ml 10% aqueous acetic acid 5 ml Distilled water make up to 100 ml

36

Plant Techniques: Theory and Practice

L. Strong Chrom-Acetic: This mixture is used for woody materials and tough, leathery leaves: 10% aqueous chronic acid 1 ml 10% aqueous acetic acid 10 ml Distilled water make up to 100 ml

Fixative Mixtures Giving Basic Fixation Images A. Zirkle-Erliki Fixative (Zirkle, 1934): This mixture preserves mitochondria, but dissolves all chromatin. The specimen is fixed for 48 hrs and then washed with water: Potassium dichromate 1.25 g Ammonium dichromate 1.25 g Cupric sulphate 1.00 g Distilled water 200 ml B. Marengo-S Fixative (Marengo, 1952): It is used to fix cytoplasm. The material is fixed for 24 hrs and washed thoroughly in running water before dehydration: 10% formalin 100 ml Normal sodium hydroxide 1 ml Pyrogallol 7g C. Zirkle’s reduced Chromic Acid: It is used for mitochondria and vacuoles (Zirkle, 1932). The specimen is fixed in this mixture for 48 hrs and later the fixed material is washed with running tap water: Chromium sulphate 5g Cupric oxide slight excess Formalin 10-50 ml Distilled water 90-50 ml The total amount of mixture should be 100 ml Table 3.1. Fixative mixtures used for specific purposes Fixatives

Specific purpose of the fixative

Pre-fixation

Post-fixation

1

Glutaraldehyde 1.6% + ruthenium red in 0.08 M cacodylate buffer (pH 7.3), fix for 8 hrs at 2oC

1.67% osmium tetroxide + 0.066% ruthenium red in 0.066 M cacodylate buffer (pH 7.3) for 2 hrs at 20oC

Good preservative for most of the organelles including microtubules (Cass et al., 1985). For anatomical and morphological studies.

2

*Formalin-acetic acidalcohol (FAA) for 18 hrs and

_

For anatomical and morphological studies

Botanical Microtechnique and Microtomy 37 later transferred to 70% ethyl alcohol for permanent preservation _

Good for bryophytes and pteridophytes

3% glutaraldehyde and 3% acrolein (1:1) buffered in 0.3 M cacodylate (pH 7.2) for 2 hrs at 4oC, later wash in buffer

Aqueous 6% potassium permanganate for 20 minutes at 4oC, later wash in buffer

Cell organelles are well-preserved and show high contrast of membrane system

5

2% glutaraldehyde in 0.14 M sodium chloride, 0.005 M sodium acetate and 0.005 M calcium chloride at pH 7 or 2% acrolein without stabiliser

_

6

4% aqueous potassium permanganate for 2-4 hrs

1% osmium tetraoxide aqueous or buffered solution for 2-12 hrs.

7

*Carnoy’s fluids for 15 minutes to 1 hr, later washed twice with 9% ethyl alcohol

8

3

*Formalin-propionicalcohol (FPA) for 18 hrs

4

Membrane systems are well-preserved (Carstenson et al., 1971)

Both membrane systems and lipid deposits are wellpreserved.

_

For cytological studies

2% acrolein and 2% glutaraldehyde in 50% aqueous dimethyl sulfoxide (DM 50) buffered with 0.02 M 3-collidine (pH 7.4) for 7 hrs. in cold, later wash with buffer

3% osmium tetroxide in veronal acetate buffer (pH 40)

General structure and lipids are well fixed in yeast (Schwab et al., 1970)

9

*Gilson’s fluid for 18-20 hrs. later washed with 50% ethyl alcohol

_

Good for gelatinous fungi

10

0.6% buffered potassium permanganate (pH 7.4-7.6) for 15 minutes -12 hrs 0oC, later wash in cold ethanol 25% and bring to room temperature

_

Preserves well and improves the contrast of membrane systems (Luft, 1956)

11

3% Glutaraldehyde, 1.53% paraformaldehyde and 1.5% acroleinin

Potassium permanganate 1-2% for 15-30 min at 0oC

Excellent preservation for protein bodies, vacuoles and (Contd.)

38

Plant Techniques: Theory and Practice Table 3.1. (Contd.) Fixatives Pre-fixation

Specific purpose of the fixative Post-fixation

5-collidine buffer for 1-2 hrs.

membranes in seeds (Millenhauer and Totten, 1971)

12

*Petrunkevitsch’s fluid for not more than 20 hrs and wash in 50% ethyl alcohol

_

13

6 ml of 10% glutaraldehyde + 3 ml of 10% acrolein + 5 ml of 6% paraformaldehyde + 5 ml of 0.2 M buffer + 1 ml distilled water

2% osmium tetroxide

14

*Taylor's chromosmoacetic fluid

15

1% (w/v) solution of malachite green in a standard solution containing 2% glutaraldehyde in 75 ml sodium dihydrogen phosphate buffer at 7.3 pH with 0.25 mM calcium chloride; fix for 90 min at 4oC, later wash in the same buffer

16

*Fleming’s chromosmoacetic mixture; since it blackens the tissue, it is necessary to bleach the fixed material

_

Chromosomes wellpreserved

17

50% aqueous solution of glutaraldehyde, 4% aqueous osmium tetroxide, and 0.1 M collidine buffer are mixed in the ratio of 1:2:5 respectively at pH 7-7.1; fix for 1 hr at 0-4oC

_

Reduces cell shrinkage, lipid extraction, and chromatin clumping which usually happens by routine glutaraldehyde and osmium tetroxide procedures (Trump and Bulger, 1966)

_

Good fixative for angiospermic megagametophytes Seeds rich in protein bodies and lipids are well-preserved

Preserves chromosomes effectively

1% osmium tetroxide Bacterial ultrastructure including lipid bodies in veronal acetate buffer (pH 6.0) for 90 are well-preserved min at 4oC

Botanical Microtechnique and Microtomy 39 18

Zirkle-Erliki fixative for 48 hrs later wash with water

_

Preserves mitochondria effectively but dissolves all chromatin

19

*Marengo-S fixative for 24 hrs later wash with water

_

Fix the cytoplasm

20

*Strong chrom-acetic mixture

_

Used for woody materials and tough, leathery leaves

(also refer to Bhandari, 1997) * See text for the composition of various ingredients

3.2 Dehydration and Infiltration After washing the fixed plant material, it is necessary to remove the water from the tissues since water is not miscible in paraffin wax in which tissues are embedded for smooth cutting. The removal of water from tissues is known as dehydration. Dehydration is also necessary for making permanent preparations, so that sections may not be infected and stored for quite a long time. Various dehydrating agents in different concentrations are used here. It is a gradual process and utmost care should be taken at this stage. Otherwise, plasmolysis takes place and leads to damage to tissues. Dehydration should be done in lower temperatures, so that mechanical distortion of specimens may be minimised. Some of the dehydrating schedules are given in Table 3.2 and these are based on the medium employed here (also refer to O’Brien and McCully, 1981; Bhandari, 1997). Table 3.2. Dehydrating Schedules and Types of Embedding Media Embedding media

Appropriate dehydrating schedules

Paraffin wax

Ethanol series; butanol/ethanol/water series; methyl cellosolve series; TBA series; diaxon (diethylene dioxide); tetrahydrofuran

Polyester wax

Methyl cellosolve series up to n-propanol

Glycol methacrylate (GMA)

Methyl cellosolve series; ethanol series; GMA itself can also be used but it is expensive

Epoxy resins

Methyl cellosolve series; ethanol or acetone series; propylene oxide series are recommended for all epoxy resins after ethanol or acetone except for Spurr’s resin

In general, the material to be infiltrated in paraffin is dehydrated through either ethanol-xylene series or ethanol-tertiary butylalcohol series. For resin-

40

Plant Techniques: Theory and Practice

embedded tissues, a closely graded ascending series (25% onwards) of ethanol or acetone is used for dehydration.

3.2.1

Dehydration

The following procedures are generally employed for dehydration: A. Ethanol Series (Chamberlain, 1932) Tissues are passed through the following ethanol/water mixture. If the plant specimen is washed with water, tissues should pass through from 2%, 5, 7, 10, 15, 20, 30, 40, 50, 70, 85, 95 to 100% ethanol. If the specimen is fixed in 70% ethanol, commence the dehydration at the same ethanol concentration. Sass (1967) proposed the following ethanol series for dehydration: from 5, 10, 15, 20, 25, 30, 35, 40, 45, 50, 60, 70, 80, 95 to 100% ethanol. Sass further suggested that acetone or isopropyl alcohol can also be substituted for ethanol. Acetone is superior to ethanol, especially for specimens infiltrated in epoxy resins because neither it reacts with osmium tetroxide which might have been left after washing the specimen nor with epoxy monomers. Chayen and Gahan (1959) recommended rapid acetone dehydration and infiltration in ester wax for retention of lipids in plant tissues. All the above steps may not be necessary for delicate specimens. Two hours of immersion of the specimen in each grade is enough. In the case of woody specimens, several hours of immersion are required. There are some disadvantages to using ethyl alcohol as a dehydrating agent. These are (a) the schedule is time-consuming, (b) if the fixed schedule is not suited to the tissue, excessive hardening and shrinkage of the tissue takes place and (c) as alcohol does not mix with paraffin or celloidin, an intermediate ‘clearing’ agent is needed which unduly lengthens the process (Sharma and Sharma, 1972). So, several alternative dehydrating agents have been used from time to time, some of which are miscible with the embedding material, so that using the clearing agent is not necessary. B. Ethanol-xylene Series It is commonly employed for paraffin embedding tissues. Fixed material is passed through the following grades: (i) (ii) (iii) (iv) (v) (vi)

25% ethanol 50% ethanol 70% ethanol 90% ethanol 95% ethanol 100% ethanol

(vii) (viii) (ix) (x) (xi) (xii)

3:1 ethanol and xylene 1:1 ethanol and xylene 1:3 ethanol and xylene Xylene I Xylene II Xylene III

In each grade, the specimen is kept for 6-24 hrs depending on the hardness of the material.

Botanical Microtechnique and Microtomy 41

C. Ethanol-Tertiary Butyl Alcohol (TBA) - Water Series It is also employed for paraffin embedding tissues. Johansen (1940) claimed that it was the least drastic of all dehydrating agents. Materials are kept in the following grades: (i) (ii) (iii) (iv) (v) (vi) (vii) (viii) (ix) (x)

25% ethanol 50% ethanol 5:4:1 ethanol (95%): 5:2:3 ethanol (95%): 5:3.5:3 ethanol (95%): 1:1 ethanol (95%): 1:3 ethyl alcohol: 100% anhydrous: ,, ,,

TBA: water for 2-4 hrs at room temperature TBA: water TBA: water TBA for 2-4 hrs at room temperature TBA TBA I II III

Material is kept for 12-24 hrs in each grade. According to Johansen (1940), this treatment removes every trace of unbound water. D. Butanol-ethanol - Water Series The following series of dehydration has been recommended for wax embedding tissue (Sass, 1967): (i) (ii) (iii) (iv) (v) (vi) (vii) (viii)

1: 2: 7 1.5:2.5:6 2.5:3:4.5 4:3:3 5.5:2.5:2 7:2:1 8.5:1.5 100%

butanol: ,, ,, ,, ,, ,, ,, butanol

ethanol (95%): ,, ,, ,, ,, ,, ,,

water ,, ,, ,, ,, ,, ,,

E. Dioxane (Diethyl Dioxide) - Water Series Dioxane mixes with water, alcohol and xylene and dissolves paraffin wax and balsam. Hence, it may be used as a substitute at any stage of the usual ethyl alcohol-xylene dehydrating and clearing schedule. Direct treatment is also possible. Later it was also used by LaCour and Maheswari (quoted in Sharma and Sharma, 1972) and Johansen (1940). The specimen is directly transferred from water to 60% dioxane in distilled water and subsequently as follows: (i) (ii) (iii) (iv) (v)

60 ml dioxane + 40 ml distilled water 70 ml dioxane + 30 ml distilled water 95 ml dioxane + 5 ml distilled water Pure dioxane I Pure dioxane II

42

Plant Techniques: Theory and Practice

Since it does not need any clearing agent, this can be followed immediately by embedding in paraffin wax. Though dioxane is very effective in short-term schedules, particularly for animal tissues, it should be used very cautiously because it has a markedly toxic and cumulative action and is an injurious one that should be handled with extreme care. It is also non-specific in its action and heavier than melted paraffin; therefore, it is very difficult to remove from tissue during infiltration (Sharma and Sharma, 1972). F. Methyl Cellosolve (2-Methoxy Ethanol) Series (Feder and O’Brien, 1968) This series is recommended for the plant material embedding in paraffin wax as well as epoxy resins: (i) Methyl cellosolve (v) n-propanol (ii) Methyl cellosolve (vi) n-propanol (iii) Ethanol (vii) n-butanol (iv) Ethanol (viii) n-butanol Tissues may be stored in the refrigerator. If the material is fragile, dehydration should be done in four concentrations of methyl cellosolve: 25%, 50, 75 and 100%. Generally, cellosolve has got remarkable power of dissolving diverse substances including cellulose nitrate and acetone; hence it is called cellosolve. It causes less shrinkage and hardening than ethanol. Since cellosolve is a less violent dehydrating agent than ethanol, there is no need to pass the plant material through graded series of a mixture of cellosolve and water. 3.2.1.1 Clearing with Chloroform and Xylene

Since paraffin does not mix with many dehydrating agents, an intermediate medium also known as ‘ante-medium’ which is miscible with both the dehydrating agent and paraffin is generally necessary. Most of these reagents render the tissue translucent because their refractive indices are close to that of the proteins of the tissue and, light rays can pass through without refraction. Hence, they are called ‘clearing’ agents and the process ‘clearing the tissue’. The ideal ante-medium should have rapid penetrating power and should mix equally well both with the dehydrating and embedding agents. The clearing is also as gradual as dehydration. Chloroform and xylene are working as clearing agents. After passing through the methanol series and butanol-ethanol-water series, the specimen should also be passed through the following series of a mixture of chloroform and absolute alcohol. (i) 2:1 absolute alcohol: chloroform (ii) 1:1 ,, ,, (iii) 1:2 ,, ,, (iv) 100% chloroform I (v) ,, II

Botanical Microtechnique and Microtomy 43

In each grade, the specimen should be kept for one hour in a corked specimen tube. Xylene hardens the tissue, sometimes it causes plasmolysis. Instead, chloroform is used since it has a slight hardening effect. Johansen (1940) also suggested Bergamot oil and cedar wood oil as a clearing agent for bulkier pieces of tissue. With cedarwood oil, 2-3 hrs of treatment in each grade is quite enough. The only limitation of these two oils is that they are not good solvents for paraffin and, therefore, they are not easily replaced from the tissue by paraffin. 3.2.1.2 Chemical Dehydration

Chemical dehydration is an alternative approach in the botanical microtechnique. The DMP (2, 2 dimethoxypropane) has also been used as a dehydrating agent for plastic embedding tissue. The specimen is dehydrated in acidified DMP, and the process is completed when no further cooling is evident (endothermic reaction). Later two changes of acetone are given, and the material is infiltrated in the usual manner. Lin et al. (1977) also employed acidified DMP and DEP (2, 2- diethoxypropane) as dehydrating agents. Three drops of 0.1N HCl are added to 25 ml DMP or DEP. After dehydration for 15 minutes, three changes of dry acetone are given, 15 minutes each. Complete dehydration of plant material is accomplished in a single step with these reagents. DEP and DMP react with water and produce acetone and methanol (Bhandari, 1997). Inorganic dehydrants such as HCl, CaCl2, anhydrous Na2SO4 and molecular sieves are also being used (O’Brien and McCully, 1981).

3.2.2 Infiltration After dehydration and clearing, the material becomes transparent. Now the material is ready for infiltration-either in paraffin wax or any resin. Infiltration is the process of replacing the dehydrating agent from the dehydrated tissues with an embedding medium. It is also a gradual process. During this process, intercellular spaces are filled with paraffin wax, so that it helps in getting the smooth cutting of tissue without any damage to the microtome. A. Paraffin Wax Method The paraffin wax method is widely followed for anatomical, embryological and cytological studies: (i) The dehydrated material is given two changes in chloroform or xylene or tertiary butyl alcohol (according to the method employed). In each change, the material is kept for 12 hrs. (ii) Add shavings of paraffin wax (M.P. 52°C) to the specimen tube containing the material and solvent. (iii) Specimen tubes are placed on the upper shelf of the oven in which the temperature should not be more than 56°C.

44

Plant Techniques: Theory and Practice

(iv) Go on adding the shavings of wax gradually in the specimen tube till some wax remains undissolved. It may take 3 hrs to saturate. (v) Pour off the mixture of solvent and wax and replace it with fresh molten wax. (vi) Give two changes of pure molten wax at an interval of 6 hrs. (vii) The process is continuously repeated till the traces of smell of solvent are not found. If any traces are unremoved, it leads to the crystallisation of paraffin wax after embedding. (viii) If it is a delicate specimen, do not keep it in the oven for more than 2 hrs during the entire process of infiltration. (ix) Pour off the paraffin and replace it with a 56-58°C tissue mat (a commercial formulation with excellent cutting properties). It is allowed to stand for three days (O’Brien and McCully, 1981). Now the tissue is ready for embedding. Other waxes such as ester and polyester wax are also used for the infiltration of plant tissues; these tissues can be embedded in tissue mats (Chayen and Gahan, 1959; Steedman, 1960; Bonnet and Torrey, 1965). B. Parowax Method This method is employed for the materials dehydrated in dioxane. With dioxane as dehydrating agent, little chips of parowax are added gradually to the dehydrated material, contained in pure dioxane. The mixture is kept in a warm bath till dioxane (not a good paraffin solvent) is saturated with parowax. The latter steps are similar to the paraffin wax method. C. Glycol Methacrylate (GMA) Method (Feder and O’Brien 1968; Bhandari, 1997) Monomer mixture Purified GMA (Purified by adding 4 g activated charcoal to 100 ml of the monomer mixture to eliminate certain inhibitors of polymerisation) Polyethylene Glycol (plasticizer) 2, 2’-Azobis (2-methyl propionitrile) (catalyst)

92.2 ml

7.5 ml 0.3 g

Bring the specimen into n-butanol to which a complete GMA monomer mixture is added. Keep it for 12 hrs for infiltration. Pour off the solution and replace it with the pure monomer mixture. Give two changes in 24 hrs. If it is necessary, infiltrate the material with mild agitation at 6°C or at room temperature for 1-4 weeks (Jurand and Ireland, 1965; Steinbrecht and Ernst, 1967; Kajtar et al., 1970). Keep these vials in the dark to reduce premature polymerisation. Proper infiltration is necessary to get thin sections of 0.5-2.0 µm thick for light microscopy and 0.05-0.1 µm thick for electron microscopy. Inadequate infiltration of the cell wall is one of the factors for unsatisfactory sections.

Botanical Microtechnique and Microtomy 45

3.3 Embedding and Sectioning 3.3.1

Embedding

Medium which converts liquid to solid form easily can be used as an embedding medium. The liquid penetrates the tissue and converts it into a solid. This conversion may involve hydrogen bonding, covalent linkage, and crystallisation or polymerisation (Baker, 1969). Baker (1969) suggested that embedding media may be divided into two groups: those that penetrate the cells and those that merely surround them. Those fixatives which coagulate the proteins in the form of a network leave the tissue in a state that favours the entry of the embedding medium. Those which fix homogeneously, without coagulation, do not favour the entry of the embedding medium into internal parts. The embedding medium enters intercellular spaces and gives considerable mechanical support during sectioning. Steedman (1960) divided the embedding media into two groups: ribboning and non-ribboning depending upon whether the successive sections are adhered to one another to form a ribbon. Flemming (1873) and Wilcox (1898) used soap as a ribboning medium, but its adhering capacity is very poor. Later it was replaced by paraffin wax. 3.3.1.1 Waxes

A. Paraffin Wax After infiltration, embedding is done with paraffin wax. Embedding involves the pouring of the molten wax into the paper tray, properly arranging the infiltrated plant material in it, cooling the entire mass quickly and solidifying within the paper tray. Paraffin wax certainly has some advantages. The process of embedding is quick and simple; embedded material can be stored for quite a long time in dry conditions, and it forms ribbons in which sections are present while cutting with a microtome. Due to these qualities, paraffin embedding is widely used for embryological, anatomical and cytological studies. The chief drawback of paraffin embedding is that despite all precautions, the lengthy procedure of dehydration, clearing and embedding in molten paraffin wax causes some amount of shrinkage and distortion. The same thing is also encountered in the celloidin method. The plant material should be dehydrated thoroughly and soaked in fluids that are easily miscible with an embedding medium. Ethanol and cello solve are particularly suitable for paraffin embedding. Paraffin wax is a derivative of crude petroleum. It is a saturated and long-chain hydrocarbon of the methane series (Baker, 1969). Commercially, paraffin wax is available at three melting points, 45°C, 52°C and 60°C. Paraffin wax with a melting point of 52°C is widely used. Hard waxes are more suitable for taking thin sections. Some of the waxes and their properties are given in Table 3.3.

46

Plant Techniques: Theory and Practice Table 3.3. Types of Waxes Melting point

Properties

1 Fibro wax

Name of Wax

57-58oC

Consists of paraffin wax and plastic polymers Difficult plant parts can be cut with this wax Its compression is less while sectioning

2 Ester Wax

47-48oC

Low melting point Harder than paraffin wax Its compression is less than the paraffin wax during cutting Sections with less than 5 µ can be achieved It should be kept in the molten stage longer than 12 hrs because it gets damaged

3 Paraplast

56-57oC

Similar to fibro wax

o

4 Polyester wax

38 C

Soluble in ethyl alcohol Used for embedding delicate specimens

5 Carbowax (Polyethylene glycol)

56oC

Wax-like substance soluble in water

Embedding receptacle (or Tray): Several types of embedding receptacles are being used for block making. Generally, stiff paper trays and L-shaped trays are used. With the help of two L-shaped metal pieces (Fig. 3.2), molten wax is poured into the paper boat and the required size of blocks is prepared.

Fig. 3.2. Embedding Receptacle.

For preparing paper boats, the below-given procedure is followed: (i) Required size of a rectangular piece of paper is taken. Stiff and glazed paper is used here (porous is avoided as paraffin penetrates in it).

Botanical Microtechnique and Microtomy 47

(ii) As is shown in Fig. 3.2, the first fold is made over CC’ and DD’.

Fig. 3.3. Making Paper Tray.

and the width of the fold is decided by the thickness of the material to be embedded. The width should be a few mm more than actual need. (iii) Next fold along AA’ and BB’, which is double the width of CC’ and DD’. Now fold back along the middle of each of these two flaps as indicated by EE’ and FF’. (iv) Make short diagonal creases along EG, E’G’ and FH, F’H’. (v) Bring one end and one side perpendicular with fold at short diagonal crease and turn the resulting flap at the back of the end wall. (vi) Bring up the opposite side wall and fold its flap back. (vii) Now fold down upper flap of end wall backward, thus securely locking the entire end. (viii) Follow the same procedure for opposite end also. Trays may also be prepared from thick cellophane (Johansen, 1940). One may use small porcelain trays or even solid watch glasses for preparing wax blocks. Wax-block preparation: Quick work and skillful manipulations are required for wax-block preparation.

48

Plant Techniques: Theory and Practice

(i) Smear the glycerine on the inner side of the paper tray or L-shaped metal pieces to avoid sticking wax to it. (ii) Slightly wet the bottom surface of the paper tray with water and place it on a glass sheet. (iii) Pour molten wax from a beaker into the paper tray about 3/4 full. Wait till the wax forms a thin congealed layer at the bottom of the tray. (iv) Transfer the specimens from the infiltration bath and arrange them by giving proper spacing in congealing wax. Warm spatula and warm needles are used here to transfer the specimens. One must work quickly at this stage. (v) To avoid air bubbles and cracks in the specimen area, melt the thin skin of wax that forms at the top surface of the block with a warm spatula and carefully pour the molten wax in the region containing the specimen which is cooling quickly. Keep the upper surface molten till the area around the specimen should be solidified. (vi) Sometimes a crust of solid wax may be carried down along with the specimen. It is essential to melt this wax with a warm spatula otherwise, it creates problems during section cutting. (vii) Now the paper tray is allowed to float on the water surface for cooling till the surface of wax becomes sufficiently solidified. After complete solidification of paraffin wax, the paper tray is peeled off and taken out of the wax block which contains sufficiently spaced specimens. Now, these blocks are kept in the air for some time. If the blocks have been kept dry for longer periods, it is necessary to soak the blocks in the mixture of glycerol and alcohol for atleast 24 hrs before section cutting. It is not advisable to cut the freshly prepared blocks immediately. The blocks should be left atleast for 24 hrs to get the temperature stabilised and proceed with section cutting. Inspite of its advantages, paraffin wax has three major faults, these are crystalline nature, hydrophobic and relatively high melting point (50°-60°C). Numerous attempts have been made to overcome these problems, such as heat treatment of wax or additives like asphalt, rubber, ceresin and other waxes are used to reduce crystallinity, and stearic acid or glycols are added to decrease hydrophobicity (O’Brien and McCully, 1981). B. Carbowax Carbowaxes are polyethylene glycols and these are soluble in water. These can be used for both dehydration and infiltration. The procedure followed by Riopel and Spur (1962) in potato tissues is given below: (i) The fixed tissues are passed through the 5, 10, 15, 20, 30, 40, 50, 60 and 70% carbowax400 in water. Keep them for 12 hrs in every step. (ii) Fill 2/3 of the vial by adding enough pieces of carbowax 1540 to the 70% carbowax400. Allow it to stand overnight.

Botanical Microtechnique and Microtomy 49

(iii) Remove the vial caps and put all vials in an oven at 56°C for 24 hrs. Gently shake the contents twice during 24 hrs period. (iv) Transfer the tissues to the mixture of carbowax1540 and carbowax 400 (19:1) and keep them in an oven at 56°C for 12 hrs. Give two changes with this mixture. (v) Maintain the supply of carbowax 1540/400 (19:1) mixture in an oven at 50-51°C. Transfer the tissues in this infiltration mixture at 56°C in an oven for 5-10 mins. (vi) Embed the material in a 51°C mixture at room temperature. Allow it to cool on a pan of ice water. Since it is soluble, embedded material should not be submerged in water (Wade, 1952). Riopel and Spurr (1962) used porcelain dishes or plastic boxes for embedding. C. Celloidin Celloidin embedding is adopted for woody material which is not possible to cut in paraffin embedding. Celloidin causes minimum shrinkage and distortion. It is a nitrocellulose substance alternatively known as a soluble gun-cotton or pyroxylin (Clayden, 1962). It is highly inflammable but non-explosive. Celloidin is available in the form of tablets or as shreds. The shredded form is stored in water. A very good alternative for shred celloidin is the Necol celloidin solution formulated by Imperial Chemical Industries Limited, Dorset. Celloidin embedding is done in the following manner (Johansen, 1940; Clayden, 1962): (i) As usual, after killing, the plant material should be dehydrated in the ethanol series or in the TBA series. In the TBA series, the material is transferred to the mixture of 75% tertiary butyl alcohol and 25% absolute ethyl alcohol. From this mixture, it is again transferred to a mixture of equal parts of tertiary butyl alcohol, absolute alcohol and ether. Now the specimen is ready for infiltration. (ii) Mixture of equal parts of ether and absolute alcohol is taken. Celloidin solutions of 2, 4, 6, 8 and 10% are prepared in this mixture and kept separately in tightly stoppered bottles. (iii) Dehydrated material is taken in a mixture of equal parts of ether and absolute alcohol, and then transferred to a wide-mouth bottle containing 2% celloidin. Later the specimen is passed through the different grades of celloidin up to 10%, allowing one too many days in each grade depending on the nature of the material. It requires several weeks for proper impregnation and embedding of the material. Some technicians apply heat to hasten the impregnation process. (iv) Now celloidin blocks are prepared with the help of thick paper. Fold the paper along dotted lines and prepare a mould or container (Fig. 3.3).

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Plant Techniques: Theory and Practice

(v) Place the tissue in this mould, with the surface to be cut facing upwards. (vi) Fill the mould with 10% celloidin. Care should be taken to see that a sufficient margin of celloidin is allowed all around the specimen, so that there may not be any problem while cutting the celloidin blocks. (vii) Place the mould on a flat surface and then transfer it to a moderately deep glass bowl with a well-fitted lid. (viii) Expose the mould to ether vapours for about three days. Ether vapours disperse any bubbles in the celloidin solution. (ix) Later replace the ether with chloroform and leave for 1 to 2 days until celloidin becomes thickened and hardened. (x) Trim off the excess celloidin leaving a good margin around the specimen. Now celloidin blocks are ready for cutting. These blocks can be stored in 70-80% alcohol or in a mixture of glycerine-95% alcohol (equal parts) indefinitely without damage to the specimen. Various types of microtomes are made for cutting celloidin sections. Leitz base sledge is a heavier microtome, and it is better for cutting hard tissue. In this microtome, the knife is fixed at an angle, whereas the tissue on the holder moves (Fig. 3.4). Reichert sledge microtome is a lighter machine, and it is good for soft tissues. In this microtome, the tissue remains stationary whereas the knife moves until the tissue is cut. Celloidin sections are stained directly without removal of the celloidin matrix, which does not interfere with staining. The slides with the sections or the individual sections themselves are brought down to water from 70% alcohol and then stained. If necessary, the matrix may be removed before final mounting in Canada balsam. 3.3.1.2 Plastics

Embedding in plastics gives good results when compared to paraffin wax (Steedman, 1960; Sidman et al., 1961; Feder and O’Brien, 1968). Glycol Methacrylate (GMA) Glycol methacrylate (2-hydroxyethyl methacrylate), a monomer mixture, is a stable and slightly viscous liquid miscible with almost all organic solvents and water. GMA infiltrated specimen is embedded in the same medium and this can be sectioned conveniently at 4 µm with an ultramicrotome. The sections are quickly and easily stuck to glass slides and staining is done in a simple pattern. Since the embedding matrix is hydrophilic, it is quite permeable to aqueous stains. Without removing the plastic, sections are conveniently stained and examined under a microscope. Embedding is done in the following manner: (i) Place the infiltrated specimen in a gelatin capsule (no. 00). (ii) Fill the capsule with GMA monomer mixture (see section 3.2.2C

Botanical Microtechnique and Microtomy 51

for ingredients). Capsules are tightly capped (oxygen inhibits polymerisation). (iii) Place the fitted capsules in an oven at 40°C for 1-2 days; the monomer mixture starts polymerisation and gives a hard block; place the block in an oven at 60°C for an additional day to complete hardening (this step may not always be necessary). (iv) GMA blocks are trimmed and sectioned with a dry glass knife on an ultramicrotome. Sections of 0.5-4 µ thickness are generally the most useful. (v) Place the section on a drop of distilled water on a microslide. Allow the water to evaporate and stain the section the next day. For certain enzymological investigations, where polymerisation at higher temperatures is unwanted, another plastic JB-4 (Polysciences Inc., Washington, PA) is used. Namba et al. (1983) proposed the cold temperature glycol methacrylate embedding medium available under the trade name JB-4. The procedure is as follows: (i) Infiltrate the specimen overnight in non-polymerising solution A with a catalyzer. (ii) Transfer into embedding moulds filled with polymerising plastic consisting of: (a) Solution A (b) Catalyzer (c) Solution B (Polymeriser)

100 ml 0.90 g 4.0 ml

Polymerization is carried out overnight at 2°C or at room temperature (23°C). Cold embedding gives good results since temperature affects enzyme localisation quantitatively. 3.3.1.3 Resins

Epon mixture is the commonly used resin as it penetrates faster than araldite of its low viscosity. The other resins which are being used are given below: A. Durcupan (Staubli, 1960; Kushida, 1964) Since Durcupan is soluble in water, dehydration is carried out in an ascending series of the resin. Embedding medium: Mixture A Durcupan Dodecenyl succinic anhydride (DDSA) Mixture B Durcupan

100 ml 234 ml 100 ml

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Plant Techniques: Theory and Practice

Nadic methyl anhydride (NMA) 124 ml Mixtures A and B are combined in the ratio of 8.2:7.3 at 20°C (Kushida, 1964). B. Vestopal W (Ryter and Kellenberger, 1958) Vestopal W is employed for preparing very hard blocks (harder than epon blocks). It is characterised by rapid penetration and polymerisation. The mixture consists of: Vestopal W Benzoyl peroxide (initiator) Cobalt naphthenate (activator)

100 ml 1 ml 0.5 ml

The initiator and activator must be stored in a cold and dark place to prevent decomposition, but they should be replaced after every few months. Vestopal ‘N remains good for several months at 4°C. Vestopal W is not miscible in ethanol but readily mixed with acetone and styrene. For dehydration and embedding, use 30, 70, 95 and 100% acetone, again 100% acetone (5-10 mins in each grade) and Vestopal mixture: acetone (1:3,1:1,3:1,30 mins in each) and finally in Vestopal mixture for 1 hr. The polymerisation is completed in 12 hrs at 60°C. C. Polystyrene (Frangioni and Borgioli, 1979) Polystyrene is employed for both light and electron microscopy. Polystyrene solution: Polystyrene 200 g Toluene 800 cc Benzyl alcohol 50 cc Dibutyl phthalate 5-10 cc First, take the toluene and then add polystyrene (BDH Chemicals Ltd.) in a one-liter bottle, seal this hermetically and shake vigorously at regular intervals until the solution becomes homogeneous. To this, add benzyl alcohol and dibutyl phthalate, and shake again until the solution becomes homogeneous. The solution which should be free from air bubbles, transparent and colourless is now ready for use. The entire process takes two days. The hardness of the polystyrene block depends on the percentage of dibutyl phthalate added to the solution.

Procedure (i) After fixation and dehydration, tissues are placed in toluene for 0.5-1 hr. (ii) Transfer the tissues to the polystyrene solution, spacing them out with a needle as soon as they sink to the bottom of the container. Polyethylene ice trays are suitable containers for forming the blocks since they are toluene permeable for speedier evaporation and flexible for easy removal

Botanical Microtechnique and Microtomy 53

(iii) (iv) (v) (vi)

of the hardened blocks. Keep the containers in the oven at 58°C for 48-72 hrs; toluene evaporates, and blocks become solid but not completely dry. Blocks should be transparent and colourless, and the embedded tissue translucent. Remove the containers from the oven and cool them, and store them in the refrigerator, if desired. Remove the block from the container and cut it into pyramidal pieces containing tissue fragments with a razor blade. With the help of a drop of polystyrene solution or with Plexiglass affix each pyramid to a rigid support.

Frangioni and Borgioli (1979) found that toluene is the most suitable solvent for embedding solution when compared to others such as benzene which shows high toxicity while xylene, chloroform, amyl acetate, ethyl acetate, and other solvents have a slow rate of evaporation. Benzyl alcohol increases the penetration of polystyrene into the tissues and dibutyl phthalate renders the block plastic which facilitates smooth cutting. 3.3.1.4 Frozen Section Technique

The frozen section technique is based on the principle of freezing the tissue directly to harden it, and cutting the sections while the tissue is frozen. This technique has several advantages. These are: (a) it is a time-saving method as compared to paraffin and celloidin methods; (b) since the tissue is not dehydrated, the cells retain a life-like appearance with little shrinkage; and (c) tissues can be -sectioned directly without any fixation. However, this technique has some drawbacks such as the serial sections cannot be taken and there is no satisfactory process of holding loose tissues before freezing. Special models of microtome are available to cut the frozen sections. In older models, ether was used for freezing and the tissue was usually soaked for some hours in gum, dextrin or sugar solutions to prevent ice crystal formation. In the modern technique, CO2 jets are used on the microtome knife for cooling. The sections are affixed to the slide by using some glue. A freeze-drying apparatus, devised with liquid nitrogen, permitted section cutting within 5 hrs after the fresh tissue was obtained (Sharma and Sharma, 1972). Another microtome is designed for cutting frozen sections in which the knife moves across the tissue, in contrast to the sledge type in which the tissue is moved while the knife is stationary. In a modification for obtaining thin sections from unfixed (fresh) tissues for histochemical staining, a microtome with an apparatus for simultaneous cooling of the knife with the freezing stage is used. For cutting frozen sections on a paraffin microtome, the specimen is fixed on the object holder in a drop of water by freezing it in dry ice in a box. Chips of dry ice are wedged between the metal disc and the object clamp of the microtome.

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Plant Techniques: Theory and Practice

3.3.2

Sectioning and Microtomy

For section cutting, rocking microtome for celloidin sections and rotary microtome for paraffin sections are recommended. Generally, modern microtomes are precision instruments that have been designed to cut the sections uniformly for detailed examination under the microscope. Microtomes can be classified into manual microtomes, semi-automatic microtomes and automatic microtomes (Mohammad and Mohamed, 2012). Manual microtomes, such as rocking microtome, rotatory microtome, sledge microtome, freezing microtome, vibrating microtome, cryostat, saw microtome, and hand microtome whereas automatic microtomes including laser microtome, computer microtomes and ultra-thin computer microtome are all commercially available. Since the variety of microtomes available in the market selecting the right instrument is very important for producing the best results we expected. Some of the commonly used microtomes are described below: 3.3.2.1 Rotary Microtomes

The rotary microtome was originally designed by Late Professor Minot and is sometimes called the Minot microtome. These microtomes are manufactured by various companies, viz., Leitz, Reichert, Spencer, Measuring and Scientific Equipment (M.S.E.), Baird & Tatlock, etc. Rotary microtome is made with stationary knife holders and a moving block holder clamp. The adjustment which controls the thickness of sections is usually situated on the front panel of the microtome (Fig. 3.4). These machines are heavy and ideal for cutting hard tissues and taking serial sections. The section thickness of 0.5 to 0.6 µm can be taken with the rotary microtomes. Sectioning of paraffin wax embedded tissues is normally cut within the range from 3-5 µm whereas resin sections are attained between 0.5 to 1 µm (George and James, 2004).

Fig. 3.4. Rotary Microtome.

Botanical Microtechnique and Microtomy 55

3.3.2.2 Sledge Microtome

Sledge microtome is used for cutting celloidin sections and routine paraffin sections. The two most popular machines of this type are manufactured by Leitz Ltd. (London) and M.S.E. (London). These microtomes are also known as base sledge microtomes. They are heavier and fitted with a heavy sledge carriage carrying block holder which is movable whereas the knife is stationary, held by two clamps. The block-holder is mounted on a steel carriage that slides backward and forward against a fixed horizontal knife. The knife-holding clamps are adjustable and allow the tilt and the angle (slant) of the knife of the block to be adjusted easily (Ellis, 2000). While cutting on a Leitz microtome the block is raised automatically when the carriage is moved forward and moves a short lever fitted on one side of the base (Fig. 3.5). In the MSE model, the block is lifted when a handwheel is turned manually. The thickness of sections can be set at 1-2 µ. Since the sledge micro tomes are heavy and rigid, these are good for cutting the hard tissues.

Fig. 3.5. Leitz Base Sledge Microtome.

3.3.2.3 Ultramicrotome

For ultrathin sectioning (0.1-0.01 µm), several models of ultramicrotome are currently available. The earliest attempt was made in ultrathin sectioning by Von Ardenne dating back to 1939 (Sharma and Sharma, 1972). Wachtel et al. (1966) gave a detailed historical account and principles of various ultramicrotomes. The first suitable model was manufactured by Ivan Sorvall Inc. in 1953 commonly known as the Porter-Blum microtome (Sorvell-MT

56

Plant Techniques: Theory and Practice

-1). The principle of its operation was based on a system of screw thread, lever arm and proper bearings. Due to its simplicity of working, it did not require a complicated maintenance process. In the slightly improved model, the specimen holder, which is of a collet type screwed at the free end of the aluminium rod, is moved vertically across a glass or diamond knife. On the return stroke, the specimen end of the aluminium rod is allowed to follow the trajectory of a parallelogram. Coarse and fine adjustments are provided for thick sectioning whereas for ultrathin sections, a mechanical device is arranged to advance the block towards the knife. To induce thermal expansion of the aluminium rod, an electric lamp is used. In the latter model viz., Sorvall-MT-2, the whole operation is much more compact; the knife stage allows controlled motion of the knife, permitting the proper cutting of ultrathin sections. Two controls are provided to adjust the thickness of sections between 100A to 4 I-1m. In the Huxley model (1959, Cambridge Instrument Co.), steel leaves are employed for hinging the arm. It is based on a mechanical advance with a double-leaf spring suspension system. In this, though the gravitational pull is responsible for section cutting, one oil-filled dashpot controls the rate of downward movement of the cutting arm (Watchtel et al., 1966). In the ultramicrotome of LKB-Producter AB-Stockholm, fluctuation in thickness of sections is eliminated to a significant extent; a thermal advance system is operating in this instrument. A cantilever arm is the principal moving part, one end of the arm holds the specimen block, and the other end is attached to a leaf spring joined to the base of the microtome. This spring causes the up and down motion of the bar. Thermal control of the cutting arm guides the advance of the block against the knife. The gravitational force controls the cutting stroke, and a motor regulates the motion and the upward movement. An electromagnetic force, which acts during the return stroke, causes the flexing of the base below the knife holder necessary to ensure the by-pass of the cutting surface and knife-edge during the stroke. Sections can be cut as thin as 10 nm with the help of DuPont Sorvall JB-4 microtome, USA. 3.3.2.4 Cryostat Microtome

Cryostat microtome is a device used to maintain low cryogenic temperatures, under which samples can be put and taken into the sections (Fig. 3.6). A deep freeze cabinet in which microtome (rotary type) is fitted. The cabinet is also equipped with a fluorescent light and a fan to ensure the circulation of cool air. The temperature is regulated between -10oC to -40oC. This can be used as an alternative to a freezing microtome for rapid sectioning of 2-16 µm thickness of sections. Block holders, on which tissue is frozen, are transferred to the microtome where the cutting is performed. After sectioning, sections are taken onto the warm microslide, then staining and mounting are performed more rapidly. Cryostat microtome has been used mostly for animal tissues

Botanical Microtechnique and Microtomy 57

and plant tissues as well. The microtome may be adjusted to cut sections at the range of 2-16µm (Bancroft and Gamble, 2008).

Fig. 3.6. Cryostat Microtome.

3.3.2.5 Computerised Microtome

The computerised microtome is equipped with an advanced rapid thermostatic switch, semiconductor freezing, cryoscalpel and cryoplate (Fig. 3.7). This microtome can perform the rapid freezing sectioning or routine paraffin sections (dual purpose). The temperature of cryoscalpel and cryoplate is ranging from 0ºC to -18ºC and -10 ºC to -45ºC respectively. It can take sections of 1-25 µm thickness.

Fig. 3.7. Computerised Microtome.

A variety of microtomes and knives are commercially available in the market and choosing the appropriate microtome is of prime importance to achieve good results without distortion of tissue of the biological samples.

4 Stains (Dyes) Stain is a chemically defined substance that is used to colour biological specimens, whereas dyes are crude preparations. In other words, stains are specified dyes used to make the structural details of cells and tissues more precise and accurate. In biochemistry, a variety of stains has been used to stain the DNA, Proteins, lipids, and carbohydrates. Classification of stains: Stains are classified based on their source and nature. Some stains are derived from mineral sources. For example, Ruthenium red (for Pectin stain) is an ammoniated oxychloride of ruthenium. Organic stains are divided depending on whether they are artificially synthesised or obtained from some natural sources. Most of the synthetic dyes are coal-tar dyes and these are again divided based on the molecular configuration of their chromophores which are responsible for giving colouration. Major kinds of stains are given below: 1. Nitroso stains: In these stains’ nitroso group (-NO) attaches to benzene ring [(C6H6); Fig. 4.1A], e.g., Fast green O, Naphthol green B, etc. 2. Nitro stains: The nitro group (-NO2) is a chromophore, attached to benzene ring (Fig. 4.1B), e.g., Picric acid, Aurantia, Naphthol yellow, etc. 3. Azo stains: The azo group (-N = N-) is found in these dyes (Fig. 4.1C). All the dyes of this group are the derivatives of azobenzene. 4. Quinone-imine stains: In the quinone (C6H4O2), two bivalent oxygen atoms may replace two monovalent hydrogen atoms (Fig. 4.1D). In these stains, quinoid benzene ring is attached to nitrogen atom. i. Indamines: In the indamine group (-N=), two benzene rings are attached to the nitrogen atom; one of these being in the quinoid form (Fig. 4.1E), e.g., Bindschedler’s green, Toluylene blue, etc. ii. Indophenols: Two benzene rings are attached to nitrogen atom; one of these benzene rings is phenolic and the other is in quinoid form (Fig. 4.1F). iii. Thiazins: In these, two benzene rings are further joined together by a sulfur atom, forming three closed rings of atoms (Fig. 4.1G). iv. Oxazins: (Fig. 4.2A) – e.g., Gallocyanin, Nile blue sulphate, Celestian blue B.

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A. Nitroso phenol

B. Nitro phenol

C. Azobenzene

D. Quinoid benzene

E. Indamine

F. Indophenol

G. Thiazin

Fig. 4.1. A-F. Molecular configuration of stains.

5. Azins: A typical molecular configuration is given in Fig. 4.2B. v. Amino azins, e.g., Neutral red, Neutral violet, etc. vi. Safranins, e.g., Safranin O, Azocarmine O, etc. vii. Indulins, e.g., Fast blue and Nigrosin W. 6. Phenyl methane stains: In these, methyl group is attached to benzene ring. i. Diphenyl methanes (Fig. 4.2C), e.g., Auramine O. ii. Diamino triphenyl methanes, e.g., Fast green FCF. iii. Triamino triphenyl methanes - e.g., Acid fuchsins, basic fuchsins, aniline blue, methyl green, etc. iv. Hydroxy triphenyl methanes - e.g., Aurin. v. Diphenyl naphthyl methanes - e.g., Victoria blue R and nile blue. 7. Methane stains i. Pyronins (Fig. 4.2D) - e.g., Acridine red 3B, pyronin Y and pyronin ii. Rhodamines - e.g., Rhodamine B and rhodamine 6G. iii. Fluorescein derivatives - e.g., Fluorescein isothio-cynate, Cosin and erythrosin.

Stains (Dyes)

A. Oxazin

61

B. Azin

C. Diphenyl methane

E. Anthraquinone

D. Pyronin

F. Thiazole

G. Quinoline group Fig. 4.2. A-G. Molecular configuration of stains.

iv. Phthalein derivatives (Phenolphthaleins and sulpho-naphthaleins) e.g., phenolphthalein, bromothymol blue and bromophenol blue. v. Acridines - e.g., Acridine orange, acriflavine and phosphine 3R. 8. Anthraquinone stains: The anthraquinone dyes include derivatives of anthracene, through its oxidation product anthraquinone (Fig. 4.2E). 9. Thiazole stains: A small group of dyes of rather complex formula contain the thiazole ring (Fig. 4.2F). 10. Quinoline stains (Fig. 4.2G) Quinoline yellow, pynacynol, etc. 11. Phthalocyanine: These are complex chemical compounds, similar in structure to chlorophyll with C6H4C2N2 surrounding a central metal atom, usually copper.

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Based on the type of study of the plant material, there are large number of stains under use. For example, a combination of safranin and fast green (double staining) is prepared to study the anatomical and embryological aspects, crystal violet for chromosomal studies, methylene blue for bacteria and delafield’s haematoxylin with counterstain of eosin for zoological and clinical works. Most of the dyes used nowadays in biological techniques are prepared from coal tar.

4.1 Natural Dyes Natural dyes are still used by botanists. These are derived from plants belonging to Caesalpiniaceae and from cochineal insects.

4.1.1

Natural Dyes from Plant Origin

A. Brazllin: This dye is obtained from ‘brazil wood’ especially from Caesalpinia crista and C. echiniata (Johansen, 1940). Chemically it is closely related to haematoxylin (Fig. 4.3A). It is neither so active nor so strong. It reacts only after mordanting with ferric ammonium sulfate. It is used to stain smears.

A

B

Fig. 4.3. A. Brazilin (C16H14O5), B. Brazilein (C16H12O5).

Brazilin 70% ethyl alcohol

0.5 g 100 ml

Allow the above stain to ripen for a week. Keep the container well stoppered and it should be away from light and air. Brazilin solution is colourless, but it becomes red when it is exposed to air and oxidised into the dye brazilein (Fig. 4.3B, Lillie, 1977). B. Haematoxylin: It is obtained from the heart woods of Haematoxylon campechianum Linn. In fact, it is a colourless solid extract, but it is readily oxidised by atmospheric oxygen and forms reddish acidic dye haematein (Baker, 1969; Lillie, 1977; O’Brien and McCully, 1981).

Stains (Dyes)

63

Haematoxylin is sold in a partially oxidised form as a brown powder and it is a mixture of haematoxylin and haematein. It is homologous to brazilin, but it contains additional hydroxyl group (Fig. 4.4A, B). Haematein is one of the most important stains in plant microtechnique since it forms complexes (chelates) with the complex cations formed when salts of iron or aluminium are dissolved in water. The chelates bind to acidic sites in the tissues so that chelates behave like a basic dye. These salts of iron and aluminium are known as mordants because they provide sites for dye attachment. The stained sites are most resistant to ethanol during post-staining dehydration. So, it is necessary to prepare the stain with salts like iron (in ferric form), aluminium and copper. Based on type of salts used, haematoxylin solutions are of the following five types.

A

B

Fig. 4.4. A. Haematoxylin (C16H14O6), B. Haematein (C16H12O6).

(i) Heidenhain’s iron haematoxylin: It is prepared in three ways: (a) Simply a solution of 0.5 g of haematoxylin dissolved in 100 ml of distilled water. (b) A stock solution is prepared by dissolving 10 g of haematoxylin in 100 ml of absolute alcohol. From this, 5% of solution in distilled water is prepared for staining. The solution is kept for few days to oxidise (ripe) into haematein. The ripening process may be hastened by placing the solution in very wide evaporating petridishes and exposing to a powerful quartz mercury-vapour arc for about 45 minutes. Petridishes are placed 2 ft away from arc and the solution is stirred frequently during exposure. This stock solution may be spoiled at higher temperatures. Metallic film is formed on the surface of the solution as, it indicates spoilage of solution. It should be discarded when it turns to brown in colour. (c) The following solution is prepared which is more stable than above two:

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Haematoxylin 10% absolute alcohol Methyl cellosolve Distilled water Tap water containing calcium compounds

0.5 g 5 ml 100 ml 50 ml 50 ml

The solution is shaken to get rich wine-red colour. If the colour does not appear, add a pinch of sodium bicarbonate and shake vigorously. Ripening occurs immediately in this solution. It does not spoil at high temperature and may be stored for quite a long period. In Heidenhain’s iron haematoxylin, ferric ammonium sulphate is used as mordant. This stain is commercially available and gives good results in morphological studies. (ii) Delafield’s haematoxylin: In this, dye and mordant are combined. It is prepared in the following manner: Solution A Solution B

100 ml of saturated aqueous solution of aluminium ammonium sulphate. 1 g haematoxylin crystals added in 6 ml of 95% ethyl alcohol.

Now solution B is added to solution A drop by drop. It is exposed to light and air for four days and filtered afterwards. Later, 25 ml methyl alcohol and 25 ml glycerol are added. The solution is exposed to air for at least two months till the colour is sufficiently dark. It is filtered and stored in tightly stoppered bottle. (iii) Harris’ haematoxylin: It resembles the Delafields in its colour effects, but its preparation is entirely different: Haematoxylin crystals Aluminium ammonium sulphate 50% ethyl alcohol

5g 3g 1000 ml

Both dye and aluminium ammonium sulphate are dissolved in ethyl alcohol by heating. Later 6 g of mercuric oxide (red powder) is added and boiled for 30 minutes. It is cooled and filtered. Now it is brought to original volume of 1 liter by adding 50% ethyl alcohol. Later 10 drops of hydrochloric acid are added to acidify the solution. (iv) Mayer’s haematoxylin: It is recommended for nuclei of filamentous algae and fungi: Haematoxylin crystals 1g Distilled water 1000 ml Sodium iodide 2g Aluminium potassium sulphate 50 g Thymol (to prevent growth of mould) 1 crystal

Stains (Dyes)

65

The above ingredients are mixed thoroughly. Excellent results are obtained by diluting the stain 10 times with distilled water. This diluted stain is allowed to stand overnight and later it is used. (v) Ehrlich’s haematoxylin: It has been extensively used along with safranin for woody stems: Haematoxylin 1g Aluminium Potassium sulphate excess amount Glacial acetic acid 5 ml Glycerine 50 ml Absolute ethyl alcohol 50 ml Distilled water 50 ml The solution is prepared and kept in dark place until the colour becomes deep red or it may be ripened by exposing to quartz mercury lamp for 3 or 4 hours. Wood sections are initially stained with safranin (counterstain). Later these are washed with acid water and tap water. Now the sections are transferred to Ehrlich’s haematoxylin and allowed to remain for 5-30 minutes. Later sections are washed with 35% alcohol and passed through alcoholic series for dehydration.

4.1.2

Dyes from Animal Origin

Cochineal and its derivatives: Cochineal is a yellowish-red powder obtained by grinding the dried bodies of female cochineal insects (Coccus cacti), a tropical American Homoptera living on the plant Opuntia coccinellifera. Carmine is bright red in colour, and it is obtained by adding alum to cochineal. It is commonly used in cytological studies. The oxidised form of carmine is known as carmein. Another derivative of cochineal is carminic acid. It is used as an indicator, yellow in acid, red-purple in alkali (Long, 1961). Carminic acid (Michrome no. 214) is obtained by extracting cochineal with boiling water followed by treatment with lead acetate and decomposition of lead carminate with sulphuric acid (Gatenby and Beans, 1950). This dye belongs to the anthraquinone group and has the formula C22H20O13, the molecular weight being 492.38 (Fig. 4.5).

Fig. 4.5. Carminic Acid (C22H20O13).

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Plant Techniques: Theory and Practice

The chromophoric property is attributed to its quinonoid linkage and auxochromes are also present. It is soluble in water in all proportions (Gurr, 1960), and is a dibasic acid and claims to be nearly insoluble at its isoelectric point, pH 4-4.5 (Baker, 1950). If it is dissolved on the acid side of its isoelectric point, it acquires a positive charge, behaves like a basic dye and stains chromatin. But if it is dissolved in alkaline solution, it behaves as an acid dye (see also Sharma and Sharma, 1972).

4.2 Coal-Tar Dyes Some of the important coal-tar dyes are given below alphabetically by mentioning their chemical nature, acidic and basic nature of dye, and solubility in water and 95% ethyl alcohol.

4.2.1

Acid Fuchsin

Acid fuchsin is a acidic stain. It is a sulfonated derivative of basic fuchsin and belongs to triamino-triphenyl methane group (rosanilins). The generally accepted formula of one of the homologs present in acid fuchsin, namely the disodium salt of rosanillin trisulfonic acid, is C20H17N3O9S3Na2 (Fig. 4.6). A 1% solution in 70% alcohol is preferable and 0.5% to 1% solution in distilled water is equally good. It is used to stain the cortex, pith parenchyma, cell walls (cellulose), mitochondria, etc. If the stain is prepared in 70% alcohol, it is essential to differentiate (removal of excess stain) the tissue in a saturated solution of picric acid in 70% alcohol for a minute and then rinse in 70% alcohol until a bright red colour is obtained (Johansen, 1940). Post-staining dehydration is done very quickly because colour of stain vanishes in higher grades of alcohols.

Fig. 4.6. Acid Fuchsin (C20H17N3O9S3Na2).

Stains (Dyes)

4.2.2

67

Aniline Blue WS

Aniline Blue is a acidic stain (Fig. 4.7A, B). It belongs to triamino-triphenyl methane group (Lillie, 1969). In fact, the commercially available Aniline blue WS is a mixture of water blue and methyl blue (cotton blue). A 1% solution in 90% ethyl alcohol is preferred. Another method of its preparation is to dissolve enough stain in methyl alcohol followed by addition of clove oil and absolute alcohol as diluents. Then clear the material with clove oil and wash it with xylol and mount it in Canada balsam (Johansen, 1940).

A

B Fig. 4.7. A. Aniline Blue (C32H2N3O3S3), B. Methyl Blue (C37H27N3O9S3Na2).

It is served as a good counterstain with safranin for plant tissues. It is good for staining filamentous algae and mycorrhizal fungi.

4.2.3

Basic Fuchsin (Pararosanilin)

Basic fuchsin is a basic stain and belongs to triamino-triphenyl methane group. Its solubility in water is 0.26% and 5.93% in ethyl alcohol (see Table 4.1). A saturated aqueous solution is prepared for staining.

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It is a powerful nuclear dye and stains mucin (mucopolysaccharides) and bacteria.

4.2.4

Chlorazol Black E

Chlorazol black is an acidic dye of the trisazo group and has a molecular weight of 781.738 (Michrome no. 92). Its formula is C34H25N9O7S2Na2 (Fig. 4.8). It is highly soluble in water and sparingly in alcohol. Being an acidic dye, the basis of its stainability with chromosomes is not clear, but it is probable that it stains the protein component (Sharma and Sharma, 1972). A solution of chlorazol black E in alcohol has been applied by Nebel (cited in Sharma and Sharma, 1972) as an auxilliary stain for chromosomes, along with acetic-carmine, this dye was applied after fixation prior to aceticcarmine staining and proved effective for Rosaceae members where ordinary acetic-carmine stain was ineffective. It is employed singly and is effective for the study of root-tip chromosomes of plants.

Fig. 4.8. Chlorazol Black E (C34H25N9O7S2Na2).

4.2.5

Congo Red

Congo red is an acidic stain and belongs to azo group (Fig. 4.9). It is not soluble in water and its solubility in ethyl alcohol is 0.19%. A solution of congo red is prepared in weak alcohol. It should be used last when it is employed in combination with other stains. The post-staining dehydration of tissue is done as rapidly as possible.

Fig. 4.9. Congo Red.

Stains (Dyes)

69

Congo red is a cytoplasmic stain in plant technique. It also stains cellulose and mucin.

4.2.6

Crystal Violet (Gentian Violet)

Crystal violet is a basic stain (Fig. 4.10). This dye is a hexamethyl pararosanilin. Its solubility in water is 1.68% and in ethyl alcohol 13.87%. It serves as an indicator in the pH range 0.0 to 1.8 changing from yellow in the acid to the usual violet-blue of aqueous solutions (Eastman, 1966). A 1% solution in distilled water is prepared. It is better to use freshly prepared stain. Since the crystal violet is quickly washed out in dehydrating alcohols, one should either mordant the stain or dissolve the stain in clove oil. The clove oil solution is kept in a dropping bottle. Another method of preparation is to make a saturated solution in clove oil and add a few drops of this solution to a staining dish which is full of xylene. This mixture is very unstable; hence it needs addition of more drops of stain to maintain the strength of stain.

Fig. 4.10. Crystal Violet (C25N30N3Cl).

It is the best stain for cytological studies and widely used in bacteriology.

4.2.7

Erythrosin Bluish

Erythrosin B is an acidic dye (Fig. 4.11). It is a tetraiodo compound corresponding to the tetrabromo compound of typical eosin (Conn, 1936). Its solubility in water is upto 11.10% and in ethyl alcohol 1.87%. Botanist used this dye in place of eosin. Like eosin, it serves as a fluorescent indicator. It is also employed as a contrast stain for haernatoxylin and certain blue and violet nuclear stains. It is a good counterstain to crystal violet in certain cytological methods (Johansen, 1932). Use 1% solution in 95% alcohol or clove oil. It stains very quickly – 10 in clove oil and 3 mins in alcohol. It is an excellent stain for gelatinous sheath of Nostoc.

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Fig. 4.11. Erythrosin (C20H6O5I4Na2).

4.2.8

Fast Green FCF

Fast green FCF is an acidic stain and belongs to diarnino-triphenyl group (Fig. 4.12). Its solubility is 16.04% in water and 0.33% in ethyl alcohol. A 0.5% fast green is prepared in a mixture of equal parts of methyl cellosolve, absolute alcohol and clove oil (Johansen, 1940). A 0.5% fast green is also prepared in 95% ethyl alcohol (Sass, 1967) or in 90% ethyl alcohol. It is a dark-green solution and used in differential staining in combination with safranin.

Fig. 4.12. Fast Green FCF (C37H34N2O16S3Na2).

4.2.9

Fluorone Black

Fluorone black was previously used in acid alcoholic solution by Turchini et al. (1944) as a specific reagent for deoxyribonucleic acid. It was also used in ammonical solution by Lipp and Ratzenbock (1963) as a specific stain for

Stains (Dyes)

71

haemoglobin and erythrocytes. It serves as an excellent nuclear stain which reacts with basic nucleoprotein (see also Lillie et al., 1975).

4.2.10 Lacmoid Lacmoid (also known as resorcin blue) is a blue acidic dye of the oxazine seties (Fig. 4.13). Its empirical formula is C12H6NO3Na, the molecular weight being 235.173. It can be obtained by heating resorcinol with sodium nitrate until the smell of ammonia is no longer present. Like carmine, it can be used as an acid-base indicator and, when dissolved in acetic acid, it behaves as a basic dye. Unlike carmine, it is soluble in water and alcohol.

Fig. 4.13. Lacmoid (C12H6NO3Na).

Darlington and La Cour (1968) used it in place of carmine and acetic lacmoid solution, and it has been found to be very effective for the chromosomes of root-tips, embryo sacs and pollen grains. For comparatively compact tissues like root-tips, heating in acetic lacmoid-HCl mixture is needed prior to squashing for dissolution of the middle lamella.

4.2.11 Light Green SF Light green SF is a derivative of brilliant green, which is sulfonated; hence it is an acidic dye. It belongs to diamino-triphenyl methane group. Its solubility in water is 20.35% and in alcohol 0.82%. A 0.2% solution is strong enough to stain the tissue. Staining is rapid and, therefore, it should not be allowed to react for long time. In differential staining, this stain reduces safranin. Hence it should not be allowed to react for too long. There are many methods of Light green SF preparation (Johansen, 1940): i. Light green SF solution is prepared in 95% ethyl alcohol. ii. The stain is dissolved in absolute alcohol and diluted with clove oil. iii. A saturated solution of light green SF is prepared either in water or alcohol and later it is acidulated with hydrochloric or acetic acid. It works as good differential stain for lignified cell walls. Light green SF is a good cytoplasmic stain, extensively employed for cellulose walls and some filamentous algae. By combining this stain with alcoholic Sudan IV, suberised and cutinised tissues are differentiated from lignified tissues.

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4.2.12 Malachite Green (Emerald Green or Light Green E) Malachite green is a weak basic stain, and it belongs to diamino-triphenyl methane group. A 0.5% solution is prepared in distilled water or ethyl alcohol. Its chemical formula is C23H252Cl2 and molecular weight 364.926 (Fig. 4.14). It is used singly or in combination with 1% aqueous safranin. If it is used singly, the specimen is kept in this stain for one minute. If double, stain is used, the specimen is kept in this stain for 20 seconds followed by 20 minutes in safranin.

Fig. 4.14. Malachite Green (C23H25N2Cl2).

Malachite green stains all histological elements such as cell walls, endodermis, cytoplasm, nuclei and chloroplasts. Double staining is used to study the pathological specimens.

4.2.13 Methyl Green (Light Green) Methyl green is a basic dye. Its chemical formula is C26H33N3Cl2 and molecular weight 458.488 (Fig. 4.15). Combining with acid fuchsin, it is

Fig. 4.15. Methyl Green (C26H33N3Cl2).

Stains (Dyes)

73

useful for staining the xylem. Generally, 1% aqueous solution is prepared for staining. It stains chromatin, nuclei and lignified tissues.

4.2.14 Methylene Blue (Loeffler’s Methylene Blue) Methylene blue is a basic dye and theoretically it is a tetramethyl thionin (Fig. 4.16). Its solubility in water is 3.55% and in alcohol 1.48%. The methylene blue of commerce is generally a double salt, the chloride of zinc and methylene blue (Lillie, 1977). However, zinc salt is toxic, so zinc-free methylene blue chloride is recommended for medicinal purposes. The zinc double salt is less soluble, particularly in alcohol, so it is less desirable: Methylene blue 1% Potassium hydroxide solution Alcohol Distilled water

0.5 g 1 ml 30 ml 100 ml

Fig. 4.16. Methylene Blue (C16H18N3SCl).

Slightly warm water is taken to which add methylene blue and stir it. Other ingredients are added later, and solution is filtered for use. Methylene blue is an important bacteriological stain and nuclear stain. It also stains yeast (Clayden, 1962).

Acidic/ Basic

Molecular Solubility Weight

Acid fushsin

Acidic

585.560

Soluble in water and in ethanol

Acridine

Basic

302

Aniline blue

Acidic

Auramine O

Alsborption maxima (nano-meters)

Diachrome (or) Fluorochrome

Metachromatic

Important Biological substances stained by the dye

546

Fluorochrome giving red fluorescence

Metachromatic

Cell walls (cellulose), mitochondria

Soluble in water (5%) and glycerol (4%) at 15oC

497

Fluorochrome

Metachromatic

DNA, RNA

737.756

Water soluble

600

Giving yellow fluorescence

--

Callose

Basic

303.844

Soluble in ethanol at 26oC

434

Fluorochrome giving yellow green fluorescence

--

Cutin, waxes

Azure B

Basic

305.840

Soluble in water, sparingly in ethanol

648

Diachrome

Basic fuchsin Pararosanilin

Basic

323.834

Soluble in water (0.26%) and in absolute alcohol (5.93%) at 26oC

545

Magenta I

Basic

337.861

Soluble in water (0.39%) and in absolute alcohol (8.16%)

550

--

Diachrome

Metachromatic Nucleic acids, lignin, chitin, etc. --

--

--

Polysaccharides, nucleic acids, lignin

Plant Techniques: Theory and Practice

Dye

74

Table 4.1. Properties of Some Common Stains

Magenta II

Basic

351.888

Bromophenol blue

Acidic

581.101

Chlorazol black E

Acidic

782

Congo red

Acidic

696.696

Crystal violet

Basic

Eosin B

--

554 585-590

Diachrome

--

Proteins

Soluble in methyl cellosolve

598-602

Diachrome

--

Cellulose, hemicelluloses

Soluble in ethyl alcohol, insoluble in water

497

Diachrome

--

Cytoplasm, cellulose, mucin

407.996

Soluble in ethyl alcohol (8.75%), water (9%) and methyl cellosolve (7.5%)

590

Diachrome

--

Cytological studies

Acidic

624.098

Water soluble at 15oC

516-519

Diachrome

--

Used as counterstain for haematoxylin

Erythrosin B

Acidic

879.890

Soluble in water (11.10%), glycerol (12%) and ethyl alcohol (1.875)

15-520

Diachrome

--

Gelatines sheath of Nostoc

Fast green FCF

Acidic

808.87

Water soluble, sparingly soluble in ethanol

622-228

Diachrome

--

Proteins, cellulose, histones

75

(Contd.)

Stains (Dyes)

Soluble in water, methanol, ethanol and benzene

Acidic/Basic

Molecular Weight

Solubility

Alsborption maxima (nano-meters)

Diachrome (or) Metachromatic Fluorochrome

Important Biological substances stained by the dye

Haematoxylin Basic

356.336

Soluble in water (10%) ethanol (10%), glycerol (20%)

440

Diachrome

--

DNA, Phospholipids

I acmoid (Resorcin blue)

Acidic

235.173

Soluble in organic solvents and in water

611

Diachrome

--

Callose

Malachite green

Basic

364.926

Soluble in water (105, clove Oil (10%), alcohol (8.5%) and cellosolve (8.5%)

617-619

Diachrome

--

DNA

Methyl green

Basic

--

629

Giving and fluorescence

--

DNA

543-556.5

Giving pink fluorescence

--

Lipids

520

Diachrome

Metachromatic (orange)

Water soluble

Rhodamine B Basic

479.029

Soluble in ethanol (1.5%) and water (0.8%) at 26oC

Safranin O

350.861

Soluble in ethyl alcohol (3%), cellosolve (5%) and glycerol (3.5%)

Basic

Lignin, suberin, chromosomes, nucleoli, centrosomes, etc.

Plant Techniques: Theory and Practice

Dye

76

Table 4.1. (Contd.)

Sudan dyes Sudan IV

Acidic

380.456

Soluble in ethyl, alcohol (0.09%); insoluble in water)

522-529

Diachrome

--

Fats

Acidic

422.492

--

--

Diachrome

--

Myelin, lipofuscins, ceroid, chromate retinal lipids and frozen sections of adrenal, cortex lipids

Oil and EGN

Acidic

394.483

Soluble in acetone, benzene and toluene, ethyl alcohol, linseed oil; sparingly soluble in ethanol, insoluble in water

--

Diachrome

--

Histological stain

Thrinine

Basic

263.759

Soluble in both water and alcohol (1%), and cellosolve (10%)

610

Diachrome

Metachromatic

Chromosomes of animal cells

Toluidine blue O

Basic

305.840

Soluble in water and ethanol

626

Diachrome

Metachromatic

Nucleic acids, lignin, phenolics, sulphonated and carboxylated polysaccharides

Stains (Dyes)

Acetyl sudan IV

77

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4.2.15 Rhodamine B Rhodamine B is a basic dye. It is similar to pyronins (characterised by presence of two alkylated amino groups parallel to the central methane carbon on the two benzene rings) except that there is a third benzene ring attached to the third carbon atom and attached to this ring is a carboxyl group in the ortho position (Fig. 4.17). Its solubility in alcohol is 1.47% and in water 0.78% (Table 4.1). It is used to stain glandular tissue, epidermis and golgi apparatus (Lille, 1969).

Fig. 4.17. Rhodamine B.

4.2.16 Ruthenium Red Ruthenium red is an ammoniated ruthenium oxychloride prepared by making an ammonical solution of ruthenic chloride, RuCl3 (Lillie, 1977). It is employed with or without addition of acetic acid to stain the pectin. It is thought to be specific for this purpose. But Jensen (1962) opined that the method is rather insensitive and less specific than the hydroxylamine-FeCl2 reaction. It is formulated as (NH3)n.Ru-o-Ru-o-Ru(NH3)n and as Ru2(OH)2 Cl4·7NH3·3H2O.

4.2.17 Safranin O The common safranins of commerce are mixtures of dimethyl and trimethyl phenosafranin. The type safranin O proves to be the best biological dye. It is a basic stain and dissolves easily in strong alcohol than in water. The chemical configuration of safranin O is given in Fig. 4.18. Various preparations of safranin solution have been followed by different workers. i. A common preparation is to dissolve 1 g safranin in 70% ethyl alcohol. It is widely used by botanists. ii. A stock solution is prepared by dissolving 2.25 g safranin in 225 ml 95% ethyl alcohol. A part of this solution is diluted with equal volume

Stains (Dyes)

79

of distilled water and used for staining. If it is still concentrated, again dilute it with 50% alcohol. iii. Johansen (1940) prepared safranin stain in the following manner. 4 g safranin is dissolved in 200 ml methyl cellosolve. Later 100 ml 95% ethyl alcohol and 100 ml distilled water are added. Then it is followed by an addition of 4 g sodium acetate and 8 ml formalin. Here acetate intensifies the stain and formalin works as a mordant. Slides are kept in the stain for 24-48 hours to get good contrast. Invariably safranin overstains the tissue and therefore it requires destaining (differentiation). It is destained by HCI dissolved in 70% ethanol. After tissues are stained with safranin, the excess stain should be washed away with tap water.

Fig. 4.18. Safranin O.

Safranin is an excellent stain for morphological and cytological studies. It stains lignin, cutin and suberin as well as chromosomes, nucleoli and centrosomes. It is also used as a protein stain and can be used to stain spore coats (Lillie, 1977).

4.2.18 Sudan Dyes Sudan IV is a weak acidic dye and belongs to the azo group. It is insoluble in water but its solubility in ethyl alcohol is 0.09%. It is a specific fat stain. Specimen is stained in saturated alcoholic dye solution for about 10 mins. Since alcohol is fat solvent, specimen should be washed very rapidly in alcohol. Lecithin, resins, latex, wax and cuticles are stained by Sudan IV; chloroplasts are stained a dull red (Johansen, 1940). Acetyl Sudan IV is made in the laboratory by acetylation in 40% acetic anhydride pyridine mixture, precipitation in water and collection by filtration. Its chemical formula is C26H22N4O2 and molecular weight 422.494. It is an excellent dye for myelin, lipofuscins, ceroid, chromated retinal lipids and frozen sections of adrenal cortex lipids, but failed to stain neutrophil leucocytes (Lillie, 1977).

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Oil Red EGN is also a close relative of Sudan IV. Oil red OB and oil red N-1700 are coming under oil red EGN. The dye oil red N-1700 is now formulated as o-tolylazo-2,5-xylylazo-2-naphthol. The chemical formula of oil red EGN is C25H22N4O and molecular weight 394.483. Oil red EGN is insoluble in water, slightly soluble in ethanol and in paraffm, soluble in ethyl acetate, linseed oil, oleic and stearic acids and turpentine, and very soluble in acetone, benzene and toluene (Lillie, 1977).

4.2.19 Thionin Thionin consists of two amino groups; hence it is a strongly basic dye and belongs to thiazin group (two benzene rings joined by a sulfur atom) (Lillie, 1977). Its chemical formula is C12H10N3SCl and molecular weight 263.759. Its solubility in both water and alcohol is 0.25%. Due to its metachromatic properties (the ability to impart different colours to different histological and cytological structures), it is widely used to stain animal tissues. Saturated aqueous solution or weak alcohol solution is employed to stain chromosomes (Johansen, 1940).

4.2.20 Toluidine Blue O Toluidine blue O is closely related to thionin and to methylene blue in structure, and to azure A (Lillie, 1969). Its chemical formula is C15H16N2S2Cl and molecular weight 305.840 (Fig. 4.19). Its solubility at 26°C water is 3.82% and in alcohol 0.57%. It is a valuable nuclear stain and employed in 0.3 to 1 per cent aqueous solutions. It is used as a chemical reagent in the standardisation of heparin and as histochemical reagent in determining DNA. It is also used for staining the cells, cartilage and certain acid mucins.

Fig. 4.19. Toluldine Blue O (C15H16N2S2Cl).

4.3 Other Substances Acting as Stains 4.3.1

Iodine

Iodine in combination with potassium iodide gives blue colour to starch. The colour reactions of iodine on fresh sections are as follows (see also Johansen, 1940):

Stains (Dyes)

Biological material Alkaloids Callose Cellulose Cork Cutin Inulin deposits Pectin Proteins Saponin Starch

4.3.2

81

Colour Brown Yellow Brown Yellow Yellow Brown Yellow Brown Blue Blue

Phloroglucinol

Phloroglucinol is not a dye. It is a trihydric phenol isomeric with pyrogallol and hyodoxyquinonol (Gurr, 1965). Aqueous phloroglucinol solution with few drops of conc. HCl gives pink colour to xylem but this colour fades soon. In plant science, various biological stains have been employed to study the structure of plant cell and to detect the secondary metabolites, such as proteins, fats, mucilage, etc. As different stains react to different parts cell tissue, they are useful in the identification of specific areas within the biological material which needs focus (Table 4.2). Table 4.2. Biological Materials and various Dyes Biological materials

Dyes

Bacterial cell wall

Graim stain; positive-Staphylococcus, Streptococcus Bacillus subtilis (Crystal violet – Negative E. coli, Nisseria

Cutin

Acid fuchsin, Auramina O, Crystal or Methyl violet, Erythrosin, Methyl green, Methylene blue, Safranin (specific)

Cellulose

Acid fushcin, Aniline blue, Bismarck brown Y. Calcofuor MZR, Chlorzol black E, Congo red, Delafield’s haematoxylin, Fast green FCF, Light green

Lignin

Crystal violet, Iodine green, Methyl green, Methylene green, Phloroglucinol, Safranin

Suberised cell walls

Safrinin, Sudan III or IV (specific)

Middle lamellae

Iron haematoxylin, Ruthenium red (specific)

Collogen and connective tissue

Methyl blue

Cutin

Safranin

Plant mucin

Bismarck brown Y, Congo red, Pararosanilin

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Cytoplasm

Acid fuchsin, Aniline blue, Eorsin Y, Erythrosin B, Fast green FCF, Indigocarmin, Light green, Malachite green, Methyl orange, Nigrosin, Orange G, Phloxine

Nuclear material

Acridine orange, Carmine, Cresyl violet, Crystal and Methyl violets, Haematoxylin, Iodine green, Methyl green, Methylene blue, Pararosanilin, Safranin, Thionin, Toluidine blue O

Chromosome

Brazilin, Carmine, Carminic acid, Haematoxylin, Iodine green, Methyl green, Safranin

Callose

Aniline blue, Azure B, Resorcinol blue (specific)

Mitochondria

Acid fuchsin, Aurantia, Crystal violet, Iron haematoxylin, Janus green B9vital)

Plastids

Crystal or methyl violet, Iron haematoxylin

Fats

Sudan III or IV (specific)

Proteins

Acid fuchsin, Aniline blue-black, Comasie blue R, Safranin

Laticifers

Dansyl choride, Oil red O

5 Staining Techniques When a specimen is stained with a particular dye, its constituents react differently with various structures of the specimen. There is a differential uptake and varying intensities of colouration that serve to distinguish the various structures of the specimen. This can be achieved by selecting a proper dye or combination of dyes. A dye is an unsaturated aromatic organic molecule that possesses one or more chromophores which are responsible for giving colouration when the specimen is observed under a microscope. Venkataraman (1952) introduced the term ‘Colligator’ to avoid confusion about auxochrome and chromophore. Both auxochrome and chromophore are responsible for giving colouration. Colligator is a special kind of auxochrome which is able to unite chemically with tissues or with certain other dyes (Gurr, 1965). Based on the type of colligators present, dyes are of two types: (A) Anionic dyes contain acidic colligators (negatively charged) such as OH, COOH, SO3, etc. These are also known as acidic dyes, e.g., fast green FCF, aniline blue. Acid dyes are attracted by positively charged objects in the specimen. Such objects are known as acidophil because they are easily stained by acid dyes (Baker, 1958). In other words, the objects of basic nature are stained by acid dyes. Dyes having both acidic and basic colligators may be regarded asamphoteric, that is, capable of combining with both acidic or basic elements, of the tissues. Certain acid dyes are amphoteric and stain the acidic as well as basic elements of tissues without the aid of mordants. Gurr (1965) called these acidic dyes pseudoamphoteric, with respect to their reaction towards tissue elements. (B) Cationic dyes contain basic colligators (positively charged) such as NH2, NH, N+, e.g., safranin, crystal violet, haematoxylin, methyleneblue, etc. These are also known as basic dyes which are attracted by negatively charged objects in the specimen. Such objects are called ‘basiphil’ which means objects that are acidic in nature and stained by basic dyes (Baker, 1958).

5.1 Types of Staining There are mainly three types of staining viz., Progressive staining, Regressive staining and Vital staining.

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5.1.1

Progressive Staining

When a specimen is stained with a dye, the intensity of staining progressively increases from time to time. This can be noticed under a microscope. After reaching the desired intensity, the dyeing is arrested. This is known as progressive staining. Phloroglucinolin combination with 5% conc. HN03 is the best example for progressive staining which is used to stain lignified cell walls.

5.1.2

Regressive Staining

The tissue is over-stained and later it is destained or differentiated with some other reagent until a satisfactory intensity is reached. It is called regressing staining and is a common practice in most of the staining schedules.

5.1.3

Vital Staining

When some non-toxic dyes are used in dilute concentration to stain living tissues, it is called vital staining.

5.2 Differentiation Differentiation of tissue should be done when the stain is used regressively. This means, if the tissue is over-stained, its excess stain is removed (destained) by other reagents. This process is referred to as differentiation. Acidified alcohol (2% HCl dissolved in 70% ethyl alcohol) is used as differentiating agent after staining with alcoholic safranin. Crystal violet and Heidenhain’s haematoxylin are differentiated with clove oil and iron alum respectively.

5.3 Mordants A mordant is a substance that brings about a particular staining reaction. Mordant comes from the Latin word “mordere” which means “to bite”. Mordant is used to make a dye to ‘bite’ the tissue or stain the tissue. There are salts of different metals which help the tissues to retain the stain. The use of mordants is called mordanting. The greatest advantage of mordanting is that the colour is not removable by neutral fluids, whether aqueous or alcoholic (Baker, 1969). The chief mordants used with carminic acid and haematein are the salts of aluminium and of ferric iron. Some of the mordants which are commonly used are acetic acid, barium chloride (2-4% solution) and silicotungstic acid (4% aqueous solution). Barium chloride is used as a mordant for acid dyes and silicotungstic acid for basic dyes. Other mordants are aluminium hydroxide (upto 3% solution), aluminium potassium sulphate (potash alum) (upto 4%

Staining Techniques 85

solution), ammonium chromate (upto 4%), ammonium dichromate (upto 3%) and Potassium permanganate (1%). Keep the tissue in these mordants on the slide for about 5-10 minutes. Later the mordants should be washed thoroughly in water before staining. It has been found that the addition of lithium is responsible for improving the action of basic dyes; iodine and picric acid are used as mordants for violet dyes viz. crystal violet and methyl violet (Johansen, 1940). Generally, mordants are used in three ways. These are as follows: i. Before staining: Ferric ammonium sulphate is used as mordant before staining the tissue with Heidenhain’s haematoxylin. ii. After staining: Picric acid and iodine are used after staining the tissue with violet dyes. iii. Along with staining: Mordant mixed with dye is employed to stain the tissues, e.g., aluminium ammonium sulphate mixed with Delafield’s haematoxylin and Harris’ haematoxylin; aluminium potassium sulphate in Mayer’s haematoxylin and Ehrlich’shaematoxylin. The selective retention of stain is influenced by the pH of washing fluids used for differentiation. In general, the basic dyes should be washed with acidified solutions and most of the acid dyes used for counterstaining should be washed with alkaline solution (Johansen, 1940).

5.4 Clearing (Bleaching) The sections cut from the plant material may be excessively blackened by the fixatives. For example, prolonged treatment of osmic acid (OsO4) turns the plant material into black. Some of the stems and roots of woody plants may contain resins and gums. These specimens should be bleached by some bleaching agents before staining. When xylene or benzene are used as clearing agents, the tissues become transparent. Hence this effect is called clearing. There are some tissues which are originally darkened, for example sclerenchyma and fibres do not need bleaching and these can be stained nicely by selecting a proper dye. One should take care about the action of bleaching agents. Sometimes it may damage the tissue. After reaching the desirable level of bleaching, the tissue should be thoroughly washed with water. Some of the bleaching solutions are given below: Hydrogen Peroxide Full strength or diluted hydrogen peroxide may be used. Dilute and prepare 50% hydrogen peroxide dissolving either in water or in 50% or 70% ethyl alcohol. It washes out blackening caused by osmic acid.

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Peroxide Ammonia Ammonia water accelerates the bleaching action of hydrogen peroxide (Johansen, 1940): Hydrogen peroxide (10% strength) 10 ml Water 200 ml Ammonia water 1 ml After bleaching is completed, the sections should be thoroughly washed with water. Sodium Hypochlorite A weak aqueous sodium hypochlorite solution is used to bleach the sections containing gums and resins. Potassium Permanganate If the sections are too blackened, these may be treated with 1% aqueous solution of potassium permanganate for a minute. Later, these sections are washed with 1% aqueous solution of oxalic acid and finally in water before staining. Chlorine Gas Chlorine gas fumes bleach the tissue excellently (Johansen, 1940). Enough crystals of potassium chlorate are taken in a coplin jar and a little dilute dhydrochloric acid is poured on it. As soon as greenish vapours appear, fill the jar with 50% ethyl alcohol. Then immerse the slides in the jar for 20 minutes to accomplish bleaching effect. Later the slides are thoroughly washed before staining. Stockwell’s Variation Stockwell’s procedure is employed for bleaching the sections which are too dark and contain too much phlobaphene. For example, root tips of Quercus, buds of Dudleya and other Crassulaceae members contain much phlobaphene. These sections should be first run down to water and bleached overnight in the following solution: Potassium bichromate Chromic acid Glacial acetic acid Water

1g 1g 10 ml 90 ml

The principle involved in this procedure is that the chromic acid dissolves the tannin, the acetic acid removes them, and the dichromate thereupon catalyses the tissues (Johansen, 1940). Now these sections are ready for sharp staining which is done in triple combination (safranin, crystal violet and orange G).

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Xylene Xylene is cheap and rapid in action. Sections become transparent and it removes the alcohol (de-alcoholisation) in the dehydrated tissues. However, it causes excessive shrinkage of delicate tissues. Since it does not dissolve the celloidin, xylene is also employed in celloidin sections (Clayden, 1962). Benzene Benzene has less hardening effect on tissues when compared to xylene (Clayden, 1962). Tissue becomes transparent with benzene treatment. It is highly inflammable and toxic. Other aspects are similar to xylene. Boric acid and ammonium persulphate are also used as bleaching agents. Chloral hydrate is also employed to clear the leaves.

5.5 Staining Schedules 5.5.1

Freehand Sections

5.5.1.1 Temporary Preparation

i. Before cutting the freehand sections, the material should be kept in 70% ethyl alcohol for five minutes and then the sections should be cut. If the material is already preserved in 70% ethyl alcohol, there is no need to keep it in 70% alcohol again; the sections can be directly taken. ii. Thin sections are stained with safranin (1 g safranin in 100 ml of 70% ethyl alcohol). iii. Destain the sections in acid water (3% HCl). iv. Then wash the stained sections with tap water to remove traces of acid. v. Mount in a drop of glycerine and observe under a light microscope.

Fig. 5.1. Water storage cell with multispiral cellulosic thickenings in Dendrobium bicameratum stem maceration.

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Macerations are made from the small pieces of stem sections (free hand sections). These small pieces of stem with 10% Potassium hydroxide are slightly boiled, and finally stained with only safranin. Cellulose thickenings in water storage cells found in the Dendrobium bicameratum are clearly seen with safranin staining (Fig. 5.1). 5.5.1.2 Permanent Preparations

Double staining is done for permanent preparation of freehand sections. This can be carried out in the following manner. A. Safranin-fast Green: This combination is widely used by botanists. It is suitable for all plant materials except algae (Johansen, 1940): i. Wash the plant material thoroughly to remove fixative and take thin and uniform sections. ii. Keep the sections in a watch glass and stain them with 1% safranin for 3-5 min. iii. Excess stain is removed by keeping the sections in acid water. iv. Then wash the sections with tap water to remove the acid traces. v. Pass the sections through graded series of alcohol from 30, 50, 75 to 90%. vi. Counter stain the sections in 0.5% fast green for 15 sec. It is a powerful stain and sections should not be kept in this stain for more than 15 sec. vii. Wash excess stain in a mixture of equal parts of absolute alcohol and xylene. viii. Clear the stained sections for few seconds in a mixture consisting of 50 parts of clove oil, 25 parts absolute alcohol and 25 parts xylene. ix. Transfer the stained sections in pure xylene. Give two changes in pure xylene. After becoming free from moisture, the sections are mounted in DPX or Canada balsam. Transverse section of leaf of Bulbophyllum khasyanum is taken and stained with safranin-fast-green, and finally made it as permanent slide. In this section, the adaxial epidermal cells that are 3 times larger than abaxial ones are clearly appeared with this double stain (Fig. 5.2)

Fig. 5.2. Upper Epidermal Cells (Hyaline) of Leaf in Bulbophyllum khasyanum(ade= adaxial; abe = abaxial).

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Frequently, milky foam appears when sections are transferred to xylene. It indicates that the sections are incompletely dehydrated or are so due to xylene exposed to air. If this happens, sections are transferred to absolute alcohol again and allowed to undergo complete dehydration. Care should be taken that xylene bottles are tightly screwed, and it should not be exposed to air. The safranin appears in brilliant red colour in nuclei, chromosomes, lignified and cutinised walls whereas fast green appears green in cytoplasm and cellulose cell walls. B. Safranin and Delafield’s Haematoxylin or Harris’ Haematoxylin: This combination is useful for semi-woody tissues (Johansen, 1940):Keep the material in 70% alcohol for 5-10 minutes, so that the material is smoothly cut. Later take uniform sections. i. Sections are over-stained with safranin. Excess stain is washed with tap water. ii. Destain the sections carefully in 50% alcohol slightly acidulated with hydrochloric acid and see that xylem appears bright red and cellulose walls deep pinkish in colour. iii. Wash thoroughly in water for 5 min. iv. Transfer the sections in haematoxylin and allow them to remain in it for 15 min till the sections turn deep purple in colour. v. Later treat the sections with acid water for few seconds. When the sections turn reddish, transfer to tap water. vi. Sections are thoroughly washed with tap water for about 20 min to remove traces of acid. If haematoxylin is not sufficiently blued by the tap water, the sections are dipped in the water which contains few drops of ammonia. vii. Then the stained sections are passed through the graded series of tertiary butyl alcohol i.e., 30, 50, 75, 85 and 90%, keeping them in each grade for 3-5 min. Later the sections are kept in absolute alcohol for 10 min. viii. Then transfer the sections in the mixture of alcohol and xylene (1:1) and keep them in this mixture for few minutes. ix. Later keep the sections in pure xylene. Give two changes in pure xylene and finally mount with Canada balsam. C. Zimmerman’s Stain: Zimmerman’s stain can be used to stain lignified tissues, it contains the following ingredients: Iodine green 0.1% aqueous solution Acid fuchsin saturated aqueous solution

9 parts 1 part

The uniform sections are kept in this stain for 10 mins. Later destain the sections in absolute alcohol containing 1.0% acetic acid and 0.l% iodine. Later

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sections are washed with absolute alcohol. Finally, these are rinsed twice with xylene and mounted in Canada balsam.

5.5.2

Microtome Sections

5.5.2.1 Paraffin Wax-embedded Tissues

Serial paraffin sections are taken on slides and staining is performed on staining dishes or coplin jars. Coplin jars are provided with grooves in which slides are kept. After the staining is completed, the slides are transferred to another container with enough graded alcohol or liquid for removal of excess stain, dehydration and clearing. During this entire process, the container should be kept closed with a lid so that the moisture of the atmosphere is not absorbed by the solution kept in the container and the solution may not be evaporated. A. Safranin-fast Green Safranin-fast green schedule is more preferred for all types of plant material except algae. The authors prefer to pass the sections through down and up series of different grades of alcohol. It is performed in the following manner: i. By using glue, paraffin sections are mounted on microslides and these slides are kept in the coplin jars containing xylene for 5 mins or until paraffin is dissolved (dewaxing). Here, do not keep slides back-toback in coplin jars; use a staggered arrangement which allows fluids to spread on both surfaces of slide. ii. Later paraffin dissolved slides are transferred to another coplin jar containing equal parts of xylene and alcohol (1:1) and keep them for 5 mins. iii. Keep slides in pure alcohol for 5-10 mins. iv. Now pass the slides through downward series of different grades of alcohol i.e., 90, 80, 70 and 50% for anatomical and embryological sections. v. Stain the sections in safranin (1 g in 70% alcohol) for 1-12 hr (depending on specimen) or in safranin prepared with methylcello solve and ethyl alcohol combination for 24-48 hr to get good contrast. vi. Wash excess of stain in water. vii. Now differentiate in 50 and 70% acidulated alcohol, and then transfer to 90% alcohol, 10 seconds in each grade; if slides have been left in stain for long period, it is necessary to wash in picroalcohol (90% alcohol containing 0.5% picric acid) for more than 10 seconds. viii. Counter-stain the sections in fast green (0.5 g in 90% alcohol) for 10 seconds. ix. Transfer the slides to 90% and then pure alcohol.

Staining Techniques 91

x. Later keep the sections in a mixture containing equal parts of xylene and alcohol (1:1). xi. Give two changes of pure xylene and mount in DPX or Canada balsam. Result: Single layered velamen in root transverse section of Bulbophyllum trimulum is lignified and clearly seen with safranin stain (Fig. 5.3A). Similarly fibrous mats, exodermis, and cortical cells in Dendrobium nobile are stained red with safranin (Fig. 5.3B). Fast-green is equally important in staining the cytoplasm and cellulosic cell walls. Both safranin and fast green are durable and show no signs of fading even after six years. Crystal violet and aniline blue can also be used as alternatives to fast green.

Fig. 5.3. Transverse section of root. (a) Bulbophyllum trimulum showing single layered velamen and vascular cylinder; (b) Dendrobium nobile indicating the fibrous mats, exodermis, and cortical cells, all are lignified and stained with safranin (exo= exodermis; fm= fibrous mat; pc= passage cell; v= velamen).

B. Safranin and Harris’ (or Delafield’s) Haematoxylin (Johansen, 1940) i. Follow 1-7 steps as described in Safranin-Fast Green method. ii. Stain in haematoxylin for 15-20 mins till the sections appear deep purple in colour. iii. Later transfer into acidulated water for few seconds. iv. As soon as the sections appear reddish, transfer to tap water. Here itis essentially important to remove every trace of acid and it takes atleast 20 mins. If haematoxylin is not sufficiently blued by the tap water, dip sections in water containing few drops of ammonia. v. Pass the slides through upward series of graded alcohol, i.e., 50, 70, 90% and pure alcohol, 3-5 mins in each grade. vi. Later keep sections in a mixture containing equal parts of xylene and alcohol (1:1). vii. Give two changes in pure xylene and mount in DPX or Canada balsam.

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C. Tannic Acid-Ferric Chloride-Safranin Triple Staining (Jensen, 1962; Lutman, 1946) i. After dewaxing, hydrate the sections by passing through 90, 70, 50 and 30% ethanol to distilled water, 5 mins in each step. ii. Place hydrated sections into 1% aqueous tannic acid for 10 mins. iii. Wash thoroughly in running tap water for 5 mins. iv. Transfer the sections to 3% aqueous ferric chloride for 5 mins. Rewash with tap water and examine. If the colour is not dark enough, repeat 2-4 steps. v. Stain with safranin for 1-24 hrs (stock solution of 1% safranin in 95% ethanol is prepared, dilute stock with equal volume of water before use). vi. Later pass the sections through 90 and 100% alcohol, and then to mixture containing equal parts of ethanol and xylene. vii. Finally, three changes of xylene are given and mount in resin-based mountant. Result: Lignified walls, chromatin and nucleoli become red and remaining blue-black to grey. D. Heidenhain’s Iron Haematoxylin and Orange G Staining (Jensen, 1962) i. After dewaxing, hydrate the sections by passing through 90, 70, 50and 30% ethanol to distilled water, 5 mins in each step. ii. Transfer hydrated sections to mordant solution (5% ferric ammonium sulfate) for 30 mins to 2 hrs. iii. Wash thoroughly in running tap water for 5 mins. iv. Stain sections in ripened haematoxylin (0.5 g haematoxylin dissolved in 10 ml absolute alcohol, makes up to 100 ml with distilled water and allow it to ripen for 2-3 weeks). v. Wash thoroughly with running tap water for 5 mins. vi. Destain in mordant solution. Observe under microscope periodically for colour development of chromatin. vii. Again, wash in running water for 5 mins. viii. Dehydrate sections by passing through 30, 50, 70, 90% ethanol and 100% ethanol. ix. Stain with Orange G for 1-5 mins (0.5% Orange G in clove oil, dissolve by shaking for 4 hr and filter it before use; counterstain may not be always necessary). x. Differentiate in a mixture containing equal parts of clove oil, xylene and ethanol for 5-15 mins and observe under microscope. xi. Give three changes in xylene and mount with DPX or Canada balsam. Result: Chromatin, nucleoli, mitochondria and plastids appear dark blue whereas cytoplasm and cell walls appear orange.

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5.5.2.2 Glycol Methacrylate Embedded Tissue

Working with xylene for dewaxing, hydration and dehydration of paraffin sections are tedious and time-consuming method. Besides xylene causes respiratory problems and irritation to eyes. Alternatively, staining of glycol methacrylate (GMA) sections is very easy and speedy, and it is proved to be particularly attractive. The plastic, GMA is freely permeable to aqueous stains, so there is no need to remove plastic before staining. Artifacts associated with the removal of the embedding medium from paraffin sections, and time lost in dewaxing and hydrating the sections are avoided in sections of GMA embedded tissue (O’Brien & McCully, 1981). In addition to following recipes, various other staining methods for GMA sections have been described in the chapter “Histochemical Methods”.

Staining with Acid Fuchsin (O’Brien & McCully, 1981) i. Stain the sections in 1% aqueous acid fuchsin for 1-5 mins. ii. Rinse in water for 5 mins or till the plastic is free from stain. iii. Allow to air dry and mount with DPX or Canada balsam. Result: Mitochondria, plastids and nucleoli are usually stained very strongly when warm dye solution is used. But starch, polysaccharides of cell wall and lignin are unstained. A variant differential staining procedure for mitochondria, plastids and nucleolus is based on the mixture recommended for staining yeast cells (Robinow and Marak, 1966). Stain the section in 0.005% acid fuchsin in 1% acetic acid. Later rinse in 1% acetic acid, air dry and mount (see also O’Brien and Thimann, 1965). 5.5.2.3 Epoxy Embedded Tissues

Due to the impermeable nature of plastic, the conventional staining methods are almost useless with epoxy sections. However, so far, the point of impermeability of this plastic has never been established (O’Brien and McCully, 1981). The first successful procedures for staining epoxy sections were those of Richardson et al. (1960) and Trump et al. (1961).Their works indicate that the use of cationic dyes like azure II, methylene nblue or toluidine blue, at high pH is more successful for epoxy sections. However, Fisher (1968) recommends aniline blue black (anionic dye). When O’Brien and McCully (1981) worked with the sodium methoxide method of Mayor et al. (1961) and the ethanolic NaOH method of Lane and Europa (1965) which removes the plastic, they found that the plastic removed sections stain better with acid fuchsin and fast green, and give beautiful fluorescent staining with aniline blue. However, such sections show very poor staining with basic dyes. A. Staining with Azure II/Methylene Blue (Richardson et al., 1960) i. Epoxy sections are treated with 1% periodic acid for 5 mins.

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ii. Rinse in water. iii. Stain sections in freshly prepared mixture consists of equal volumes of 1% aqueous azure II and 1% methylene blue in 1% sodium borate. Staining is performed on hot plate at 60°C for 1-5 mins. Staining is usually adequate when the edges of stain solution just begin to dry. Do not allow to dry further. iv. Rinse in water and dry with hair dryer. v. Mount with immersion oil or some other mountant. Result: Lignified walls and tannins appear green. Some cell walls may stain meta-chromatically, i.e., reddish purple. All other components show various shades of blue. B. Del Rio Hortega’s Silver Stain (Goldblatt and Trump, 1965) Stock Solutions (1) 10% silver nitrate (AgNO3) (2) 5% aqueous sodium carbonate (Na2CO3) (3) 0.88% ammonia (NH4OH) (4) Silver amino carbonate 5 ml (A) 10% AgNO3 (B) 5% Na2CO3 20 ml Combine A and B to which add NH4OH drop-wise till the precipitate disappears. Dilute to 45 ml and filter. This mixture can be stored for several weeks in a brown bottle. Procedure i. Stain the sections (glutaraldehyde/OsO4 fixation) in silver amino carbonate for 0.5-2 hrs or until you get walnut colour at 60°C. ii. Rinse the sections for 2 mins in distilled water and dry them on hotplate at 60°C. iii. Reduce in 0.4% HCHO for 30-60 secs. This step may be omitted ifthe tissue shows sufficient differentiation. iv. Wash in distilled water for 1-2 mins. v. Fix in 5% Na2S2O3 for 5 mins. This step may be omitted for epon sections but it is essential to prevent fading with araldite sections. vi. Wash in distilled water for 2 mins. vii. Dry and mount in DPX or other synthetic resin. Result: Tracheary elements are very intensely stained, and nuclei and plastids are also stained. C. Coomassie Brilliant Blue for Plasmodesmata (Fisher, 1968; O’Brien & McCully, 1981) Stock solution: 0.25% Coomassie brilliant blue in 7% acetic acid.

Staining Techniques 95

Procedure i. Stain the sections (glutaraldehyde/OsO4 fixation) with 0.25% coomassie brilliant blue (an anionic dye) in 7% acetic acid for 10 mins at 50°C. ii. Rinse in 7% acetic acid and dry with hair dryer. iii. Counterstain (if necessary) with 1% safranin O for 1 min at 60°C. iv. Rinse with water, air dry and mount. Coomassie brilliant blue is an excellent stain for plasmadermata. D. Haematoxylin-Safranin Staining for Semi-thin Epoxy Sections (Johansen, 1940; Warmke and Lee, 1976) Following are the staining solutions: (a) Oxidant (5% H2O2) 30% Hydrogen peroxide Distilled water (b) Mordant (3% ferric ammonium sulfate, acidified) 2-Methoxy ethanol (Methyl cellosolve) Distilled water Glacial acetic acid Sulfuric acid Ferric ammonium sulfate (c) Haematoxylin 2-Methoxyethanol (Methyl cellosolve) Tap water Distilled water 10% haematoxylin in absolute ethanol

5 ml 25 ml 50 ml 50 ml 1 ml 0.12 ml 3g 50 ml 25 ml 25 ml 10 ml

Allow to ripen at room temperature for about two weeks. (d) Safranin O Safranin 0.2 M Tris buffer (pH 9.0)

1g 100 ml

Procedure i. Plant materials like anthers, leaves and roots are killed and fixed in glutaraldehyde or glutaraldehyde-paraformaldehyde, post-fixed in osmium tetroxide. ii. Dehydrate the tissues in alcohol and acetone and embed in various epoxy resins like Epon, Epon-Araldite, Spurr, Maraglass-cardolite, etc. iii. Sections of 1 µ thick are cut with diamond knife on ultramicrotome. Later sections are transferred to adhesive coated slides. iv. Now immerse slides in solution A at room temperature for 10 mins. Rinse thoroughly with tap water.

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v. Slides are transferred to solution B in screw-capped coplin jars and placed in oven at 60°C for 30-60 mins depending on the resin, material and section thickness. vi. After-rinsing, the slides are transferred to solution C in coplin jar and placed in an oven at 60°C for 30-60 mins. vii. Again, rinsing sections, these are stained with solution D (preheated to 60°C) for 5 mins. viii. After rinsing again with tap water and distilled water, slides are dried on a warming table at 60°C. ix. Clear by flooding sections in a 1:1 mixture of clove oil and xylene for 30 secs. x. Finally, slides are passed through the pure xylene. Three changes are given in pure xylene. Later slides are mounted in permount or similar resin. Staining is performed here without removing the plastic matrix. The haematoxylin requires prestain oxidation and later it is used as a stain. It binds to the typically basophilic tissue constituents and stains them light grey to black. Safranin stains cellulose walls and secondarily deposited materials such as cutin, lignin, callose and sporopollenin. Use of methyl cello solve in mordant and in haematoxylin stain improves penetration and prolonged life of solution. This is one of the excellent procedures for plastic embedded plant tissues. Haematoxylin which is used here has the advantage of being stable, insoluble in common counterstaining, dehydrating and mounting fluids, and is not likely to form precipitates. E. Methylene Blue-Azure A-Safranin for Semi-thin Epoxy Sections (Humphrey and Pittman, 1974; Warmke and Lee, 1976) Staining solutions (a) Methylene blue-azure A Methylene blue Azure A Glycerol Methanol 0.1 M phosphate buffer (pH 6.9) Distilled water (b) Safranin O Safranin O 0.2 M Tris buffer (pH 9.0)

0.13 g 0.02 g 10 ml 10 ml 30 ml 50 ml 0.1 g 100 ml

Procedure Follow (i)-(iii) steps described in Haernatoxylin-safranin for semi-thin epoxy sections method.

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i. Immerse slides in solution A (preheated) in coplin jars at 60°C for 30 mins. ii. After rinsing, transfer the slides to solution B in coplin jars and keep them at 60°C for 2 mins. iii. After rinsing once again with tap water and distilled water, dry the slides on a warming table at 60°C. iv. Finally, the slides are given three changes in pure xylene. Later these are mounted in permount or similar resin. Clearing is not required in this procedure. In both procedures (Haematoxylin-safranin and methylene blue-azure A-safranin methods), staining is progressive, that is without over staining in any of the solutions and without requiring differentiation (destaining). Probably the most striking feature of this combination is the bright blue colour imparted to the exine and Ubisch bodies of older anthers. Safranin generally stains cellulose walls and secondarily deposited materials. The methylene blue-azure A combination tends to be metachromatic and shows more colour variety and sharper contrast than haematoxylin. It does not form precipitations. F. Differential Staining for Tannin in Epoxy-embedded Plant Tissue (Parham and Kaustinen, 1976) Differential staining for tannin localisation is done in the following manner: i. Small pieces of callus or cell clusters from suspension cultures of Pseudotsuga menziesii and Pinus taeda are fixed in the mixture of 2% glutaraldehyde: 2% acrolein in sodium cacodylate buffer (0.05 Mat pH 7.0) for 12 hrs at 4°C. ii. After several rinses and gentle centrifugation in buffer, the samples are post-fixed in buffered 1% OsO4 for 12 hrs at 4°C. iii. After several rinses in buffer, the specimen is passed through the graded acetone series at room temperature. iv. Later the specimen is embedded with low-viscosity epoxy resin (Spurr, 1969). v. For light microscopy, semi-thin sections (0.5 µm thick) are taken; ultrathin sections are cut for electron microscopy. vi. Semi-thin sections are stained with sudan black B (Bronner, 1975) and mounted with Karo syrup; ultra-thin sections are sequentially stained with uranyl acetate and lead citrate for TEM studies. Result: Tannin deposits are stained brownish-orange colour and these are distinguished from lipid bodies of similar size which stain dark blue or black colour. Tannin deposits are also demarcated from starch grains which are unstained. Since tannins are strongly osmiophilic, they appear black in electron micrographs.

6 Centrifugation The centrifugation technique involves the sedimentation of particles present in the solution. If the particle density is higher than the solvent, then the particles will sink (sediment); where particles are higher than the solvent, then they will float to the top. When a solution of particles is allowed to stand, the particles will tend to sediment according to various parameters such as molecular mass, shape and density, under the influence of gravity. The greater the difference in density, the faster they move. If there is no difference in density (isopycnic condition), the particles stay steady. To take advantage of differences in density to separate various particles in a solution, gravity can be replaced with powerful centrifugal force applied by a centrifuge. Svedberg and Nicols in 1929 employed a centrifuge for the first time to increase the gravitational force to speed up the rate of sedimentation for the purpose of measuring particle sizes. Since then, a variety of centrifuges, including ultracentrifuges have been developed. The basic components of a centrifuge are a metal rotor with holes in which test tubes are placed and a motor that can spin the rotor at a selected speed. During centrifugation, force is applied to sediment the particles more rapidly. This force (centrifugal force) is greater than the gravitational force of the earth thus increasing the rate of sedimentation. Particles of different sizes, shapes, density are separated as they are sedimented at different rates (sedimentation rate) in a centrifugal force. The sedimentation rate is directly proportional to the applied centrifugal force.

6.1 Basic Principle of Sedimentation The rate of sedimentation is dependent on the applied centrifugal force (G) which exerts radially outwards. It is determined by the square of the angular velocity of the rotor (ω, in radians s-1) and radial distance (r, in centimeters) of the particle from the axis of rotation. G is expressed with the following equation: G = ω2r Since one revolution of the rotor is equal to 2π radians, its angular velocity, in radians S-1, can be readily expressed in terms of revolution per minute (rev min-1), the common way of expressing rotor speed is:

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ω=

2π rev min-1 60

The centrifugal field (G) in terms of rev min-1 is then: G=

4π2 (rev min-1)2 r 3600

It is generally expressed as the multiple of the earth’s gravitational field (g = 981 cm-2), i.e., the ratio of the weight of the particle in the centrifugal field to the weight of the same particle when acted on by gravity alone, and it is then referred to as the relative centrifugal field or more commonly as the ‘number times g’. Hence, RCF is as follows: RCF =

4π2 (rev min-1)2 r 3600 × 981

The above is shortened as RCF = (1.118 x 10-5) rev min-1)2 r

6.2 Instrumentation As we mentioned earlier that the metal rotor and motor are the basic components of the centrifuge. The centrifugal force created by a spinning rotor faces a lot of stress on rotor material. In low-speed centrifuges, rotors are made up of brass, steel, or perspex. Whereas in high-speed centrifuges in which higher stress forces are generated, rotors are made up of aluminium alloy or titanium alloy. Rotors made of titanium alloy have a greater strength-to-weight ratio and are therefore capable of withstanding nearly twice the centrifugal force of rotors made from aluminium alloy. They are more resistant to chemical corrosion (Wilson and Walker, 2010; Hofmann and Clockie, 2018). All rotors are given a durable protective coating to the metal surface either by applying the epoxy paint or by anodising in the case of aluminium rotors. Some of the preparative rotors and their design are given below:

6.2.1

Swinging Bucket Rotors

The swinging buckets rotor has buckets that start off in a vertical position but during acceleration of the rotor they swing out to a horizontal position; in this process during centrifugation, the solution in the tube is perpendicular to the axis of rotation, and parallel to applied centrifugal free. During deceleration, the tube returns to its original position. Since the centrifugal field is axial, particles in a centrifugal field fan out radially from the centre of rotation rather than sedimenting in parallel lines (Fig. 6.1).

Centrifugation 101

Fig. 6.1. Centrifuge with swinging bucket rotors.

Fixed-angle rotors – In these rotors, the test tubes are located in holes of the rotor body set at a fixed angle of between 14° and 40° to the vertical. Centrifugal force is exerted at an angle to the tube well, particles move radially and have a short distance to travel before precipitating on the outer wall of the centrifuge tube. A region of high concentration is formed that has a density greater than that of the surrounding medium, with the result that the precipitates inks and collects as a small compact pellet at the outermost point of the tube. It is proved to be valuable for differential centrifugation separating the particles differing significantly in their sedimentation rates.

6.2.2

Vertical Tube Rotors

These are regarded as a zero-angle fixed rotor in the tubes aligned vertically in the body of the rotor. During the operation of the rotor, the solution in the

Fig. 6.2. Vertical tube rotor: (a) cross-sectional image of vertical tube rotor. (b) The centrifuge tube is filled with a gradient; the sample is layered on top and then placed in the rotor. (c) As the rotor accelerates, the sample and gradient begin to reorient. (d) The sample and medium reorientation are complete. (e) Sedimentation and separation of particles occur during centrifugation. (f) Reorientation of separated particles and gradient occurs during the rotor deceleration. (g) Finally,the rotor is at rest; bands of separated particles and gradient are fully reoriented (Redrawn from Wilson & Walker, 2002).

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tube reorients through 90° during rotor acceleration to lie perpendicular to the axis of rotation and parallel to the applied centrifugal field and return to its original position during deceleration of the rotor. Since sedimentation of a particle occurs across the diameter of the tube, the vertical rotor presents the shortest possible path length (Fig. 6.2). This sedimentation is quicker than any other types described earlier.

6.2.3

Zonal Rotors

Zonal rotors may be batch or continuous flow types. The former one is extensively used, and it is designed to minimise the wall effect encountered in swinging-bucket and fixed-angle rotors and to increase the sample size. Lowspeed batch rotors with 5000 rpm are made up of aluminium, having a thick transparent Perspex top and bottom to permit direct examination of particle sedimentation during the course of centrifugation. High-speed batch rotors with 60,000 rpm are made up of aluminium or titanium alloy. The body of typical batch-type rotors is either a large cylindrical container or a hollow bowl, in which rotor volume varies with the square of the radial distance from the centre of rotation. The centre of the rotor has a core to which attached a vane (broad blade) assembly that divides the rotor internally into four sector-shaped compartments and minimises swirling of the rotor contents. The rotor core may be of two main types. The commonly used standard core permits the loading and unloading of the rotor while it is spinning (dynamic method) whereas the second core type (the reorienting gradient core) is designed to allow the motor to be loaded and unloaded with a reorienting gradient while it is at rest (static method).

6.3 Mechanism of Centrifugation There is two type of centrifugation; these are differential centrifugation and density gradient centrifugation

6.3.1 Differential Centrifugation In differential centrifugation, various cell components are separated at different speeds at different times. The cellular components of rat liver cells can be isolated by differential centrifugation (Fig. 6.3). The liver is homogenised by adding sucrose solution. The homogenate is subjected to a low speed of 700 G for 10 minutes in a centrifuge. The sediment is formed (pellet). As it is the first sediment, it is called pellet-1 or nuclear fraction. It contains nuclei, unbroken cells, and large fractions of the plasma membrane. The solution is called supernatant-1. Now, supernatant-1 is transferred to another tube, and it is centrifuged at 10,000 G for 20 minutes. The pellet-II (mitochondrial fraction)

Centrifugation 103

is formed. This fraction contains mitochondria, lysosomes, and peroxisomes. Here supernatant-II is in the centrifuge tube. Repeat the above step; the supernatant-II is transferred to another tube and centrifuged at 1,00,000 G for one hour. Now pellet III or microsome fraction is formed. This fraction contains ribosomes and small fragments of the Golgi complex, endoplasmic reticulum, etc. The solution remaining in the tube is supernatant-III. The supernatant-III is also called cytosol which contains proteins, soluble nucleic acids, soluble polysaccharides, lipid droplets, etc. These are separated by electrophoresis or chromatographic techniques. Liver homogenate Centrifugation 700 G for 10 minutes

Pellet-I (Nuclear fraction) Pellet-II (Mitochondrial fraction) Pellet-III (Microsome fraction)

Supernatant-I Centrifugation 10,000 G for 20 minutes Supernatant-II Centrifugation 1,00,000 G for 1 hr Supernatant-III (Cytosol consists of proteins Lipids, carbohydrates)

Fig. 6.3. Liver homogenate - differential centrifugation. (Adopted from Wilson and Walker, 2010).

6.3.2

Density Gradient Centrifugation

In density gradient centrifugation, components are separated based on density gradients. In the density gradient column, the homogenate, say sucrose, is poured on the top of the column for centrifugation. During centrifugation, the highest-density sucrose is settled at the bottom, medium-density sucrose solution in the middle whereas the lowest density is at the top. The higher the concentration of sucrose, the greater the density of the solution. The materials are collected at their static positions in bands according to their isodensities in the gradient and each band is isolated separately and carefully. This technique is useful to separate particles of same size with different densities, for example, RNA can be separated from DNA, and similarly, fragments of rough endoplasmic reticulum are separated from the smooth endoplasmic reticulum.

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6.4 Sedimentation Co-efficient A solution is a mixture of one or more solutes and solvents when the solution is allowed to stand, the solutes settle at the bottom and sedimentation takes place. Sedimentation is accelerated by the movement of the solution in a centrifuge. The speed at which solute sediments depend on the mass of the molecule, the speed of rotation, density, viscosity, and temperature of the medium, and the shape of the solute. The sedimentation rate or velocity of a particle can be expressed in terms of its sedimentation rate per unit of the centrifugal field; it is commonly referred to as its sedimentation coefficient, s. V = sω2r (or) V dr/dt s= 2 = ωr ω2r Here, V ω r dr dt

= = = = =

Velocity of the particle Angular velocity of the centrifuge radial distance of the particle from the axis of rotation density of the particle density of the medium

According to the above equation, the sedimentation coefficient, s, is the ratio of the velocity of the particle to the centrifugal acceleration.

6.5 Classification of Centrifuges and their Uses Centrifuges are classified into four major groups, such as small bench centrifuges, large capacity refrigerated centrifuges, high-speed refrigerated centrifuges, and ultra-centrifuges (preparative and analytical ones).

6.5.1

Small Bench Centrifuges (Portable Centrifuges or Table-Top Centrifuges)

Small bench centrifuges are the simplest and least expensive ones. Generally, they have a maximum speed of 4000 to 6000 revolutions per minute (rpm), with maximum relative centrifugal fields of 3000 to 7000 G. Mostly operate at ambient and room temperature. Some of the modern designs have refrigerator systems to keep the rotor cool. Some microfuges are available, having instant acceleration to a maximum speed of 8000 to 13000 rpm and developing fields of approximately 10,000 G. These instruments are extremely useful for sedimenting small volumes of material (250 mm3 to 1.5 cm3) very quickly (1

Centrifugation 105

or 2 min). These centrifuges have been useful for the rapid sedimentation of blood samples and synaptosomes (isolated synapticterminals from neurons).

6.5.2

Large-Capacity Refrigerated Centrifuges

Large-capacity refrigerated centrifuges have a maximum speed of 6000 rpm and a maximum relative centrifugal field of about 6500 G. They have a refrigerated rotor chamber and vary only in their maximum carrying capacity; they have a variety of interchangeable swinging-bucket and fixedangle rotors enabling separation to be achieved in 10, 50 and 100 cm3 tubes. Larger capacity (4-6 dm3) centrifuges are also available, and their rotors can accommodate bottles of large capacity. In these centrifuges, the rotors are mounted on a rigid shaft. Therefore, it is very important that the contents of the centrifuge tubes are balanced accurately, and they should not be loaded with an odd number of tubes.

6.5.3

High-Speed Centrifuges

High-speed centrifuges with refrigerated rotors are available with a maximum speed of 25,000 rpm,generating a relative centrifugal field of about 60,000 G. They have a total capacity of up to 1.5 dm3 and a range of interchangeable fixed angle and swinging-bucket rotors. These centrifuges are commonly used in biotechnology laboratories to collect cell debris, large cellular organelles (nuclei, mitochondria, lysosomes, etc.), precipitates of chemical reactions, and immunoprecipitates.

6.5.4 Ultra Centrifuges There are two types of ultracentrifuges, these are preparatory centrifuges and analytical centrifuges. Centrifugation for the isolation of and purification of components is known as preparatory centrifugation, whereas centrifugation for the characterisation of these components is known as analytical centrifugation. 6.5.4.1 Preparatory Ultracentrifuges

In preparatory ultracentrifuges, the rotor spinning is at the maximum speed of 80,000 rpm and relative centrifuged field up to 6,00,000 G. At such speed the friction between air and spinning rotor generates an excess amount of heat. To minimise the heat, the rotor chamber is refrigerated (0°-4°C) sealed and evacuated by two pumps working in tandem. Apart from that, an infrared temperature sensor has been employed to monitor continuously the rotor temperature and control the refrigeration system. The drive shaft (made up of aluminium or titanium alloy of high tensile strength) on which the rotor is mounted is merely 1/16 inch in diameter. This small diameter allows the shaft to flex during rotation accommodating a certain degree of rotor imbalance without spindle damage.

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All centrifuges have an over-speed device to prevent the rotor from operating at maximum high speed. Operation of the rotor at excessive speed can result in an explosion of the rotor. To contain such damage, the rotor chamber is always enclosed in a heavy armour plate. Table-top, an air-drives preparatory centrifuge, known as airfuge, is also available; it can accelerate a magnetically suspended 37 cm diameter rotor, accommodating 6 x 175 mm3 tubes on a vertical friction-free cushion of air in a non-evacuated chamber, with a maximum speed of 1,00,000 rpm (1,60,000 G). An airfuge is very useful in biochemical and chemical research where the small number of samples available require high centrifugal forces; for example, steroid hormone receptor assays separation of the major lipoprotein fractions from plasma deproteinisation of physiological fluids for amino acid analysis and macromolecules ligand binding-kinetic studies. 6.5.4.2 Analytical Centrifuges

Analytical centrifuge is almost similar to that of the preparative centrifuge. It also has a refrigerated and evacuated chamber. However, it differs from the others in having a different type of rotor and in possessing a specialised optical system to monitor the progress of centrifugation. The rotor is elliptical, and it has two holes for holding two centrifuge cells. One cell is known as the analytical cell and the other is the counterbalance or the counterpoise cell. The rotor holds these cells vertically, while it is at rest or rotating position. The analytical cell is sector-shaped and can hold a liquid column about 14 mm high. It has the capacity to accommodate about 1 cm3 of the sample. The upper and lower planes of analytical cells are transparent having synthetic sapphire windows. The windows are provided for the passage of light to monitor the progress of centrifugation. The optics used here are either Schlieren optics or Rayleigh interference optics. A portion of the solution in the analytical cell that contains micromolecules will have a refractive index that is higher than the rest of the solution, which is the basis on which the Rayleigh interference functions. At the beginning of sedimentation, the peak of refraction would be at the meniscus. As the sedimentation progresses, the macromolecules move down the cell, the peak also shifts giving direct information about the sedimentation characteristics of the macromolecules. The entire optical information is continuously photographed. Analytical centrifugation has been useful to measure the sedimentation coefficient and molecular weights of macromolecules. Molecular weights of various biological molecules, when they are in a gross mixture, have been measured by this technique.

7 Colorimetry and Spectrophotometry Colorimetry deals with the measurement of light absorption by coloured compounds in solutions. The instrument that measures the intensity of colour is known as Colorimeter. The instrument has been used to test the concentration of coloured substances by measuring its absorbance of a specific wavelength of light. Colorimeters are relatively simple instruments designed to function only in the visible range (400-700 nm) whereas spectrometers are equipped to operate both in the visible and UV range. Visible Light and Electromagnetic Radiation Light is a form of energy and made of particles called photons. The normal light is only a collection of waves whose wavelengths are in the narrow range between 4000-8000 A (1 Angstrom = 10-8 cm) or 400-800 nm, whereas the energy spectrum or electromagnetic spectrum covers wavelengths of the order of meters (radio waves) to less than angstrom (e.g., gamma rays). It is well known fact that energy increases as the wavelength decreases. An atom consists of electrons, protons and neutrons, and orderly arrangement of atom forms molecules. In all these cases, the movement or motion of the particles is intrinsically involved. When the energy is supplied to an atom or molecule, the various movements are affected to varying extent, depending on the level of energy input. For instance, if a molecule is subjected to infrared radiation (0.8-2.0), only the vibrations of the atoms in the chemical bonds are affected whereas, ultraviolet radiation with, a greater energy (2000 – 4000 A) affects the electrons in the atoms. Interaction with Matter Electromagnetic phenomenon exhibits energy, frequency, wavelength and intensity. All these are interrelated and explained in terms of wave forms of particles (photons or quanta). This phenomenon can be explained by considering the electronic spectra. Electrons in either atoms or molecules may be distributed in several energy levels. When an electron is promoted to higher level (excited state) from the ground state, some energy is put into the system, and this gives rise to absorption spectrum. Only the exact amount of energy equivalent to the difference in energy level will be absorbed. This is termed one quantum of energy for a single electron transition, and the absolute

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magnitude of each quantum will differ according to the difference in energy levels involved. When an electron falls from a higher to lower energy level, then exactly one quantum of energy is emitted from the system giving rise to an emission spectrum.

7.1 Colorimetry As already stated earlier, colorimetry is a form of photometry, measuring the intensity of colour of the substances in solutions. It is a known fact that the visible light (white light) is consisting of seven colours, such as violet, indigo, blue, green, yellow, orange and red (VIBGYOR). The colour of the substance depends primarily on its light absorption properties. A red colour object, for example, absorbs all colours of the white light except the red region. So, the object looks red. This is universal phenomenon for other colours as well. Interestingly white coloured objects consist of all the colours whereas black coloured objects have none of the colours of the spectrum. The intensity of such coloured substances can be measured with colorimeter. The white coloured substances which do not show any significant absorption properties could be reacted significantly with some other reagents to form coloured product. This coloured product can be measured colorimetrically. The principle involved in colorimetry is that when a beam of incident light passes through a coloured solution, the coloured substances in the solution absorb a part of light and, remaining part of radiation comes out; the intensity of this transmitted light is measured by colorimeter. The intensity of transmitted light is always less than the incident radiation. As the number of light absorbing molecules increases, the intensity of light coming out of the medium decreases exponentially, and vice-versa. The difference in intensities between the incident and transmitted light reflects the number of absorbing molecules or in other words, the concentration of the absorbing molecules in that solution. The light absorbing molecules are called chromophores and this property is attributed to the chemical structure of these molecules. Some of colorimetry assays are showed in Table 7.1.

7.1.1 Quantification of Absorbing of Light The absorption of light by biological molecules/chemical substance can be quantified based on two basic laws propounded by Beer and Lambert, also known as Beer-Lambert law. 7.1.1.1 Beer-Lambert Law

As already stated that Beer-Lambert law is a combination of two laws, viz., Beer law and Lambert law, each dealing separately with absorption of light, related to the concentration of the absorber (the substance responsible for ‘absorbing the light’) and the path length or the thickness of the layer (related

Colorimetry and Spectrophotometry 109 Table 7.1. Some colorimetric assays* Substance

Reagent

Wavelength (nm)

Inorganic phosphate

Ammonium molybdate; H2SO4; 1,2,4-amino-nephthol; NaHSO4, Na2SO4

600

Protein

Folin Biuret BCA reagent (bicinchoninic acid) Coomassie brilliant of blue

660 540 562 595

Aminoacids

Ninhydrin Cupric salts

570 620

Peptide bonds

Biuret (alkaline tartarate buffer, cupric salt)

540

Carbohydrates

Phenol, H2SO4 Anthrone (anthrone, H2SO4)

Varies, e.g., glucose 490, xylose 480, 620 or 625

Reducing sugars

Dinitrosalicylate, alkaline tartrate buffer

540

Pentoses

Bial (Orcinol, ethanol, FeCl3, HCl) Crysteine, H2SO4

665 380-415

Hexoses

Carbazole, ethanol, H2SO4 Cysteine H2SO4 Arsynomolybdate

540 or 440 380-415 Usually 500-570

Glucose

Glucose oxidase, Peroxidase, O-dianisidine, phoshphate buffer

420

Ketohexose

Resorcinol, thiourea, ethanoic acid, HCl, Carbazole, ethanol, Cystcine, H2SO4 Diphenylamine, ethanol, ethanoic acid, HCl

520 560 635

Kexosamines

Ehrlich (dimethylaminobenzaldehyde,etha nol, HCl)

530

DNA

Diphenylaine

595

RNA

Bial (Orcinol, ethanol, FeCl3, HCl)

665

Sterols

Libermann—Burchardt reagent (Acetic anhydride, H2SO4, chloroform)

425

Cholesterol

Cholesterol oxidase, peroxidase, 4-aminoantipyrine, phenol

500

*(Courtesy: Wilson and Walker, 2010)

to the absolute amount of the absorber); provided an absorbing substance is partially transparent and it will transmit a portion of the incident radiation. Beer’s law propounds that when a parallel beam of monochromatic light passes through an isotropic, light absorbing medium, the amount of light that

110 Plant Techniques: Theory and Practice

is absorbed is directly proportion to the number of light absorbing molecules in that medium, nothing but the concentration of the chemical substance in that medium. This is indicated as follows: A ∝ C; here ‘A’ is the absorbance or optical density, and ‘C’ is the concentration of light absorbing chemical substance in the medium. Lambert’s law states that when a parallel beam of monochromatic light passes through the isotropic, the amount of light absorbed is directly proportional to the length of the medium through which light passes, nothing but the amount of chemical substance in that medium. This is indicated as follows: A ∝ L; here ‘A’ is absorbance of light and ‘L’ is the length of the medium (or thickness). Hence the measurement of light absorption depends on both the laws (Beer-Lambert law); comprehensive equation is given below: A ∝ C × L (or) A= eCL; here ‘e’ is known as Extinction co-efficient. also Ii 100 A = log I = log T = log 100 − log T = 2 − log T t

Here, ‘Ii’ is the intensity of incident light and ‘It’ is the intensity of transmitted light. Molar-absorbance Coefficient (or Molar Extinction Coefficient)

In the above equation, if the concentration is l M i.e., one mole per litre, and path length of light is l cm, then ‘e’ is known as molar extinction coefficient (ΣM). ΣM is constant for a particular compound at a particular wavelength and has a maximal value when the compound is in purest state. Specific extinction coefficient

If the molecular weight of compounds such as nucleic acids, proteins, etc., is not known, then the term “Specific extinction coefficient” can be conveniently used. This is nothing but the extinction of a 1% (W/V) solution of the compound when the light path is 1 cm. Limitations of Beer-Lamberts Law

Beer-Lamberts law has some limitations. It holds true for parallel, monochromatic light in an isotropic medium i.e., the light absorbing molecules are randomly oriented in the solution. If these conditions are not met, it deviates from the law. This deviation may be due to the instrument or due to sample as well.

Colorimetry and Spectrophotometry 111

Beer-Lambert’s law deviates due to the instrument. The law holds only for the monochromatic light, i.e., light of single wavelength. But it is not realised in practice. In colorimeters, filters of broad band of light have been used. Red filter of 640 nm, for example, passes light ranges from 625 nm, i.e., 640 ± 15 nm. Same is the case in other filters also. Therefore, deviation takes place. Further, the Beer-Lambert’s law deviates due to the sample. This is as follows: 1. According to law, the relationship between concentration and absorbency is linear. But in practice, such linearity is observed up to certain concentration, i.e., threshold concentration. After the threshold concentration, the ionic species are close enough to interact one another and absorb less light. Therefore, light absorption does not follow the linearity of the law. 2. When the absorbing species contain acidic or basic groups, they dissociate or associate in solution to form ‘acidic-base’ pair with various absorption properties. In such case pH varies, hence deviation takes place. Even if absorbing species are in equilibrium, change of temperatures could cause deviation in the absorption of light. 3. Deviation could be possible due to polymerisation of absorbing molecules at high concentration leading to turbid suspension, which might increase or decrease the apparent extinction. 4. In some cases, the absorbing molecules may be physically absorbed on the walls of cuvette and thus decrease their effective concentration in solution. 5. Deviation could also arise due to sample fluorescence. That means, the absorbed light is emitted as fluorescent radiation of longer wavelength in all directions. But only the small portion of emitted radiation which passes through the direction of the detector is measured. This results into low or even negative absorbance values. Finally, 6. Deviation may arise due to dilution of coloured solutions. For instance, potassium dichromate, when it changes its colour from orange to red upon dilution with water can cause deviation with two absorption maxima. Cr2O7+H2O-→ 2CrO4+2 H+ dichromate chromate (Orange) (Yellow) ~ 340 nm ~ 770 nm Therefore, it is not at all possible to eliminate every factor that causes deviation from Beer-Lambert’s law. Hence, a calibration curve (standard graph) is plotted and from which the concentration of unknown is determined from its absorbancy value (in the linear range) and taking into consideration proper controls, if necessary, apart from blank.

7.1.2

Colorimeter

Colorimeter is the simplest instrument to measure the intensity and wavelength of light after it passes through an absorbing medium (coloured solution). Instrument consists of light source, a condensing lens to render the light

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rays parallel, a filter to regenerate monochromatic light, a sample holder, a photocell with a detector to measure the light intensities and a digital display galvanometer to measure the electrical energy generated. The light from a light source (tungsten lamp composed of wavelengths 400-900 nm) passes through a slit, a condensed lens, a filter and finally emerges as a parallel beam of monochromatic light. The monochromatic light passing through the sample solution is projected on to photocell. The photocell converts the transmitted light into electrical energy and it is amplified and measured by galvanometer. The galvanometer is calibrated to read the absorbance/transmittancy directly. 7.1.2.1 Complimentary Colours and Complimentary Wavelengths

The coloured substance absorbs the light from a particular colour of the filter to a maximum extent, that colour is called complimentary colour to the colour of the solution (Table 7.2). Table 7.2. Complementary colour for some coloured solutions Colour of the solution Violet Blue Green Yellow Orange Red Purple red

Range of the wavelength (nm) 400-465 465-482 498-530 576-580 587-610 617-660 670-720

Complementary or subtraction colour Greenish colour Yellow Red purple (magenta) Blue Greenish blue Bluish green Green

When a coloured solution absorbs light maximally at a particular wavelength, then the wavelength is called complimentary wavelength (λ max) of the solution. Complimentary colour and complimentary wavelength have been used to estimate the concentration of the substances. The given substance absorbs maximally at a particular wavelength and contributes to high intensity to the estimations and less interference from other substances.

7.1.3 Protocols The following has been taken up to determine the complementary colour (wavelength) of a test solution and to verify Beer-Lambert’s law, and to carry out NADH analysis as well. 7.1.3.1 To find out the Complementary Colour/Wavelength (Palanivelu, 2016)

(i) Prepare 0.01% of potassium dichromate and 1% copper sulphate solutions in distilled water.

Colorimetry and Spectrophotometry 113

(ii) Now, switch on the colorimeter and warm up for 15-20 minutes, with one of the filters in position or set the spectromic-20 at minimum wavelength of 340 nm. (iii) Place the cuvette containing the blank (distilled water) in the cuvette holder and adjust the readings to 100% transmittance and switch off the galvanometer. (iv) Take the test solution in the cuvette and place it in the cuvette holder. (v) Switch on the galvanometer and record the absorbancy and transmittance of the test solution, viz., potassium dichromate or copper sulphate. (vi) Repeat steps (iii) and (v) for each filter or for every 5 or 10 nm wavelength. (vii) Find out the colour of the filter, which absorbs maximally. This colour is the complimentary colour to the colour of the solution. (viii) To determine the complementary wavelength plot the graph absorbancy vs wavelength. The peak gives the λ max of the solution. Note: Switch off the galvanometer whenever changing the sample and filters. 7.1.3.2 Verification of Beer– Lamberts Law (Palanivelu, 2016)

(i) Prepare 0.05% Potassium dichromate and 5% copper sulphate solutions in distilled water. (ii) Now switch on the colorimeter and warm it up for 15-20 minutes. (iii) Take clean, dry test tubes and number them 0 to 10 in triplicates. (iv) Pipette 0.5 ml up to 5.0 ml with a gradual increase of 0.5 ml and keep a blank as well. (v) Add distilled water to make a final volume of 5 ml and mix thoroughly. (vi) Using the complementary coloured filter or complementary wavelength (from the previous experiment) adjust the blank to 100% transmittance and then read each tube and record the absorbance and transmittance. (vii) Plot absorbancy (transmittance) against concentration and find out up to what concentration, the concentration obeys the Beer – Lambert’s law. The standard graph can also be plotted to find out unknown concentrations of Potassium dichromate solution. 7.1.3.3 Nicotinamide Adenine Dinucleotide (reduced form - NADH) Analysis

(i)

(ii)

Turn on the colorimeter and set the wavelength 340 nm. After 1-2 minutes, switch on the UV (or deuterium) lamp and allow the instrument to warm up for 15-20 minutes (follow the Manual supplied by the manufacture company). By taking blank (buffer alone i.e., 3 ml of 0.05 M Tris-HCl) in the silica cuvette, adjust the instrument to ‘zero’ absorbance (or 100% transmission).

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(iii)

(iv) (v) (vi) (vii)

In another cuvette take 0.1 ml of 1 mM solution of NADH and 2.9 ml of 0.05 M Tris – HCl buffer, so that the total volume in the cuvette is 3 ml. Mix thoroughly and record the absorbance of test sample (equivalent to 33 µM NADH or 0.1 µ moles in 3 ml). Discard the solution from the sample cuvette, rinse it with distilled water. Add 0.2 ml of NADH and make the final volume to 2.8 ml with 0.05 M Tris-HCl buffer, and then record its absorbance. Repeat step (iv) by taking 0.40 ml of NADH solution and 2.6 ml of 0.05 M Tris-HCl and note down its OD. Repeat step (iv) by taking 0.6 ml of NADH solution and make the volume to 3 ml with 0.05 M Tris-HCl buffer and record the absorbance. The above results are given in Table 7.3. Table 7.3. Results of NADH analysis Sample No.

NADH solution (in ml)

Conc. of NADH (µM)

Fold Conc. of NADH

A340

Ratio*

1 2 3 4

0.10 0.20 0.40 0.60

33.3 66.6 133.3 200.0

1.0 2.0 4.0 6.0

-

1.0 -

* Calculate the ratios

A 340 of sample 2 A 340 of sample 1

for samples 2, 3 and 4.

(viii) Check whether the calculated values of the ratio for samples 2, 3 and 4 correspond with the increase in concentration of NADH in these samples as compared to sample 1. If the ratio increases with the concentration of NADH in the samples, it indicates that the absorbance of light by the sample is directly proportional to the concentration of NADH. (ix) Prepare a standard graph of A340 against concentration of NADH in the sample. A straight line indicates proportional increase in the A340 with increasing concentration of NADH in the sample. By using this standard graph one can determine the unknown concentration of NADH present in the test sample. (x) Calculate the molar extinction coefficient (ɛ340 nm) for NADH; it is as follows: For example, 133.3µM solution of NADH gives A340 of x1µM. NADH corresponds to (x/133.3) absorbance, 1M solution NADH gives A340 values of (x/133.3) × 106; this value is ɛ340 for NADH. In this way, the concentration of NADH in unknown test sample can be directly calculated from its A340 by using the molar extinction coefficient of NADH.

Colorimetry and Spectrophotometry 115

7.2

Spectrophotometry

Spectrophotometer measures light absorption as a function of wavelength in the UV as well as visible regions. It is possible to make quantitative analysis of molecules depending on how much light is absorbed by coloured compounds. Spectrophotometer with photometer can measure light beam intensity as a function of its colour. The absorption of light is due to interaction of light with the electronic and vibrational mode of molecules. Each molecule has its own individual set of energy levels associated with the makeup of its chemical bonds and nuclei, and thus will absorb the light of specific wavelength, resulting in unique spectral properties.

7.2.1 Spectrophotometer Components Spectrophotometer has various parts such as light source, conducting lens, monochromater, sample holder, detector and recorder. Spectrophotometer has two light sources, such as tungsten lamp for visible light and deuterium lamp for UV light. The light from light source has wide range of wavelengths. This is known as Polychromatic or Heterochromatic. This polychromatic light reflected using a plane mirror, passes through an entrance slit, a condensing lens and falls on a monochromater. The monochromater disperses the light and desired wavelength is focused on the exit slit using the wavelength selector. Monochromaters that produce radiation of single wavelength, are based either on refraction by a prism or diffraction by a grating. Prism is made up of glass for visible light and of quartz or silica for UV light, whereas grating has ruled lines (about 2000 lines per millimetre) on a transparent or reflecting base. Resolving the power of the grating is directly proportional to the closeness of these lines. Gratings are more superior to prisms as they yield linear resolution of the spectrum for the entire range of wavelengths. The efficacy of monochromation is enhanced by double monochromaters, in which a selected part of the spectrum from the first grating is further resolved by a second grating, resulting in the bandwidth of as low as 0.1 mm. The standard cuvettes are made up of quartz and have an optical path of 1 cm and hold a volume of 1-3 ml. Quarts does not absorb UV light and hence they are used for both UV and visible light measurements. A photocell (photoelectric device) converts the light energy into electrical energy, and then it is amplified, detected and recorded (Fig. 7.1). In photocell cathode and anode are present. When photons strike the semi-cylindrical photo-emissive cathode in vacuum, it causes the emission of electrons, which is proportional to the intensity of radiation. When a potential difference is applied across the electrodes, the emitted electrons, flows to the anode wire generating a photocurrent. This current is amplified and measured.

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Fig. 7.1. A schematic representation of photocell.

A photomultiplier tube is similar to that of photocell. It also has a cathode with photo emissive surface (a selenium layer) and a wire anode. Besides that, photo cathode contains a circular array of 9 additional cathodes called dynodes. Dynode I with potential of 90 V is more positive than that of photo-emissive cathode and hence, the emitted electrons from photoemissive cathode are accelerated towards it. Upon striking the dynode 1, each photoelectron causes emission of several additional electrons within turn accelerated towards dynode 2, which is 90 Volts more positive than dynode 1. Again, several electrons are emitted from each electron. This process is repeated for 4 times and therefore each photon produces 106-107electrons. These amplified electrons flow to the anode generating a much larger electric current than the photocell.

7.2.2 Types of Spectrophotometers A variety of spectrophotometers are available; these are single-beam, double-beam, recording and multi brand instruments. In a single-beam spectrophotometer, the variations in source intensities are not compensated whereas in double-beam, these are connected automatically by equally dividing the monochromatic light between reference and sample at a given time. 7.2.2.1 Single-beam Spectrophotometer

As already mentioned in earlier section, the spectrophotometer has two light sources; an UV light (to measure light absorption from ~200 to ~400 nm) and a white light (to measure from ~400 to ~900 nm). With the help of a shutter only one of the lights is allowed to fall on a silver mirror (SM). The reflected light from the mirror passes through an entrance slit and a condensing lens. The lens projects the parallel beam of light on to monochromater (grating).

Colorimetry and Spectrophotometry 117

The monochromater disperses the light into its component wavelength using the wavelength selector, the desired wavelength is selected. Now this selected beam (single beam) of light again passes through the lens and falling on to the cuvette in which sample is kept. In turn, this transmitted light falls on a photo multiplier (PMT). This PMT converts the light energy into electrical energy, which is amplified and measured and recorded on the digital reader (Fig. 7.2).

Fig. 7.2. Single beam spectrophotometer.

7.2.2.2 Double-beam spectrophotometer

In Double-beam spectrophotometer, the monochromatic light, after passing through the lens, splits into two halves by half-silvered mirror (HSM) which is placed on its path. Now 50% of the light passes through the mirror and falls on the reference cuvette; remaining 50% of the light is allowed to fall on the simple cuvette. The difference in intensities of the transmitted lights of reference and sample are computed, measured, amplified and sent to the digital reader (Fig. 7.3).

Fig. 7.3. Double-beam spectrophotometer.

118 Plant Techniques: Theory and Practice

7.2.2.3 Ultraviolet - Visible (UV-Vis) Spectrometer

Ultraviolet-visible (UV-Vis) spectrophotometry is primarily a quantitative analytical technique concerned with the absorption of near ultraviolet (180390 nm) or visible (390-780 nm) radiation by chemical compounds in a solution or in the gas phase. The near-ultraviolet and the visible regions of electromagnetic spectrum provide energy that give rise to electronic transitions. The various colours of visible light and the complimentary colours of solutions absorbing a particular colour are given in Table 7.2. Because of the superimposition of vibrational and rotational transitions, UV-Vis spectra of analytes in a solution show little fine structure, however, it has been widely used for quantitative analysis. UV-Vis spectrophotometer could also be used to study the environmental impact of the treated wastewater, especially presence or absence of chlorides. 7.2.2.4 Infrared Spectrometer (IR Spectrometer)

Infrared spectrometer involves the interaction of infrared radiation with the sample, mostly based on absorption spectroscopy. Sample may be solid, liquid or gas. It has been used for identification of functional group and structural elucidation of chemical compounds. Entire region is divided into two, known as group frequency region and fingerprint region. Range of group frequency is 4000-1500 cm-1 while that of fingerprint 1500-400 cm-1. In group frequency region, the peaks corresponding to different functional groups can be observed. According to corresponding peaks, functional group could be determined. Then each atom of the molecule is connected by bond and each bond requires different IR region, so characteristic peaks are observed. This region of spectrum is known as fingerprint region of the molecule. It can be determined by characteristic peaks. As usually IR spectrum can be visualised in a graph of infrared light absorbance (transmittance) on the vertical axis versus frequency (wavelength) on the horizontal axis. Infrared spectrophotometer has broad range of industrial applications, especially in pharmaceutical industry for identification of raw materials and final product release as well. It has been employed in forensic analysis.

7.2.3 Applications Spectrometric methods are useful for quantitative analysis of a variety of biological compounds. For example, the presence of colourless substances such as proteins, nucleic acids, NADPH, etc., can rapidly be examined by measuring the absorbance at a chosen wavelength which is characteristic and unique for that substance. Most of the analytical methods for quantitative estimation of coloured biomolecules such as chlorophylls, carotenoids, anthocyanins, haemoglobin, etc., are colourimetric or spectrophotometric and are based on measurement at a specific wavelength.

Colorimetry and Spectrophotometry 119

Another important application is the study of enzyme activity in the test sample. The method can be employed for any enzymatic reaction in which the product, substrate or cofactor shows absorbance at a unique wavelength. By simply monitoring the change in absorbance at a particular wavelength, the rate of utilisation of the substrate, formation of the product or changes in the concentration of the coenzyme can be followed spectrophotometrically. For example, nicotinamide adenine nucleotides undergo reduction or oxidation during enzymatic reaction. Though both forms of these nucleotides show intense absorbance at 260 nm but, only the reduced form exhibits specific absorbance at 340 nm. This provides a convenient system for monitoring activities of NADPH-linked oxido-reductases. 7.2.3.1 Nucleic Acid Applications

Spectrophotometer has been used to estimate DNA and RNA, and to analyse the purity of these preparations. Typical wavelength for measurement is 260 nm and 280 nm; besides, measurements at 230 nm and 320 nm can provide further information. Purines and pyrimidines in nucleic acids naturally absorb light at 260 nm. For pure samples, it is well established that for a path length of 10 mm, absorption of 1A unit is equal to a concentration of 50 µg/ml DNA and 40 µg/ml for RNA. For oligonucleotides the concentration is around 33 µg/ml, but this may vary with length and base sequence. So, for DNA, concentration (µg/ml) = Abs260 × 50; these values are known as conversion factors. Several other substances which also absorbs light at 260 nm could interface with DNA values, artificially increasing the result calculated from the absorption readings. To compensate for this a selection of ratios and background corrections have been developed to help eliminate false readings. There is a wide absorbance peak at 260 nm preceeded by a dip at 230 nm. Therefore, to measure the DNA absorption, the 260 nm DNA peak must be distinguishable from the 230 nm reading. If the reading at 230 nm is similar to 260 nm, then DNA cannot be measured accurately; there must be some contaminations in the sample. There should also be a rapid Tail-off from 260 nm down to 320 nm. In this case, 320 nm is often used to measure the background. 7.2.3.2 Direct UV Measurement

A260/A280 ratio The most common purity check for DNA and RNA is A260/A280 ratio. Any protein contamination will have maximum absorption at 280 nm. So, the ratio of these two is taken to check the purity. For DNA, the ratio would be greater or equal to 1.8 that indicates good in terms of purity in the sample, whereas for RNA, this reading should be 2.0 or above. Results lower than this value indicates the impurities of samples.

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A260/A230 ratio An increase in absorbance at 230 nm can also indicate contamination, which in turn affects the reading at 260 for DNA and RNA sample. Many substances absorb at 230 nm, as this is the region of absorbance of peptide bonds and aromatic side chains. Many buffer components (EDTA) exhibit strong absorbance at 260 nm and therefore, alter the results of samples. Other contaminations in the sample, such as proteins, phenols or urea, absorbing at 230 nm, cause the altered results. Phenol concentration also increases sample absorption at 280 nm and therefore can be identified through a lower A260/ A280ratio. An A260/A230ratio of 2 or above is an indicative of pure sample. A320 Background correction Background correction is a process whereby the absorption at a point on the spectrum unrelated to the sample being analysed is also measured and the reading subtracted from the peaks. Absorption at A320 could be due to light scattered caused by particles or precipitation in the sample. Damaged and dirty cuvettes can cause absorption at 320 nm. Contamination with chaotropic salts, such as NaCl, can also cause increased light scattering. Therefore, measuring and correcting for the reading at 320 nm removes any interference from light scattering from cuvettes or other particles present in the samples. Background correction is particularly important, using the small volume cuvette cells and specialist small volume spectrophotometers.

8 Common Adhesives and Mounting Media

8.1 Common Adhesives Adhesives are used for affixing the paraffin ribbons to the microslides. Some of the common adhesives used in plant microtechnique are described below:

8.1.1

Haupt’s Fluid

Haupt’s fluid is the best adhesive for the paraffin ribbons and is also used for affixing unicellular and many colonial algae to the microslide: Plain gelatin Distilled water Phenol crystals Glycerine

1g 100 ml 2g 15 ml

Dissolve gelatin in distilled water at 300 e and then add phenol crystals and glycerine. Stir well and filter it.

8.1.2

Mayer’s Adhesive

It is prepared from the white part of the egg as follows: White of egg Glycerine Sodium salicylate or crushed Thymol crystals

20 ml 20 ml 1g

To the white part of the fresh egg, add glycerine and sodium salicylate. Shake well and filter through sterile cotton or cheesecloth. It is the older standard adhesive. When the solution is kept for more than a month, it loses its adhesive quality. Mayer’s adhesive has less holding quality than Haupt’s fluid and it has got the annoying property of absorbing coal-tar dyes.

8.1.3

Gum Solution

Gum solution is very commonly used by Indian plant technicians:

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Gum arabic Potassium dichromate Water

1g 1g 98 ml

Dissolve gum in water and add potassium dichromate crystals. Depending on the thickness of the sections the concentration of gum in the solution may be increased.

8.1.4

Egg-albumin

Mix equal parts of the white yolk of an egg, water, and glycerine; then add 0.5% sodium benzoate and 1% sodium salicylate. Homogenise the mixture in a blender, filter, and store it in a dropping bottle by keeping it in a refrigerator. Place a drop of the mixture on the slide and smear it into a thin film. Flood the slide with water before floating the sections.

8.1.5

Celloidin

For harder materials (woody sections and sections of certain marine algae) a spray of celloidin is used. Celloidin of 1 or 2% (in equal parts of absolute alcohol and ether) is used to coat the slides after the sections are affixed with Haupt’s adhesive and finally the slides are dried.

8.2 Mounting Media Mounting media preserve the sections in a sufficiently transparent condition for microscopic investigations. A good mounting medium should have the following characteristics as ascertained by Bhandari (1997): i. It should have the correct refractive index (1.48-1.5 or high; see also Table 8.1). ii. It has to be transparent; should not be turned yellow or any other colour when exposed to light or stored for quite a long time. iii. It has to be dissolved in media like xylene or toluene. iv. It should not take too long to harden. v. It should remain amorphous. vi. It should possess good adhesive properties to the glass slide. vii. It should be free from acidity. viii. It should not have a too low melting point (especially when slides are projected). ix. It should be easily available in the market and not too expensive.

Common Adhesives and Mounting Media 123 Table 8.1. Refractive indices of mounting media* Resins

Solvents

Water-soluble Media 1.367

Abopon

1.4372

1.580

Apathy-Lille

1.4189

1.5300 Castor oil Balsam Cedarwood oil (60:40 in xylol) (thickened)

1.490 1.520

Distilled Water 1.3360 Farrant’s glycerol 1.4417 gum Arabic

1.5352 Cinnamon oil Clarite X (60:40 in xylol)

1.567

1.5317 Clove oil Dammar resin (60:40 in xylol)

1.533

Fructose syrup

1.4362

Diaphane (colourless)

1.4777 Creosote

1.538

Glycerol jelly

1.4353

Euparal

1.483

Harleco resin (in xylol)

1.5202 Liquid paraffin

1.471

Sea water

1.343

Lucite (in xylol)

1.4962 Methyl alcohol

1.323

solution of white of egg

1.350

Mahady

1.4879 Methyl benzoate

1.517

Micromount (solid resin)

Oil of aniseed Oil of bergamot

1.557 1.473

Permount (solid resin)

1.5376 Oil of turpentine

1.473

Polystyrene (in xylol 1:1)

1.473 1.5378 Olive oil polyvinyl alcohol 1.386

Polystyrene (solid resin)

1.6279 Terpinol Toluene Xylol

Balsam (dry) Balsam (in xylol)

1.535

Absolute ethyl alcohol

1.5322 Aniline oil

Glycerol

1.4674 Glycerine (50% aq. sol.)

1.484 1.4956 1.4982

* Refractive index of crown glass 1.518.

8.2.1

Mounting Media for Temporary Preparations

8.2.1.1 Glycerine Jelly

Glycerine Jelly is commonly used for temporary preparations: Gelatin Glycerine Water

1 part 7 parts 6 parts

1.397

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Warm gelatin in water for 2 hrs and pour glycerine, stir it for 15 mins. To this mixture add 1% phenol as a preservative. Glycerine Jelly’s refractive index is 1.47. 8.2.1.2 Lactophenol

Lactic acid Phenol Glycerine Water

100 ml 100 g 100 ml 100 ml

Dissolve phenol in water without heating to prevent oxidation. Later add glycerine and lactic acid. For special purposes, other ingredients are also added such as dyes to stain the colourless tissues and copper acetate to preserve green pigments. 8.2.1.3 Calcium Chloride (Herr, 1992)

Herr (1992) has used 20% calcium chloride as a mountant for a specific purpose. The procedure is as follows: 1. Cut fresh sections (120 µm thick) on microtome from stems of Psilotum nudum, Coleus blumoi, and Pelargonium peltatum, and roots of Setereasea purpurea. 2. Fix the sections in FPA (formalin: propionic acid: 50% alcohol in a ratio of 5:5:90 v/v) for 24 hrs and store in 70% ethanol. 3. Transfer the sections to water directly or through the ethanol series. 4. Later sections are put in a mixture containing 1% phloroglucin (250 mg in 25 ml of 20% aqueous calcium chloride) + 4 ml cone. HCl. Lignin stains instantly. 5. Now each section is transferred to the microslide with a few drops of phloroglucin-calcium chloride solution or fresh 20% calcium chloride solution with HCl. 6. Other weak and strong acids may be substituted for HCI.

8.2.2

Mounting Media for Permanent Preparations

8.2.2.1 Natural Resins

Natural resins are mixtures of terpenes, carboxylic acids, and their esters. Their acid number is high (usually above 50), so they diffuse cationic stains. Among several media composed of natural resins, dammar balsam has been reported to cause the least fading of haematoxylin (Vijayaraghavan and Shukla, 1990). While xylene and toluene are the usual solvents for resinous media, benzene has been suggested to be a better solvent for Canada balsam as it offers lesser oxidation sites in its molecule than xylene. Chloroform and ethanol have also been used as solvents for Canada balsam. The addition of 2,6-di-tert-butyl-p-

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cresol (to 1% concentration) into the mountant has been found to inhibit the oxidation and the consequent acidification of the media.

A. Canada Balsam It is extracted from the bark of Abies balsamea. It is a viscid yellowish or greenish substance and soluble in xylene, dioxane or trichloroethylene. Its refractive index is 1.54. B. Dammar Balsam or Gum Dammar It is obtained from Shorea viesnerii of Dipterocarpaceae. It is a vivid yellowish substance prepared by dissolving 100 g gum dammar in 100 ml xylene. Dammar balsam is completely soluble in alcohol and terpentine; refractive index 1.52. Other natural resins like Oregon fir balsam (from the red fir, Pseudotsuga menziesii of Pinaceae), Styrax (Liquidamber orientalis and L. styracifolia of Hamamelidaceae), Tolu balsam (from Myrospermum toluiferum of Leguminosae) and Yucatan gum elemi (from the gumbo limbo, Bursera simaruba of Burseraceae) are dissolved in suitable organic solvents and used as mountants. Tolu balsam (RI = 1.64) and styrax are preferred when fine details of diatoms are needed. 8.2.2.2 Synthetic Resins

A. Euparal This mixture is commonly used and consists of the following ingredients: Camsal Gum sandarac Eucalyptol Dioxane Paraldehyde

10 ml 40 g 20 rnl 20 ml 10 ml

Camsal is obtained by mixing equal quantities of camphor and phenyl salicylate. This mountant has the property of intensifying haematoxylin stain. It is employed in 95% alcohol, refractive index 1.48.

B. Styrax, Hyrax and Naphrax These are commonly used for mounting diatoms. Their refractive indices are 1.6, 1.8 and 1.8, respectively. C. Clarite X It is a naphthalene polymer, colourless, strictly neutral, homogeneous and dries quickly. It is usually employed in 60-70% toluene; refractive index 1.56. However, it cannot be used for acetic-carmine preparations and becomes milky with ethanol (Sharma and Sharma, 1972). The tissues have to be thoroughly washed and passed through ethanol-xylol grades before mounting in clarite X.

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D. Noel’s Staining cum Mounting Medium Solution A 0.5% Aniline blue in lactophenol Solution B 0.5% Basic fuchsin in lactophenol Solution C Iodine 3 g, potassium iodide 6 g, lactophenol 1 L. Prepare the medium by mixing four parts of solution A, one part of solution B and five parts of solution C.

E. Dibutylphthalate Polystyrene Xylene (DPX) Mixture This mixture is also known as Kirkpatrick and Lendrum’s DPX and it is prepared by mixing 7.5 cm3 of tricresylphosphate with 40 cm3 of xylol to which add 10 g of distrene. Since the DPX mixture is completely colourless, it preserves the colour of basic dyes and also medium, so that it provides clear visibility. The pH of the mixture does not change upon long storage. 8.2.2.3 Light-polymerising Plastics

Light-polymerising plastics were tested by Silverman (1986). These plastics harden quickly when exposed to UV light and they do not require solvent evaporation for hardening. Methacrylate-based ultraviolet polymerising plastics, namely Impruv Potting Compound 363 and Impruv Sealant/Adhesive 365, tested by Silverman (1986) are superior to ordinary mounting media because they harden very rapidly and eliminate the formation of air bubbles under the coverslip. The optical features of the plastics were found to be useful for both light and fluorescence microscopy,

8.3 Substitute for Immersion Oil in Microscopy Immersion oils of commercial importance are recommended for microscopy since their refractive index, i.e., 1.515 matches the oil immersion objectives. Thickened cedarwood oil is prone to drying and it is difficult to remove from the slide. In some cases, commercial, immersion oils contain polychlorinated biphenyls (PCB’s) which are highly toxic (Nashel and Fischmann, 1983). An immersion medium should have a refractive index of 1.515 to 1.520 and it should be non-drying and sufficiently viscous. Methyl salicylate is one of the substitutes for immersion oil. Synthetic methyl salicylate has got a refractive index of 1.522 (Weast, 1970). It is viscous, non-volatile, nondrying, and less expensive. Other organic compound substitutes for cedar oil are mineral oil, mineral oil and bromonaphthalene, methyl benzoate, glycerine oil of resin, linseed oil, oleum recini, or different combinations of these oils.

9 Plant Collection and Herbarium Techniques The herbarium (plural: herbaria) is an anglicised term, herbar, that refers to the collection of dried, pressed, and preserved specimens mounted on sheets, and these are arranged based on accepted classification, further, these are a ready reference to plant taxonomists (Narayana et al., 2016). The collection of plants and field preparation of specimens are the fundamental aspects of plant systematics. The collection of live plant specimens from the wild, sometimes referred to as plant hunting, is an activity that has been happening over the centuries. Similarly, Herbarium specimens and permanent records of plant species of a particular locality at a given time are playing a greater role in plant identification. Therefore, the plant specimens should be carefully collected and herbarium specimens properly prepared and preserved for future generation studies.

9.1 Plant Collection and Preservation The following points should be taken into consideration while collecting plant specimens: i. Collect entire, vigorously growing typical specimens. ii. Select such plant individuals that represent all phases of natural population. iii. Avoid collecting insect damaged and infected specimens. iv. For herbaceous perennials, collect underground parts such as roots, tubers, rhizomes, bulbs, etc. v. Collect those specimens of flowering plants that contain flowers, buds, fruits and seeds, as the taxonomic key is constructed based on these plant parts. vi. If the specimen is larger than the size of the herbarium sheet, then it should be cut into 2 or 3 parts and pressed on the series of sheets. vii. Collect plant individuals with leaves in-tact, as different kind of foliage prove helpful in identification of species. viii. For woody plant, collect bark and wood samples.

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ix. For palms, several herbarium sheets are necessary to show the various portions of leaf, inflorescence, and fruit. Photographs of the tree and of each part are essential for the palm tree. x. Spiny plants must be placed under a board first and stood on before pressing to prevent tearing the paper. xi. Succulent plants need to be killed first by soaking in methylated spirit for 15-20 minutes and later mounted on aherbarium sheet. xii. Aquatic plants must be floated onto the dish of water, and later taken onto the blatters, excess water is drained off; then press the plants on the piece of waxed paper. xiii. Tall rosette plants and grasses may be pressed completely by bending them once or more into the shape of V. N or M (Australian Natural Herbarium, 2016). Avoid collecting rare or uncommon plants; if necessary, collect only seeds and grow them in the botanical garden, finally preserved on the herbarium sheet.

9.1.1

Field Equipment

The following are some commonly used field equipment for plant collection and preparation of herbarium specimens: i. Field plant press typically consists of hardwood frames (for rigidity, Fig. 9.1), corrugated cardboard ventilators (to allow air to flow through the press), a blotter paper (to absorb moisture), and a folded old newspaper (to contain the plant material). The plant press is tightened using straps with buckles or bolts with wing nuts. The objective of plant pressing is to remove moisture in a shortest period of time, while preserving the morphological characters of plant intact, whereby it is readily mounted on herbarium sheet (an acid-free cardstock for long storage). The standard herbarium sheet should be not more than 48 cm long x 28 cm width. ii. Field notebook with ruled pages should be carried by plant collector, to record the field data such as date, place, locality, habitat, elevation, local name, collector number (numerical series starting with 01 and continuing through out the lifetime of the collector), collector name, etc. iii. Collecting bags are plastic bags and close them with rubber bands; small brown paper bags should be used for collecting fruits, seeds and bryophytes. iv. Tie-on tags should be made of water-proof material and are large enough to take your name and field number. They may also be used to label collecting bags. v. Digging and clipping tools such as trowel, diggers, hammers, pruning shears, garden clippers, geological pick, heavy sheath knife, etc., are all used for digging and clipping of plants (Fig. 9.1).

Plant Collection and Herbarium Techniques 129

Fig. 9.1. Plant press.

vi. Vasculum made up of aluminium sheet with a hinged light lid, should be carried away by the plant collector. Plants that are not pressed in the field, should be placed in the vasculum to preserve their freshness for some time. vii. Liquid preservative such as Formalin-acetic-alcohol (FAA) is used to preserve the flower buds and vegetative parts such as roots, tubers, leaves, stems, etc., for anatomical and embryological studies. A mixture of chloroform, 95% ethyl alcohol and glacial acetic acid (6:3:1) has been used for cytological studies. viii. Collection bottles (glass or plastic bottles) with leak-proof screw cap are used to collect small-sized materials to be preserved in liquid preservatives. ix. Topographic maps are necessary for locating position and determining the altitude. A GPS (Global Positioning System) unit is equally important for fixing accurate latitude and longitude of the locality. x. A safety gear with long-sleeved shirt and long trousers to keep sun off, a jumper and waterproof raincoat to keep the cold and rain off, and first aid kit are necessary during field trip. Apart from above, other tool, such as waxed paper (used for pressing viscid or weak-looking plants), felt-tipped pens (for numbering the bryophyte collections), Hand lens of 10× (for field observation and identification) card board storage boxes (to store dry materials) and camera should be carried during plant collection from wild.

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9.1.2

Field Notes

During the time of collection, a numbered tag should be tied on the specimen. The best system of giving number is to provide the consecutive numbers, right from 01 and then go up. All duplicates of a collection should bear same numbers. The collection is recorded in the field notebook along with the information about the collection, such as exact locality including latitude and longitude, altitude, nature of the habitat (soil type, topography, slope), vegetation type and associated species, date of collection, collectors name, etc. Apart from this, record other features which would not be evident from the pressed specimens, e.g., whether it is herb, shrub or tree, height, branching, root system, colour of the stem and flower, colour and odour. Some ecological data should be collected; it includes insect damage, weather damage/lightning strike, visits by insects/animals (pollination), usage by local people, common name or other forms of cultural knowledge with respect to that particular plant specimen.

9.1.3

Pressing and Preserving the Specimens

After collection, specimens should be pressed as quickly as possible. If not possible, they may be stored in plastic bags, preferably wrapped in old newspapers (dry paper). Bags should be packed tightly and should be kept cool and moist. Make sure that each bag correctly labelled its place of collection (locality) and name of plant (if known). The specimen can be placed carefully in a fold of several sheets of newspaper and put it in the press with numbered tie-on tag attached. If necessary, occasionally insert a sheet of corrugated cardboard in between the newspapers as ventilator. All specimens are placed for pressing in such a way, so that the entire bundle is ultimately of almost uniform thickness in the middle and sideways. Once the plant specimens are arranged, close the press and tightly bind the press with the straps or ropes to prevent wrinkling of the specimens; now the press with specimen is ready for drying (Fig. 9.1). Inspect the presser daily and change the newspaper during first few days, until plants get dried. Delicate plants and petals may be lost during changing and these should be kept in tissue-paper folders throughout changes. Properly dried plant specimens are brittle and fragile; therefore, all plant parts are carefully preserved.

9.1.4

Drying of Specimens

Usually during drying, the presser containing the specimens should be kept in the sunlight for about 24 hours. After that, the presser is opened, and specimens are placed in fresh blotters. Now presser is again tightly roped. The wet blotters are dried in the sun for reuse. For three to five days wet blotters or driers are changed daily as mentioned above, until the specimens are completely dried. Artificial heat may also be used for the drying press.

Plant Collection and Herbarium Techniques 131

But specimens should not be dried in an oven. In humid regions, or in rainy season, the plants may be dried in a drier. A drier in the form of wooden box, 5 feet length, 18 inches in breadth, is made up of 1/10-inch-thick boards. Light bulbs of 60 watts are fitted inside for producing heat. Small openings should be made at the bottom of the box for the entry of air which gets heated and dries the plants. In the process, it is possible to dry some 100 specimens bundled in a presser within a short time about 8-12 hours. An excellent drying chamber may also be made by using wooden or mental box with an open top that accommodates a presser. This drying chamber is equipped with an electric heater with a fan.

9.1.5 Plant Identification For plant identification, flora books from various regions across the globe are available. Besides, world-class herbaria such as Royal Botanic Gardens (Kew), Missouri Botanical Garden (Saint Louis), Singapore Botanic Gardens, Hortus Botanicus Leiden (Netherlands), Botanical Research Institute of Texas (BRIT, USA). Botanical Survey of India (BSI, Calcutta) among others are rendering yeomen service for the identification and characterisation of plants. Try to find a book that contains an illustrated glossary to assist in defining terms that encounter when using the dichotomous key to identify the plant collection. Make sure that the text botanist is using, should be relevant to that particular geographical area from where plant has been collected. Some expert botanists are needed for identification of particular group of plants. For example, orchid plant collected from North-Eastern Himalayas (India); for this, expert orchidologist of that region is needed for identification. Botanical experts, sometimes, receive email photographs (plant images), based on which plant identification is made. However, physical specimen is usually required for definitive identification of plants. The common practice is to send the plant specimen as a gift to herbarium with which the botanist is associated and get the definitive identification, if he wishes to confirm it. Local herbaria can be particularly useful in providing connections to local botanists who can also assist with identification (Franklin Rahman, 2015).

9.1.6

Mounting and Labelling of Plant Specimen

Before mounting, dry plant specimen is dipped in a saturated solution of mercuric chloride dissolved in ethyl alcohol to prevent infection of fungi, insects and others. Then mounting is performed. The process through which a dry specimen and a label are glued on to the hard board paper (herbarium sheet) is called mounting. The standard size of herbarium sheet is 28.75 × 41.25 cm. The sheet should be long lasting and durable; use of acid-free paper of good quality (100% cotton rag) is advisable for it. Mounting is performed with help of glue (Fevicol or Elmer’s glue). The label is always glued down at the right-hand corner of the sheet. The excess glue may be removed by placing

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a newsprint paper/blotter by gently pressing it over the specimen. Place a sheet of wax paper over the entire specimen, and then place weights or heavy books on the top of the specimen until the glue dries. Fruits and seeds are collected in polythene pockets, properly sealed and mounted at one corner of the herbarium sheet. During drying specimen becomes very brittle, sometime parts may be detached from the main stem, all these parts should be collected and preserved in the polythene pockets and mounted in the herbarium sheet. It is also suggested the use of quick drying liquid plastic as a good medium for mounting of specimens. The combination of ethyl cellulose and resin is dissolved in a solvent (methyl/alcohol and toluene) to give a thick syrup used for mounting. This liquid is sprayed on the lower surface of the dry specimen with the help of sprayer and paste on the sheet which dries very quickly. This is the quick method of gluing the specimen on to the herbarium sheet.

9.1.7

Labelling of Plant Specimens

A label, usually of 10.5 × 6.5 cm dimension, is pasted on to the lower right-hand corner of the herbarium sheet; label should contain the definite information about the plant specimen (Table 9.1). These herbarium sheets are carefully preserved in wooden or steel cabinets/almirahs is a separate room (Fig. 9.2). Finally, identification of pressed, dried and labelled specimens has been performed with the help of various floras as mentioned in previous section. Table 9.1. Specimen label Scientific name of the plant: Name of the family: Local/common/vernacular name (in English): Locality of the specimen: -Plant name (village) -Country -State -Altitude -Latitude -Longitude Habit & Habitat: Diameter at breast height (DBT), 1.3 m above soil line: Flower Colour & Odour: Additional information: -Colour of leaves and stems, associated plants, economic importance Collector’s name: Date of collection: Collection number:

Plant Collection and Herbarium Techniques 133

Fig. 9.2. Herbarium cabinet with herbarium sheets (Courtesy: Botany collections - The National Herbarium of Canada).

9.2 Herbarium An herbarium (Herbaria in plural) is a collection of dried and pressed plant specimens arranged according to a classification system, and associated data used for scientific and taxonomic studies. While most of the early herbaria were prepared with sheets bound into books, Carl Linnaeus came up with the idea of mounting the plant specimens on free sheets that allowed their easy reordering with the cabinets. Plant specimens are usually mounted on a sheet to hard board as described in earlier section. Properly dried, pressed and identified plant specimens are placed in thin paper folders (species covers) which are kept together in thicker paper folders (genus covers) and in turn all these are kept in the herbarium cabinets. Succulents are preserved in liquid preservatives instead of being pressed and dried. Bulky plant parts, such as fruits, cones of several gymnosperms, etc., are dried without pressing and stored in special boxes. A herbarium containing specimens collected locally and from different parts of the world, is housed in a big building. These herbarium sheets are kept in the herbarium cabinets forever. However, periodically we have to check herbarium specimens and ensure that they should be free from pathogens. Herbaria in different countries remain associated with universities, scientific societies, research in states, botanical gardens, arboretas, colleges and some well-funded government organisations. In addition to their taxonomic importance, herbaria are commonly used in the fields of plant anatomy, morphology, ecology, conservation biology, biogeography, ethnobotany and palaeobotany. The herbarium sheets provide biogeographic information, which in turn is useful in locating the rare and

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endangered species and to trace the expeditions of explorers and plant collectors. In fact, specimens are the best sources of genetic material for DNA analyses and pollen for palynological studies.

9.2.1

History of Herbarium

The concept of herbarium was developed by Italian plant taxonomist, Luca Ghini during 1490-1556 as he had collected plants, dried and affixed them on paper in the form of herbarium specimen. Later Gherardocibo and Falconer learnt the art of herbarium development from their teacher, Ghini, and collected and preserved the specimens in 1553. According to Holmgren et al. (1981), the great herbarium of the museum of Natural History, Paris was established in 1653. In olden days, during 17th and early 18th centuries, there was a usual practice of preparing a herbarium by mounting the specimens and binding them into volumes. The plants were usually sewn by thread. Later Linnaeus during 1707-1778 started the current practice of mounting the specimens on a single sheet of hard paper and preserving them carefully in the herbarium boxes. From that simple beginning of Linnaeus, various herbaria have been developed and housing millions of specimens in steel boxes across the globe. The herbarium of the Indian Botanical Gardens, Calcutta, known as Central National Herbarium,was established in 1832. In USA, the oldest herbarium was started at Salem College in 1772. The Herbarium of Academy of Sciences, Philadelphia was started in 1812, University of Michigan in 1838, Missouri Botanical Garden in 1857, and US National Herbarium in 1868. In early days, herbaria were confined with plant specimens of local or regional significance. In the course of time, they have been preserving the specimens of tropics and subtropics across the globe. Now herbarium have become the centres of advanced research in the field of plant taxonomy. Herbaria of Paris, Kew, Leningrad (St. Petersburg, Russia) and Calcutta are big centres of taxonomic research.

9.2.2

Role and Functions of Herbaria

i. Preserved specimens in herbaria are useful for scientific and taxonomic research. ii. Herbaria should be useful for student education and impart training to students in herbarium practices. iii. Herbaria should have the specimens of regional and global scale. iv. A complete information about the species of genus/genera of the family would get from the herbarium only. v. List of endangered species of any region is prepared only by herbarium specimens. vi. Herbaria are useful in preparing the monograph of the particular genus or family.

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vii. Unknown plant collection could be identified by the preserved specimens in herbaria. viii. The present knowledge of the distribution of plants and evolution is based mainly on the herbarium specimens available across the globe. ix. Herbarium specimens are useful in resolving the several taxonomic disputes of the species. x. Information regarding the exact area, region or location of the plant may be collected from herbarium. xi. Herbaria preserve type specimens, and thus serve as a repository of experimental voucher specimens and more helpful in chemotaxonomic and chromosomal studies. xii. Herbaria should publish the catalogues periodically. xiii. Herbaria should have the responsibility to report the adverse effects of plants (any poisonous plants) on livestock and sensitise the public on disease epidemics. xiv. Herbaria preserve the national plant wealth and provide the scientific information to the public with respect to the plant that are useful to mankind. xv. Herbarium material is used to study the anatomy, embryology, palynology and chemical aspects of plants. xvi. Herbaria should provide the specimen as a loan to other institutions for scientific research but not for commercial propose.

9.2.3

Precautions While Using the Herbarium

Certain precautions should be taken while using the herbarium sheet. These are as follows: i. As the dry specimens are brittle and easily damaged, utmost care should be taken while handling the plant specimens. ii. Support the specimens with a ventilator when carrying them over short distances. iii. If the specimen is damaged due to pests and insects, please inform immediately to the curator and he will disinfect the herbarium sheet to prevent further damage. iv. Specimen borrowed from other institutions, should be carefully handled, and kept in fire-proof, dust-free and pest-free mental boxes. v. Specimen taken on loan from other institution should be returned before the loan period is over.

9.2.4

Major Herbaria in the World

Some of the largest herbaria in the world are Royal Botanic Gardens (K) Kew, England (65 million collections), National d’HistoireNaturelle (Pl), Paris, France (65 million), Komarov Botanical Institute (LE), St. Petersburg, Russia

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(5 million), New York Botanical Garden (NY), Bronx, New York, USA (4.3 million), Missouri Botanical Garden (MO), St. Louis, USA (2.9 million), University of Tokyo Herbarium (1.7 million), etc. Further, some of the major Indian herbaria with the number of their specimens deposited are given below (Sharma, 1993): 1. Central National Herbarium, Botanical Survey of India, Calcutta 2. Herbarium of Forest Research Institute, Dehradun 3. Botanical Survey of India (BSI) i. ii. iii. iv. v. vi. vii. viii. ix. x. xi. xii. 4. 5. 6. 7. 8. 9.

Indian Botanical Garden (CAL), Howrah, Calcutta Northern Circle, Dehradun Southern Circle, Coimbatore Western Circle, Pune Eastern Circle, Shillong Central Circle, Allahabad Andaman & Nicobar circle Arid zone circle, Jodhpur Sikkim Himalayan circle Arunachal Field station Herbarium of Industrial Section, Indian Museum Deccan Circle, Hyderabad (started in 2006)

.. 25,00,000 .. 3,00,000 .. .. .. .. .. .. .. .. .. .. .. ..

12,000 1,11,188 2,33,000 1,50,000 2,50,000 48,000 20,000 16,500 10,000 10,000 50,000 300

Blatter Herbarium, Bombay National Botanical Research Institute (NBRI) Herbarium Herbarium of Rajasthan University, Jaipur Herbarium of Delhi University Herbarium of Jiwaji University, Gwalior Herbarium of School of Plant Morphology, Meerut College, Meerut

.. 1,00,000 .. 80,000 30,000 .. 15,000 .. 15,000 ..

25,000

In addition to above, BSI also maintains the Orchidaria (to protect and preserve the orchids) such as National Orchidarium, Yercaud (Tamil Nadu) and another one at Shillong (Meghalaya). Some other well-known herbaria in India are Herbarium of the Division of Botany, Indian Agricultural research Institute (IARI), New Delhi; Regional Research Institute (Ayurveda), Medicinal Plant Herbarium, Bangalore (Karnataka); Fischer Herbarium, Institute of Forest Genetics and Tree Breeding (ICFRE), Coimbatore (Tamil Nadu); Herbarium of Calicut University, Calicut (Kerala); Delhi University Botany Herbarium; New Delhi; National Bureau of Plant Genetic Resources (NBPGR), National Herbarium of Cultivated plants, New Delhi; Herbarium, French Institute of Pondicherry, Pondicherry; Rapinat Herbarium, St. Joseph’s College, Tiruchirapalli (Kerala); JNTBGRI Herbarium, Trivandrum (Kerala); Lloyd Botanic Garden Herbarium, Darjeeling (West Bengal), among others.

10 Biostatistics

Introduction Plant biology encompasses diverse areas of research using plant materials that include, study and analysis of various phenotypic and biochemical parameters. It helps in understanding the range and quantum of genetic diversity among plants and plant groups. The measurements of diversity may vary significantly and considerably at both inter- and intraspecific levels, resulting in challenges of identification and resolving them into distinct plant groups. Both continuous and discontinuous variations are noticeable in many of the metric traits in plants. Seed numbers, crop yield and fruit size are just only a few characters to mention wherein diversity can be significantly observed. Often, most of such research-based investigations are designed, keeping the objectives of plant breeding and crop improvement, which generate a vast amount of information. Irrespective of the type of study undertaken, a meaningful and concrete conclusion is extremely difficult to deduce from raw data, unless certain appropriate biostatistical analysis are adopted for meaningful interpretations. Thus, bio-statistical tools have become an indispensable part for accurate analysis and precise interpretation of data (Peace and Cannon, 1988; Compton, 2006). The basic framework of any biostatistical study involves determination of appropriate measurable parameters to be considered, after which, a population of interest is to be selected for observations and data collection. Four types of basic studies can be undertaken, such as: survey, observations, experiments, and meta-analytical treatise. Experimental studies are of maximum relevance wherein whole or part of the samples is measured for various traits and comparative observations are made, both before and after the experimental procedures (Compton, 2012). Conventionally, statistical parameters like measures of central tendency, dispersion, standard deviation, variance, correlation coefficients, bivariate, partial and multiple regression, etc., are methods in-vogue (Steel and Torrie, 1980; Gomez and Gomez, 1984). In addition, several other statistical methods such as ANOVA (Analysis of Variance), Regression and path analysis, etc. are also used for understanding the diversity among plant genotypes and populations.

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10.1 Basic Statistics (Descriptive Statistics) The experimental data are often classified into four major types: (a) Categorial Data, (b) Nominal Data, (c) Ordinal Data, (d) Numerical Data. Specific characteristic traits such as “flower colour”, “fruit colour”, “seed shape (round/ wrinkled)”, etc. are some examples of categorial data. Although typically, these characters are represented by non-mathematical parameters, however, sometimes, a mathematical value maybe assigned to depict each variation of the trait being examined. On the other hand, nominal data represent discrete values or labels used for denoting variables. However, these variables have no quantitative value and can be depicted in any order. Examples of such variables can be “flower colour”, “pod shape/colour”, etc.

10.1.1 Experimental Designs Any experiment, particularly in plant biology, is designed keeping in mind, the type of sample(s) to be analysed, its distribution and frequency of occurrence in its natural habitat and the principal objective of the study, which impact the outcome and will yield results. Depending upon all the parameters, and type of experiments, the samples being investigated can be grouped into categories. Experimental designs can typically be of the following kinds: 10.1.1.1 Completely Randomised Block Design

Completely Randomised Block design (CRD), assigns treatments following no distinct pattern, rather randomly, to the experimental samples (Fig. 10.1). Each sample gets equal chance of receiving a particular treatment and the outcome will have no selection bias.

Fig. 10.1. Illustration of completely randomised block design: Six experimental samples were assigned random treatments (1-6).

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Randomised Block design It is widely used in agricultural research experiments. Such designs categorise similar experimental units together into blocks. However, the experimental subjects are selected randomly for a treatment (Fig. 10.2).

Fig. 10.2. Illustration of randomised block design: Six experimental samples were assigned into specific blocks (blue or green); however, treatments are assigned to the samples randomly.

10.1.1.2 Latin Square Design

Latin Square Designs assign Latin symbols to each experimental treatment. Each of such treatment when assigned to a particular sample should be of equal duration and all the experimental subjects should receive the treatment for the same number of times (Fig. 10.3).

Fig. 10.3. Illustration of Latin square design. Treatment α signifies treating the plantlets with Hormone A while Treatment ß signifies treatment with Hormone B. The plant samples in each group are treated for the same time duration.

10.1.1.3 Factorial Design

Factorial experiments are favoured when an experimental design involves simultaneous analysis of more than one factor, each of which may result in multiple observations.

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10.1.1.4 Split Plot Design

It is a statistical experimental design, wherein, the several key factors that may impact the outcome are dependent on each other and cannot be replaced (Fig. 10.4).

Fig. 10.4. Example of split plot design to study the effects of irrigation pattern and fertilisers on plant growth. Four experimental plots were chosen to grow either paddy and wheat. Each of these fields were subjected to either manual irrigation (I1) or drip irrigation (I2). All the plots were further split into two, to test whether organic fertilizer (F1) or chemical fertiliser (F2) were suitable for healthy growth of the crops.

Average In statistical terms, average maybe defined as the value that is central in a particular data set. It is typically obtained by dividing the total of the values of all the data in a set by the total number of observations. Average has great significance in analysis of biological data, as it gives an idea about the central value occurring in each set of observations, which represents that particular trait. Mean and average are statistical terms, often used interchangeably, and refers to the value representative of a set of data. Mean, Median and Mode are commonly used measures of central tendencies in data analysis. Median, on the other hand, is the value that occurs in the middle, when a set of observations are arranged either in ascending or descending order. The value occurring most frequently in a dataset is referred to as Mode. Use of all the three tools yields clear and accurate information from the raw data. Mean can be applied to all the numerical data obtained for investigating a particular trait/ characteristic of plant samples. Mathematically, it is represented as: ∑n Mean (µ) = —— n where, n: number of observations

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Arithmetic Mean The mean is a measurement of unit most frequently used to describe a frequency distribution of same type. The arithmetic mean or average is calculated by dividing them sum (total) of all the individual values of data series by the total number of items. — X1 + X2 + X3 + X4 + X5 ........ + Xn Arithmetic Mean or X = n — ∑x or X = —— n where ΣX is sum of all Xs from 1 to n and n = number of items. Example: The heights of 10 plants are found to be 15, 21, 17, 19, 21, 16, 14, 17, 15 and 13 cm. Calculate the arithmetic mean of the plant height. Solution: ∑x Sum of the values of all items — Arithmetic Mean X = —— i.e., n Number of items ΣX = X1 + X2 + X3 + X4 + X5+X6 + X7 + X8 + X9 + X10 = 15 + 21 + 17 + 19 + 21 + 16 + 14 + 17 + 15 + 13 = 168 cm n = 10 — 168 X = —— = 16.8 10 So, the arithmetic mean of the plant height is 16.8 cm or the average plant height is 16.8 cm. In case of many observations, i.e., more than 10, the arithmetic mean can be calculated by preparing a frequency distribution table. Frequency is the number of individuals representing a class. 1. When the data are discrete and unclassified but tabulated, the arithmetic mean can be calculated in the following way: Item

X1

X2

X3

X4

-------

Xn

Frequency

f1

f2

f3

f4

-------

fn

f1x1 + f2x2 + f3x3 + f4x4 ....... fnxn _ Arithmetic Mean (x) = fn ∑fx ∑fx —— or x = —— = ∑f n Example: The seed yield (in grams) per plant for 30 maize plants was found to be 110, 115, 112, 108, 110, 120, 115, 1128, 108, 115, 121, 115, 110, 112,

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108, 121, 110, 110, 115, 110, 115, 112, 110, 115, 112, 112, 115, 110, 112 and 110. Calculate the average yield per plant from the given data. Solution: From the given data, there are some figures which appear more than _ once, for example seed yield 110 g/plant has been recorded in nine plants. So, the frequency of plants with 110 g. seed yield in this case is 9. The frequencies of other values can also be calculated in the same way and the data can be tabulated as follows: Class

Frequency (f)

Seed yield per plant x

fxX

No. of plants representing

108 110 112 115 120 121

3 9 7 8 1 2

324 990 784 920 120 242

Total

30

Σfx = 3380

∑fx _ 3380 = —— = 112.66 gm/plant Arithmetic Mean (x) = —— 30 n So, the average yield per plant is 112.66 grams. II. When the data of continuous series are classified, then only the frequency of individuals falling in the class appears. In such cases, for calculating the arithmetic mean, the mid value of each class is taken into consideration and the subsequent steps of computation are like those of discontinuous series. Example 1: Calculate the arithmetic mean from the following data: Class Frequency

10-20

20-30

30-40

40-50

50-60

6

10

12

18

4

Solution: Class

Mid value of class (x)

Frequency (f)

fx

10-20 20-30 30-40

15 25 35

6 10 12

90 250 420

40-50 50-60

45 55

18 4

810 220

Total

Σf = 50

Σfx = 1790

∑fx _ 1790 = —— = 35.8 Arithmetic Mean (x) = —— 50 ∑f

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Example 2: Calculate the arithmetic mean of flowers per plant from the following data: Class (flowers/Plant) Frequency (No. of plants)

0-10

10-20

20-30

30-40

40-50

3

10

15

12

8

Solution: Class

Mid value of class (x)

Frequency (f)

fx

0-10 10-20 20-30 30-40 40-50

5 15 25 35 45

3 10 15 12 8

15 150 375 420 360

Σf = 48

Σfx = 1320

Total

∑fx _ 1320 = —— = 27.5 Arithmetic Mean (x) = —— 48 ∑f So, the average number of lowers per plant is 27.5 or 28. Merits of Arithmetic Mean 1. It is a mathematical mean. 2. It is easy to calculate. 3. It is always definite. 4. It establishes a simple relation among the values of data series. Demerits of Arithmetic Mean 1. It cannot be determined by simple visual observation of data. 2. It cannot be presented by chart or graph. 3. It is not suitable for qualitative studies. 4. Sometimes it does not show representation of series.

10.1.2 Median When the values of all items of a series are arranged in increasing (ascending) or decreasing (descending) order it is usually called an array and the middle item of an array is called median. The median divides the series into two groups; one group in which the values of items are less than the middle value and the other group in which the values of the items are greater than the middle item. Mean is denoted by Me or Mdn. The methods of calculating the median are comparatively simple. The value of median is not affected by change in extreme values. If the amount of data in a series is odd, the median is the middle value. But if the amount of data in a series is even, the median is the average of the two middle values.

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10.1.2.1 Methods of Determining Median

1. For unclassified and untabulated data: To calculate the median, the data are first arranged in increasing or decreasing order and then the following formula is used: i. For odd number of data in the series n+1 Me = —— , where n = number of items or data. 2 Example: The heights (in cm) for 9 plants are given below. Find out the Median Height 67, 65, 70, 68, 62, 63, 64, 63, 66. Solution: The height measurements can be arranged in ascending order as follows: 62, 63, 64, 65, 66, 67, 68, 69, 70. The number of the data in the above array n = 9, n+1 9+1 Media (Me) = —— = —— = 5 (i.e., value of the 5th data in the array) 2 2 = 66 ii. For even number of data in the series. The median is calculated as follows: _n th item + _n + 1 2 2 Me = 2 Suppose, for example, there are 10 data in a series So, Me = =

(102 )+ (102 + 1) 2

=

5 + (5 + 1) 5 + 6 = 2 2

Value of 5th data + Value of 6th data of the array

2 Example: The number of flowers recorded on plants are: 15, 10, 8, 12, 13,7, 11, 14, 9, 16. Find out the median value of flowers per plant. Solution: The given numbers of flowers on 10 plants can be arranged in ascending order as under: 7, 8, 9, 10, 11, 12, 13, 14, 15, 16. Total amount of data in the above array n = 10 (even number) _n + _n + 1 2 2 Me = 2 _ 10 _ 10 5th value + 5 + 1 value 2 + 2+1 = = 2 2 =

11 + 12 23 = 2 = 11.5 flowers 2

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Example: Calculate the median of the following series of data obtained by measuring the height or 16 plants: 9, 10, 10, 8, 9, 7, 8, 11, 7, 12, 14, 12, 11, 14, 15, 13. Solution: The given data of plant heights are arranged in ascending order as follows: n = 16 (even number)

Me =

_n + _n + 1 2 2

=

_ 16 _ 16 2 + 2+1

2 2 10 + 11 = 2 = 10.5 So the media height = 10.5 cm

=

Value of 8th item + Value of 9th item 2

For grouped data (i) Discontinuous or Discrete series of data. To calculate the median for discrete grouped data, first of all _2n + _2n + 1th the cumulative frequency of whole series is obtained. The value of data against

n+1 th 2

cumulative

frequency will be the median for odd number of data and the mean of values against cumulative frequencies will be the median for series containing even number of data. Example: Calculate the median of the following data obtained by counting the number of flowers on 19 plants: Class (No. of flowers/Plant)

1

2

3

4

5

Frequency (No. of plants)

3

4

6

3

3

Calculation: Class (No. of flowers/plant)

Frequency

Cumulative frequency

1 2 3 4 5

3 4 6 3 3

3 7 13 16 19

n = 19 (odd number) + 1 cumulative frequency Media (Me) = Value of data against n—— 2 19 + 1 = Value against —— cumulative frequency 2 19 + 1 20 = = = 10 2 20

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Since cumulative frequency 10 is included in 13 which represent the class value 3, therefore, for cumulative frequency 10, the class value will be 3 which is the median. Example: Calculate the median for the following data recorded for heights (in cm) of 80 plants: Class (Plant height)

119

120

121

122

123

124

125

4

9

14

18

15

13

7

No. of plants

Solution: Class [Plant height (in cm)]

Frequency (No. of plants)

Cumulative frequency (Cf)

119 120 121 122 123 124 125

4 9 14 18 15 13 7

4 13 27 45 60 73 80

n = 80 (even number)

Class value against _n th + _n + 1th cumulative frequency 2 2 Media (Me) = 2 =

80 th + 80 + 1 cumulative frequency 2 2 2

The class values for cumulative frequencies 40 and 41 are included in the class value of cumulative frequency 45 which is 122. Therefore, Mean (Me) 122 + 122 = 122 2 For classified grouped data. The median is determined in the following way: =

(a) first, the cumulative frequency of all the classes is obtained from the given frequencies. (b) median class value is determined which is class. (c) then that class is ascertained whose cumulative frequency precedes that of median class (c.f.) (d) the median is calculated by the following formula:

Biostatistics 147

N + 1− F Media (Me) = L + 2 ×i f N − CF 2 Media (Me) = L + ×i f Where, L = N= Cf = F= i=

lower limit of median class sum of all the frequencies (N = Σf) cumulative frequency of class preceding that of median class frequency of median class class interval of the median class (i.e., upper limit - lower limit or L2 - L1).

Example: The number of seeds produced by 55 plants of plot are given in the following table. Calculate the median seed number of a plant. Class interval

40-50

50-60

No. of plants

5

8

60-70 70-80 80-90 8

15

90100

100110

110120

4

3

2

7

Solution: Class

Frequency (f)

Cumulative frequency (Cf)

40-50 50-60 60-70

5 9 9

5 14 23 = c.f. of the class preceding that of median class

70-80

15(f)

38

8 4 3 2

46 50 53 55

80-90 90-100 100-110 110-120

Σf = 55

To find out median class we use formula n 2 55 So, Median class = 2 = 27.5 Now, the median class value falls within the cumulative frequency of class interval 70-80, therefore, the median class is 70-80. The width or class interval (i) = L2-L1 = 80 - 70 = 10 Now the median is calculated by the following formula:

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n − cf 2 Me = L + ×i f 27.5 − 23 = 70 + × 10 15 45 45 = 70 + × 10 = 70 + 15 15 = 70 + 3 = 73. Merits of Median 1. It is calculated easily and located exactly. 2. It is not affected by abnormally large or small values. 3. Its size cannot be changed much by adding a few more items. 4. Median can be used in quantitative measurements. Demerits of Median 1. The median of two or more series cannot be calculated by using the median of the component series. 2. It may not be represented in central data. 3. It cannot be used where weightage is given to some items.

10.1.3 Mode Mode is another unit of measurement of central tendency which is used to obtain a quick estimate of central tendency. Mode is defined as the value which appears again and again for maximum number of times in a set of observations. In other words, mode is that variety or item in the series which shows highest frequency. It is denoted by Mo or Z. Sometimes, in a series of data all the varieties have equal frequency. In such a series there is no mode. In some series more than one data have equal frequency. In such series there may be more than one mode. Such a series is said to be multimodal series. That series is analysed properly to get the genuine mode. 10.1.3.1 Method of Determining Mode

For different types of data, the methods of computation of Mode are different. i. For Non-grouped data. If the values of all the items are given separately then the mode is the value of that item which shows largest frequency. Example: Find out the mode size of shoe from the following numbers of shoes 2, 7, 4, 1, 7, 7, 6, 7, 7, 6, 5, 3, 2, 2, 7, 7, 5 Mode = 7. ii. For discrete series of data. Mode of discrete series of data is determined easily because the figure with highest frequency is mode of the series.

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Example: Determine the mode from the following data regarding the number of flowers per plant: Number of flowers/plants

1

2

3

4

5

6

7

8

9

10

No. of plants

4

7

10

19

3

25

20

5

3

1

Solution: The above Table of data distribution indicates that 5 flowers per plant is represented 35 times out of 129 plants which is the largest frequency. Therefore, the value of Mode is 5 flowers per plant. For grouped data, the mode can be calculated by identifying the class with highest frequency by grouping method which is detailed as under: i. Grouping Method: When in a given series two or more classes show same frequency, mode cannot be determined by inspection method. Sometimes, the frequencies of neighbouring classes affect the mode determination in such a way that a class other than the true mode class with highest frequency is treated as mode. In order to avoid such mistakes, grouping method is used to determine mode which is the best method. Grouping method is outlined below: Grouping Table: For this, table is made in which first column is meant for observed items or classes and the next 6 columns are meant for frequency. In the first frequency column, the frequency of each item or class is mentioned. In the second frequency column, the given frequencies are grouped in pairs starting from first and the combined values of paired frequencies are mentioned. In the third frequency column, the groups of two frequencies each are made starting from the frequency of the second item or class and the combined values are noted against different groups. In the fourth frequency column groups of three frequencies each are made starting from the first frequency and the combined values of three frequencies in different groups are recorded. In the fifth frequency column, the frequencies of first column are made into groups of three each starting from the second frequency and combined values of 3 frequencies of different groups are recorded. In the sixth frequency column, the frequencies of first column are bracketed into groups of three each starting from the third frequency and the combined value of three frequencies is noted against each group. ii. Analysis Table: This is made to record the items or classes represented by highest frequency of different columns. iii. Determination of Mode: The value of item or class which is represented by highest in the analysis table is mode. The grouping method for determining mode is illustrated by taking the example of the following data:

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Example: Find out the mode of the following series: No. of fruits per plant

30

31

32

33

34

35

36

37

No. of plates

3

9

5

4

8

7

6

2

(i) Grouping Table Frequency column

Number of fruits per plant

1

2

30 31 32 33 34 35 36 37

3 9 5 4 8 7 6 2

]12 ]9 ]15 ]8

3

5

4

]17

]14 ]12 ]13

]19

] 18 ] 21

6

]17 ]15

(ii) Analysis Table Frequency column

Items or class with highest frequency 30

1 2 3 4 5 6 Total

31

32

33

31 31

32

2

32 2

33 33 2

34

35

34

35

34 34 34 4

35 35 3

36

37

36 2

From the above analysis table, 34 fruits/plant appeared four times which has the highest frequency. Therefore, the mode (Mo) = 34. (iii) For Classified Data. Firstly, the class with highest frequency, i.e., Mode class is identified either by inspection method or grouping method and then the mode is calculated with the help of the following formula: Mode (Mo) = Lt + Since L2 – L1 = i Mo = L1 +

fm- f1 2fm – f1 – f2

fm- f1 2fm – f1 – f2

× (L2 – L1)

×i

Where, L1 = Lower limit of mode class fm = Frequency of mode class f1 = Frequency of class preceding the mode class

Biostatistics 151

f2 = frequency of the class succeeding mode class i = Width of the mode class (i.e., upper limit – lower limit or L2 - L1 of mode class) Let us take the following classified data table for calculation of mode here: Class

40-50 50-60 60-70 70-80 80-90 90-100 100-110 110-120

Frequency

5

9

9

15

8

4

3

2

From the above data table, it is apparent that the class 70-80 has highest frequency, i.e., 15. Therefore, 70-80 is the mode class. The mode can now be calculated with the help of the following formula: fm- f1 Mo = L1 + ×i 2fm – f1 – f2 Here, L1 = 70, fm = 15, f1 = 9, f2 = 8 and i = 10 15 - 9 × 10 2× 15 – 9 – 8 6 70 + × 10 30 – 9 – 8 6 6 70 + × 10 = 70 + × 10 13 30 – 17 60 70 + = 70 + 4.61 13 74.61

∴ Mo = 70 + = = Mo = Mo =

Merits of Mode 1. It is based on all values and can be subjected to further algebraic treatments. 2. It represents group as well as extreme items and is not affected unless they are in modal class. Demerits 1. It requires intricate calculation. 2. It cannot be used for further calculations and interpretations.

10.1.4 Range Statistical range is the difference between the highest and lowest values in a data set. It is a standard measure of dispersion or variability among samples. The value of range can never be negative; however, it can be positive or equal to zero. In terms of significance, range is often considered to be unreliable as its value is dependent on two extreme observations, and not the entire data set

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as a whole. However, it is a rather convenient way to express the quantum of variations existing among different samples for a particular parameter under study. Typical parameters that make use of range in depicting data pertaining to plant samples are, chromosome numbers of a particular genus, plant height, length of seedling, etc. Mathematically, range of a given set of data is obtained by the following formula: Range = Highest Observation – Lowest Observation This is the simple measure of dispersion. The image of distribution is the difference between the largest and the smallest data or items of a series. The range of data provides certain amount of variability. Suppose that the heights (in cm) of ten individual wheat plants in a plot are: 70, 50, 60, 90, 80, 65, 40, 75, 55 and 85. The range of plant height (R) = Highest value – Lowest value or R = H – L = 90 – 40 = 50 i.e., plant height ranges from 40 cm to 90 cm Co-efficient of Range =

Highest value – Lowest value Highest value + Lowest value

Example: Find out the range and co-efficient of range for the following series of data of plant heights (in cm): 17, 15, 12, 16, 10, 18, 20, 14, 11. Solution: Range (R) = Higher value – Lowest value R = H–L Here, H = 20, L = 10 So, R = 20 –10 = 10 cm H – L 20 – 10 10 1 Co-efficient of Range = = = = H + L 20 + 10 30 3 1 Co-efficient of R = 3 Example: Calculate the range and co-efficient of range of plant height from the following data: Plant height (in cm)

10-15 15-20

20-25 25-30 30-35 35-40 40-45 45-50

Number of plants

5

20

10

40

35

Solution: Range (R) = H – L Here, H = 50 and L = 10 ∴ Range = 50 – 10 = 40 cm H – L 50 – 10 40 2 Co-efficient of R = = = = = 0.66 H + L 50 + 10 60 3

30

25

15

Biostatistics 153

Merits of Range 1. It represents the two extremes of observations. 2. Very easy to calculate. 3. It indicates two extremes between which the data or items of the series are dispersed or spread. Demerits 1. Range cannot be considered as reliable measure of variability because it is based on only two extremes of observation, i.e., highest and lowest values. 2. It is extremely influenced by the variation in sampling. 3. Range is much sensitive to size of sample or population. 4. It is not based on frequency of data.

10.1.5 Standard Deviation Mean Deviation Deviation refers to the difference between the calculated means (arithmetic mean, median and mode) and the value of individual data. Mean deviation is the mean of the deviations of all data of a series from the real mean value. Theoretically, the mean deviation is a true measure of dispersion. In the calculation of mean deviation all the values (+ and –) of deviation are taken as positive. When the mean deviation is divided by the mean of the series, the value so obtained is called co-efficient of mean deviation. Computation of Mean Deviation 1. When the data are unclassified and untabulated ∑(dx) (i) Mean deviation (ox) from arithmetic mean = n

δx (ii) Co-efficient of Mean deviation or Co-efficient of δx = x ∑(dm) (iii) Mean deviation from Median or δM = n (iv) Co-efficient of Mean deviation from Median (Co-efficient of δM) δM = M ∑(dZ) (v) Mean deviation from Mode (M⸰ or Z) or δZ = n and

(vi) Co-efficient of deviation from Mode (δZ) = Where, d͞ x = deviation from arithmetic mean dM = deviation from Median dZ = deviation from Mode n = number of items or data in a series

∑(dZ) Z

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x͞ = Arithmetic mean M = Median Mo or Z = Mode Example: Calculate the mean deviation and co-efficient of mean deviation from the arithmetic mean from the following data on seed weight: 4.8, 5.5, 6.5, 7.1, 7.3, 7.5, 8.0 and 8.5. Solution: S.No.

1 2 3 4 5 6 7 8 X=8

Seed Weight (x)

4.8 5.5 6.5 7.1 7.3 7.5 8.0 8.5

Arithmetic (Mean(x)

x=

55.2 8

= 6.9

55.2

Deviation from the Arithmetic Mean (all in +) 4.8 5.5 6.5 7.1 7.3 7.5 8.0 8.5

d = x-x – 6.9 = 2.1 – 6.9 = 1.4 – 6.9 = 0.4 – 6.9 = 0.2 – 6.9 = 0.4 – 6.9 = 0.6 – 6.9 = 1.1 – 6.9 = 1.6

∑ dx = 7.8

∑(dx) 7.8 n = 8 = 0.975 0.975 Co-efficient of mean deviation from Arithmetic Mean = dx x = 6.9 = 0.141 2. When the data are ungrouped and tabulated The mean deviation of ungrouped data is calculated with the help of the following formulae: Mean deviation from Arithmetic Mean dx =

∑ fdx n ∑ fdM (ii) δM = , and n ∑ fdZ (iii) δZ = n Where, f = frequency (i) δx =

3. When Data are Classified and Tabulated The mean deviation of tabulated and cdata is also calculated with the help of same formulae as are used for ungrouped data. Example: 100-seed weight of maize recorded for 100 plants are presented in the following Table. Calculate the mean deviation and co-efficient of mean deviation from the data table.

Biostatistics 155 100-Seed weight (in gm)

10-20

20-30

30-40

40-50

50-60

Number of plants

10

25

42

15

8

100-seed Mid-value Frequency Weight of the (f) of the class class (x) class 10-20 20-30 30-40 40-50 50-60

15 25 35 45 55

10 25 42 15 8

fx

͞X=

150 625 1470 675 440

∑fx n

3360 — 100 =33.60

n =100 Σfx = 3360

Deviation d͞ =x-͞x 15-33.6 = 18.6 25-33.6 = 8.6 35-33.6 = 2.6 45-33.6 = 12.6 55-33.6 = 22.6 ͞ = 65.0 Σfdx ͞ = Σfx

fd͞ x

186 215.0 109.2 189.0 180.8 880

∑ fdx 880 n = 100 = 8.8 δx 8.8 Co-efficient of δx = x = = 0.261 33.6 The statistical parameter, Standard Deviation (S.D.), as the name suggests, is a measure of the extent of dispersion of a particular set of data from its mean. The mean or average of a dataset reflects the central value, but does not provide any information on the outliers, which may have significant bearing on information analysed from the dataset. Mathematically, Standard Deviation is depicted by the symbol “σ” (Lee et al., 2015). For a given set of data, if the value of S.D. is low, it reflects that all the values are clustered around the mean, while a higher S.D. indicates how far a particular value is spread out from the mean. Mathematically, Standard Deviation is deduced by: δx =

Standard Deviation (σ) =

√

n

∑ (xi – µ)2 i=1

n–1 Standard deviation is routinely used for statistical analysis of biological data as it yields information on whether a dataset is far apart/closely scattered from the mean. It basically accounts for the frequency of variations occurring in the data, which helps in predicting consistency of a particular parameter/ trait being investigated among various plant samples assessed. The value of standard deviation also gives an idea of the extent of heterogeneity in the samples. It is the positive square root of mean of deviations of individual values of a data series from the arithmetic mean of the series. In other words, the square of standard deviation is equal to mean of the square of deviations of individual observations from the arithmetic mean. It is also called mean square deviation from the mean and is denoted by a Greek symbol σ. Let us suppose that n is the total number of observations (Say, X1, X2, X3, X4 ....... Xn) and the sum of the values of all observations is S.

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S Thus, the mean ( ͞x ) = n The deviations of individual observations can now be expressed as follows: X1 – ͞x,X2 – ͞x,X3 – ͞x,X4 – ͞x ……Xn – ͞x The square of standard deviation SD2 or σ2 =

(x1 – ͞x )2 + (x2 – ͞x )2 + (x3 – ͞x )2 + (x4 – ͞x )2..... (xn – ͞x )2

∴S.D. or σ =

n

√

(x1 – ͞x )2+(x2 – ͞x )2+(x3 – ͞x )2+(x4 – ͞x )2.......(xn – ͞x )2 n

If the values of observed data are equal, the mean deviation will be zero which represents its lowest value. The value of standard deviation will increase with the increase in deviations of individual data from their arithmetic mean. To make the standard deviation comparable, co-efficient of standard deviation is calculated which is the ratio between standard deviation of observation ∑ d2 series and its mean, i.e.., standard deviation (S.D. or σ) = n Standard Deviation Co-efficient of SD = Arithmetic Mean =σ x

√

10.1.5.1 Methods of Calculating Standard Deviation

Generally, the following three methods are used for calculating standard deviation:

1. Direct Method. 2. Short Cut Method. 3. Step Deviation Method. i. Direct Method: In this method, first the arithmetic mean (͞x) of the series is calculated. the deviations of individual values from the mean are calculated (d = X – ͞x) which may be either positive or negative number. The value of d2is always a positive figure. Then the standard deviation (S.D. or σ) is calculated as follows: (i) When the data are unclassified, the following formula is used to calculate the standard deviation: S.D. or σ = Where n = number of data

√

∑ d2 n

Biostatistics 157

Example: Calculate S.D. from the given data collected on 10 plants: 55, 58, 60, 62, 68, 65, 66, 72, 75, 69. 650 Solution: Arithmetic mean = 10 = 65.0 Data

Mean x

Deviation d = X-͞x

d2

Σ d2

1

55

55 – 65 = –10

100

358

2 3 4 5 6 7 8 9 10

58 60 2 68 65 66 72 75 69

650 — 10 = 65.0

58 – 65 = –7 60 – 65 = –5 62 – 65 = –3 68 – 65 = +3 65 – 65 = 0 66 – 65 = +1 72 – 65 = +7 75 – 65 = +10 69 – 65 = +4

49 25 9 9 0 1 49 100 16

S. No.

√

358 10 = 6.0 (ii) When the data are ungrouped and tabulated, the S.D. is calculated with the help of the following formula: S.D. or σ =

√

∑ fd2 , where f = frequency of data n (iii) If the data are classified and tabulated then also the same formula is used in the calculation of S.D. as is used for ungrouped data. But the difference here is that, in case of classified data, the mid-values of different classes are used to calculate the mean (x) and then the deviation (d) of each class is calculated by subtracting the mean from mid-value of each class, i.e., d = (X – x). The deviation squares (d2) are then multiplied by the frequency (f) of the respective classes to get (fd2). The standard deviation is calculated with the help of the following formula: ∑ fd2 S.D. or σ = n S.D. or σ =

√

ii. Short Cut Method for Calculating S.D. The method involves the following steps: 1. Any value of an unclassified series or mid-value of any class in case of classified series is treated as assumed mean (A). 2. The deviation (dx) of individual item or class is calculated by subtracting the assumed mean from the individual data (in case of unclassified series) or mid-value (X) of each class (in case of classified series) dX = X – A.

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3. The deviation values are squared to get dx2 and then for ungrouped data and for classified data separate formulae are used to calculate standard deviation: Formula for calculation of S.D. of ungrouped data σ =

√

1 or σ = n

∑ dx2 ∑ dx – n n

2

√ n ∑ dx – (∑ dx)

2

2

Formula for calculation of S.D. of tabulated classified data σ =

√

1 or σ = n

∑ fdx2 ∑ fdx – n n

2

√ n ∑ fdx – (∑ fdx) 2

2

Example: Find out the standard deviation of the following observations regarding the number of flowers per plant: 3, 2, 1, 5, 7, 9, 11, 13, 14, 15 80 Solution: Arithmetic mean = =8 10 ∴ (x) = 8. S. No. 1 2 3 4 5 6 7 8 9 10

No. of flowers per plant 3 2 1 5 7 9 11 13 14 15

n =10

S=8

Deviation (d) – (x-͞x) 3 – 8 = –5 2 – 8 = –6 1 – 8 = –7 5 – 8 = –3 7 – 8 = –1 9 – 8 = +1 11 – 8 = +3 13 – 8 = +5 14 – 8 = +6 15 – 8 = +7

Square of deviation (d2) 25 35 149 9 1 1 9 25 16 49 Σ dx2 = 240

240 SD2 or σ2 = 10 = 24 SD = √24= 4.889 Example: Calculate the S.D. of the following data on the heights (in cm) of 10 plants: 73, 75, 80, 42, 57, 65, 52, 42, 47, 67. Solution: Suppose that the assumed mean of the given series is 57.

Biostatistics 159 Height in cm (x)

Deviation dx = (X-A)

dx2

1 2 3 4

73 75 80 42

73 – 57 = 16 75 – 57 = 18 80 – 57 = 23 42 – 57 = –15

256 324 529 225

5 6 7 8 9 10

57 (A) 65 52 42 47 67

57 – 57 = 0 65 – 57 = +8 52 – 57 = –5 42 – 57 = –15 47 – 57 = –10 67 – 57 = 10

00 64 25 225 100 100

Σ dx = 30

Σ d2 = 1848

S. No.

n =10

√

1 n ∑ dx2 – (∑ dx)2 n 1 = 10 √10 × 1848 – (30)2 1 = 18480 – 900 10 √ 1 17580 = 10 √ 1 = (132.58) ∴S.D. = 13. 258 10

S.D. (σ) =

Example: Calculate the standard deviation of data presented in the following table: Height of plants of mutant population (in cm)

50- 6060 70

7080

8090

90100

100- 110- 120- 130- 140- 150110 120 130 140 150 160

Frequency

5

18

26

35

25

7

40

20

16

6

2

Solution: Suppose 105 is the assumed mean of plant height. Class (plant MidFrequency height in value of (f) cm) Clas (x) 50-60 60-70 70-80 80-90 90-100 100-110 110-120

55 65 75 85 95 105=A 115

5 7 18 26 35 25 40

Deviation dx = x-A

fdx

dx2

Fdx2

55 – 105 = –50 65 – 105 = –40 75 – 105 = –30 85 – 105 = –20 95 – 105 = –10 105 – 105 = 00 115 – 105 = 10

–250 –280 –540 –520 –350 00 400

2,500 1,600 900 400 100 00 100

12,500 11,200 16,200 10,400 3,500 00 4,000

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120-130 130-140 140-150 150-160

125 135 145 155

20 16 6 2

125 125 145 155

– – – –

105 105 105 105

n = 200 2

∑ fdx2 (∑ fdx) – n n

√ 94800 – – 320 = √ 200 200

S.D. (σ) =

=

= = = =

20 30 40 50

400 480 240 100 Σ f dx = -320

400 900 1,600 2,500

8,000 14,400 9,600 5,000 Σ f dx2 = 94.800

2

√ 474 – 2.56 = √ 471.77

σ = 21.71 Merits of Standard Deviation 1. Standard deviation is mathematically computed and reliable. Therefore, it is used in quality research work. 2. It is based on every individual data of a series. 3. It is least affected by sudden deviation of any type. 4. It is a definite measure of dispersion. Demerits of Standard Deviation 1. Method of its calculation is complicated and so inconvenient. 2. In this measure, higher values are given more importance than the lower values which affect the S.D. value.

10.1.6 Standard Error of Mean Like Standard Deviation, Standard Error of Mean (SE) also reflects the extent of variation in a set of data. However, a principal difference exists among the two (Lee et al., 2015). While standard deviation gives an idea about the extent of variation prevalent among individuals in a data set, standard error of mean reflects the variation present in a population. A higher value of SE is expected with a lower value of n (where n = population size), and thereby a higher value of SD. Similarly, the value of SE decreases, with the higher value of n and thus a smaller SD. Mathematically, Standard Error of Mean is given by: σ Standard Error of Mean (σx̄ ) = SE = √n Standard Error of Mean is another statistical parameter that is indispensable in accurately assessing variations in a given data set. It basically provides information among the extent of dispersion among the population mean. A lower SE is often associated with a bigger sample size which is representative

Biostatistics 161

of the entire population. On the contrary, a lower sample set will lead to observations with higher variations, thereby, a higher SE which may result in inaccuracies and lead to false conclusions. If instead of considering one sample, several samples are taken from the same population, the standard deviations of different samples may vary. This variation is measured by standard error or standard error of mean. Standard error is a measure of dispersion of different hypothetical means around the grand mean. It is a statistical constant which measures the dispersion of sample means around population mean. Standard error is calculated by the following formula: Standard deviation or S.D. or σ ____________________________ √n Where S.D. or σ = Standard deviation of individual sample n = Total number of observations Standard Error (S.E.) =

Standard error thus gives an idea as to how a mean of any sample differs from the true mean of population. Therefore, while making a statement on some measurements one should state the average + standard error, say for example, the height of plants of particular species measures 105 + 0.13 cm. Here, 105 cm is the average height and 0.13 is the standard error. Sometimes instead of standard error, probable error (P.E.) is used which is 2/3 or 0.6745 time the standard deviation ∴ P.E. = 2/3 or 0.6745 or Example: In a sample plot of 1 square metre, the height of 100 plants was measured and the standard deviation was calculated to be 5.0. What will be the standard error of the sample? Solution: σ Since, S.E. = √n 5.0 ∴ S.E. = √100 5.0 = 10 = 0.50

10.2 Analysis of Variance (ANOVA; Components of Mean) Variance It is a very useful measure of dispersion which is commonly used for population data. The arithmetic mean of the squares of deviations obtained from the mean is referred to as variance. It is based on standard deviation and the square of the standard deviation is termed as variance.

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V = σ 2 or (S.D.)2 = or V =

∑ d2 ∑ fd2 n or n

∑(X– ͞x )2 n

Most of the research experiments by biologists are designed on the basis of a key hypothesis, whose relevance is reflected in the results obtained (Kaufmann and Schering, 2007). However, the dynamic nature of biological specimens, especially plant systems lead to a great deal of confusion as it is difficult to determine whether the variations obtained in the observations are a result of effect of experimental factors or due to random factors. In general, for any plant-based research experiment, the significance of the data obtained is enhanced if the sampling is broad and comprises of enough individuals representing a population. Sampling of individuals should be carried out with respect to their geographical location, characteristics and other relevant factors prior to setting up of research experiment. As described earlier, the raw experimental data often do not yield the true picture as multiple variables influence the results. Analysis of Variance (ANOVA), first described by Fischer (1925), is one of the most widely used statistical tool, which has utility in a wide range of experiments conducted in biological science for assessing the range and quantum of variation present in them (Edwards, 2005; Lazic, 2008). ANOVA finds applicability in a vast majority of experimental designs in plant biology and biotechnology. It relies on the basic methodology that the raw data obtained after experimental procedure is structured and the variabilities are accounted for, so that the homogeneity in data is regained in subsets of subjects and heterogeneity if any, is attributed to the relevant factors (Kaufmann and Schering, 2007). It basically makes use of the sum of squares of deviation observed from the means of respective samples (as component of mean). Analysis of variance of each trait for the randomised block design can be estimated with the following model equation (Panse and Sukhatme,1978): Yij = m + ai + bj + eij

Effect of jth block Random error associated with the ith genotype under jth block General mean Observed for the ith genotype in jth block

Biostatistics 163 Table 10.1. ANOVA forms assumed for data from randomised block design experiments Source of variation

Degree of freedom

Sum of squares

Mean sum of squares

Expected mean sum of squares

Replication

r-1

SSr

MSr

σ 2 e + g σ 2r

Genotype

g-1

SSg

MSg

Error

(r-1) (g-1)

SSe

MSe

Where, r = g= σ 2e = σ2g = σ2r =

σ 2 e + g σ 2g σ2e

number of replications number of genotypes error variance genotype variance replication variance

From ANOVA, standard error and critical difference between two means (Mean and Expected sum of squares) can be calculated as: Standard error (SE) = (MSe/r)1/2 Standard error of differences (SEd) = (2 MSe/r)1/2 Critical difference (C.D.) = (2 MSe/r)1/2 * t

10.3 Null Hypothesis In statistical terms, a null hypothesis conjectures that the data resulting from an experiment will have no significant bearing on variations from the control population. An alternative hypothesis that contradicts the claim is also accompanied with the null hypothesis. The null hypothesis assumes that any kind of difference between the chosen characteristics that is seen in a set of data is due to chance. Hypothesis testing provides a method to reject a null hypothesis within a certain confidence level only. Null hypothesis may be estimated as: H01: the replication effect is null (replication homogeneity) H02: the genotype effects are null (genotype homogeneity) The two null hypotheses are mutually independent. Choice of significance level (α) = 0.05 and 0.01.

10.4 T-test When a set of data follows normal distribution pattern, it is referred to as parametric data (Hazra and Gogtay, 2016). Parametric tests are carried out based on certain criteria such as: (a) the data comprises of numerical values that follow normal distribution pattern and have homogeneity of variances, (b)

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the observations in a data set are drawn independent of each other, and (c) the samples under investigation are randomly selected from a larger population. T-test is a type of statistical analysis performed on parametric data, which compares two mean values that belongs to different groups and may even be unrelated (Kim, 2015). In analysis of biological data, such a test is applied when there are two distinct groups of observations, in response to a particular experiment. It is popularly termed as Student’s t-Test, and it has three variants, depending upon the type of analysis. Each of the three variants are described below: a. One sample t-Test This variant of t-Test is used when the sample mean differs significantly from the population mean. If the exact value of population mean is not known, its value is derived hypothetically. b. Two sample t-Test This variant of t-Test is also referred to as the independent sample test and is used to determine if the population means of two separate groups are equal or not. c. Paired t-Test It is also sometimes denoted as “dependent sample t-Test” and is a statistical measure to assess if the mean difference between pairs of measurements is zero. The data for each sample in paired t-Test is measured twice, resulting in two sets (paired) of observations.

10.5 Coefficient of Variation Co-efficient of variation is a relative term which measures the relative magnitudes of variations present in observations and is related to the magnitude of their arithmetic mean. Since the standard deviation is independent measure of dispersion obtained from a single series of data, the comparison of variability of two different series of data is not possible directly by standard deviations. To make it possible Karl Pearson used co-efficient of variation. Co-efficient of variation is defined as the ratio of standard deviation to arithmetic mean and is expressed in percentage. σ Co-efficient of variation or CV(%) = ͞x × 100, where σ = standard deviation of a series ͞x = arithmetic mean of series. Thus, when standard deviation of a series is compared to arithmetic mean of the series in terms of percentage, we get co-efficient of variation. It expresses the relative variability of each data series. The co-efficient of variation can be used to compare the homogeneity, consistency and stability of data of two or more series. The data series with high value of co-efficient of variation is said to be unstable and the series with low values of co-efficient of variation are stable and they show good degree of homogeneity of data.

Biostatistics 165

Example: Compare the relative variability of two samples with the following data: Sample A – Arithmetic mean (͞x) = 30 and S.D. = 1.5 Sample B – Arithmetic mean (͞x) = 20 and S.D. = 2.5 Solution: (i) For example A, co-efficient of variation (CV) = S.D. or σ × 100 mean = 1.5 × 100 = 5.0% 30 σ (ii) For sample B, CV = × 100 mean 2.5 = × 100 = 12.50% 20 Example: Calculate standard deviation, variance and co-efficient of variation from the following classified data regarding number of pods per plant recorded on 50 plants in a plot: Number of pods/plant

1-3

3-5

5-7

7-9

9-11

11-13

13-15

No. of Plants

2

6

7

16

11

5

3

Solution: Class (No. of pods per plant) 1-3 3-5 5-7 7-9 9-11 11-13 13-15

Mid-value of class (x)

No. of Plants (f)

Dx = (x-A)

dx2

f.dx2

f.dx

2 4 6 8=A 10 12 14

2 6 7 16 11 5 3

2 – 8 = –6 4 – 8 = –4 6 – 8 = –2 8–8=0 10 – 8 = 2 12 – 8 = 4 14 – 8 = 6

36 16 4 00 4 16 36

72 96 28 00 44 80 108

-12 -24 -14 00 22 20 18

Total

Σ.fdx2 = 428

n =50

Arithmetic mean (͞x) = A + x͞ = 8 +

n 10 = 8 + 0.2 = 8.2 50

∑ fdx2 ∑ fdx – n n

√ 428 – = √ 50

S.D. or σ =

Σ f.dx2

10 50

= √ 8.52 = 2.92

2

2

= √ 8.56 – 0.04

Σ f dx =10

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Variance = σ2 2 = ( √ 8.52 )

σ Co-efficient of variance = x × 100 2.92 or C.V. = × 100 8.2 or C.V. = 292 = 35.61% 8.2 Meaningful comparison and quantification of various morphological and other metric traits can be achieved by using coefficient of variation (Pélabon et al., 2020), a very robust statistical tool. It is often used to decipher and critically analyse the variations of traits among plant samples. Mathematically, it is obtained by multiplying 100 with the ratio of average upon square root. The coefficient of variation (CV) is a statistical measure which shows the level of variability of data in a sample. It measures the dispersion of the data points in a data series, around the mean of the population. It is highly useful in understanding the degree of variation of one data series from the other, even if the arithmetic means differ greatly. The genotypic coefficient of variation (GCV) and the phenotypic coefficient of variation (PCV) can be calculated by the formula given by Burton (1952): (σg2)1/2 σg × 100 GCV = × 100 or x x where,

σp (σg2)1/2 PCV = × 100 or × 100 x x

x is the grand mean of the trait σg is the genotypic standard deviation of the trait σp is the phenotypic standard deviation of the trait

10.5.1 Components of Variance: Genotypic-, Phenotypic- and Environmental Variance: Genotypic and / or phenotypic coefficient of variation can be determined using the formula of Burton (1952) by dividing the square root of genotypic variance and / or phenotypic variance by the population mean and multiplying the resultant with 100. Phenotypic variance was taken as the sum of error and genotypic variance, the later being calculated by subtracting the error mean square from the varietal mean square and dividing the remainder by number of replications, as follows: σ2e = MSSe σ2e = MSSg – MSSe / r σ2p = σ2g + σ2e

Biostatistics 167

Where σ2g, σ2p and σ2e are genotypic, phenotypic and environmental variances, respectively and r is the number of replications of the experiment. MSSg and MSSe are genotypic- and error- mean sum of squares, respectively.

10.6 Heritability Heritability is used in plant breeding studies, to express the proportion of total phenotypic variance among individuals in a population, that can be attributed to the heritable genes, which also determines the degree of resemblance between relatives (Holand et al., 2003; Schmidt et al., 2019). It can be simply defined as the extent to which a phenotype is determined genetically. If genotypic values are considered, we refer to broad sense heritability (H2) and if breeding/phenotypic values are considered, we refer to narrow sense heritability (h2). Heritability in broad sense was calculated on percent basis by the formula suggested by Hanson (1956). h2 = σ2g / σ2p σg2 h2 % = 2 = 100 σp

10.7 Genetic Advance Genetic advance and heritability are usually considered together, as their combined estimate is more meaningful and reliable. Genetic advance (or response to selection) is a measurement of gain obtained from a phenotypic selection for a trait (Ogunniyan et al., 2014). In other words, it is the genetic improvement of the progeny through selection. High genetic advance with high values of heritability is the most preferred condition for selection. Genetic advance or genetic gain or genetic improvement per generation, denoted by G, was computed using the formula given by Johnson et al. (1955): G = i.h2.σp where ‘i’ is the standardised selection differential h2 is the heritability in broad sense σpis the phenotypic standard deviation Expected genetic advance (E ∆ G) represented as genetic advance in percent of mean; Expected genetic advance (E ∆ G) = G × 100 x

10.8 Correlation Genetic correlation is defined as the proportion of variance shared by two different traits, due to genetic causes. Genetic correlations can be either

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positive or negative, which is denoted by a number in the range –1 to +1 (Hill, 2013). The most common cause of correlation is pleiotropy and linkage disequilibrium among genes, each affecting one trait only. Bivariate correlations: Bivariate correlations is a statistical measurement to determine whether any relationships exist or not, between two different variables. It is computed using the following formula: Co Vij rij = (Vi Vj)1/2 where, rij is the correlation coefficient between the traits i and j Co Vij is the covariance between i and j Viis the variance of i Vj is the variance of j

10.8.1 Genotypic, Environmental and Phenotypic Correlations Genotypic, environmental and phenotypic correlations were computed using the formulae suggested by Al-jiburi et al. (1958) and Falconer (1960, 1981): rg(ij) = σ2g (ij) / (σ2g) (i) ×σ2g (j)1/2 re(ij) = σ2g (ij) / (σ2e) (i) ×σ2e (j)1/2 rp(ij) = σ2p (ij) / (σ2p) (i) ×σ2p (j)1/2 where, rg(ij) = genetic correlation coefficient between ith and jth character re (ij) = environmental correlation coefficient between ith and jth character rp (ij) = phenotypic correlation coefficient between ith and jth character σ2g (i), σ2p (i) and σ2e (i) are genotypic, phenotypic and environmental variances respectively for ith characters σ2g (j), σ2p (j) and σ2e (j) are genotypic, phenotypic and environmental variances respectively for jth characters

10.9 Path Analysis Path analysis is often used to determine the cause-and-effect relationships of a dependent variable, with two or more independent ones. It is a form of multiple regressional statistical analysis. In biological experimental models, it finds wide usage in evaluating the effects of a set of variables on a pre-set outcome through several pathways. Statistically, it is represented by structured, linear equations. The exact inter-relationships among them can be depicted with the help of a path diagram, which is essentially a graphical presentation of the cause and effect of the variables involved in a particular pathway. r (x1y) = a + r (x1x2)b + r (x1x2) b + r (x1x3)c ……… + r (x1xn)k

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r (x2y) = a r (x2x1) + b + r (x2x3)c …………………+ r (x2xn)k r (x3y) = a r (x3x1) + r(x3x2)b + c …………………..+ r (x3xn)k r (xny) = a r (xnx1) + r (xnx2) b + r (xnx3)c………….+k where, x1, x2, x3 …………xn are plant traits studied y is seed yield per plant a, b, c ……. k are the direct effects of the traits x1, x2, x3………xn, respectively r(x1x2), r(x1 x3), r (x2 x3) and r (x3xn), etc. are the genotypic correlation coefficients of different pairs of plant traits. These simulations equations are put in the form of matrix and values of direct effects were found as follows: r (x1y) = r (x1x1) r (x2y) = r (x2x1) r (x3y) = r (x3x1) r (xny) = r (xnx1)

r (x1x2) r (x1x3) …. r (x1xn) a r (x2x2) r (x2x3) …. r (x2xn) b r (x3x2) r (x3x3) …. r (x3xn) c r (xnx2) r (xnx3) …. r (xnxn) k

a = r (x1y) b = r (x2y) c = r (x3y) k = r (xny)

r (x1x2) r (x2x2) r (x3x2) r (xnx2)

r (x1x3) …. r (x1xn) -1 r (x2x3) …. r (x2xn) r (x3x3) …. r (x3xn) r (xnx3) …. r (xnxn)

C11 C21 C31 Cn1

C12 C22 C32 Cn2

a = r (x1y) b = r (x2y) c = r (x3y) k = r (xny)

r (x1x1) r (x2x1) r (x3x1) r (xnx1) =

C13 C23 C33 Cn3

….C1n ….C2n ….C3n ….Cnn

where, C11, C12………Cnn are the respective elements of inverse matrix, then, a = C11 r (x1 y) + C12r(x2 y) + C13 r(x3 y) ………… C1nr(xn y) b = C21 r (x1 y) + C22r(x2 y) + C23 r(x3 y) ………… C2n r (xn y) c = C31 r (x1 y) + C32r(x2 y) + C33 r(x3 y) ………… C3n r (xn y) k = Cn1 r (x1 y) + Cn2r(x2 y) + Cn3 r(x3 y) ………… Cnn r (xn y) The indirect effects of the trait via different traits were obtained in the following way: The indirect effect of ith trait via jth trait = r (xiyj) × p where ‘p’ is the direct effect of jth trait.

10.10 Residual Effect The residual effect was obtained by the following formula: h = (1 – ∑ Piy riy)1/2

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where, h = residual effect Piy = path coefficient from x1 to y riy = correlation coefficient between x1 and y

10.10.1 Regression Analysis ANOVA for dose-response data, simple linear regression and curvilinear regression using quadratic equation were computer by the method of Gomez and Gomez (1984). For the dependent variable (Y) the percentage values have been used. The following equations and formulae are used in the regression analyses: Simple linear regression analysis: i) Regression equation Yc = a + bX where, Yc = computed Y from the regression equation a = intercept/level of the line b = linear regression coefficient/slope of the line ii) Calculation of a and b

iii) Correlation coefficient (r)

a = y – b – bx ∑ xy b= ∑ x2 × ∑ y2 r=

where,

∑ xy ∑ x2 × ∑ y2

X=X –X Y=Y –Y where, X = independent variable X = mean of X Y = dependent variable Y = mean value of Y iv) Standard error of estimate (Syx) ∑ (Y – YC)2 Sxy = √ N where, y = dependent variable

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YC = computed value of Y N = number of variables Curvilinear regression analysis: This was estimated using a quadratic regression equation as given below. Y= αD + βD2 + C where Y= computed value of the dependent variable α = linear coefficient of dependent variable per unit of dose β = quadratic coefficient of dependent variable per unit of (dose)2 C = intercept constant D = dose point

Conclusion These above-described statistical tools are of great importance and utility in analysing biological data obtained from various field experiments related to genetics and plant breeding aimed at crop improvement. Incorporation of such tools for the analysis of data not only enhances the authenticity of observation, but also helps in arriving at reliable and meaningful conclusions. In addition, they also aid in minimising errors in the data and eliminate background noise, for enhanced accuracy and meaningful interpretation of biological data.

Part B: Recent Advances in Plant Techniques

11 Histochemical Methods Histochemistry is a combination of histology and analytical biochemistry. In histochemical methods, especially for plant material, various histochemical stains and qualitative reactions have been employed (Table 11.1). Here some important techniques to localise various chemical substances have been described.

11.1 Preparation of Sections for Histochemical Studies Strictly fresh material should be employed for histochemical studies. Generally, slightly thick sections are best for microchemical tests. Sections can be cut with safety razor blades or sharp scalpels with thin blades. Clean the blades before the operation to avoid contamination. In most tests, sections are to be placed directly in the reagent, and in some cases, sections are placed in distilled water before keeping in the reagent.

11.2 Carbohydrates Carbohydrates are classified into Monosaccharides (simple sugars contain aldehyde or ketone groups, e.g., glucose, fructose, etc.), Disaccharides (combined product of two monosaccharides, e.g., sucrose) and Polysaccharides. Mono and Disaccharides are difficult to localise through histochemical methods. Therefore, the following text deals with mainly polysaccharides. The polysaccharides are high molecular weight molecules, and these are divided into homopolysaccharides which yield only one type of monomer upon hydrolysis and heteropolysaccharides which yield not only monomers containing C, H and O but also monomers containing N and S additionally.

11.2.1 Cellulose Cellulose is a homopolysaccharide and occurs abundantly in plants. It is the main constituent of the cell wall and maintains the shape and structural rigidity of plant cells. Cellulose, on total hydrolysis, yields glucose. Numerous β-Dglucose residues are linked together linearly by β-1-4 linkages in cellulose.

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Table 11.1. Some commonly used stains used in histochemical localisation of some plant chemical substances (Khasim, 2002; Badria and Aboelmaaty, 2019) Stains

Chemical substances

Calcofluor MZR

It stains the cellulose and fluorescing intensely under a fluorescent microscope.

Ruthenium red

It stains acidic mucilages, pectin and nucleic acids and appears magenta or red in colour.

Alcian blue

Also stains acidic mucilages, pectin and nucleic acids and gives light blue colour.

FASGA (Fucsina, Aldan blue, Safranina, Glicerina, Agua) complex stain

Under fluorescent microscope, starch granules appear bright-greenish yellow.

Lugol’s reagent

Starch granules appear dark blue to dark.

Aniline blue

Callose stains clear blue.

Resorcinol blue

Callose stains Cobalt blue.

Lacmoid blue

Callose stains greenish-blue.

Azure B

Sieve plate callose stains reddish

Mercuric bromophenol blue

Proteins stain blue

Aniline blue black

Protein stain blue-black

Coomassie blue

Protein stains blue and gives similar result of Aniline blue black

Ferric chloride

It highlights the phenolic compounds through iron precipitation producing a dark colour.

Potassium dichromate

Phenolic compounds appear in brown or redbrown colour.

Vanillin-Hydrochloric acid

This stains tannins, giving red in colour.

Phloroglucinol-Hydrochloric acid stain

It gives red colouration to lignin.

Dragendorff’s reagent

It marks alkaloids in red colour.

Wagner’s reagent

It also stains alkaloids red or red-brown.

Sudan Black

Lipids appear dark blue or black in colour.

Nile blue

Acidic lipids stain blue whereas neutral lipids pink in colour

Azure B

DNA stains blue-green whereas RNA dark blue

Acidic orange

DNA fluoresces bright-yellow and RNA flame red.

Toluidine Blue O

RNA stains purple while DNA blue or bluegreen.

Cresyl violet

RNA stains violet.

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The linear chains of the monomer form microfibrils or bundles of parallel chains that are held together by hydrogen bonds. 11.2.1.1 Zinc-Chlor-iodide Method (Rawlins and Takashi, 1952)

Stock Solution Zinc chloride Potassium iodide Distilled water

50 g 16 g 17 ml

The above three are mixed and iodine is added to them to make a saturated solution. Allow this solution to stand for several days. Collect only the supernatant in brown bottles and use it. The other procedure to prepare the staining solution is to dissolve commercial chloroiodide of zinc in an equal volume of water and add enough iodine crystals to give the solution a deep brown colour. Gahan (1984) prepared this solution by mixing 30 g zinc chloride, 5 g potassium iodide, 1 g iodine and 14 ml distilled water.

Procedure i. Generally fresh sections are taken. ii. Keep the sections in the chloroiodide of zinc solution for a few minutes and observe under the microscope. Result Cellulose turns blue to violet. Biological Reaction Iodine is said to be accumulated within the cellulose when the latter’s basic structure is disrupted. This happens when glucose strands are separated by the removal of hydrogen bonds by treating the specimen with acid or a strong ionic salt solution (Krishnamurthy, 1988). The test is non-specific because hemicelluloses are also coloured blue; if lignin occurs, colour is not developed. So, the removal of lignin, cutin and suberin will enhance the specificity of the reaction. 11.2.1.2 Chlorazol Black E and Lignin Pink Method (Robards and Purvis, 1964)

i. Fresh or FAA fixed tissue is taken; section the tissue, deparaffinise (for paraffin-embedded tissue) and bring down to the water. ii. Keep the sections in distilled water for 5 mins. iii. Stain in 1% aqueous lignin pink for 5 mins. iv. Wash the sections in 95% ethanol to fix the lignin pink. v. Dehydrate twice in absolute ethanol, 2 mins each.

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vi. Stain the sections in 1% chlorazol black E in methyl cellosolve for 5 mins. vii. Dehydrate in absolute alcohol, clear in xylene and mount in Canada balsam.

Result The gelatinous layer which is stained by chlorazol black E becomes black to black grey in contrast to the pink colour taken up by the lignin. When the single stain is desired, then after deparaffinising, sections are directly stained in chlorazol black E and observed under a microscope. 11.2.1.3 Calcofluor MZR Method (Gahan, 1984)

i. Fresh or frozen sections are taken. ii. Stain the sections in 0.1% aqueous solution of calcofluor MZR New for 30 secs. iii. Examine under fluorescence microscopy using UV light.

Result Cellulose and extracellular mucilages are fluorescing intensely under a fluorescent microscope. The above stain is also useful for staining the sections of paraffinembedded and GMA-embedded tissues (Hughes and McCully, 1975). 11.2.2 Starch Starch is the most important reserve food for plants. It occurs as granules in membrane-bound cytoplasmic organelles, like plastids. It is a homopolysaccharide and yields only D-glucose upon hydrolysis. However, starch consists of two basic molecules, amylose and amylopectin. Amylose glucose units are linked in a simple chain α1→4 with a regular branching of the main chain by secondary α1→6 linkages. Starch differs from cellulose in possessing the linkage between glucose units α1→4 and not β1→4, and in possessing some random branching in the chain. 11.2.2.1 Iodine-Potassium Iodide Test (Johansen, 1940; Krishnamurthy, 1988)

Procedure i. Fresh, frozen, or chemically fixed and paraffin-embedded tissues may be taken. ii. Section the material, deparaffinise (for paraffin-embedded tissue), and bring down to the water.

Histochemical Methods 179

iii. Stain the section in iodine-potassium iodide solution (solution is prepared by dissolving 2 g potassium iodide in 100 ml water and adding 1 g iodine flakes to potassium iodide solution).

Result Starch appears blue to black in a few minutes. Newly formed starch may appear red to purple. Biological Reaction The accumulation of iodine in the centre of the helical starch molecule gives colouration in this reaction. Shorter molecules appear red while longer molecules are blue. 11.2.2.2 Periodic Acid-Schiff (PAS) Stain (Feder and O’Brien, 1968)

For staining the tissues with PAS, it is necessary to perform the ‘aldehyde blockade’ which covers the Schiff-positive groups (probably aldehyde groups). It is introduced especially for tissues fixed in acrolein. PAS stain is also useful for GMA sections (Bennet et al., 1976). Aldehyde blockade is performed by using blocking agents like dime done (5,5-dimethyl cyclohexane-1,3-dione) and DNPH (2,4-dinitrophenylhydrazine).

Saturation solution of dimedone Dimedone 5g Distilled water 100 ml Stir it at least for 5 hours and filter. 2,4-dinitrophenylhydrazine (DNPH) Solution DNPH 5g 15% acetic acid in water 100 ml Stir it for an hour and filter. Procedure i. Aldehyde blockade is done either in dime done or in DNPH solution. Slides are left in dimedone at room temperature for 16-24 hours or in DNPH solution for 10 mins. Dimedone treated sections do not show colour and the aldehyde blockade is not reversed by treating the section with acid. DNPH treated sections show a yellow colour, often useful as counterstain. ii. Oxidise the sections in 1% periodic acid for 5-10 mins. iii. Wash the slide with running tap water for 5-10 mins. iv. Keep in Schiff’s reagent (Feulgeri’s fuchsin) for 10-30 mins. (Schiff’s reagent is prepared in the following manner: add 0.5 g basic fuchsin to 100 ml of boiling distilled water. It is followed by shaking for 5 min.

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After cooling to 25°C, filter the solution and add 30 ml. of HCl and 3 g potassium metabisulphate. Later store the solution in dark for 24-48 hrs. Now add 1 g decolourising charcoal to the solution and shake well. Allow the preparation to stand for about half an hour before filtering. Note that the red colour disappears, and a completely colourless solution is formed at the end; see also Gurr, 1965. v. Transfer the slides quickly and directly to three successive baths of 0.5% sodium metabisulphite, 2 mins each. vi. Rinse in running water for 5-10 mins. vii. Sections may be counterstained with haematoxylin or toluidine blue, or without counterstain sections are examined under a light microscope.

Result Starch reacts very strongly and appears red. Some complex polysaccharides, especially in the compound middle lamella, stain a purplish red to magenta colour. In Carthamus tinctorius L., epidermal and endothecial cells of the mature anther, as well as pollen grains, are filled with PAS-positive substance (Fig. 11.1A, B).

Fig. 11.1. A-B Transverse section of anther in Carthamus tinctorius L. (A) Pollen grains filled with carbohydrate reserves and endothecial cells possess PAS-Positive fibrous bands; (B) Epidermal cells possess distinct PAS-Positive outer and radial walls and these cells are covered by extracellular PAS- positive substance indicated by arrow mark. (c = connective; e = epidermis; Bar = 20 µm) (Courtesy: Dodappa, 2005)

Biological Reaction The oxidant, periodic acid, is acted upon the 1,2-glycol linkage within the sugar molecule of carbohydrate (Fig. 11.2). As a result, two free aldehyde groups are produced. These aldehydes react with leucofuchsin resulting in highly coloured complexes.

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Fig. 11.2. Schematic representation of PAS test for polysaccharides (Adopted from Vijayaraghavan and Shukla, 1990).

11.2.2.3 FASGA (Fucsina, Aldan Blue, Safranina, Glicerina, Agua) Staining for Starch Granules Using Fluorescence Microscopy (Revila et al., 1986)

Procedure F.A.S.G.A. staining technique is performed in the following manner: i. The cotyledons of lentil seeds (Lens culinarisi are soaked in distilled water for 4 hrs. This material is then processed in two different ways. a. After the material is fixed in 7% acrolein for 20-24 hours at 4oC, the tissue is then washed thrice in distilled water at 30 mins. per wash. Sections of about 12-16 µm are cut with a freezing microtome. b. Fixation in FAA for 18-24 hrs. Fixed tissue is dehydrated in ethanol series. Then the dehydrated tissue is embedded in paraffin and sections of about 10-12 µm thickness are cut. ii. The sections are stained with FASGA complex stain for 10 mins. Stock solutions of stains are prepared in distilled water. These are:

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Solution A: 1% safranin B: 1% a1cian blue C: basic fuchsin Glycerine Distilled water up to

4.5 ml 9.0 ml 1.0 ml 50.0 ml 250 ml

After mixing of above solutions, the resultant one is filtered and used for staining the sections. iii. Later the stained sections are dehydrated briefly by passing in 96% and 100% ethanol, in xylene and mounted in Canada balsam, or simply by washing the stained sections in water and mounting with water. Specimens are observed under an epifluorescence microscope.

Result Under epifluorescence microscope, starch granules appear very bright greenish-yellow and the cytoplasm is full of dark spherical structures which are identified as a protein with TEM (Revila et al., 1986). It is possible to observe both intact and degradative starch granules in this procedure. When sections are observed with visible light, the cellulosic cell walls are stained blue and, the nucleus and lignified tissues are stained red. The starch granules are not stained under these conditions. FASGA staining procedure permits observing other cell structures along with intact as well as degradative starch granules. Sections stained with this complex stain are long-lasting and do not show loss of colour even after three years. 11.2.3 Callose Callose is a homopolysaccharide and it has β-1, 3-Glucon subunits. It forms a major component of cell walls of pollen tubes, microspore and megaspore mother cells, intine of pollen grains, sieve elements, and cell walls which are immediately adjacent to the freshly formed wounds. Many botanists believe that callose is deposited in response to wounding. Callose can be localised by aniline blue which exhibits deep yellow fluorescence under a fluorescence microscope. It is also stained with lacmoid blue (resorcin) which is more specific than aniline blue. 11.2.3.1 Aniline Blue Method (Johansen, 1940)

Procedure i. Fresh tissues are preferred. If fixed in FAA, the material should be thoroughly washed with water. ii. Section the material, deparaffinise (if necessary), and bring it down to the water.

Histochemical Methods 183

iii. Stain the sections in a dilute aqueous aniline blue (0.005%) for an hour or in 0.005% aniline blue in 50% ethanol for 4-24 hrs. iv. Excess stain is removed by treatment with glycerine. v. Mount in gelatin; for paraffin-embedded sections, dehydrate, clear and mount in DPX.

Result Callose stains a clear blue. 11.2.3.2 Aniline Blue Fluorescence Method (Smith and McCully 1978a, 1978b)

Sections of fresh tissues or aldehyde fixed, or GMA embedded tissues are used for this method. This method is also adopted for epoxy embedded material from which resin should be removed (Sutherland and McCully, 1976). Peterson and Fletcher (1973) used a modified procedure to stain cleared whole mounts.

Procedure Section the material and mount the sections in 0.05% aqueous aniline blue. Use freshly made staining solution (in 0.067 M phosphate buffer, pH 8.5) or store in a brown bottle in the refrigerator. Discard when the blue colour changes to greenish. Stained sections of embedded material are permanently mounted by gently air drying the sections (do not wash out the stain before drying) and then mount in immersion oil of low fluorescence. The gentle air-drying ensures that some water molecules that remain present are essential for the formation of the fluorescing complex, especially in sieve plates and new cell walls (O’Brien and McCully, 1981). Result Sieve plate callose fluoresces in a yellow colour under a fluorescence microscope. The fluorescence fades during observation but this can be restored by thoroughly washing the section with tap water for 12 hours followed by restaining. 11.2.3.3 Resorcinol Blue Method (Eschrich and Currier, 1964)

Preparation of the Stain Resorcinol blue is prepared by dissolving 3 g pure white resorcinol in 200 ml distilled water, to which add 3 ml 0.88 NH3 and heat without boiling for 10 mins on a steam bath. Store reddish-brown solution, stoppered with cotton till a dark, blueish colour appears. Heat on steam bath further for 30 mins, filter this hot solution, and heat again till no significant NH3 escapes (check with moistened red litmus paper). Store stock solution in dropping bottles. Fresh sections are mounted with three drops of stock solution freshly diluted with 10 ml of water. Callose stains cobalt blue. Other features, like

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lignified walls, also stain blue but it turns to red when the sections are mounted in buffer at pH 3.2 whereas the callose remains cobalt blue (see also O’Brien and McCully, 1981). 11.2.3.4 Lacmoid Blue Method

i. Fresh, frozen or aldehyde fixed, and paraffin-embedded tissues are used here. Section the material, deparaffinise (for paraffin-embedded tissue), and bring down to the water. ii. Stain the sections for 15-72 hours in 0.17-0.25% lacmoid prepared in 30% ethanol. iii. Wash the stained sections thoroughly with tap water. iv. Place the sections in 1% sodium bicarbonate in 50% ethanol for a few mins. v. Dehydrate, clear, and mount with DPX.

Result Callose stains greenish-blue. Control Callose is dissolved by treating the sections with 5% sodium hydroxide or saturated calcium chloride (Reynolds and Dashek, 1976) or stannous chloride or by treating with glycerine heated to 280°C (Johansen, 1940). 11.2.3.5 Azure B Method (Jensen, 1962)

Procedure i. Plant material may be fixed in Carnoy’s fluid and embedded in paraffin wax. ii. Section the material, deparaffinise and bring down to the water. iii. Stain the sections for 2 hours in azure B [0.25 mg/ml of stain in Mcllvaine buffer (24.6 ml of 0.1 M citric acid + 15.4 ml of 0.2 M disodium phosphate adjusted to pH4)] at 50°C. vi. Rinse in tertiary butanol for 30 mins. v. Give two changes in xylene and mount in DPX. Result Sieve plate callose stains reddish. However, Krishnamurthy (1988) observed that callose present around the microspore mother cells and microspore tetrads was stained only green. This indicates three alternatives suggested by him. These are (1) The chemical composition of callose of microspore mother cells and microspore tetrads may be different and, therefore, it does not respond to this staining schedule; (2) The callose around the microspore mother cells and microspore tetrads is in a bound form with other substances such as glycoproteins; such type of callose was reported in some pollen tubes by Reynolds and Dashek (1976); The bound form may not respond to this

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staining procedure. (3) The procedure itself is non-specific, therefore further studies are needed regarding the chemical nature of callose. 11.2.3.6 Chlorantine Fast Green BLL for Callose (Pearce, 1986)

Procedure i. Bark includes all tissues from cambium whereas rhytidome is collected from mature oak trees (Quercus suber L.) and fixed in acrolein and embedded in polyethylene glycol (water-soluble wax). ii. Transverse sections of about 15 µm are cut from oak bark. These are stained in a 0.25% aqueous solution of chlorantine fast green BLL for about 5 mins. iii. Later stained sections are rinsed and mounted in water. iv. For comparison, adjacent sections are stained with resorcinol blue (Eschrich and Currier, 1964), or aniline blue WS as a fluorochrome (O’Brien and McCully, 1981). Control Sections are immersed in 0.75 ml aliquots of laminarinase (1 mg/ml in 0.05 M citrate buffer, pH 5.0); or in zymolase 5000 (5 mg/ml in 0.05 M phosphate buffer, pH 6.5). Sections are incubated in these enzymes for 5 hrs at 35°C. The enzymes laminarinase and zymolase are known to remove callose (Shimomura and Dijkstra, 1975; Allen and Friend, 1983). After incubation sections are rinsed and stained with chlorantine fast green BLL, resorcinol blue, or acriline blue. Control sections are also incubated with buffer alone. Result Sieve plates and cell wall appositions in the phloem tissues are stained deep green with chlorantine and fast green BLL in both fixed and fresh sections. The location of stained material corresponds exactly with the location of material stained with resorcinol blue or aniline blue. Treatment of sections with either laminarinase or zymolase-5000 (β-l,3-glucan callose degrading enzymes) removes all material staining with chlorantine fast green BLL. No change in staining properties is observed in sections incubated in buffer alone.

11.3 Proteins Proteins occur throughout the plant body in all types of tissues. They are high molecular weight biopolymers, composed of a-amino carboxylic acids, commonly known as amino acids. These amino acids are linked together with peptide bonds and form polypeptides. At a certain size, the polypeptide is, considered to be a protein. Generally, a polypeptide with 50 amino acid residues and a molecular weight of 6,000 is considered a protein. Proteins are classified into simple and conjugated proteins. Simple proteins yield only amino acids upon hydrolysis, e.g., histones whereas conjugated

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proteins yield not only amino acids but also non-protein substances, e.g., nucleo-proteins (proteins conjugated with nucleic acids), glycoproteins (with carbohydrates), metalloproteins (with metals), lipoproteins (with lipids), etc.

11.3.1 Total Proteins Vijayaraghavan and Shukla (1990) briefly outlined some of the commonly used procedures for the localisation of total proteins. Later, Wilson (1992) reviewed the use of amido black, coomassie blue G, and coomassie blue R in protein staining. 11.3.1.1 Ninhydrin/Alloxan-Schiff’s Method (Yasuma and Ichikawa, 1953)

Preparation of Stains i. 0.5% w/v solution of ninhydrin in absolute alcohol. ii. 1% w/v solution of alloxan in absolute alcohol. iii. Schiffs reagent (see 9.2.2B). Staining Procedure Fresh or chemically fixed tissue is used here. Fixation in 10% formalin is found to be ideal (Krishnamurthy, 1988). The chemically fixed tissue may be embedded in paraffin, i. Deparaffinise the sections and bring them down to the water. ii. Place the sections in 0.5% ninhydrin or 1% alloxan solution for 20-24 hrs at 37°C. iii. Rinse twice with absolute alcohol, later in distilled water. iv. Now place the sections in Schiffs reagent for 10-30 mins. v. Rinse in water and place in 2% sodium bisulphate for 1-2 mins. vi. Wash the sections in running tap water. vii. Dehydrate in TBA, clear in xylene, and finally mounted in DPX. Result Proteins appear reddish-purple. Control Before treatment with ninhydrin or alloxan, the tissue should be deaminated or acetylated to prevent the formation of aldehydes from polysaccharides. Deamination Keep the tissue in a mixture containing 200 ml of 60% sodium nitrite and 60 ml of 1% acetic acid for 2-24 hrs at room temperature. Acetylation Keep the tissue in a 10% solution of acetic anhydride in pyridine for 2-20 hrs at room temperature. If colour appears in either of the controls, the reaction is localising some other compound other than proteins having α-amino and α-carboxyl groups in the constituent amino acids.

Histochemical Methods 187

Biological reaction On reacting with ninhydrin or alloxan, the proteins yield an aldehyde (produced by an α-amino, α-carboxyl amino acid) which in turn reacts with Schiff’s reagent to produce a reddish-purple colour. 11.3.1.2 Mercuric Bromophenol Blue (Mazia et al., 1953; Ruthmann, 1970; Chapman, 1975; Dodappa, 2005)

The mercuric bromophenol blue method is employed for the localisation of total proteins in anthers of Carthamus tinctorius L. It is a very effective method in the localisation of even minute quantities of proteins. The dye binds to the basic protein even when mercury is present and with other proteins when mercury is absent (Dodappa, 2005).

Preparation of Stain Reagents Bromophenol blue 10 mg dissolved in 100 ml of 10 % mercuric chloride solution in 95 % alcohol. Staining Procedure (i) Deparaffinisined sections were brought to absolute alcohol and incubated for 15 minutes in mercuric bromophenol blue at room temperature. (ii) Sections were rinsed in 0.5 % acetic acid for 5-10 minutes and differentiated in tap water until sections were blue. (iii) Sections were air-dried, cleaned in butanol, and then xylol and mounted with DPX. Result Proteins stain blue (Fig. 11.3A, B); in sites of protein concentration, reddishblue may appear resulting in metachromasia.

Fig. 11.3. A-B. Transverse section of anther in Carthamus tinctorius L. (A) Prior to meiosis meiocytes and tapetum are rich in total proteins; (B) Pollen grains in fertile anther rich in total proteins. (Courtesy: Dodappa, 2005)

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11.3.1.3 Dinitrofluorobenzene (DNFB) Test (Burstone, 1955)

Preparation of DNFB and other reagents i. Dissolve 0.5 g of DNFB in a mixture of 5 ml of 0.2 N sodium hydroxide and 95 ml of 95% alcohol. ii. 5% w/v aqueous solution of sodium hyposulphite (or) 20% sodium dithionite (dissolve in water at 40°C). iii. Nitrous acid is prepared by mixing 5 ml of 4N sulphuric acid (1.09 ml of H2SO4 in distilled water to make up to 10 ml) with 100 ml of freshly prepared cold 5% w/v sodium nitrite (at 4-5°C). iv. Dissolve 2 g of H-acid (1-amino-8-naphthol-3,6-disulphonic acid) in 100 ml of 0.1 barbitol/Hf.l buffer at 9.2 pH (or) dissolves 0.25 g of Hacid in 1% aqueous solution of sodium bicarbonate. Staining Procedure Fresh or chemically fixed and paraffin-embedded tissue may be used. i. Section the material, deparaffinise and bring down to absolute ethanol. ii. Place the slides in DNFB reagent for 20-24 hrs. The sections should become yellow. iii. Wash slides thoroughly in 95% ethanol for 2-3 mins give three changes in ethanol. Later rinse in distilled water. iv. Place the sections in a 5% solution of sodium hyposulphate for 40 mins. at 40-45°C. The sections should lose their yellow colour or almost completely fade. Later rinse in distilled water. v. Place the sections in freshly prepared cold nitrous acid for 5 mins at 4-5°C. vi. Rinse the slides in cold distilled water. vii. Later place the slides in a cold H-acid for 15 mins. viii. Rinse in distilled water, dehydrate in TBA. ix. Finally clear in xylene and mount in DPX or any other mountant. Result The sites of proteins containing free a-amino groups-, E-amino groups of lysine, phenolic-OH groups of tyrosine, imidazole groups of histidine, and sulphahydryl groups appear a deep reddish-purple. Control Deamination and acetylation are performed before placing the tissue in the DNFB reagent (see Ninhydrin/Alloxan Schiff’s reagent procedure). Biological Reaction 2,4 Dinitofluorobenzene reacts with a-amino groups, ε-amino groups of lysine, phenolic-OH groups of tyrosine, imidazole groups of histidine, and sulphahydryl groups and produces the yellow colour. This colour is not intense enough and so this protein-DNFB complex is treated with the reducing agent

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to reduce the nitro groups of DNFB to amino groups. These amino groups are diazotised with nitrous acid. The diazotised product, when treated with H-acid, results in a deep reddish-purple colour (Krishnamurthy, 1988). 11.3.1.4 Aniline Blue Black Method (Vijayaraghavan and Shukla, 1990; Vijayaraghavan et al., 1972; Maze and Shu-Chang Lin, 1975).

Preparation of Stain Aniline blue-black 1g 7% aqueous acetic acid 100 ml Dissolve the dye in 100 ml of 7% aqueous acetic acid. Staining Procedure Chemically fixed tissue embedded in paraffin may be used here. i. Deparaffinise the sections and bring them down to the water through xylene alcohol, alcohol-water graded series. ii. Stain the sections in 1% dye solution for 10-15 mins at 50-60°C. iii. Differentiate by dipping the slides in 7% acetic acid. iv. Mount the sections in glycerine containing 5% acetic acid. Result Protein stains blue-black. 11.3.1.5 Coomassie Brilliant Blue (CBB) Method

(Fisher, 1968; Knox and Heslop-Harrison, 1970; Cawood et al., 1978; Eklavya, 1979)

Preparation of Stain Dissolve 0.25 g coomassie brilliant blue (R 250) in 100 ml of 7% acetic acid. Staining Procedure Fresh tissue or tissue fixed chemically in 2.5% glutaraldehyde and paraffinembedded may be used here. i. Section the material, deparaffinised, and hydrate through xylene-alcohol, alcohol-water graded series. ii. Stain the sections in Coomassie brilliant blue (R250) for 3 mins at 30°C. iii. Rinse in 7% acetic acid. iv. Dehydrate in 98% and absolute ethanol, 5 mins in each grade, and finally mount in glycerine containing 5% acetic acid. Result Protein stains blue to bluish-black. Control Protease treatment or pepsin digestion before staining in CBB.

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11.3.1.6 Brilliant Sulfaflavine-Fluorescence Method (Ruch, 1970)

Preparation of Stain i. Take 200 ml Mcllvaine citrate-phosphate buffer (pH 2.8). ii. Dissolve 0.2 g Brilliant sulfaflavine in the above buffer. Staining Procedure Plant material fixed in acetic acid-alcohol and paraffin-embedded is used here. i. Sections are deparaffinised and brought down to the water. ii. Keep the slides in citrate-phosphate buffer (pH 2.8) for 2 mins. Later stain in Brilliant sulfaflavine solution for 30 mins. iii. Rinse in the same buffer for 5 mins. iv. Dehydrate in TBA thrice. v. Clear in xylene (two changes) and mount in DPX (or) another suitable mountant. vi. Observe under a fluorescence microscope (UV excitation range 360-400 nm). Result Proteins fluoresce yellow. 11.3.1.7 Biuret Reaction (Gahan, 1984)

Fresh tissue or chemically fixed tissue embedded in paraffin may be used: i. Section the material, deparaffinise and bring down to the water. ii. Warm the sections on a slide in 10% aqueous potassium hydroxide. iii. Add a drop of 1% copper sulphate solution. iv. Mount the slide in water.

Result Violet colour indicates peptide linkages in higher proteins whereas red colour indicates peptide linkages in lower protein forms. 11.3.2 Basic Proteins 11.3.2.1 Alkaline Fast Green Method (Prento and Lyon, 1973)

Preparation of Stain i. Take 100 ml of 5% w/v trichloroacetic acid. ii. Dissolve 50 mg Fast Green FCF in 50 ml of 0.01 M Phosphate buffer (pH 8.0). Staining Procedure Chemically fixed tissue in buffered neutral formalin and paraffin-embedded may be used.

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i. Section the material, deparaffinise and bring down to the water. ii. Rinse in distilled water at 60oC iii. Keep the sections in trichloroacetic acid and then in 0.01 M phosphate buffer (pH 8.0) for 90 mins at 60oC. iv. Immerse in distilled water and then wash in 70% ethanol giving three changes of 10 mins each. v. Rinse in distilled water thoroughly and stain in Fast Green FCF for 30 mins. vi. Rinse in 0.01 in phosphate buffer (pH 8.0) for 1 min. vii. Dehydrate in 95% rectified alcohol, later in 100% ethanol. viii. Rinse in ethanol: xylene (1:1) ix. Clear in xylene I and II, and mount in DPX

Result Basic proteins, nuclei, and chromosomes stain green to bluish-green. 11.3.2.2 H/ydroxy-3-Naphthoic Acid Hydrazide Method for Proteins Containing α-acylamide Carboxyl Groups (Barrnett and Saligman, 1958)

Preparation of 0.1% 2-hydroxy-3-naphthoic acid hydrauzide Hydrazide 50 m Hot glacial acetic acid 2.5 ml 50% ethanol 47.5 ml Hydraulic is dissolved in hot glacial acetic acid and added to 50% ethanol.

Staining Procedure Frozen tissue or chemically fixed tissue embedded in paraffin may be used here: i. Section the material, deparaffinise, and rehydrate. ii. Keep the sections for one hour in a 1:1 mixture of acetic anhydride and anhydrous pyridine at 60oC. iii. Wash in absolute ethanol (aldehyde free). iv. Later keep the sections for two hours in 0.15 2-hydroxy-3-naphthoic acid hydrazide at room temperature. v. Wash in four changes of 50% ethanol, 10 mins each. vi. Keep the sections for 30 mins. in 0.5 N hydrochloric acids at room temperature. vii. Then rinse in distilled water. viii. Keep the sections for 2-5 mins in a solution of 0.006 M phosphate buffer (pH 7.6) mixed with an equal volume of absolute ethanol, and stir in diazo blue B (tetrazotizeddiorthoanisidine, 1 mg/ml). ix. Wash in several changes of distilled water. x. Dehydrate and mount in DPX.

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Result Proteins containing α-acylamido carboxyl groups will appear red to blue. The red indicates widely spaced groups whereas blue indicates numerous closely spaced groups. Control Tissue is treated with 0.1 N HCl in methanol for 24 hrs at 60°C. This esterifies all carboxyl groups. Biological Reaction The α-acylamido carboxyl groups are converted to methyl ketones by acetic anhydride and absolute pyridine. The methyl ketones then react with 2-hydroxy-3-naphthoic acid and a naphthol group is linked to the protein. These naphthol groups are coupled with diazo blue B, so that red to blue colour develops. 11.3.2.3 DDD Test for Proteins Containing Sulphydryl-Disulphide Groups like Cysteine, Cystine, and Methionine (Barrnett and Saligman, 1952; Roberts, 1960).

i. Dissolve 100 mg of 2,2’-dihydroxy-6-6’-dinaphthyl-disulphide (DDD) in 60 ml of absolute ethanol. Immediately before using take 15 ml of stock solution and add 35 ml of barbital buffer (pH 8.5). ii. Dissolve 50 mg of diazo blue B (tetrazotizeddiorthoanisidine) in 50 ml of Sorensen phosphate buffer (pH 7.4).

Staining Procedure Frozen tissue or formalin-fixed tissue embedded in paraffin may be used. i. Section the material, deparaffinise and bring down to the water. ii. Keep the tissue in a fresh aqueous 0.2-0.5 M solution of thioglycolic acid adjusted to pH 8.0 with 0.1 N sodium hydroxide for 1-2 hrs at 50°C or keep the sections in a thioglycerol solution prepared by dissolving 3 m1 of thioglycerol in 40 ml of water and adding 10 ml of 0.5 M borate buffer (pH 9.1-9.5) for 1-2 hrs at 50°C or overnight at room temperature. iii. Wash in several changes of distilled water. iv. Keep the sections in DDD solution for one hour at 56°C and bring them to room temperature. v. Rinse the sections briefly in distilled water and wash for 5 mins. each in two changes of distilled water adjusted to pH 4.0 with acetic acid. vi. Dehydrate, wash in absolute ether for 5 mins. and again hydrate. vii. Keep the sections in a freshly prepared diazo blue B for 2-5 mins at room temperature. viii. Wash in running tap water. ix. Dehydrate in acetone and mount in DPX.

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Result Sites of sulphydryl groups appear blue. If diazo red RC is used instead of diazo blue B the localisation would appear red. Control Sulphydryl blocking agents are used here. Keep the sections in a 0.1 M N-ethyl maleimide solution in a phosphate buffer (pH 7.4) for 4 hrs at 37°C or keep sections in a 0.1 M iodoacetate solution adjusted to pH 8.0 with sodium hydroxide for 20 hrs at 37°C or keep sections in a 0.001 M solution of iodine in n-propanol for 4 hrs at 20°C. Biological Reaction Sulphydryl groups undergo a reaction with naphthols present in DDD. The resultant product is colourless but when coupled with diazo dye it produces a visible colour.

11.4 Detection of Ions in Plant/Animal Tissues There are some minerals, such as ferric and calcium, useful for the growth and development of plants. Their ion levels could be visualised by histochemical methods.

11.4.1 Perls’s Reaction This method is particularly important for the detection of ferric ion levels in plant tissues and animal tissues like spleen and bone marrow. In this technique, ferric ions present in the tissue will combine with ferrocyanide resulting in the formation of a pigment called Prussian blue (ferric ferrocyanide). In plants, the technique is based on the conversion of ferrocyanide into insoluble crystals (Prussian blue) in the presence of ferric ions under acidic conditions. Prussian blue formed here, is bright blue which indicates the presence of ferric ions.

Staining Procedure In Perls’s reaction technique, a known positive control tissue is used as control while 10 percent formalin is used as a fixative. Requirements Microwave oven, Acid clean glassware, non-metallic forceps, 5% potassium ferrocyanide, 5% hydrochloric acid, and sample plant section (4 µ thickness of plant organ tissue). It is important to use gloves, goggles, and a lab coat because some of the chemicals used for this procedure can cause irritation. Procedure i. Deparaffinise the section and hydrate with distilled water.

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ii. Microwave for about 50 seconds and allow the specimen to expose to acid ferrocyanide for about 5 minutes in the fumed wood. Then section can be treated with the same solution for about 10-30 minutes. iii. Rinse the section with 0.5% aqueous neutral red or 0.1% nuclear fast red for nucleus staining. iv. Rapidly wash the section using distilled water. v. Dehydrate the section, clear it, and mount it on the microslide for viewing under the microscope.

Result When viewed under a microscope, blue parts are indicative of iron while red and pink indicate the nuclei and background respectively. 11.4.2 Von Kossa’s Technique (1901) It is a more sensitive technique to detect calcium deposits on cyst fluids, ductal ectasia papillomatosis. Although this technique is used to demonstrate the presence of calcium, it demonstrates an anion rather than the calcium ion itself. The principle involved here is the section is treated with the silver nitrate solution and calcium (if present) is reduced by the strong light and replaced with deposits of silver. It is visualised as metallic silver.

Requirements A central sample (a tissue with known calcium deposits, 10% formalin (fixative), glassware, 60-watt lamp, a mirror or a foil, 5% silver nitrate solution, and 5% hypo (sodium thiosulfate). Procedure i. Deparaffinise and hydrate the section using distilled water. ii. Place the solution in the silver nitrate solution (in a glass jar) and place it in bright light. Then place on a mirror or a paper foil behind the jar to reflect the light. Leave it for one hour or until the calcium turns black. iii. Rinse the section in distilled water. iv. Stain the section with hypo for about 5 minutes. v. Wash the section and rinse in distilled water. vi. Stain the section with nuclear-fast red for a few minutes. vii. Wash using distilled water. viii. Dehydrate and observe under the microscope. Results The black colour in the section indicates the presence of calcium; nuclei appear red in colour while the cytoplasm pink.

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11.5 Nucleic Acids Nucleic acids are high-molecular-weight polymeric molecules and are composed of nucleotides. Nucleotides consist of purine (adenine and guanine) and pyrimidine (uracil, thymine, and cytosine) bases, both linked with a pentose sugar which is esterified with phosphoric acid. The pentose sugar may be ribose or deoxyribose, depending on which the nucleic acids are known as ribonucleic acid (RNA) or deoxyribonucleic acid (DNA). The non-esterified forms of the nucleotides are called nucleosides. The nucleotides of DNA are linked together and form polynucleotides whose general form is a double-stranded helix (Watson and Crick model). The two strands are connected by specific hydrogen bonds always between adenine and thymine in DNA, adenine, and uracil in RNA, and between guanine and cytosine in both DNA and RNA. DNA is present in the chromatic material of the nucleus and, in chloroplasts and mitochondria of the cytoplasm. In the nucleus, heterogeneous RNA and messenger RNA are present. While in the cytoplasm, ribosomal RNA and transfer RNA are present. The former two types are essentially present in the nucleolus whereas the latter two are in the ribosomes.

11.5.1 Azure B Method (Flax and Himes, 1952; Jacqward et al., 1972) Preparation of Stain and Enzymes i. Azure B: Take 0.25 mg/ml of stain in McIlvaine’s buffer (24.6 ml of 0.1 M citric acid plus 15.4 ml of 0.2 M disodium phosphate adjusted to pH 4). ii. DNase enzyme: Take 0.25 mg/ml DNase in 0.03 M magnesium sulphate, pH 6.5. iii. RNase enzyme: Take 0.25 mg/ml RNase in 0.03 M magnesium sulphate, pH 6.5. Tissue fixed in Carnoy and embedded in paraffin is used here. i. Section the material, deparaffinise and bring down to the water. ii. Digest DNA by DNase enzyme to localise RNA alone. Sections are incubated in the enzyme solution for 4 hrs at 37°C. Later rinse the slides twice in distilled water, 5-10 mins for each rinse. iii. Digest RNA by RNase enzyme to localise DNA alone. This is carried out by treating the slides in RNase solution for 3 hrs at 37°C (pH of 6.5 is adjusted with 0.1 N sodium hydroxide). iv. Stain the slides in Azure B for 2 hrs at 50°C.

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v. Rinse in tertiary butanol for 30 mins. to variable time depending on the type of material. vi. Later clear in two changes of xylol. vii. Mount in DPX or any other suitable mountant.

Result RNA stains dark blue whereas DNA is blue-green. Biological Reaction The reaction is based on metachromasia. The colour of the dye solution is called orthochromatic shade, while the second colour formed in the cells, when the dye is used, is called a metachromatic shade. Here metachromatic shade is blue-green while the orthochromatic shade is dark blue-green. RNA is stained blue and DNA blue-green. The dye-binding site is probably the phosphates. Presumably, the arrangement of the phosphates is tighter in DNA than in RNA resulting in DNA becoming metachromatic. Control Both RNA and DNA are extracted and stained with Azure B. 11.5.2 Toluidine Blue O Method for RNA (Chayen et al., 1973; Dodappa, 2005) Toluidine blue of 0.50 mg is dissolved in 100 ml of 0.05 M Citrate Phosphate buffer at pH 4.4.

Stock Solutions A. Citric acid solution 0.1M (19.21 g in 1000 ml) B. Disodium phosphate 0.2 M (53.65 g of Na2HPo4.7H20 or Na2HPo4 12H20 in 1000 ml) A and B solutions are mixed by taking the 27.8 ml of ‘A’ and 22.2 ml of ‘B’ diluted with distilled water to a total volume of 100 ml.

Staining Procedure (i) Deparaffinised and hydrated sections were immersed in 0.05 % toluidine blue for 5 minutes. (ii) Sections were rinsed in distilled water, air-dried, cleaned in xylol, and mounted with DPX. Result RNA stains purple while DNA blue or blue-green (Fig. 11.4 A-C). The pH should be very carefully maintained; otherwise, the ability of the dye to bind to nucleic acid diminishes (Khasim, 2002).

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Fig. 11.4. A-C. Transverse section of anther in Carthamus tinctorius L. (A) Putative spores and binucleate tapetal cells are rich in RNA; (B) Fibrous bands in endothecial cells stain green with toluidine blue (solid arrow). Similar stained layer lines the inner face of the endothecium (arrowhead). Pollen grains rich in RNA; (C) Plasmodium rich in RNA (m = microspore; t = tapetum; e = endothecium; Bar = 20 µm). (Courtesy: Dodappa, 2005)

11.5.3 Acridine Orange Method for DNA and RNA (Gahan, 1984) Preparation of Stain Prepare 0.05% acridine orange solution in hydrochloric acid-acetate buffer (pH 2.1). Staining Procedure Fresh, frozen, or chemically fixed tissue embedded in paraffin may be taken here. i. Section the material, deparaffinise and bring down to the water.

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Immerse the sections in hydrochloric acid-acetate buffer (pH 2.1). Stain the sections in Acridine Orange solution for 30 mins. Wash the sections in the veronal-acetate buffer for 15 mins. Mount in the above buffer and examine with blue or ultraviolet light.

Result DNA fluoresces bright yellow and RNA flame red. Control DNA and RNA extraction is performed according to the procedure given in the Azure B staining schedule. Biological Reaction The dye binds to DNA by intercalating between adjacent base pairs. When a sufficient number of dye sites become saturated in this way, the DNA-acridine complex shows fluorescence. It is important to note that the dye-binding and induced fluorescence are indicators of molecular size and shape only and not qualitative markers of two nucleic acids. In other words, acridine orange does not distinguish between DNA and RNA. DNA has a higher molecular weight and so fluoresces yellow, while RNA has low molecular weight and so fluoresces red (see Krishnamurthy, 1988). 11.5.4 Feulgen Reaction for DNA (Feder and O’Brien, 1968; Krishnamurthy, 1988) Preparation of Stain Basic fuchsin Boiled distilled water

0.5 g 100 ml

Basic fuchsin is dissolved in boiled distilled water, cooled to 50°C, and filtered. To the filtrate add 10 ml of N hydrochloric acid and 0.5 g of potassium metabisulphite. Shake thoroughly, close tightly, and store in dark until use.

Staining Procedure Fresh, frozen, or chemically fixed tissue with paraffin embedding may be used here. i. Section the material, deparaffinise and bring down to the water. ii. Rinse in cold N hydrochloric acid. iii. Hydrolyse in N hydrochloric acid for 4-5 mins at 60°C. iv. Again, rinse in N hydrochloric acid. v. Keep the sections in Schiffs reagent (Feulgen’s fuchsin). vi. Drain and pass quickly through three closely arranged Coplin jars each containing 5 ml n HCl + 5 ml 10% potassium metabisulphite + 100 ml distilled water, 10 mins in each jar. vii. Dry and apply coverslip and observe under the microscope.

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Result DNA stains purplish red. The nucleoli and most of the cytoplasm is stained green. Biological Reaction Treatment with warm HCl hydrolyses the purine-deoxyribose linkages and releases free aldehyde groups. These aldehydes form a coloured complex with fuchsin. Control Pearse (1968, 1972) suggested that before hydrolysis a blocking of aldehydes in a saturated solution of dimedone (5,5-dimethyl-cyclohexane-1,3-dione) for 16-24 hrs should be performed. Later hydrolysis is to be carried out. 11.5.5 Cresyl Violet Method for RNA (Ritter et al., 1961; Cecich et al., 1972) Preparation of Stain Prepare 0.1% aqueous cresyl violet solution adjusted to pH 4.2 with 0.1 N hydrochloric acid. Staining Procedure Tissue chemically fixed in Carnoymplex or other fixatives and embedded in paraffin is taken. i. ii. iii. iv.

Section the material, deparaffinise and bring down to the water. Digest DNA by DNase treatment (see Azure B method) and wash. Stain the sections in aqueous cresyl violet solution for 2 hrs. Dip in absolute ethanol and remove the ethanol periodically until ethanol remains colourless. v. Dehydrate, clear, and mount.

Result RNA stains violet.

11.6 Lignin, Suberin and Cutin 11.6.1 Lignin Lignin is a phenolic polymer present in the cell walls of tracheary elements, xylem and phloem fibres, and sclerenchyma. Approximately 20% of the cell wall materials of the secondary xylem of dicotyledons and 35% of that of gymnosperms constitute the lignin. It has a molecular weight of Ca 11000. On oxidation, it yields three simple phenolic aldehydes: p-hydroxybenzaldehyde, vanillaldehyde, and syringaldehyde in various proportions. Because of the

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presence of double bonds, lignin strongly absorbs UV light, and it can be readily distinguishable from other cell wall components by its f1uorescence. 11.6.1.1 Phloroglucinol Method (Gahan, 1984)

A slightly modified technique of Gahan (1984) is adopted here. The phloroglucinol method is suitable for studying the structure of nodes. Fresh material is preferred.

Preparation of Stain Dissolve 10 g of phloroglucinol in 100 ml of distilled water. Take 10 ml of concentrated HNO3 in 90 ml of water. Staining Procedure i. Section the material and keep in 10% aqueous solution of phloroglucinol for 1 to 3 mins. ii. Drain the stain and add a few drops of 10% concentrated HNO3, wait for a few minutes till it develops colour. iii. Mount in 75% glycerine. Result Lignin gives a red colouration. Biological Reaction The basis of reaction is not very clear. However, phloroglucinol appears to react with the coniferyl and cinnamaldehyde end group of lignin to yield a cationic chromatophore (Wardrop, 1981). Comment Phloroglucinol test is not specific for lignin since it stains hemicellulose and suberin also (Reeve, 1974; Shah and Babu, 1986). It does not stain lignin in some monocots and protoxylem in certain dicots (Krishnamurthy, 1988). 11.6.1.2 Maule’s Reaction (Johansen, 1940)

Fresh tissue is preferred. However, chemically fixed and paraffin-embedded tissue can also be used. Even herbarium specimens are used successfully after revival with warm water (with some detergents added) treatment.

Staining Procedure i. Section the material and sections are put in 1% neutral potassium permanganate solution for 5-20 mins. ii. Wash the sections in distilled water. iii. Decolourise with 2% hydrochloric acid. iv. Wash thoroughly in water. v. Treat with a few drops of ammonium hydroxide or sodium bicarbonate solutions.

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vi. Observe under the light microscope.

Result Lignin takes red colour due to syringyl type of lignin and brown colour due to guaiacyl type of lignin which is present in angiosperms and gymnosperms respectively. Krishnamurthy (1988) observed that in some plants lignin does not respond indicating the presence of a different type of lignin other than the two mentioned above. Biological Reaction Potassium permanganate is reduced to manganese dioxide deposited on the lignified walls. The hydrochloric acid then reacts with manganese dioxide and a chlorination reaction causes the colouration of lignin. 11.6.1.3 Toluidine Blue O Method (O’Brien et al., 1964; Fisher, 1985)

For Preparation of Stain (see Toluidine blue O method for Nucleic Acids see section 12.5.2). Staining Procedure Fresh or frozen or chemically fixed and paraffin-embedded tissue may be used. i. Section the material, deparaffinise and bring down to the water. ii. Stain the sections in 0.05% aqueous toluidine blue O solution at pH 4.4. iii. Wash in tap water, clear, and mount in DPX or another suitable mountant.

Result Lignin stains green-blue. Biological Reaction Though the biological reaction is not known clearly, this method is useful in differentially staining the gelatinous cell layer of fibres of reaction wood. In these fibres, lignin stains green blue, whereas inner gelatinous layer stains reddish-purple. It is questionable whether the gelatinous layer is exclusively cellulosic as was claimed by several investigators. The reddish-purple colour shown by this layer with toluidine blue O indicates that this layer contains sulphated and carboxylated polysaccharides (Feder and Wolf, 1965; O’Brien et al., 1964). 11.6.2 Suberin Suberin is commonly present in cork tissue and root caps of some plants. It is composed of two-thirds aromatic phenols and one-third long-chain aliphatic acids. Due to the presence of aliphatic acids, suberin shows lipidlike properties. It is resistant to strong acid treatment, but it is dissociated by strong alkali treatment.

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11.6.2.1 Cyanin and Potassium Hypochlorite Method (Chamberlain, 1932)

Preparation of Stain Cyanin 1% is prepared in a mixture of 50% ethanol and an equal volume of glycerine. Staining Procedure i. Fresh tissue is taken and sections the material. ii. Treat the sections in potassium hypochlorite to destroy tannins and to make lignified walls lose their staining capacity. iii. Stain in cyanin solution. 11.6.2.2 Phosphate Buffer Method (Gahan, 1984)

i. Fresh or frozen tissue is taken and sections the material. ii. Mount the sections in phosphate buffer, 0.02 M (pH 9.1), and observe under the fluorescence microscope using UV light.

Result Yellow fluorescence indicates suberin. Biological Reaction The mechanism is not very clear. The phenol component of suberin may be responsible for this optical property. Control Mount the sections in concentrated sulphuric acid. Resistance to this treatment is indicative of suberin. 11.6.3 Cutin Cutin is not a true fat component but is closely related to it. Cutin forms the non-cellular cuticle which is present on aerial parts of higher plants. It strongly resists acid attack and decay, but readily dissolves in strong alkalis to produce a mixture of fatty acids and dihydroxyacids, mainly with C16-C18 chains. 11.6.3.1 Auramine O Method (Heslop-Harrison, 1977; Malti and Shivanna, 1984)

Preparation of Stain Prepare 0.01-0.2% auramine O in water or 0.05 M tris-HCl buffer (pH 7.2). Staining Procedure Fresh or frozen tissue is taken.

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i. Keep the tissue in chloroform for 5 mins. to remove lipids, later section the material. ii. Stain the sections in auramine O. iii. Mount in glycerine and observe under a fluorescence microscope.

Result The cuticle fluoresces under the microscope. Control The material is pretreated with repeated washing in 1:1 ether-methanol before proceeding to the staining schedule. This treatment removes the cuticle.

11.7 Cytokinins and Auxins Plant growth regulators such as cytokinins and auxins play an important role in plant growth and development. The accumulation of indole-3-acetic acid (IAA), an auxin, has been demonstrated in insect galls, using Ehrlich’s reagent (Bedetti et al., 2014). Gall-inducing insects manipulate host plant signals and redirect normal plant development toward gall morphogenesis (Giron et al., 2016; Richardson et al., 2017). The development of galls commonly involves hyperplasia and cell hypertrophy, and it requires these growth regulators even at low concentrations (Yamaguchi et al., 2012; Azizi et al., 2015).

11.7.1 Histochemical Localisation of Cytokinins and IAA (Zhang et al., 2014; Bedetti et al., 2018) In this technique, histochemical localisation of cytokinins and IAA has been performed in plant tissues and it is validated by immunocytochemistry based on methodology developed by Bedetti et al. (2018) with slight modification.

Materials and reagents - Roots of Phaseolus vulgaris (Fabaceae). - Lenticular gall morphotypes (both lenticular concave–convex and lenticular plane-convex) induced on pinnulas of Piptadenia gonoachantha (Fabaceae). - Silica plates for Thin Layer Chromatography (TLC) for cytokinins (zeatin). - Rotatory microtome. - Zeatin (80% trans form). - Silver nitrate 0.5% in Ethanol (solution A). - Bromophenol 0.2% and 0.15% silver nitrate in 1:1 ethanol: ethyl acetate (solution B). - Solution C: EDAC (1-ethyl-3(3-dimethylaminopropil) carbodiimide

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hydrochloride) /100 mM phosphate-buffered saline (PBS) at pH 7.0, 91% + Paraformaldehyde 4% + Glutaraldehyde 5%. Alcohol: Xylene series. Paraffin wax. Ehrlich’s reagent (1% p-dimethylaminobenzaldehyde in 1M HCl w/v). Tween-20. Glycine 15%, 50%. 5% bovine serum albumin (BSA). Blocking solution consists of 93.4% 10 mM PBS, 0.1% Tween-20, 1.5% glycine and 5% bovine serum albumin (BSA). Salt rinse solution (RSR) containing 99% 10 mM PBS, 0.1% Tween-20, 0.9% BSA. Primary rabbit anti-IAA polyclonal antibody (As09 445; Agrisera, Vännäs, Sweden). Primary rabbit polyclonal antibody to trans zeotin riboside. Secondary antibody anti-rabbit IgG (whole molecule)-FITC goat antibody (F9887; Sigma Aldrich).

Protocol Thin layer chromatography (TLC) of cytokinins 1. Load the zeatin on silica plates and 9:1 chloroform: methanol used as mobile phase. 2. Upper separation, the plates are sprayed with 0.5% silver nitrate in ethanol (solution A). 3. Heat at 100oC for 5 min, sprayed with 0.2% bromophenol blue plus 0.15 silver nitrate in 1:1 ethanol: ethyl acetate (solution B) and heat it at 100oC for additional 10 mins. 4. Chromatogram is photographed using digital camera. Free-hand sections 5. Free-hand sections (transverse and longitudinal) of roots of P. vulgaris and lenticular morphotypes of Piptadenia gonoachantha are taken. Histochemical localisation of cytokinins and IAA 6. For cytokinins detection, take root section and stain with solution A and B and incubated at room temperature; in solution A for 5 min, followed by solution B for 5 min. 7. After incubation, sections are washed in absolute ethyl alcohol. 8. Finally mount the sections on microslide. Colour appeared in root tissues as well as gall tissue should be compared with the coloured developed by the standard. 9. For IAA, incubate sections in Ehrlich’s reagent at room temperature for 5 min. 10. Stained sections are observed under the light microscope and photographed.

Histochemical Methods 205

IAA and Cytokines immunolocalisation 11. Three samples each of root of P. vulgaris and galls of Piptadenia gonoachantha are taken and prefixed in EDAC/100 mM PBS. 12. Then the above samples are post-fixed in the solution C at 4oC for 15 h. However, to detect cytokines, the post-fixation is omitted because paraformaldehyde links to cytokines bases, N6-isopentenyladenine, DHZ and Zeatin in plant tissues. 13. After post-fixation, above material is dehydrated in alcohol: xylene series and embedded in paraffin wax (Johansen, 1940; Khasim, 2002). 14. Using Rotary microtome, longitudinal sections of roots and transverse section of galls are obtained. 15. Above sections are deparaffinised using alcohol and xylene series and washed in 10 mM PBS. 16. Later the above sections are incubated in blocking solution for 45 min. 17. After incubation, these sections are washed in regular salt rinse solution for 5 min, then treated with 99.1% 10 mM PBS, 0.8% Triton X-100 and 0.1% BSA. 18. Sections then incubated separately in either the primary rabbit anti-IAA polyclonal antibody (Agrisera, Vännäs, Sweden) diluted 1:40 in PBS or the primary rabbit polyclonal antibody to trans-zeatin riboside for 2h in the dark. See that control sections should not be treated with these antibodies. 19. After the primary antibody treatment, sections are washed in saltrinse solution and PBS an incubated at room temperature for 2 h with the secondary antibody anti-rabbit IgG (whole molecule)-FITC goat antibody diluted 1:40 in PBS. 20. After washing thrice in salt-rinse solution and PBS the sections are mounted in 50% glycerin. 21. The analyses are performed using Confocal Zeiss 5 LIVE microscope (Oberkochen, Germany) at 488 nm excitation and 505-530 nm emission filter.

Results Cytokinins are detected as blue spots in both TLC plates as well as tissue sections stained with bromophenol blue/silver nitrate. The zeatin riboside antibody labeled the cytokinins in sites similar to those obtained by histological stains of root and gall tissues. In root tissue of P. vulgaris, xylem cell walls and nuclei are the main sites of cytokinins accumulation. In the gall sections, the cytokinins are detected in the central inner cortical cells. In case of IAA, Anti-IAA antibodies are weakly labelled accumulation in xylem of roots and Ehrlich’s reagent could not detect IAA. Whereas IAA is detected in gall tissue by the anti-IAA antibodies and by the Ehrlich’s reagent in the vacuoles and the cytoplasm of outer cortical cells. However, IAA is not detected in the central inner cortex of any galls.

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11.8 Latex and Rubber Latex is a colloidal substance consisting of various substances such as carbohydrates, organic acids, salts, alkaloids, sterol, fats, tannins, and mucilages. The dispersed particles commonly present are essential oils, balsams, resins, camphor, carotenoids, and rubber (Bonner and Galston, 1947). Among these substances, the resins and rubber (C5H8)n are the characteristic components in the latex of many plants. Latex may contain a large number of proteins (Ficuscallosa), sugars (Compositae), or tannins (Musa). The bestknown latex is produced by various rubber-yielding plants (Hevea braziliensis, Ficus elastica, etc.). When the latex is released from the plant, the rubber particles clump together, i.e., the latex coagulates. This property is useful in the commercial separation of rubber from latex (Esau, 1993).

11.8.1 Staining Technique to Localise Rubber in Guayule (Parthenium Argentatum Gray) (Jayabalan and Shah, 1986) Before staining, sections have to be deresinated and total lipid extraction should be performed in the following manner.

Deresination Cross-sections of guayule stem about 20-40 µm thick are taken; sections are deresinated with a minimum of three changes of preheated acetone-water solvent (88:12) at 50°C for 45 mins (Kay et al., 1980). Extraction of Lipids Total extraction of lipids is carried out at room temperature in the following way: (a) Sections are kept either in a mixture of 1:1 chloroform and methanol for 30 mins (Galanos and KaPoulas, 1962) or in a solution of 2% sodium methoxide in equal volumes of methanol and butanol. (b) Then lipids are completely removed by agitating the sections with 5% NaOH in 95% ethanol for 30 seconds.

Staining with Dansyl Chloride Alone i. Deresinated sections are treated with a proteolytic enzyme, Pepsin pH 6.0 for 6 hrs to remove cellular proteins (Pearse, 1972). ii. To remove precipitated proteins, sections are briefly agitated in 10% trichloroacetic acid (TCA). iii. Treat with 70% ethanol for a minute. iv. Wash thoroughly with distilled water. v. Rinse sections with 1% sodium bicarbonate (pH. 8.0-9.0). vi. Immerse in dansyl chloride stain (1 mg/4 ml acetone) for 2-3 mins.

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vii. Now stained sections are washed in distilled water and mounted in 1% sodium bicarbonate solution to avoid background during photomicrography.

Staining with both Oil Red O and Dansyl Chloride i. Deresinated and deproteinated sections are treated with oil red 0 for 3-5 mins (Pearse, 1968). ii. Counterstain with dansyl chloride (1 mg in 4 ml 60% isopropanol). iii. Rinse the sections in distilled water and mount in 1% sodium bicarbonate. Result Stained sections are observed under a Zeiss epifluorescence microscope. White fluorescence is observed in the parenchyma cells treated with dansyl chloride. The combination of oil red O and dansyl chloride gives better fluorescence of rubber globules than using these stains separately (see Jayabalan and Shah, 1986). Biological Reaction The rubber globules stained with oil red O or oil blue N are extracted by common fat solvents like acetone, diethylene glycol, or xylene (Lillie and Fullmer, 1976). This indicates that the stain is due to the dye molecule reacting with a hydrophobic matrix of the rubber. The fluorescence of rubber stained with dansyl chloride, auramine O, or neutral red is different from that of these substances in solution by themselves. These dye molecules may form chemical bonds through the residual double bond of rubber. 11.8.2 Oil Red O-Dansyl Chloride Method for Laticifers (Inamdar et al., 1987) Procedure i. Fixation of plant material is done in FPA (formalin-propionic acidalcohol) (Johansen, 1940). ii. Freehand sections are taken with the help of a razor blade. iii. Clear the sections in a 1:1 mixture of hydrogen peroxide (30%) and glacial acetic acid by keeping them in a hot air oven for 1-3 hrs. iv. Later sections are taken out from the oven and washed with 70% ethanol followed by two or three changes of distilled water. v. Then sections are kept in 1% toluidine blue O for 2-3 minutes to reduce autofluorescence of lignified walls (O’Brien and McCully, 1981). vi. The sections are first stained with Oil Red O and followed by dansyl chloride and mounted in 1% sodium bicarbonate (pH 7-9) to reduce background interference fluorescence. vii. Later sections are thoroughly washed with deionised water and mounted in 60% glycerine and observed under a fluorescence microscope.

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Comments When sections are stained with oil red 0 alone, faint red fluorescent and UV illumination are observed. It is enhanced following treatment with dansyl chloride. Deproteinised sections used as controls here are shown as weak fluorescence. Also deresinated and lipid extracted controls show very faint staining with oil red O under a bright-field microscope. Generally, oil red O is used as a fat stain (Lillie, 1977) and dansyl chloride for proteins (Ringertz, 1968). The fluorescence is due to the high number of proteins, fats, and resins present in laticifers (Bruni et al., 1977).

12 Electrophoresis Histochemistry is a combination of histology and analytical biochemistry. In histoThe term electrophoresis defines as the migration of charged particles in a medium under the influence of an electric field. Many biological molecules such as nucleic acids, nucleotides, amino acids, peptides, and proteins possess ionisable groups and at any given pH in a solution, they are electrically charged either as cations (+) or anions (−). Under the influence of an applied electric field these charged particles will migrate either to the cathode or to the anode, depending on the nature of their net charge. Even if two molecules have the same charge, they may not migrate together because there is a difference in their molecular weight, they will have different charge/ mass ratios. This difference is of more use in electrophoresis on gels. These differences are brought about by the differential migration when the ions in a solution are subjected to an electric field. Modern scientists have been using this technique very widely to determine the molecular weight of proteins and DNA sequencing. Migration of an ion in an electric field When a spherical molecule of net charge ‘q’ is placed in an electric field, the electric force ‘F’ that acts upon this particle will depend upon (i) the net charge density of the molecule, and (ii) the strength of the field in which it is placed. This can be described mathematically as follows: ΔE q F = —— d ΔE = Field strength applied (potential difference between two molecules) d = Distance between two electrodes q = Net electric charge on the molecule Since the particle is suspended in a solution, one has to consider the friction occurring between the accelerating molecule and the solution to obtain a valid relationship of electrophoretic migration. The extent of friction, as described by Stoke’s equation, will depend upon (i) the shape and size of the molecule, and (ii) the viscosity of the medium, it is as follows: F = 6πrηv F = Function exerted on the spherical molecule

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r = radius of the molecule η = viscosity of the solution v = velocity at which the molecule is migrating The above frictional force will oppose the accelerating force generated by the electric field. Equating the force of acceleration with Stoke’s equation, we get: ΔE —— q = 6πrηv d If we rearrange the above relationship, we get the equation as follows: ΔEq q = 6πrηv In the above equation, the velocity (v) of the molecule is proportional to (i) the field strength (ΔE/d) and (ii) charge (q) on the molecule but is inversely proportional to the field strength, the particle size (r), and (iii) viscosity of the solution (η).

12.1 Modes of Electrophoresis The instrument required for electrophoresis has two components: a power pack and an electrophoresis unit. Electrophoresis units are available for running either vertical or horizontal gel systems.

12.1.1 Low Voltage Electrophoresis Paper electrophoresis is low-voltage electrophoresis; this could be demonstrated for undergraduate students. Filter paper has been used for the study of normal and abnormal plasma proteins. Paper of good quality will have 95% cellulose and a very slight absorption capacity. In fact, chromatography paper is suitable for electrophoresis. Usually, the electrophoretic cell contains electrodes, buffer reservoirs, support for paper, and a transparent insulating cover; the electrodes are usually made up of platinum the low voltage power pack can be used here (Fig. 12.1). The supporting medium, i.e., filter paper or cellulose acetate, must first be saturated with buffer and held on a sheet of insulating material like Perspex. The sample is then applied in the form of a small spot by using a micropipette. The location of the spot depends on the nature of the molecular mixture present in the sample; for example, if there are molecules with opposite charges, then they will separate and move towards opposite electrodes and hence the sample is applied at the centre. The two reservoirs on either side of the supporting medium are isolated from the electrodes so that any change in pH does not affect the buffer. Then the power is switched on for a required amount of time after the sample is applied. At the end of the experiment, the migrated constituents may be analysed.

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Fig. 12.1. Low voltage electrophoresis (Redrawn from Pattabhi and Gautham, 2004).

Once the run is complete, the electrophoretograms (electrophoretic papers) are dried in a vacuum oven at 100oC.

12.1.2 Vertical Slab Gel Electrophoresis Vertical slab gel unit type (Fig. 12.2) is commercially available and routinely used to separate proteins in acrylamide gels. The gel is formed between two glass plates that are clamped together but held apart by plastic spacers. Gel dimensions are typically 12 cm × 14 cm, with a thickness of 0.5 to 1 mm. A plastic comb is placed in the gel solution, and it is removed after polymerisation to provide a loading well for samples. When the apparatus is assembled the lower electrophoresis tank buffer surrounds the gel plates and it causes some cooling to the gel plates. Since several wells formed side-by-side, a number of samples can be loaded simultaneously, and these samples can be compared. This is the great advantage of this technique over the column type. This technique is extremely popular in the field of molecular biology.

Fig. 12.2. Vertical Slab Gel Electrophoresis (Redrawn from Pattabhi and Gautham, 2004).

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12.1.3 Horizontal Gel Electrophoresis In this system, the gel is cast on a glass or plastic sheet and placed on a cooling plate (an insulated surface through which cooling water is passed to conduct away generated heat). The connection between the gel and electrode buffer is made using a thick wood of wetted filter paper (Fig. 12.3). In all these systems, a power pack is used to supply a direct current between the electrodes. Electrophoresis is carried out in an appropriate buffer, which is essential to maintain a constant state of ionisation of the molecules being separated. Since pH determines the degree of ionisation of organic compounds, it can also affect the rate of migration of these compounds. An increase in pH increases the ionisation of organic acids and a decrease in pH increases the ionisation of organic bases.

Fig. 12.3. Horizontal Gel Electrophoresis (Redrawn from Wilson & Walker, 2010).

Supporting Media Electrophoresis is carried out on a porous mechanical support, which is wetted in electrophoresis buffer and in which electrophoresis of buffer ions and samples could occur. The supporting medium cuts down convection currents and diffusion so that the separated components remain as sharp zones (Wilson and Walker, 2010). Now-a-days, agarose gels and polyacrylamide gels have been effectively used to analyse macromolecules such as proteins and nucleic acids. Agarose is a linear polysaccharide (average relative molecular mass 12000) made up of the basic unit agarobiose, which comprises alternating units of galactose and 3,6-anhydrogalactose. Agarose is one of the compounds of agar and it is a mixture of polysaccharides isolated from certain seaweeds. Generally, 1% and 3% concentrations of agar have been used. Agarose gels are formed by suspending dry agarose in an aqueous buffer, then boiling the mixture until a clear solution is formed. Later, it is allowed to cool down to room temperature to form a solid gel. The pore size in the gel is controlled by the initial concentration of agarose; large pore sizes are formed from low

Electrophoresis 213

concentrations and smaller pore sizes are from higher concentrations. Agarose gels have been used for the electrophoresis of both proteins and nucleic acid. Polyacrylamide gels are also used in analysing proteins and nucleic acids. Electrophoresis in acrylamide gel is frequently referred to as PAGE (polyacrylamide gel electrophoresis). Cross-linked polyacrylamide gels are formed from the polymerisation of acrylamide monomer in the presence of smaller amounts of N, N’-methylenebisacrylomide (bis-acrylamide). In this, bis-acrylamide is essentially two acrylamide molecules linked by a methylene group and is used as a cross-linking agent. The polymerisation of acrylamide is an example of free-radical catalysis and it is initiated by the addition of ammonium persulphate and the base N,N,N’, N’-tetramethylenediomine (TEMED). TEMED catalyses the decomposition of the persulphate ion to give a free radical (a molecule with an unpaired electron). S2O2‾8 + e-- → SO2‾4 + SO‾4• Here SO‾4• is a free radical, represented as R• and M as an acrylamide monomer molecule; then the polymerisation is as follows: R• + M → RM• RM• + M → RMM• RMM• + M → RMMM• Acrylamide gels are defined in terms of the total percentage of acrylamide present and the pore size in the gel is dependent on the concentrations of both the acrylamide and bis-acrylamide. Acrylamide gels can be made with 3%30% acrylamide. Thus, low-percentage gels have larger pore sizes; this can be used to separate proteins and DNA molecules.

12.2 Detection and Quantitative Assay To detect and identify unknown components in the resolved mixture, the electrophoretogram may be compared with another electrophoretogram on which standard components have been resolved under identical conditions. In this respect, it is similar to that of chromatography. Individual compounds are usually detected and identified by their physical properties such as fluorescence and ultraviolet absorption have been used for detection. Staining – A variety of dyes have been used for detecting various components (Table 12.1). Usually, a fixative may be applied before staining; excess stain is eluted after staining.

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Plant Techniques: Theory and Practice Table 12.1. Visual and fluorescent dyes to detect components separated by electrophoresis (adopted from Upadhyay et al., 2003)

Components

Dyes

Remarks

Proteins

Bromophenol blue in acetic acid

Visual, quantitative

Nigrosine in trichloroacetic acid/acetic acid

Visual, very sensitive

Lissamine green in acetic acid

Visual, quantitative

Coomassie brilliant blue

Visual, quantitative

Dansyl chloride

Fluorescent, quantitative

Aqueous anilinonapthalene sulphonate (ANS)

Fluorescent, very sensitive

Methyl green-pyronine

DNA-blue, RNA-red sensitive

Lanthanum acetate + acridine orange in acetic acid

DNA, RNA-orange red

Toluidine blue

Visual, sensitive

Pyronine

RNA, sensitive

Methylene blue

RNA, sensitive

Ethidium bromide

Fluorescent, very sensitive

Lipoproteins

Sudan black in 60% ethanol

Visual, sensitive

Glycoproteins

PAS

Visual, quantitative

Alcian blue

Visual, sensitive

Iodine

Visual, sensitive

Nucleic acid

Polysaccharide

Detection of enzymes in situ – If resolved components are enzymes, these can be detected by special staining techniques. A paper strip of the same size as the electrophoretogram is impregnated with the substrate for the enzyme desired to be separated. This paper strip is now juxtaposed with the electrophoretogram and placed in a suitable buffer. After some time, the sheets are separated. The substrata on the paper strip develop colour upon reaction with the enzyme. The position of the colour developed on the paper strip indicates the position of the enzyme on the electrophoretogram (Fig. 12.4).

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Fig. 12.4. Detection of enzyme in situ (Redrawn from Upadhyay et al., 2003).

12.3 Discontinuous (Disc) Gel Electrophoresis Disc gel electrophoresis (discontinuous buffer employed and discoid appearance of macromolecular zones) is a modification of conventional zone electrophoresis. This allows the sample to enter the gel as a sharp band, thereby helping further resolution (Fig. 12.5). In this, macromolecular mixture is subjected to an electric field in a retarding gel support that is separated

Fig. 12.5. Schematic diagram of Disc Gel Electrophoresis (Redrawn from Upadhyay et al., 2003).

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into two sections differing in porosity and buffered at different pH. The macromolecular mixture migrates from the more porous into less porous gel, a process accompanied by change in pH. As a result, sample can be resolved into very thin, sharp bands (much higher resolution) that can be achieved in a continuous buffer. Polyacrylamide gel has been preferred in the disc gel electrophoresis. The two different porosity gels used are known as the stacking gel (high porosity) and separating gel (low porosity). Separating gel – The separating gel is prepared by using 5-10% acrylamide (the amount of acrylamide used depends upon the approximate molecular weight of the macromolecule being separated) (Table 12.2). Hence, the pores are numerous and of a smaller diameter imparting molecular sieving property to this gel. In this gel, the macromolecules subsequently separate. The buffer used in this gel is usually an amine such as Tris that is adjusted to the proper pH (i.e., 8.3) using hydrochloric acid. Table 12.2. Percentage of separating gel depending upon the molecular weight of the macromolecules Macromolecules Proteins

Nucleic acids

Molecular weight of the macromolecule to be separated

Acrylamide in separating gel (%)

10,000 - 40,000 40,000 - 100,000 100,000 - 300,000 300,000 - 500,000 >500,000

15-20 10-15 5-10 5-7 2-4

Oligomes – 10,000 10,000 – 50,000 50,000 – 200,000 200,000 – 2,000,000

15-20 10 5 2.2-3

Stacking gel – After the polymerisation of separating gel, the stacking gel is poured on top of it. Stacking gel is prepared by using 2.3% acrylamide, since highly porous, it is devoid of molecular sieving action. The buffer used here is also an amine, mostly Tris. The pH is 6.7 (less than the separating gel) and it is adjusted with hydrochloric acid. The buffer used in the sample is identical to that of the stacking gel. The buffer used in the lower reserve is identical to that of the separating gel. The buffer used in the upper reservoir is also an amine. However, it differs from the rest of the buffers in that its pH is adjusted with glycine but not with hydrochloride acid. The pH of the upper reservoir is kept slightly above that of the running gel.

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12.4 Protocols 12.4.1 Agarose Gel Electrophoresis for DNA Analysis (Chawla, 2003; Lee et al., 2012) Agarose gel electrophoresis is a simple and rapid technique of resolving the fragments of DNA and further the localisation of DNA within the gel could be determined directly by staining with ethidium bromide dye under UV light. These bands of DNA can be recovered from the gel and used for variety of cloning purposes. Agarose gel is cast by melting the agarose in the presence of a designed buffer until a clear transparent solution is obtained. The melted agarose is poured into a mould of variety of sizes, shapes and porosity and allow them to harden. Upon hardening, this forms the matrix, the density of which is determined by the concentration of agarose. When the electric field is applied across the gel the DNA, which is negatively charged at neutral pH due to phosphate along the backbone, migrates towards the anode. DNA has a uniform size mass/charge ratio, it is separated by its size within the agarose gel in a pattern such that the distance travelled is inversely proportional to the log of its molecular weight. The rate of DNA migration is determined by the size of DNA molecule, agarose concentration, DNA confirmation, voltage applied, presence of ethydium bromide, type of agarose and electrophoresis buffer (Lee et al., 2012). Various buffers such as Tris-acetate (TAE), Tris-borate (TBE), Trisphosphate (TPE) and alkaline buffer at pH 7.5 – 7.8 are available in the market (Table 12.3). All these buffers contain EDTA (pH 8.0). Double stranded linear DNA fragments migrate approximately 10% faster through TAE than TBE or TPE buffers (Chawla, 2003). Table 12.3. Electrophoresis buffers Buffer (working solution)

Stock solution concentration

Tris-acetate 50x (TAE) (1x = Tris-acetate 0.04 M + EDTA 0.01M)

50x: 242 g Tris base + 57.1 ml glacial acetic acid + 100 ml 0.5 M EDTA (pH 8.0)

Tris-borate (TBE) (0.5x: Tris-borate 0.045 M + EDTA 0.001 M)

5x: Tris base + 27.59 boric acid + 20 ml 0.5 EDTA (pH 8.0)

Tris-Phosphate 10x (TPE) (1x = Tris-phosphate 0.09 M + EDTA 0.002 M)

10x = 108 g Tris base + 15.5 ml 85% phosphoric acid + 40 ml 0.5 M EDTA (pH 8.0)

Alkaline buffer (1x = NaOH 50 mN+EDTA/mM)

1x = 5ml 10N NaOH + 2 ml 0.05M EDTA (pH 8.0)

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Equipment and reagents Agarose gel electrophoresis apparatus, UV transilluminator, gel documentation system, Eppendorf centrifuge, etc. Agarose, Ethidium bromide, Ethylenediamine tetracebate (EDTA), Tris base, glacial acetic acid, phosphoric acid (for TPE buffer), boric acid (for TBE buffer), bromophenol blue, xylene, glycerol, sucrose, molecular weight marker DNA ladder and sodium hydroxide. Stock solution of 0.5 M EDTA (pH 8.0) Dissolve 186.12 g Disodium ethylenediamine tetracelate-2H2O (Na2 EDTA.2H2O. MW = 372.24) in 800 ml water. Stir vigorously with magnetic stirrer. Adjust to pH 8.0 with NaOH and make upto 1 litre by adding water. Sterilise by autoclaving and store at room temperature. Ethidium bromide stain Dissolve 100 mg ethidium bromide in 10 ml distilled water and store at 4oC. Procedure 1. The gel casting platform should be procured along with an electrophoretic apparatus. Seal the ends of a suitable plastic gel casting platform with an adhesive tape. Get the mould either on a horizontal to the bench or on a leveller supply with an apparatus. 2. Now, prepare the 0.6-0.8% agarose gel mix, depending upon the experimental requirement, with 1xTAE or 0.5xTBE electrophoresis buffer and boil it, and stir gently till a homogeneous clear solution is formed. Then cool it to 55%. Agarose with required buffer can also be melted in a microwave oven. 3. By using the pipette, seal the edges of the mould with a small quantity of agarose and then pour it into the gel mould. Place the comb into the gel mould but the position of the comb should be 0.5-1.0 mm above the plate so that a complete well is formed when the agarose is placed. The gel should be 3-5 mm thick. Allow the gel to set for atleast 30 minutes at room temperature. 4. Remove the tape from gel casting mould and keep this gel in an electrophoresis unit containing 1xTAE or 0.5xTBE buffer; see that gel should be completely submerged in the buffer while removing the comb. 5. Take the DNA sample in the desired gel loading buffer and mix them thoroughly. Gel loading buffers are of different types. Add 1/10th volume of gel loading buffer (6x stock) to the DNA sample. Gel loading buffer would serve to increase the density of DNA sample that ensures the DNA drops evenly into the wells, adding colour to the sample and being containing dye that migrates towards the anode at predictable rates in an electric field. Bromophenol blue migrates approximately 2.2-fold

Electrophoresis 219

6.

7.

8.

9.

faster in agarose gel than xylene cyanol FF irrespective agarose gel concentration. Now load the DNA sample into the agarose gel. Generally, 100-500 ng of DNA should be loaded per 0.5 cm slot (a typical slot of 0.5 cm would hold about 37.5 µl). Also load a suitable molecular weight marker in the first and last well along with the DNA sample (~3 µl). Then run the electrophoresis unit by attaching the electrodes. The DNA will migrate towards the anode end (red electrode). Variety of DNA ladders (molecular weight markers) have been used, such as ladders of 1 kb DNA, 500 bp DNA, 100 bp DNA, DNA/EcoR 1 digest, lambda DNA/Hind III digest, lambda DNA/EcoR 1 + Hind III double digest, etc. All these ladders produce fragments of different molecules sizes. After some time, disconnect the electrophoresis unit from the current and take out the gel from it. This gel should be stained with ethidium bromide stain for 20-30 min (always wear gloves while doing the experiment). These stains should be prepared from 20 µl ethidium bromide stock (10 mg/ml) in 200 ml distilled water. Transfer the gel to the UV transilluminator and take a photograph of the gel.

After its use, ethidium bromide should be discarded carefully. As it is powerful mutagenic and toxic, handle the gel carefully by wearing gloves (Chawla, 2003).

12.4.2 SDS Polyacrylamide Gel Electrophoresis (SDS Page) for the Separation of Proteins (Chawla, 2003; Ramudu and Khasim, 2022) Glassware and other equipment Vertical gel electrophoresis apparatus Spacers (x2) Glass plates (x2) Combs Plastic trays (x4) Mortar and pestle Micropipettes Water bath Reagents Sodium dodecyl sulphate (SDS) Acrylamide Tetra methyl ethylenediamine (TMED) Glycerol Bromophenol blue dye

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Coomassie brilliant blue Glycine Tris Methanol Silver nitrate Sodium thiosulphate Sodium carbonate Formaldehyde Hydrochloric acid Glacial acetic acid N’,N’ methylene bisacrylamide Ammonium persulphate (APS) Ethanol 2-Mercaphoethanol Stock solutions i. 10% SDS 10 ml Dissolve 1 g SDS in 8 ml distilled water. Heat to become crystallised. Adjust the volume to 10 ml. Store it at room temperature. ii. 30% Acrylamide-bisacrylamide stock 100 ml Acrylamide 29 g Bisocrylamide 1 g Dissolve acrylamide and bisacrylamide in 60 ml of water. Heat the solution upto 37oC to dissolve chemicals, make up to 100 ml with distilled water and store in dark bottle at 4oC. In this, bisacrylamide is a cross linking reagent. During polymerisation, it cross links the linear molecules of acrylamide to form pore like structures. By varying the concentration of cross linker and monomer, the pore size can be increased and decreased. Precaution: Wear gloves while handling the gel. Since the gel is neurotoxin, it should not come to contact with skin. iii. Ammonium persulfate (APS) 10% APS i.e., (NH4) 2SO4 can be used to initiate the polymerisation reaction. It gives a smooth homogenous polymerisation but may react with proteins and may change artefacts. Dissolve 1g APS in 10 ml of distilled water. This solution is made afresh every time before the gel casting. iv. Tris solution 1.5 M (pH 6.8) 100 ml Dissolve 18.21 g Tris (hydroximethyl) amino methane in 60 ml water (Tris, M.W. = 121.14). Add concentrated HCl, drop-wise until pH falls to 7.4. Then, add in HCl drop-wise until pH falls to 6.8. Make up this solution to 100 ml with water and store at room temperature. Also prepare 1.5 Tris solution at pH 8.8 in the same manner.

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v. Sample loading buffer (pH 6.8) Tris (pH 6.8) 1.5 M 5 ml SDS 10% 6 ml 2-Mercaptoethanol 7.5 ml Bromophenol blue 0.1% 100 mg Glycerol 15% 15 ml Dissolve the ingredients and make total volume of 100 ml. Store it at 4oC. vi. 10x SDS-PAGE running electrode buffer (pH 8.3) 1 litre (Tris-glycine buffer) Tris base 25 mM 30.3 g Glycine 250 mM 144.1 g SDS 0.1% 10 g Dissolve in 850 ml water and make up to 1 litre with distilled water. Store at room temperature. vii. Coomassie staining solution Dissolve 2.5 g Coomassie brilliant blue in a mixture of 450 ml methanol, 100 ml acetic acid and 400 ml water. Adjust volume to 1 litre with distilled water and store it at room temperature. viii. Coomassie destaining solution Methanol (45%) Acetic acid (10%)

450 ml 100 ml

Make up this mixture to 1 litre with water and store it at room temperature. Procedure i. Assemble the leak-proof glass plates according to the instructions given by manufacturers. ii. Decide the quantity of separating and stacking gel. Desired concentration of resolving gel is prepared using the values given in the Table 12.4. Mix the components in the order shown. Polymerisation begins as soon as TEMED is added. Shake the mixture rapidly. iii. Prepare three-fourth volumes of separating gel mix and one-fourth volume of stacking gel and mix them. Use the following volumes of the gels for different spacers. Spacer 0.5 mm spacer 1.0 mm 1.5 mm

Approximate volume of Separating gel Stacking gel 4 ml 8 ml 10 ml

1.5 ml 3 ml 4 ml

iv. Pour the separating gel solution three-fourth of glass plates. Add 200250 µl of water to make the surface even.

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v. Allow the gel to solidify. It takes 20-30 minutes. Solidification can be observed as a line between gel and water. After polymerisation is completed, discard the water deposited on the top of the gel carefully. vi. Now prepare 5% stacking gel according to the table given (Table 12.5). Stacking gel is poured onto separating gel. Insert clean Teflon comb into the stacking gel solution immediately without any air bubbles. Comb must be inserted maximum half-length of stacking gel. Allow it to polymerise at room temperature. Table 12.4. Varying components and their volumes used for the preparation of different concentrations of resolving gel for SDS-PAGE (Chawla, 2003) % Gel

Components

Volume of components (ml) per gel mould volume of 10 ml

20 ml

30 ml

8%

Water 30% Acrylamide-bis-acrylamide mix 1.5 M Tris (pH 8.8) 10% SDS 10% Ammonium persulfate TEMED

5.3 2 2.5 0.1 0.1 0.008

10.6 4 5.0 0.2 0.2 0.016

15.9 6 7.5 0.3 0.3 0.024

10%

Water 30% Acrylamide-bis-acrylamide mix 1.5 M Tris (pH 8.8) 10% SDS 10% Ammonium persulfate TEMED

4.0 3.3 2.5 0.1 0.1 0.008

7.9 6.7 5.0 0.2 0.2 0.016

11.9 10.0 7.5 0.3 0.3 0.024

12%

Water 30% Acrylamide-bis-acrylamide mix 1.5 M Tris (pH 8.8) 10% SDS 10% Ammonium persulfate TEMED

3.3 4 2.5 0.1 0.1 0.008

6.6 8 5.0 0.2 0.2 0.016

9.9 12 7.5 0.3 0.3 0.024

Table 12.5. Various components and their volumes used for the preparation of

stacking gel for SDS-PAGE (Chawla, 2003)

Components

Value of components (ml) per gel mould volume of

5% stacking gel

4 ml

6 ml

Water 30% Acrylamide-bis-acrylamide mix 1.5 M Tris (pH 8.8) 10% SDS 10% Ammonium persulfate TEMED

2.7 0.67 0.5 0.04 0.04 0.004

4.1 1.0 0.75 0.06 0.06 0.006

10 ml 6.8 1.7 1.25 0.1 0.1 0.01

Electrophoresis 223

vii. Fill 1x Tris-glycine buffer in the power pack of SDS-PAGE apparatus. viii. Once stacking gel is solidified, carefully remove the lower spacer and insert gel plate into lower tank. Remove any air bubbles that became trapped at the bottom of the gel between the glass plates. ix. Add running buffer to top reservoir and carefully remove comb. x. Preparation of sample – Prepare protein sample leaf material of Coelogyne nervosa by crushing the material in 200 µl of sample buffer. Transfer it to an eppendorf tube and keep the samples in water both at 100oC for 3-5 min. Centrifuge at 12,000 g for 10 min. Collect the supernatant in a fresh eppendorf tube and store in a refrigerator. xi. Load 10 µl of each protein samples and 10 µl of molecular weight marker (ready to use) into wells (avoid spillage in between wells) with the help of a microsyringe. xii. It should be washed with buffer from bottom reservoir after each sample is loaded. Leave first and last wells, load samples from second well onwards. xiii. Now give the electric power supply to the electrophoresis apparatus. The positive electrode should be connected to the bottom buffer reservoir. Apply a voltage of 8 V/cm to the gel. After the dye front has moved into the resolving gel, increase the voltage to 15 V/cm and run the gel till the bromophenol dye front reaches 0.5 cm from the lower edge of the gel; then turn off the power pack. xiv. Remove gel plates carefully and free them apart with the help of a spatula. Mark the orientation of the gel by cutting a corner from the bottom of the gel that is closest to the no. 1 well. xv. Staining – After the electrophoresis is completed, the gel is stained for proteins. Polypeptides separated by SDS-PAGE can be stained with Coomassie Brilliant Blue-250 and simultaneously fixed with methanol: glacial acetic acid. The gel is placed in a stain and incubated at 37oC for one hour or at room temperature till bands are observed. xvi. Excess stain is removed by soaking the gel in the destaining solution of methanol/acetic acid. Change the destaining solution 3 or 4 times. The dye is sensitive and can detect up to 100 µg (0.1 mg) of protein in an acrylamide gel. xvii. The bands can be photographed and analysed at this stage. Gels may be stored indefinitely in a sealed plastic bag. Result: The distinct sharp blue bands can be seen in the gel. The intensity of these bands is variable depending on the number of polypeptides in the same protein solution loaded in the gel. Using a protein molecular weight marker, calculate the molecular weight of protein samples. The distance migrated by each protein sample can be measured and compared with the standard protein travelled distance. By taking molecular weight in Y-axis and distance in X-axis, plot a graph.

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Fig. 12.6. SDS-PAGE protein banding pattern in six populations of C. nervosa (PM – Protein marker, P1 – Yercaud, P2 – Dodabetta, P3 – Palni, P4 – Kodaikanal, P5 – Wayanad, and P6 – Munnar) Courtesy: Ramudu and Khasim, 2022.

In C. nervosa, the SDS-PAGE protein profile showed multiple bands of varied molecular weight ranging from 14 Kd to 116 Kd in six populations collected from different parts of the Western Ghats of India (Fig.12.6). Intraspecific and interspecific diversity could be studied by SDS-PAGE protein profile data.

13 Molecular Techniques Minute and cryptic genomic variations in nucleic acid sequences are crucial factors that distinguish different plant systems from one another, and such variations can be ascertained through molecular assays. In recent times, the field of plant molecular biotechnology has witnessed the emergence of a wide array of techniques, from molecular markers to monitor genetic variations, Next Generation Sequencing technologies, and genome editing tools such as conventional plant transformation or Clustered Regularly Interspaced Short Palindromic Repeats/Cas9 (CRISPR/Cas9) genome editing system. All these techniques have collectively revolutionised plant molecular biology and biotechnology. However, an important prerequisite is the extraction of genomic DNA from plant specimens, which is the starting material for any of these studies (Aboul-Maaty and Oraby, 2019). Over the years, numerous protocols have been developed and reported for the extraction of high-quality genomic DNA isolation from plants (Dellaporta et al., 1983; Saghai-Maroof et al., 1984; Doyle and Doyle, 1990; Scott and Playford, 1996; Pirttilä et al., 2001; Shepherd et al., 2002; Syamkumar et al., 2003; Masoodi et al., 2020). Although all these procedures are reportedly successful in the isolation of pure and intact plant DNA, they often require modifications/ minor alterations when applied to different plant systems (Aboul-Maaty and Oraby, 2019). This may be attributed to biochemical heterogeneity among plant samples (Heikrujam et al., 2020). Various primary and secondary plant metabolites, as well as carbohydrates, lipids, polyphenolic compounds, etc, severely affect the quality of the DNA extracted (Khanuja et al., 1999; Aboul-Maaty and Oraby, 2019). Most of these compounds also reportedly act as inhibitors in Polymerase Chain Reaction (PCR) based molecular studies (Xin et al., 2003; Heikrujam et al., 2020), hence, affecting the quality (purity) of the DNA samples isolated. Removal of biomolecular contaminants from the cells is often achieved with the help of chemicals such as CTAB (Cetyl-trimethyl Ammonium Bromide)(Saghai-Maroof et al., 1984; Doyle and Doyle, 1990) or Sodium Dodecyl Sulfate (SDS) (Dellaporta et al., 1983).Therefore, most of the protocols developed for isolation of genomic DNA from a plant cell follow the basic principle, wherein, the cell wall, cell membrane and nuclear membrane are disrupted to precipitate the nucleic acids (DNA and RNA), which is further purified to obtain the intact genomic DNA. The concentrations

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of the contents of the extraction buffer are routinely altered to increase the quality and yield of DNA isolated.

13.1 Genomic DNA Extraction 13.1.1 Choice of Plant Tissue and Function of Each Reagent Used The plant tissues from which DNA is to be isolated has to be identified prior to extraction. Young and disease-free leaves are frequently chosen, as the quality and yield of DNA is superior. The tissues are then homogenised in dry ice, liquid nitrogen or tissue homogeniser to digest the cell wall and release the contents of the cell. The sample is then exposed to a suitable extraction buffer. The extraction buffer contains chemicals such as SDS (anionic) or CTAB (cationic) which are detergents which facilitate the disruption of the cell membranes. Sometimes, chemicals such as PVP (polyvinylpyrolidine) are used in addition to CTAB (Cetyl trimethyl ammonium bromide) to denature metabolites such as polyphenols and lipids (Heikrujam et al., 2020). β-mercaptoethanol in the extraction buffer helps dissolve disulfide bonds to denature complex proteins, while Tris and NaCl protect the nucleic acids and trap the proteins bound to them. In nature, genomic DNA is protected from endogenous nucleases. During DNA extraction, EDTA (Ethylene diamine tetra acetic acid), present in the extraction buffer acts as a chelating agent for divalent cations such as Mg2+and Ca2+ions, thereby preventing endonuclease activity. With the removal of the cellular membranes, the DNA remains free in the solution but is still bound to proteins and other contaminants. Denaturation of proteins is accomplished by phenol, which is aided by chloroform which also acts as a stable barrier between the aqueous phase and the alcohol phase. Thorough mixing of the Phenol and C: I solution is necessary to ensure phase separation and maximise the yield of DNA. The addition of Isoamyl alcohol is carried out to prevent foaming and ensure proper mixing of solutions and phase separation thereafter (Khanuja et al., 1999; Ignacimuthu, 2001). The denatured proteins form a layer at the interphase. The nascent DNA in the aqueous phase is prone to wear and tear and thus should be handled with extreme care and caution. Chilled EtOH or isopropanol is used to spool out the DNA from the aqueous phase (Heikrujam et al., 2020). Purification of extracted DNA: Repeated freezing and thawing of samples should be avoided and efforts must be made to reduce its exposure to the extraction buffer. Since the genetic material isolated will comprise both RNA and DNA, the samples have to be subjected to treatment with RNase A enzyme to facilitate the breakdown of RNA. Proteinase K will further purify the samples to disrupt the peptide bonds of the proteins attached to the DNA though endopeptidase activity.

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13.2 Quantification of DNA Samples Isolated Quantification of the DNA samples involves estimating the concentration of DNA in the samples. Usually, this estimation is carried out spectrophotometrically, using a NanoDrop spectrophotometer. For an unknown sample, the DNA concentration can be measured using BeerLambert’s law. Different substances generally contaminating DNA samples absorb UV light at different wavelengths as detailed in Table 13.1. At the same time, the A260:A280 ratio can give a clue about the purity of the DNA (Wilson and Walker, 2010). Generally, for double stranded DNA, its characteristic ratio should be 1.7 to 1.9 (Ignacimuthu, 2001). A higher ratio is expected if the sample is contaminated with RNA, while a lower ratio is observed with proteins and phenolic compounds. Absorbance ratio of ~1.8 is ideally associated with high quality DNA. A260:A280 ratio thus accurately suggests the purity of DNA samples isolated. Table 13.1. Peak absorbance wavelength of biomolecules and chemicals involved in DNA extraction Peak Absorbance (in nm)

Substances

230

EDTA, Polysaccharides, Ethanol

260

Nucleic acids (DNA/RNA)

270

Phenol

280

Proteins

320

Cell debris

13.3 Quality of the DNA Samples Isolated The quality of the DNA samples isolated can be ascertained by separating them on 0.8% agarose gel. Gel electrophoresis involves migration of DNA bands on agarose gel matrix under influence of an electric field (Ignacimuthu, 2001; Chaitanya, 2013). Since most protocols for extraction involves isolation of total genomic DNA, which is high molecular weight molecule, and hence, a lower percentage of gel is used to separate it. Table 13.2 details the size of DNA fragments and the gel percentage used to separate them. Intact DNA, in its native form shows up as a sharp band (≥40 bp), while sheared and degraded samples appear smeared on the gel. Parameters such as application of voltage and agarose concentration should be critically evaluated to ensure proper migration of DNA on the gel matrix. Generally, a ladder (500 bp) or Hind III digest is used as a standard marker to evaluate the genome size of the plant (Aboul-Maaty and Oraby, 2019). Lambda DNA is the commonly used marker for gel electrophoresis and when it is cut with restriction enzyme EcoR1 and Hind III (double digest) the following marker sizes (bp) will appear in the gel (Ignacimuthu, 2001).

228

Plant Techniques: Theory and Practice Table 13.2. Percentage of agarose gel to separate DNA fragments based on their size

Agarose %

Size of fragments separated (kb)

0.5

1-30

0.7

0.8-12

1.0

0.5-10

1.2

0.4-7

1.5

0.2-3.0

2.0

0.05-2.0

13.4 Agarose Gel Electrophoresis 13.4.1 Reagents and Buffers Two different buffer systems are generally used for carrying out gel electrophoresis, viz., TAE (Tris-Acetic acid, EDTA) and TBE (Tris Boric acid EDTA) buffers. While both the systems are equally competent, TAE has comparatively lower ionic strength and is used to separate larger DNA fragments (>12 kb). This buffer system is usually efficient when the aim is gel purification and extraction of the DNA fragment. However, this buffer has to be replaced for extended run times. In contrast, TBE buffer has no such requirement, and has a much higher buffering capacity. It facilitates separation of smaller DNA fragments ( 7.5); acidic pH affects UV absorption spectrum of RNA and significantly decreases the ratio.

13.7.10 In Situ Hybridisation Techniques Cytogenetics studies have contributed immense knowledge to biology. Cytogenetics is a branch of genetics that studies the number and morphology of chromosomes. It studies the heredity and variations through morphology, structure and behaviour of chromosomes during cell division. Various cytogenetical studies have led to the revealing of unexpected details of

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chromosome morphology, behaviour and evolution. The combination of the two disciplines of molecular biology and cytogenetics has generated a new area of research, which is known as Molecular Cytogenetics. The field of molecular cytogenetics has developed to include many cyto-molecular techniques, mainly FISH (Fluorescenti in situ Hybridisation), GISH (Genomic in situ Hybridisation), etc. The use of molecular cytogenetical techniques like FISH and its variants helps in the studying of individual genomes/chromosomes or chromosomal fragments. These techniques are very helpful in analysing polyploids, hybrids, etc. Molecular cytogenetics includes various techniques such as heterochromatin banding, FISH, GISH, McFISH, Oligo-FISH, Flow cytometry, Flow sorting, Flow cytogenetics, Microcloning, Microdissection, etc. For these studies, preparation of mitotic or meiotic slides is required for analysis. 13.7.10.1 Heterochromatin Banding

Heterochromatin staining with base-specific fluorochromes is a reliable method to distinguish some types of heterochromatins in plants. Heterochromatin detection together with physical mapping of DNA sequences by fluorescent in situ hybridisation (FISH) has proved to be a useful technique to analyse the patterns of karyotype evolution. This information provides a clearer view of the possible role of epigenetic factors in influencing complex cellular phenomena. 13.7.10.2 FISH

FISH uses complementary hybridisation of labelled DNA/RNA probes. It helps in the direct mapping of the DNA sequences on the chromosomes. It is very effective for mapping single and repetitive DNA sequences present on the chromosome. FISH studies performed in the rRNA genes act as chromosome markers, which are excellent tools to improve karyotype description. The broad applications of FISH in structural, comparative and functional genomics place plant cytogenetics in a unique position to complement, accelerate, or guide plant-genome research. It has enhanced the elucidation of the numerical and complex chromosomal rearrangements (deletions, duplications, translocations, etc.). In the FISH technique, the slides are pre-treated with pepsin to remove RNA and protein contamination which may bind to the probe and increase the background noise. The removal of these contaminants helps the probes to access the target DNA easily. It also fixes the chromosomes and nuclei on the slide, so that they are not washed off during the procedure (Fig. 13.2).

Molecular Techniques 241

Fig. 13.2. Methodology of Fluorescence in situ hybridisation.

13.7.10.3 Mc-FISH (Multi-coloured FISH)

It is a variant of the FISH technique. Mc-FISH is preferred when several target sequences are to be probed on a chromosome simultaneously. Mc-FISH requires the use of several fluorophores. Different target sequences can be labelled with fluorophores which have different emission spectra (Table 13.4). Table 13.4. Fluorophores and binding sites Fluorophores

Colour

FITC (Fluorescein isothiocyanite)

Green

CY3 (Cyanine3)

Red

AMCA (7-amino 4-methyl Coumanine 3-acetic acid)

Blue

DAPI (4′,6-diamidino-2-phenylindole)

Blue

TRITC (tetramethylrhodamineisothiocyanite)

Red

Binding region 5’ end of DNA

AT-rich regions

13.7.10.4 GISH

GISH is a powerful complementary tool, which uses total genomic DNA as probes. It is very useful to study inter-specific hybrids. The DNA probe helps

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in visualising the whole genome or a portion of the genome, contributed by each parental species in interspecific hybrids. Just like FISH, the GISH also has the following steps:

1. Slide preparation Cells at metaphase (mitotic metaphase spread) stage. Denaturation is carried out on the genomic DNA of one parent and the genomic DNA of another parent. 2. DNA probes preparation Here one is nick translated genomic DNA which is labelled one from one parent; another probe is unlabelled one. Nick translation, developed by Rigby et al. (1977), is a tagging technique in molecular biology, in which DNA polymerase I is used to replace some of the nucleotides of a DNA sequence with their labelled analogues, creating a tagged DNA sequence. So that fluorochrome is effectively bound to this tagged DNA sequence (Nick translated DNA sequence).

3. Hybridisation Prepare hybridisation solution (HS) that contains autoclaved Salmon testes DNA as carrier DNA, autoclaved genomic DNA as an unlabelled competitor DNA from one parent and fluorochrome labelled nick translated genomic DNA from another parent. 4. Counter stain with DAPI (Diamidino-2-Phenyl indole) blue dye. 5. Visualisation Observe under the fluorescent microscope. Unlabelled probe is visualised with counterstaining (blue) where labelled probe with green. Therefore, the GISH is also helpful to analyse natural polyploids, hybrid plants and their back cross progenies to ascertain their origin, genomic composition and intergenomic rearrangements. 13.7.10.5 Oligo-FISH

In this technique, oligonucleotides are used as probes. Oligonucleotides are polynucleotides containing a relatively small number of nucleotides. If highly repetitive sequences are targeted, properly designed synthetic oligonucleotide probes may prove to be superior reagents for rapid FISH. They have certain advantages such as, it hybridises rapidly, it is consistent and cheap. Oligo probes can be designed easily using the Chorus software (https://github.com/ forrestzhang/Chorus). Chromosome spreads can be prepared from actively growing root tips. 13.7.10.6 Fiber-FISH

Fiber-FISH is the technique where in situ hybridisation (FISH) is performed on extended DNA fibers. A high-resolution map can be created by using the

Molecular Techniques 243

interphase nuclei as chromatin fibres extending into the nuclei because they stretch much more when compared to those in the metaphase chromosomes. It provides a fast and direct means of physical mapping at the molecular cytogenetic level. Nuclei are isolated from plant cells by destroying the nuclear membrane. The DNA fibers generated are then hybridised directly with DNA labelled probes. This procedure is used to map DNA clones at a resolution of 50-500 kilobase pairs. 13.7.10.7 Chromosome Painting

Chromosome painting (CP) is a technique to visualise the entire chromosome via fluorescence in situ hybridisation (FISH) using chromosome-specific painting probes (Pinkel et al., 1988). CP is a powerful tool to diagnose chromosome abnormalities, detecting karyotype alterations during evolution, and constructing ancestral karyotypes (Marshall and Obe, 2015; Meng et al., 2021). During the last few decades, CP probes have been mainly amplified for flow sorted or microdissected chromosomes, followed by degenerate oligonucleotide-primed PCR amplification. However, presence of repetitive DNA in the genomes of plants results in the unfavourable non-specific hybridisation signals. Technical advances in DNA synthesis have made it possible to massively synthesise oligonucleotides based on genome assembly without repetitive sequences. FISH technique in plants has shown superior resolution and versatility of oligo-based probes compared to conventional genomic clone or single-copy gene-based probes (Meng et al., 2018, 2020; Xin et al., 2018; Simonikova et al., 2019). Various steps involved in the chromosome painting are given below (refer to Meng et al., 2021): 1. Chromosome spread preparation and denaturation Root tips are taken and fixed in Carnoy’s fluid (3:1 Acetic acid and alcohol) and stored in -20 °C until use. Later, root tips are digested in an enzymatic solution consisting of 2% cellulase and 1% pectolyase; incubated for 1 hour at room temperature, then squashed with a coverslip. Afterwards slides are dehydrated in an ethanol series (70, 90 and 100%, 5 minutes each) prior to FISH assay. 2. Designing and synthesis of oligo-based chromosome 2 (from Saccharum spontaneum) painting probeThe oligo-based Painted probe of S. spontaneum chromosome 2 is designed using Chorus software. The oligos (59 nt) specific to chromosome 2 are synthesised de novo in parallel by MY microarray (Ann Arbor, MI, United States). These are labelled in biotin or digoxigenin. 3. FISH studies using oligo and rDNA probes˗ The above labelled chromosome painting probes are directly used for FISH analysis.

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˗ Rice 45S and 5S rDNA (Gong et al., 2002) are labelled with either digoxigenin-11-dUTP (Roche Diagnostics, USA) or biotin-16-dUTP using standard nick translation reactions. ˗ In FISH assay, firstly, chromosomes are denatured in 70% formaldehyde in 2X SSC at 70°C for 1 minute and dehydrated in an ethanol series again. Now, the chromosome mixture (50% formaldehyde, 10% dextran sulphate, 20X SSC, 50 ng labelled probe) is denatured at 90°C for min and applied to the denatured chromosome studies, incubated for 12 h at 37°C. ˗ Subsequently, digoxigenin-and biotin-labelled probes are detected using rhodamine-conjugated anti-digoxigenin (Roche Diagnostics, USA) and fluoroscen-conjugated avidin (Life Technologies, USA) respectively. ˗ Slides are counterstained with DAPI in VeetaShield antifade solution. 4. Visualisation of ChromosomesFISH signals are detected under the Olympus BX63 fluorescence microscope. Images are captured and merged by Cell Sens Dimension 1.9 software with an Olympus DP80 CCD Camera. The final signals are processed and adjusted by Adobe Photoshop CC software (Meng et al., 2021). Chromosome Painting has become a versatile tool in basic research disciplines ranging from radiation biology to evolutionary cytogenetics and nuclear structure studies.

13.8 Other Approaches 13.8.1 Flow Cytometry In plants, ploidy estimation is carried out by chromosome counting on microslides prepared from actively growing root-tips. Sometimes it may be complicated due to very small size of chromosomes, insufficient spreading on slides and low frequency of dividing cells. Under such conditions Flow Cytometry (flow = motion; cyto = cell; metry = measure) can be used for the determination of the nuclear DNA content in plants. To estimate the nuclear DNA content, suspension of nuclei or permeabilised cells are stained with DNA specific flourochrome and amount of light emitted by the each nucleus is quantified and the result of the analysis is displayed in the form of histogram of relative fluorescence intensity, representing the relative DNA content in the samples. In short, measuring the properties of cells while in the fluid stream is known as Flow Cytometry. However, in this technique, preparation of nuclear suspension or cell suspension suitable for flow cytometry is achieved with utmost care. Lucretti et al. (1993) studied the flow karyotyping and sorting of Vicia faba chromosomes. They sorted out the acrocentric and metacentric chromosomes

Molecular Techniques 245

using the flow cytometry. The amount of light emitted from the chromosome can be quantified and able to sort the type of chromosomes. 13.8.1.1 Instrumentation

Traditional flow cytometers: These flow cytometers have three systems such as fluidics, optics and electronic system. In fluidics system, sample is injected into a stream of sheath fluid (mostly buffered saline solution), and it is exposed to laser beam. Next one, optics system consists of excitation optics and collection optics. Excitation has laser and lens that are used to focus the laser beam to flow of sample; whereas collection optics consists of collection of lenses which collects the light emitted from samples and these signals detected by optical detector. Finally electronic system converts signals into digital form and that can be read by computer. Data analysis: Various software are available, among them FCS 3.1 software has been used for any flow cytometry analysis program. Stains: Now-a-days, different dyes available in the market with excitation and emission spectra ranging from ultraviolet to near infrared. Polymer dyes such as brilliant violet (BV), brilliant ultraviolet (BUV), brilliant blue (BB); fluorescent protein dyes including green fluorescent protein (GFP), cyan fluorescent protein (CFP), and yellow fluorescent protein (YFP), and all these are derived from jelly fish, while red fluorescent protein from mushroom anemone; other dyes for nucleic acids including propidium iodide (PI), 4,6-diamidino-2-phenylindole (DAPI), dyecyclic violet etc. have been used for flow cytometry studies. 13.8.1.2 Vicia Faba Chromosomes Sorting

Lucretti et al. (1991) classified the chromosomes according to their relative DNA content, base content, protein content and morphological parameters by using flow cytometry. A chromosome suspension from root tips is prepared and see that all cells should be at metaphase state: Synchronisation and accumulation of cells in metaphase– • Root tips are incubated in Hoagland solution containing hydroxy urea for 12 hours. • Washing and treated with amiprophos-methyl (APM) for 2 hours; all cells are accumulated in metaphase state. • Then root tips are collected, washing, fixed for 30 min at 5 ºC in 6% formaldehyde made in tris buffer (10 mM Tris, 10 mM Na2 EDTA, 100 mM NaCl, pH7.5) with 0.1% Triton X-100. • Washing; root tips cut into pieces and treated with 1 ml lysis buffer (15 mM Tris, 2mM spermine, 15mM mercaptoethanol, 0.1% Triton X-100, pH 7.5).

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• The suspension of released chromosomes and nuclei is filtered through a 50µm and 15µm nylon meshes. This chromosome suspension is separately collected and used for Flow cytometry analysis. • To remove interphase nuclei and chromosome clumps, 750 µl of the suspension is layered on 700 µl of 40% sucrose in Tris buffer in a 10 ml centrifuge tube and centrifuged at 200 rpm for 15 min. Now the supernatant is transferred to a sterile 1.5 ml Eppendorf tube. • Chromosome suspension is stained with propidium iodide (PI) at final concentration of 50 µg/ml for 30 minutes before analysis. • Now this chromosome suspension is analysed with flow cytometers and sorter. Acrocentric and metacentric chromosomes could be sorted using this technique. In karyotyping studies, the histogram of fluorescence intensity obtained from PI-stained chromosome suspension is developed. Based on the histogram data, chromosome sorting is carried out. Modelling of flow karyotypes can be done using software i.e., KARYOSTAR software. Now these sorted chromosomes are shown to be suitable for in situ hybridisation. Chromosome suspension is separately collected and used for Flow cytometry analysis.

Fig. 13.3. Methodology of analysis of various plant samples using flow cytometry. (Adapted from the website http://www.ueb.cas.cz/Olomouc1/LMCC/lmcc.html.).

Molecular Techniques 247

Flow cytometry and sorting is an exceedingly powerful technique for purifying cell sub-populations from complex cell mixtures. Flow cytometry gives us the scope to use multi-parametric analysis to identify highly specific populations. Phenotypic characteristics, DNA content of cells, RNA content, or even assess functional characteristics such as ion flux or pH or altered cell states such as apoptosis and cell death can be analysed through this technique. This technique uses metaphase chromosomes which are stain with fluorescent dyes (one that binds to AT pairs, the other to GC pairs). The stained chromosomes flow past a laser beam and a photometer sorts them based on differences in the scattering of light (Fig. 13.3). According to the method of Galbraith et al. (1983), preparation of suspension of intact nuclei for analyses of nuclear DNA content has been done by mechanical homogenisation (chopping). The nuclear suspension prepared in a nuclear isolation buffer (4-morpholine Propane sulfonate/TRIS) is then sieved using a nylon mesh to remove large debris and nuclei stained with DNA specific fluorochrome (Propidium iodide/DAPI). As flow cytometry analyses relative fluorescence intensity, absolute estimation of nuclear DNA content requires the use of a reference standard of known genome size (Fig. 13.3).

13.8.2 Microdissection and Microcloning In microdissection and microcloning technique, the target chromosome is cut by glass micro-needles or lasers in an inverted microscope for separation from other chromosomes and the isolated chromosome segment is then collected and amplified in vitro to establish specific DNA libraries. After the development PCR technique, Chromosome dissection and microcloning methods was firstly applied to the rye chromosomes by Sandery et al. (1991). Now it has become a powerful technique for the localisation and cloning of plant genes. Experimental procedure The chromosome microdissection and microcloning technique is divided into five steps: 1. Preparation of chromosome specimen 2. Identification of target chromosomes 3. Microdissection and collection of target chromosomes or chromosome segments 4. Construction of microcloning library 5. Screening and identification of microclones. Identification of target chromosomes The chromosome specimen must be prepared precisely by conventional methods. After chromosome preparation, appropriate methods must be used to identify the target chromosome. Karyotyping analysis and chromosome banding are employed here. For example, meiotic monovalent chromosomes

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in wheat trisomics lines can be identified by karyotyping analysis and successfully microdissected; similarly, B chromosome of Rye can be detected by chromosome banding technique and then microdissected. Chromosome microdissection and construction of Microcloning Library Microdissection and collection of target chromosome/segments can be carried out by glass micro-needles, laser micro-beams and flow cytometry. Currently, there are special laser beam devices available for microdissection such as Laser beam devices—Arcturus XT, USA make; PALM Microbeam, Carl Zeiss, Germany. After microdissection, the obtained segments can be amplified in vitro for subsequent studies. The construction of chromosome micro cloning libraries has mainly been conducted by two methods such as Direct cloning after enzyme digestion and PCR mediated cloning methods. Direct cloning after digestion This technique was widely used before the PCR came into existence. Here, firstly, DNA is extracted from target chromosomes/segments, later deproteination and purification of DNA is carried out. Next, the extracted DNA is digested by restriction enzyme. Finally, the DNA digested is added to ligase to link the vector. This DNA linked to the vector is transferred into host bacteria to construct the chromosome DNA library. Sandery et al. (1991) first applied this technique to rye plant; B chromosome was separated and cut with glass microneedle and successfully microcloned the chromosome after extraction, deproteinisation, purification and digestion. PCR-mediated microcloning After PCR amplification in vitro, additional target chromosomes can be obtained to meet the needs of the molecular biology experiments. Here Linker Adaptor PCR (LA-PCR) has been adopted, this is as follows:

• First, DNA from target chromosome and digested by restriction enzyme; • Second, according to the base sequence of sticky end produced after enzyme digestion, one linker and adaptor (short stretches of nucleic acid that helps in DNA ligation) are designed; and • Finally, PCR amplification is carried out by using the linker as the primer. Guo (1998)—successfully cloned the B chromosome of rye by LA-PCR with the linker-adapter system of Sau3A1 (from Staphylococcus aureus). Screening and identification of microclones The specificity of the constructed DNA library is generally screened and identified by using Southern hybridisation, in situ hybridisation and the PCR amplification. Southern hybridisation is useful to identify the source of the inserted DNA fragments in the library. Hu et al. (1998) successfully proved that microclones produced from in vitro PCR were derived from maize genome.

Molecular Techniques 249

Guo et al. (1998) successfully located the PCR products at the terminal end of the B chromosome of the rye using the in situ hybridisation. With the help of PCR technique Hu et al. (2004) identified four different DNA fragments, R1, R2, R3, R4 and these were proved to be derived from wheat 6B chromosome using SSR markers. Chromosome microdissection and microcloning are efficient tools that combine the fields of cytogenetics and molecular genetics. It is highly useful for establishing the relationships between specific chromosomes and specific linkage groups, for localising and cloning genes of interest etc.

13.9 Protocols 13.9.1 Isolation of Intact Genomic DNA, its Quality Check and Quantification Reagents for DNA Extraction

• CTAB stock: 10% CTAB solution is made. Complete dissolution of CTAB occurs at higher temperatures, therefore heating may be necessary. • IM Tris-HCl, pH 8.0: 6.057g Tris is dissolved in 50 ml double distilled water using a magnetic stirrer. • 5M NaCl: 50ml solution is prepared by dissolving 14.96g NaCl dd water. • 0.5 M EDTA, pH 8.0: 0.7306g of EDTA is dissolved in 10 ml double distilled water. All the solutions for DNA extraction should be autoclaved and stored at 4°C for future use (Ignacimuthu, 2001).

• Extraction Buffer: A typical extraction buffer contains 100mM Tris buffer, 1.4M NaCl, 10mM EDTA solution and 1% CTAB. 0.2% mercaptoethanol is added to the buffer just before use. • RNase stock: 10mg/ml solution is prepared by dissolving RNaseA in 1M Tris buffer and 5M NaCl solution. • 3M Sodium Acetate, pH 5.2: 40.81g of sodium acetate is dissolved in 100 ml of distilled water. The pH of the solution is adjusted to 5.2 with glacial acetic acid. • 1X TE: 10 mM Tris buffer + 1mM EDTA is mixed and the final volume is made up with double distilled water. • Proteinase K: For 80 mg tissue, 100 mg/ml proteinase K is used. The samples are incubated for 12-18 hours at 50°C. Step-wise procedure for DNA Extraction i. 100-150 mg of young and disease-free leaves from each sample are cleaned with EtOH. The samples are then ground to fine powder using liquid nitrogen with mortar and pestle or by a tissue homogeniser. The macerated samples are then immediately transferred to 800µL of the pre-

250

ii.

iii. iv. v.

vi. vii.

viii. ix. x.

Plant Techniques: Theory and Practice

warmed (at 60°C water bath) extraction buffer. The tubes containing the buffer and the samples are mixed by inversion to form even suspensions. The suspensions are incubated in a water bath at 60°C for 45 minutes-1 hour. The tubes are gently inverted every 15 minutes to ensure mixing. After the incubation period is over, equal volume of phenol: chloroform: isoamyl alcohol solution (25:24:1) is added to the tubes. The tubes are then placed on a shaker at room temperature for 15-20 minutes so that the contents of the tube are properly mixed and phase separation occurs. The phases, aqueous and organic are separated by centrifugation at 13,000g for 10 minutes. The aqueous phase containing nucleic acids are pipetted out to fresh tubes. The DNAis precipitated out by adding ¾ volume of ice chilled isopropanol or 2 volumes of chilled EtOH to the aqueous phase. Additionally, 3M sodium acetate may also be added to the sample to facilitate precipitation of DNA. After gentle mixing of the contents of the tube, the DNA strands maybe visualised floating in the solution. The tubes are kept at -20°C for 20-30 minutes or may even be kept overnight to increase the yield of DNA. After the requisite incubation period is over, the DNA is precipitated by centrifugation at 13,000g for 10 minutes at room temperature. The DNA is obtained at a pellet, which is subsequently air-dried and washed several times with 70% EtOH. The DNA is again pelleted after the washing step is over and the pellet is air-dried and dissolved in 50-100µL of TE buffer. The samples are then incubated with RNaseA and Proteinase K for 1 hour (at 37°C) and 12 hours (at 50°C) respectively. Quality check and quantification of DNA is further tested by agarose gel electrophoresis and estimation based on absorbance values.

Step-wise procedure for agarose gel electrophoresis i. Sealers/adhesive tapes should be used to securely seal the ends of the gel casting tray in which the gel is to be polymerised. ii. The gel casting tray is set, and combs are placed at appropriate distance to ensure the formation of properly formed wells. iii. Agarose is weighed in a balance and melted in 1X running buffer till a homogenous solution is obtained. iv. EtBr is added to the agarose solution, which is thereafter poured into the casting tray. It is allowed to settle down for the gel to polymerise. v. Once it is properly polymerised, the combs and the adhesive tapes are carefully removed. vi. The gel is then placed in the tank by submerging it in the running buffer. vii. The samples are loaded into the wells. 1/10 volume of Tracking dye (usually Bromophenol blue or SyBr Green) is added to the samples so that we can track the movement of DNA in the gel matrix.

Molecular Techniques 251

viii. A marker is added to the last lane which will serve as a standard to estimate the size of our DNA fragment. ix. The electrodes are connected with the negative terminal connected to the black electrode and the positive terminal connected to the red one. x. The power supply should be turned on and the gel is run till the samples reach 1.0 cm above the end of the gel. xi. The gel is removed from the apparatus and visualised in the gel documentation system. Result / Note: While running down the genomic DNA on agarose gel, the bands appear very clearly without any contamination. If RNA appears, that indicates the incomplete RNase treatment of DNA. Being smaller in size, the RNA moves faster than the DNA.

13.9.2 Polymerase Chain Reaction (PCR) PCR is performed on the thermocycler. As it is already mentioned earlier that it involves three steps, such as Denaturation, annealing and extension. Equipment and Chemical Reagents

• PCR machine – Thermocycler program should be set up according to the details given in Table 13.5. Table 13.5. Thermocycler for DNA amplification – various steps Temperature (oC)

Time

94

5 min

35-65

30 sec

Annealing

56

40 sec

4

Extension

72

1 min 50 sec

5

Final extension

72

15 min

S. No.

Step

1

Initial denaturation

2

Denaturation

3

No. of cycles 1 35 1

Components of Reaction Mixture (20 µl)

• Template DNA - 20-30 ng of genomic DNA that is to be amplified. • Pfu DNA Polymerase – 1 unit of this enzyme obtained from Pyrococcus furiosus and it could withstand at high temperature 98oC. • A pair of oligonucleotides (forward and reverse) primer – 10 µM. Two short fragments of single-stranded DNA with 15-30 nucleotides are specific for the target sequences and are complementary to the 3’ ends of the sense and antisense strands of the target sequence so that they have extended each other in opposite direction. • Deoxynucleotide triphosphates (dNTPs) – 2 µl of dNTPs. They include single units of all DNA – forming nitrogenous bases in their deoxynucleoside triphosphates (dATP, dTTP, dGTP, dCTP) which form the building ingredients for DNA synthesis.

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• MgCl2 – 20mM – Magnesium ion (Mg2+) in the +2 oxidation state acts as cofactor and enhances the enzyme activity and polymerisation of DNA. • 10x PCR buffer (pH 8.0) – 20 µl of this buffer host the reaction and provides congenial environment for PCR activity. • Distilled water is added to make up the final volume of reaction mixture, i.e., 20 µl. Protocol i. Thawing the working standard PCR components. ii. Gently vortex and briefly centrifuge all solutions after thawing. iii. Prepare PCR master mix in a sterile 0.5 ml PCR tube by adding all the above components according to the quantity given. Enough master mix for the number of reactions should be prepared and add one more to compensate for pipetting errors. iv. Aliquot the master mix into individual sterile 0.2 ml PCR tubes and the template DNA (genomic DNA) to it. v. Gently spin the samples. vi. Place the PCR tubes loaded with samples in the thermal cycler. Carryout the amplification according to the program set up mentioned in the Table 13.5. vii. Polymerisation should be carried out for 1 min for every 1000 bp of length of the target DNA. viii. After reaction is over, take out the tubes and run 10 µl of reaction mix in 1% agarose gel for 45 min at 60 volts. Run the sample along with the marker (1 kb ladder). ix. Visualise the PCR products in the UV light by using a gel documentation system.

Fig. 13.4. RAPD banding pattern of PCR amplified products of the mother plant and in vitro regenerated plants (L = Ladder; M- Mother plant; R1 – R6 – regenerated plants).

Molecular Techniques 253

Result: Purified DNA from the mother plant and six samples of in vitro derived plants of Dendrobium aphyllum were used for the genetic fidelity studies. Amplified PCR product with RAPD primer showed the monomorphic and polymorphic bands (Fig. 13.4).

13.9.3 RNA Extraction Extraction buffer (for RNAextraction): 8M Guanidine hydrochloride, 20mM MES (2-N-Morpholinoethanesulfonic) DEPC treated water, 20mM EDTA pH7.0, sterilise by filtration. Before use, add 50 mM MES (2-(N-Morpholino)ethane sulphonic) DEPC-treated water. Add DEPC (diethyl-pyrocarbonate) to double distilled water until saturation occurs (2 0.1-0.5%) and several drops are visible on the bottom of the tube. Shake drops vigorously and allow to stand at room temperature overnight and autoclave. Protocol i. Freeze plant material in liquid nitrogen; homogenise to a fine powder using a mortar with pestle. ii. Add 2 volumes of extraction buffer and after thawing, the powder is homogenised further in the buffer. Eventually, glassbeads can be added to improve homogenisation. iii. Transfer the extract to centrifuge tubes, and centrifuge for 10 min at 10000 rpm at 4°C. Do not allow the extract to stand too long at room temperature; especially, if the tissue is expected to contain large amounts of starch (material starts to swell; store on ice in this case). iv. Recover supernatant and transfer to fresh tubes containing I volume of phenol: chloroform: isoamyl-alcohol (12:24:1) and mix the phases by vortexing. In case the extract contains floating particles, filter through one layer of sterile muslin cloth. However, this step might be an additional risk for RNA degradation. v. Centrifuge for 15-30 min at 10000 rpm (room temperature). In case of high protein and polysaccharide contents of the tissue, an additional phenol extraction step is advisable. vi. Move supernatant to a fresh Corex tube; add 0.2 vol 1M acetic acid, and 0.7 vol ethanol (precipitation of RNA, and not of protein and DNA). Incubate overnight at –20°C, or 1 h at –70°C. vii. Precipitated RNA is pelleted at 10000 rpm for 20 min. Wash the pellet then with 70% ethanol to remove the salt, dissolve pellet in DEPC-treated water. viii. Precipitate insoluble polysaccharides. Store at –70°C. For longer storage, store as ethanol precipitate. ix. Suspend the pellet in 1 ml of ‘Q’ water or sterile dd H2O in an aliquot of 1.5 ml Eppendorf tubes. x. The total RNA is quantified by spectrophotometer.

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xi. The integrity of RNA is estimated by running the sample (1 µg RNA) on 1% (w/v) formaldehyde denaturing the agarose gel. xii. When the RNA is electrophoresed on agarose gel, it appears in different lanes; low molecular weight RNA molecules run faster than the high molecular weight molecule on the agarose gel. Result/ Note: The integrity of total RNA could be estimated by analysing 1µg RNA sample on 1% formaldehyde denaturing agarose gel (Masoodi et al., 2020). Various lanes demonstrated that the RNA has been purified and run on the agarose gel. Since it has low molecular weight, it runs faster on the gel. An absorbance of 1.0 at 260nm corresponds to approximately 40µg/ml of RNA. The A260-A320/A280-A320 ratio should be 1.8 - 2.1.

13.9.4 Protocol for Fluorescence in situ Hybridisation (FISH) Materials required: 1. Target DNA 2. A probe which binds to the target 3. Label which is attached to the probe for the detection of the target Fluorescence in situ hybridisation (FISH) on metaphase chromosomes 1. Pepsin pre-treatment of slides 2. Denaturation of chromosomes 3. Probe preparation 4. Hybridisation 5. Post Hybridisation washes 6. Counter staining the chromosomes 7. Microscopic analysis and Digital imaging Pepsin pre-treatment of slides: 1. The slides are incubated in 0.005% of pepsin in 0.01N HCl at 37ºC for 20 min 2. Washed in 1X PBS at RT for 5 min 3. Fixed for 5 min in Post fixative solution at RT 4. Washed again in 1X PBS at RT for 5 min 5. Air dried and dehydrated in 70%, 80%, 90%, 2 X 99% ethanol series at RT for 1 minute each and air dried. Denaturation of chromosomes: To allow hybridisation of the labelled probe DNA, the DNA target on the slide must be denatured to make it singlestranded. Using high temperatures in formamide is a frequently employed technique to denature chromosomes. 1. Slides are denatured in denaturation solution for 3 min and then immediately dehydrated in ice cold alcohol series (70 %, 80 %, 90 %, 2 X 99 %) for 1 minute each.

Molecular Techniques 255

2. Air dry the slides. Probe preparation: Probes are complementary sequences of nucleotide bases to the specific sequence of interest. Most commercial probes are found dissolved in hybridisation solution which includes appropriate blocking reagents. Preparation of probe mix: Probe (1 μl), Hybridisation buffer (7 μl), Distilled water (2 μl). Probe denaturation: Probe mix is denatured in a water bath at 73º C for 5 min and immediately chilled on ice for 5 min. Hybridisation: 1. 10 μl of the prepared probe is hybridised on to the slide and covered with a 22 × 22 mm cover slip and sealed with rubber gum. 2. Slides are hybridised in a humid chamber and incubated overnight at 37ºC. Post Hybridisation washes: The post hybridisation washes remove the hybridisation mixture and unbound probe. The three variables to be considered are the composition of the solutions, washing temperature and the washing time. 1. Slides are washed in Solution I for 5 minutes at 72 ºC 2. Slides are washed in Solution II for 2 minutes at RT 7.7 Counter staining the chromosomes: Counter staining reagents such as DAPI (blue) and/or Propidium iodide (PI, orange-red) are used along with the antifade solution, which prevents the fading of signals when viewed. 1. 27μl of Vectashield antifade mounting medium, DAPI or 4 μl of DAPI and 4 μl of PI are added, covered with a 22 × 50 mm cover slip and sealed with nail polish 2. Slides are stored at 4ºC till further use. Result/Note: The term ‘in situ hybridisation’ is derived from the fact that the DNA/RNA can be visualised when it is in the cell (in-situ). Several probes with different fluorescent dyes can be used simultaneously to investigate different sequences or chromosomes in plants. FISH technique has been widely used to identify the chromosomal location of human genes as well (Pierce, 2012).

13.9.5 Protocol for Oligo-FISH Preparation of plant materials: 1. Seeds were first acid-scarified by soaking them in 98% H2SO4 for 24 hrs. 2. Rinsed and sowed in pots in a greenhouse. 3. Root tips were collected from fully developed seedlings.

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4. Root tips were harvested and immediately pre-treated with 2 mM 8-hydroxyquilonine for 4 hrs in the dark at room temperature. 5. Rinsed and fixed in 4:1 (95% ethanol: glacial acetic acid) and stored overnight at RT. 6. Fixed root tips were rinsed (for 30 min) with de-ionised water (di-H2O) to remove the fixative. 7. Mildly hydrolysation (0.2 N HCl at 60 °C for 10 min and then 10 min at RT) 8. Rinsed with di-H2O followed by cold (4 °C) 0.01 M citrate buffer before enzyme digestion. 9. The enzyme digestion time varied from 24 to 35 min based on the thickness of root-tips. After chromosome preparation, slides containing good spreads were selected and used for FISH within a week or two, or stored at − 80 °C for future use. 10. Preparation of the enzyme mixture: 2% cellulase, 1% macerozyme, 2% Pectolyase, 30% cellulase, 30% pectinase and 40% 0.01 M Citrate buffer (pH 4.5). FISH with oligonucleotide probes:

• Treatment of slides with RNase-A in 2 × SSC in a water bath at 37 °C for 60 min. • Wash twice in 2 × SSC at 37 °C, 5 min each. • Dehydration through an ethanol series, 5 min each (70%, 85%, 95% and 100%) at RT and air dried overnight. • Hybridisation mixture was placed on the spread and covered with a glasscover slip without sealing. • The hybridisation mixture (25 μL/slide): • 50% deionised formamide, 10% dextran sulfate, 5.0 μg of E. coli DNA (used as blocking DNA), 25 ng of rDNA oligo probes in 2 × SSC and adjusted the volume to 25 μL with TE buffer. • Place in a pre-heated (at 80 °C) humidity chamber, and then the chromosomal DNA was denatured at 80 °C for 4 min. • The slides were cooled down for 2 to 3 mins at RT and then placed at 37 °C for incubation for 2 hrs. • After incubation, slide off the glass-coverslip using 2 × SSC. • Wash slides immediately in 2 × SSC at RT for 5 min, two washes in 0.1 × SSC at 40°C 5 min each, and then washed in 2 × SSC at RT for 5 min followed by quick rinsed in di-H2O. • Dry the slides, counter-stain with DAPI and cover with a glass-coverslip. Result/Note: Oligo-FISH technique can facilitate chromosome identification and comparative cytogenetic analysis. With the help of this technique Luo et al. (2022) distinguished the various sub-species of Hippophae rhamnoides L. (Elaeagnaceae). This taxa differentiation data is

Molecular Techniques 257

helpful in providing visual and elaborate physical mapping and guide the breeders’ utilisation of wild sources of this taxon.

13.9.6 Southern Blotting It is a molecular biological technique, used for the analysis of DNA. It can be used to detect our desired DNA from a mixture of DNA samples. In this process, DNA fragments are first separated by electrophoresis, and then transferred to a nitrocellulose filter. In the filter, the target DNA is hybridised with some ssDNA probes, which aids in their detection (Fig. 13.5). The process of Southern Blotting involves the following steps: (a) DNA extraction from the cells were performed with standard gDNA extraction and then purified. (b) Restriction digestion of the DNA sample is then performed using suitable restriction enzymes. (c) Electrophoretic separation of DNA fragments is done. (d) Depurination is done by using dilute HCl which breaks the DNAfragments into smaller fragments, thus enhancing their transfer efficiency. (e) Denaturation uses a mild alkaline solution to make the target DNA single stranded which is suitable for hybridisation. (f) The gel is put on top of a buffer saturated filter paper and nitrocellulose filter papers are put on top of the gel. On top of it, some dry filter papers are also placed. The DNA fragments are then pulled towards the nitrocellulose/nylon filter by capillary movement. (g) The nitrocellulose membrane, carrying the ssDNA bands attached to it, is baked in the oven at around 80°C for 2-3 hours to attach the DNA onto the membrane. Alternatively, UV radiation can also be used to achieve the same. (h) Hybridisation probes binds to the target DNA and helps in detection of the desired DNA segment for further studies.

Fig. 13.5. Schematic representation of Southern Blotting technique.

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(i) The membrane is thoroughly washed with a buffer to remove the excess probes. (j) An autoradiograph is used to observe the hybridised DNA molecules (for Autoradiography, refer chapter 19). Southern blotting is useful in RFLP mapping, identification of gene rearrangements and phylogenetic studies of various plants and animals.

13.9.7 Northern Blotting This technique is used to study gene expression by using mRNA transcripts. The technique of northern blotting is very similar to that of southern blot technique; with the main difference being northern blot is used to study RNA while southern blot is used for DNA analysis. The northern blot technique separates the mRNA by size, which is then transferred to a nitrocellulose/

Fig. 13.6. Schematic representation of Northern Blotting.

Molecular Techniques 259

nylon membrane, followed by hybridisation with a probe for detection (Fig. 13.6). Steps of northern blotting: (a) Extraction of RNA by using standard extraction protocols or extraction kits. (b) From the RNA extracted, mRNA is selected by using oligo dT cellulose chromatography. Probes complementary to the poly A tail is used to isolate the mRNA. (c) Agarose gels, containing formaldehyde were used to disrupt the secondary structure of RNAs, which ensures proper binding with the probes for detection. (d) The RNA separated by electrophoresis is transferred to a blotting membrane, and immobilised by using UV light or heat. (e) Probes (~25 bases long), complementary to the RNA of interest is allowed to hybridise with the RNA. The excess probes are washed off with buffer. (f) Autoradiography or chemiluminescence was then used to detect the RNA bound with the probe. While the Southern blotting cannot blot RNA due to its inability to bind with the nitrocellulose filter, the Northern blotting is useful to detect and analyse RNA in a biological sample.

13.9.8 Western Blotting Western blotting is also an analytical technique which is used to target a specific protein for study, from a mixture of proteins. The mixture of proteins is first separated by gel electrophoresis, on the basis of molecular weight. Similar to the other blotting techniques, the proteins are transferred to a membrane, which is then labelled with specific antibodies (Fig. 13.7). The following steps are followed to carry out western blot (Mahmood and Yang, 2012): (a) Cells are lysed to extract the proteins, using phosphate buffer saline (PBS) and cell lysis buffer. (b) The extracted protein sample is then prepared for electrophoretic separation. Both stacking and separation gels are prepared, and electrophoresis was performed. (c) 6 filter sheets and sponge, soaked with transfer buffer and one polyvinylidene fluoride (PVDF) (moistened with methanol) are used to make a sandwich with the gel. Sponge

3 filter papers

Gel

PVDF

3 filter papers

(d) The sandwich is covered with buffer and the electrophoresis is done at 4⁰C.

260

Plant Techniques: Theory and Practice

(e) Proteins are moved onto a solid support from the gel by electroblotting method. In this method, electric field is used to pull out the proteins from the gel. (f) Non-specific binding of antibodies to the membrane is prevented through blocking. BSA and non-fat dry milk are commonly used blockers. (g) The primary antibody, thereafter, binds to the target protein, and the membrane is repeatedly washed with antibody-buffer solution to remove the excess and unbound antibodies. (h) The membrane is subsequently exposed to a specific, labelled enzyme conjugated secondary antibody, which binds with the primary antibody. (i) A substrate reacts with the enzyme that is bound to the secondary antibody to generate coloured substance.

Fig. 13.7. Schematic representation of Western Blotting.

Western blotting has been employed to detect low-quantity proteins and, to ensure and quantify a gene product as well.

14 Plant Tissue Culture Plant tissue culture is one of the most important techniques used for growing plant cells under sterile conditions. The suitable medium composition is a specific factor for the growth and morphogenesis of plant species. The medium consists of water, solid or semi-solid support, macronutrients, micronutrients, vitamins, carbon sources, plant growth regulators or hormones, amino acids, sorbents, and other undefined organic supplements. The advantages are used for rapid clonal propagation, plant breeding, plant research, production of useful substances, pathogen-free plants, longterm conservation, and international exchanges. The potential is there to produce a significantly greater number of healthier plants in less space, with less labour, and at less cost than by other means of vegetative propagation (Kyte and Kleyn, 2001).

14.1 History Gottlieb Haberlandt (1902) did experiments on the culture of single cells,but they were not successful. He is recognised as the father of plant tissue culture. Kotte (1922) succeeded in culturing isolated root tips. Success was also achieved with bud cultures by Loo (1945) and Ball (1946). The first true plant tissue cultures were obtained by Gautgeret (1934 and 1935) from the cambial tissue of Acer pseudoplatanus. The 1940s, 1950s, and 1960s proved to be exciting times for the development of new techniques. During the 1990s, continued expansion in the application of in vitro technologies to an increasing number of plant species has been observed. Tissue culture techniques are being used with all types of plants, including cereals, grasses, etc. Plant tissue culture helps in applied plant biotechnology, as well as stimulating fundamental scientific progress.

14.2 Setup a Tissue Culture Laboratory A plant tissue culture laboratory has several functional areas, whether it is designed for teaching, research, or business and no matter what its size or how elaborate it is. Avoid locating it adjacent to laboratories that handle microorganisms or insects or facilities that are used to store seeds or other

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plant materials. Contamination from air vents and high foot traffic can be a problem. Foot traffic scuffs up the wax on floors, as well as dust which help to spread contaminants (Smith, 2000). The tissue culture area should always be kept clean. In designing a plant tissue culture laboratory, three major rooms, which are a preparation room, a transfer room, and a culture room, are divided. In a small laboratory, it may be divided into 2 parts which may combine a preparation room and a transfer room, or a transfer room and a culture room.

14.2.1 Preparation Room A media preparation room should have smooth walls and floors which are easy to clean to maintain a high degree of cleanliness, as well as enough space to prepare the media on the bench with pH meters, balances, and a sink in proximity. The reagents and stock solutions should be located alphabetically either on shelves or in a refrigerator adjacent to the bench. The equipment used in these rooms is: 1. 2. 3. 4. 5. 6. 7.

A balance A hotplate or a microwave A magnetic stirrer A pH meter An autoclave A hot air oven A refrigerator

14.2.2 Transfer Room A transfer room needs to be as clean as possible with good ventilation. It should be isolated as much as possible from outside doors to prevent contamination and minimise the opening of doors. The walls, floors, and ceilings with lighted colours should be smooth to ensure frequent cleaning with detergent and standard disinfectants. The illumination of the laminar airflow cabinet is sufficient for work. Sterilisation of the instruments can be done with glass-bead sterilisers or flaming after dipping in 90% ethyl alcohol. The culture containers should be stacked on mobile carts (trolleys) to facilitate easy movement from the medium storage room to the transfer room, and finally to the c u l t u r e room. The chair seats of the transfer operators should be comfortable, as they must work for long periods in the same place. Fire extinguishers and first aid kits should be provided in the transfer room as a safety measure. The personnel should leave shoes outside the room. Special laboratory shoes and coats should be worn in this area. Ultraviolet (UV) lights are sometimes installed in transfer areas to disinfect the room; these lights should be used only when people and plant material are not in the room. Safety switches can be installed to turn off the UV lights when regular room lights are turned on. Many workers find it more comfortable and relieve eye

Plant Tissue Culture 263

strain to see out through the windows. The transfer room contains: 1. 2. 3. 4. 5.

A laminar air-flow cabinet with comfortable chairs A dissecting microscope Gas outlets Vacuum lines Forceps, spatulas, scalpel and disposable blades

14.2.3 Culture Room Controlled temperature, lighting, relative humidity, and shelving need to be considered in planning the culture room. The culture room should have suitable lights for different growth stages and temperature (20-28 oC). Air quality should be clean without dust. High humidity in the culture room should be avoided because it increases contamination. Some culture rooms have dehumidifier and air scrubbers. There is no need for windows except when natural light is used. Overhead light sources can be minimised because the primary source of illumination is from the lights mounted on the shelves. Culture room contains: 1. Shelves with lighting on a timer and controlled temperature. 2. Incubators with controlled temperature and light. 3. Shakers for liquid medium.

14.3 Media Components and Preparation The media is composed of inorganic compounds, plant growth regulators, vitamins, amino acids and complex organic supplements, carbohydrates, water, gelling agents, and activated charcoal. Table 1 shows components of commonly used tissue culture media. Inorganic compounds are composed of macroelements (N, P, K, Ca, Mg, and S) of which all plants require in relatively large amounts and microelements (Fe, Mn, Zn, B, Cu, Mo, and Cl) in relatively small amounts. Auxins and cytokinins are two main groups of plant growth regulators that are often used in several plants (Wu et al., 2004) and are required for stages of plant tissue culture. Auxins are used for inducing plant root, stimulating shoot cell elongation and callus formation or enhance the cytokinin effects. Cytokinins are popularly used for shoot proliferation, promoting cell division, and regulating growth and development. The presence of auxins in the medium is generally essential for embryo initiation (Steward et al., 1958). Auxins play two main roles in plant tissue culture. Firstly, they can be used to induce root formation, such as indole acetic acid (IAA) and indole butyric acid (IBA). Secondly, they can initiate callus formation, such as 2,4-dichlorophenoxy acetic acid (2,4-D) and NAA. and thidiazuron (TDZ) (Chen et al., 2004). TDZ is a substituted phenyl urea with cytokinin-like activity. It is generally

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believed that TDZ is more active in stimulating shoot formation than BAP or kinetin, even at extremely low concentration (Huetteman and Preece, 1993). Disadvantage of TDZ in regeneration is the difficulty in elongation and rooting of the regenerated shoots. This problem was overcome by transferring regenerated shoots to MS medium supplemented with BAP and NAA. Vitamins have catalytic functions in enzyme systems and are required in trace amounts. Thiamine (vitamin B1) is essential for all plant tissue cultures, whereas, nicotic acid (niacin) and pyridoxine (vitamin B6) may stimulate late growth (Gamborg et al., 1976). Amino acids (mainly glycine) and complex organic supplements, such as coconut water, peptone, yeast extract, malt extract, fruit juices, and fruit pulps are used because of cheap price and less labour work, as well as other combination of known components supports growth of tissues. Carbohydrates are energy and carbon source which are added into all media because most cultures are not completely autotrophic. Carbohydrates are very important component in in vitro cultures because they have energy and carbon source, as well as an osmotic agent. They were added in any nutrient media that are essential for in vitro growth and development because photosynthesis is insufficient in in vitro culture (Pierik, 1987). Sucrose (20-40 g/l) are commonly used in plant tissue culture in addition to glucose, fructose, and sorbitol under some conditions. Water is employed in all media, including the water used during the culture procedure. 95% of nutrient medium consists of water. Distilled water is needed for research; otherwise, purified water by reverse osmosis, deionisation, and filtration are appropriate. Gelling agents are mostly used for stationary supports. Agar, agarose, filter paper, cotton, cheesecloth, vermiculite, etc. are used as gelling agents depending on the purpose and suitability of the culture. Agar is a natural product which is obtained from various types of seaweeds. Agarose is a highly purified line air galactan hydrocolloid isolated from Gelidium species of seaweed. Activated charcoal (AC) is composed of carbon that has numerous pores on the surface. AC was used as a culture component for adsorption of toxic plant metabolites, such as phenolic compound and plant growth regulators (Thomas, 2008). AC provided a dark environment during in vitro culture that affected shoot and root production. AC could effectively stimulate formation of somatic embryos on ½ MS medium (Nainget al., 2011). Graphite and AC have carbon structures, odourless, and non-toxic substances; therefore, they were used to darken the culture medium of plant tissue culture.

14.4 Explant Preparation Explant is a portion of a plant (tissue, organ, a few cells, or part of a callus mass) taken for culture in vitro. Strong and healthy, young age, season (Spring

Table 14.1. Components of commonly used tissue culture media (mg/L) Components

Gamborg et al., 1968

White, 1963

Lloyd and McCown, 1981

Vacin and Went, 1949

Modified Knudson, 1946

Mitra et al., Nitsch and 1976 Nitsch, 1969

525

180

180

250 250 500

250 150 100

250 150 100

200

200

80 6.2 0.075

0.03 0.6

200 1650 1900 440 370 170

400 2500 150 250 134 150

80 720

16.50 300 200 65

96 370 170

556

720 950 166 185 68

990 0.83 6.2 22.3

0.75 3 10

0.75 1.5 7

6.2 0.75

10 25

29.43 (Contd.)

Plant Tissue Culture 265

Macronutrients Ca(PO4)2 NH4NO3 KNO3 CaCl22H2O MgSO47H2O KH2PO4 (NH4)2SO4 NaH2PO4H2O Ca(NO3)24H2O Na2SO4 KCl K2SO4 Micronutrients KI H3BO3 MnSO44H2O MnSO4H2O

Murashige and Skoog, 1962

Murashige and Skoog, 1962

8.6 ZnSO47H2O 0.25 Na2Mo042H2O 0.025 CuSO45H2O 0.025 CoCl26H2O Co(NO3)26H2O 37.3 Na2EDTA 27.8 FeSO47H2O MnCl2 FeCC4H4O6)32H2O Vitamins and other supplements Inositol 100 Glycine 2 ThiamineHCl 0.1 PyridoxineHCl 0.5 Nicotinicacid 0.5 Ca-panthothenate CysteineHCl Riboflavin Biotin Folic acid

Gamborg et al., 1968

White, 1963

Lloyd and McCown, 1981

2 0.25 0.025 0.025

2.6

8.6 0.25 0.25

37.3 27.8

Vacin and Went, 1949

Modified Knudson, 1946 0.25 0.025 0.025

37.3 27.8

74.6 25 3.9

Mitra et al., Nitsch and 1976 Nitsch, 1969 0.05 0.05 0.05

10 0.25 0.025

0.05 37.3 27.8 0.4

37.3 27.8

28 100 2 10

3 0.1 0.1 0.5 1 1

100 2 1 0.5 0.5

0.3 0.3

0.3 0.3 1.25

100 2 0.5 0.5 5

0.3

0.05 0.05 0.3

0.05 0.5

Plant Techniques: Theory and Practice

Components

266

Table 14.1. (Contd.)

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is appropriate), size (larger is better for more nutrients and plant growth regulators) plants should be chosen without viruses, bacteria, fungi, mites, etc.) After detaching explants from the source plant, they must be cleaned. There is no rule for cleaning explants depending on how clean or dirty the explants are to begin with. The most common cleaning agent used to disinfect explants is bleach in various concentrations (pure household bleach is 5% sodium hypochlorite) commonly used concentrations are 5%, 10%, and 20% concentrations. Others are 70% ethyl alcohol, 3% hydrogen peroxide. The cleaning should be done in the media preparation room. Transport the cleaned explants to the transfer hood, still in bleach in covered containers, and apply the final rinses in the hood using sterile techniques.

14.5 Aseptic Technique A sterile environment during the culture of plant tissues must be maintained. Air currents must be avoided over the sterile area because they carry spores of contaminating microorganisms. Laminar air-flow cabinets allow a gentle flow of ultra filtered sterile air to pass across the working area; thus, preventing airborne contamination. Before the start of any sterile procedures, the working area should be thoroughly scrubbed with a tissue soaked with ethyl alcohol (70% v/v). Several techniques are employed for the sterilisation of glassware, surgical instruments, liquids, and plant materials. The goal in surface sterilisation of plant tissue is to remove all microorganism with a minimum of damage to the plant system to be cultured. Methods are dry heat, wet heat, ultrafiltration, and chemical sterilisation. Dry heat is used only for glassware, metal instruments, or other materials that are not charred by high temperatures. Cotton, paper, or plastic cannot be used. Using 160 oC (320 oF) for 4 hours is sufficient. Wet heat is used for liquids, a steam pressure of 15 lb/in2 (103 kPa) from an autoclave with vapour under pressure at 121 oC (250 oF) for 15 minutes. Do not close the escape valve until a steady stream. At the end of 20-min period, the pressure must be allowed to return slowly to the atmospheric level because rapid decompression will cause the liquids to boil out of the containers. Prolong autoclaving must be avoided because of the results in the decomposition of the chemicals present in the medium. Ultrafiltration is used for some medium components which are heat-labile and must be sterilised at room temperature, then add to cooling agar medium and still in the solution state, such as Sintered-glass filters, the Nuclepore filter, etc. Chemical sterilization is used, such as 70% ethyl alcohol for surfacesterilized, 0.5% NaOCl (sodium hypochlorite) for sterilising solution. Cleanliness is of primary importance in the transfer room. The following practices should always be observed to maintain safe and clean conditions in the laboratory:

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1. No eating, drinking, smoking, or storing food in the laboratory. 2. Wear a protective lab coat or apron when working with cultures. Long, full sleeves should be avoided because they collect dust. 3. Wear sterilises disposable medical gloves or plastic garden gloves. 4. Tie-back and cover long hair. Loose hair is a potential contaminant for cultures and risks catching fire from any open flames. 5. Sit straight and keep your head out of the laminar air-flow cabinet. 6. Set only necessary equipment in the laminar air-flow cabinet. 7. Sterilise in an autoclave or pressure cooker any jars and petri dishes containing contaminated cultures before emptying. 8. Disinfect work surfaces before and after working in the laminar air-flow cabinet. 9. Keep transfer room doors closed to avoid unnecessary drafts.

14.5.1 Sterile Technique for Explants Procedure 1. Before a start to work, turn on the ultraviolet lights for 15 minutes. 2. When to prepare the explants or any works; turn on the laminar air-flow cabinet for 10 minutes before use. 3. Wipe both hands and all things including plant material with a cloth soaked with 70% ethyl alcohol, and then put them inside the cabinet. 4. Wipe or spray inside the cabinet with 70% ethyl alcohol. Do not spray the HEPA filter. 5. Put knifes, forceps, and other equipments in a glass bead steriliser or a container (if use alcohol lamp or gas for sterilisation). 6. Flame the petri dish and open the lid. 7. Put plant material on the petri dish. 8. Soak the knife and forceps in 90% ethyl alcohol and flame (if use alcohol lamp or gas) 9. Cut plant material to be explants. 10. Flame the prepared medium container lid and open. 11. Place the explant on the medium. 12. Flame the lid and the edge of the medium container, then close the lid. 13. After finishing, take containers out and then wipe inside cabinet and close the cabinet. Sterile technique for transfer Procedure 1.-5. are the same as sterile technique for explants. 6. Flame both a medium container with explant and a new medium container and open both lids. 7. Place the explant on the new medium.

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8. Flame the lid and the edge of the new medium container, then close the lid. 9. After finishing, take containers out and then wipe inside cabinet and close the cabinet.

14.5.2 Contamination One of the most important factors in preventing contamination of the cultures is the removal of airborne contaminants in the sterile area. Always keep all windows and doors closed to avoid drafts. A minimum number of persons should be presented. There are 4 sources of infections: 1. The plant (internal as well as external), 2. The medium should be sufficiently sterilised, 3. The air, and 4. The person who works (inaccurate work). The most important is the plant, well sterilised before culture in vitro. Contamination in tissue culture operations can be costly, particularly if the contamination rate is greater than 1%. If the contaminations are observed in test tubes or bottles, the containers should not be opened and should be sterilised before discarding the contents.

14.6 Pathways of Culture Cells and Tissues Roots, shoots, and flowers are the organs that may arise de novo from plant tissue culture (Organogenesis). Embryos are not classified as organs because these structures have an independent existence. Skoog and Miller (1957) observed that relatively high auxin: cytokinin ratio induced root formation in tobacco callus,whereas, a low ratio favoured shoot production. Somatic embryogenesis is the producing embryos not from the development of the fertilised egg. Anther and pollen culture is to produce haploid plants by the induction of embryogenesis. The in vitro growth and development of a plant is determined by a number of complex factors which are: the genetic make-up of the plant, nutrients, physical growth factors (light, temperature, pH, O2, and CO2) and some organic substrates (plant growth regulators, vitamins, amino acids, coconut water, banana, etc.). There are four stages of plant tissue culture growth: 1. 2. 3. 4.

Stage I: Establishment-Prepare the explants and start culture Stage II: Multiplication-Proliferation of shoots Stage III: Rooting-Root initiation Stage IV: Acclimatisation or transferring to the greenhouse - Lower relative humidity and increase light intensity and temperature

It is not necessary that each stage needs different culture medium. A single medium may be satisfactory for the first three stages. Stage III can be skipped since rooting may stop multiplication. Instead, rooting will occur at Stage IV.

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14.7 Acclimatisation Acclimatisation is defined as the climatic or environmental adaptation of an organism, especially a plant that has been moved to a new environment (Conover and Poole, 1984). Acclimatization is required because there is a significant difference between the tissue culture or micropropagation environment and the greenhouse or field environment. In the laboratory, the cultures have low light intensity; low temperature, 90-100% relative humidity, and cuticle poorly develop. Good techniques for acclimatising tissue cultured plantlets should be applied. Acclimatisation can be started, while plantlets are still in vitro by providing higher light intensity and reducing relative humidity by loosening the lid (closure) for 1-2 weeks before moving the plantlets to the greenhouse. An abrupt change to the lower humidity and greater light in the greenhouse can be fatal to plantlets in a very short time (Kyte and Kleyn, 2001). In the greenhouse, plantlets should be covered with plastic, be watered very often, and gradually increase light intensity for a few weeks before growing normally in the greenhouse.

14.8 Commercial Production Successful micropropagation on a commercial scale for producing many quality plants largely depends on high multiplication from suitable protocols and successful acclimatisation. It is not enough to produce high quality, healthy plants; one must also produce it in a timely fashion, giving customers what they want, when they want it, for a price they can afford. It is desirable to predict the financial success of a commercial laboratory, record it in progress, and periodically evaluate it in retrospect. Good management involves constant planning, study, observation, review, and adjustment. Good marketing practices are essential for success (Kyte and Kleyn, 2001).

14.9 Culture Guide to Selected Plants The choice of medium depends mainly on plant species, the tissue, or organ to be cultured, and the purpose. It is best to start with published protocols. A starting point for tissue culture from a dicot tissue explant would be the preparation of the Murashige and Skoog, 1962 medium (MS medium) at different concentrations. MS medium is the most useful tissue culture medium and many variations have been developed. The medium is derived from White’s medium and originally developed for the cultivation of Nicotiana tabacum Calli. Compared to the White medium, the concentration of all components is increased. Due to the high concentration of minerals, MS medium is a very rich and saline medium and can be too salty to certain plant

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species. To avoid this problem, MS medium is often used with the micro elements in full concentration, but with the macro elements in respectively half or three quarters of the concentration as originally described. Monocot cultures respond to the Schenk and Hildebrandt, 1972 medium (SH medium). The two examples are micropropagation of Grammato phylum speciosum Blume, a giant orchid species (Sopalunet al., 2010) and Vanda coerulea Griff. ex Lindl., a beautiful blue vanda orchid with tessellation on petals and sepals (Wasiksiriet al., 2010) are given in section 14.14.2 and 14.14.3.

14.10 Meristem Cultures and Production of Disease Free Plants Generally apical meristems such as shoot apex, root apex and axillary buds, are free from pathogens or carry a low concentrations of viruses, bacteria or fungi. These meristems are taken as explant and raised the plantlet. Since the meristems are free from the pathogens, plantlets derived from them are also free from pathogens. Various reasons have been attributed to the escape of the meristems by virus/bacterial presence; these are: (1) actively dividing meristematic cells do not allow the virus replication which are unable to cope up with the rapid cell divisions of meristems, (2) Absence of vascular elements and plasmodesmata are possible causes of low virus concentration in meristems (3) endogenous auxin supply is high. Morel and Martin (1952) exploited the meristem culture techniques to eliminate the viral infection in plants. They were able to develop pathogenfree Dahlias by using the meristems cultures. It is also possible to eliminate other pathogens such as viroids, mycoplasma, etc. (Pierik, 1989; Kane, 2000).

14.10.1 Collection of Meristem Tips Meristems should be collected from growing buds, such as stem tips, root tips, leaf axile or from germinated seeds. Size of the explant: It is better to have the explant size as minimum as possible; may be up to 0.4 mm. Because the size of the meristems is critical for virus elimination since a gradient distribution of pathogen established in plant tissues. It has been generally observed that virus-free plants regenerated in vitro are inversely proportional to the size of meristems cultured. About 60% cassava (Manihot esculenta) plantlets regenerated from 0.4 mm meristems, collected from sprouted cuttings of infected plants happened to be diseasefree ones (Razdan, 2005). In contrast, plants regenerated from large size meristems were indexed as diseased (Kartha and Gamborg, 1975). Physiological conditions of explant: Meristem tips should be taken from actively growing buds. The percentage of virus-free plants can also depend on the season, especially trees which display periodic growth. In the temperate trees, the growth is limited to only a short period in spring, after that shoot

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apex remains dormant for quite a long period. In such cases, meristems tips should be collected and cultured during spring only. Thermotherapy: The donor plant (mother plant) from which meristem tips are collected should be given heat treatment. A careful decision should be taken for suitable temperature and duration of heat treatment. Heat treatment may be given for 5-10 weeks. It is performed through hot water or hot air. The hot-water treatment effectively eliminates virus in dormant buds, whereas hot-air treatment is recommended for elimination of virus from actively growing shoots. For hot-air treatment, actively growing plants are placed in a thermotherapy chamber adjusted at a temperature at 35-40oC for a period of a few minutes to several months. Heat treatment can kill the bacteria, virus, fungi, etc. Chemical treatment: Mother plant treated with antibiotics like actinomycin-D and cyclohexamide, can inhibit the virus replication. Further plant treated with nucleoside analogue ribavirin (syn. Virizole) was given positive effect and eliminating Potato virus X (PVX) in potato plantlets (Shepard, 1977), CMV from cucumber and alfalfa mosaic virus (Pierik, 1989),

14.10.2 Culture Medium Generally modified MS medium has been used for the meristem tip cultures. Medium (2-5% sucrose) supplemented with low concentrations of auxins and cytokinins can stimulate cell divisions. In case of Cassava (Manihot esculent) and Dahlia, GA3 can suppress the callusing and favour better growth and differentiation. Medium further supplemented with vitamins such as pyridoxin, pantothenic acid and nicotine acid, could enhance the growth. Cultures are incubated under room temperature at 21-25oC. Bulbous plant cultures can be grown under fluorescent light.

14.10.3 Virus Indexing Screening of plants for the presence or absence of virus is known as virus indexing. It is a very important step in the production of virus-free plants. Many viruses have delayed resurgence periods in cultured plants (Razdan, 2005). This necessitates performing the virus indexing periodically. Various strategies have been adopted for virus indexing. These are as follows: i. EM observations: Latent virus that do not exhibit any visible symptoms, have to be observed under Electronic Microscope. Immunosorbent Electron Microscopy (ISEM) method described by Barbara and Clark (1986) could be adopted, that combines both serology and EM observations for the detection of virus. ii. Sap transmission test: It is easily performed on commercial basis. Leaf sap is prepared from test plant (plant to be assayed for the presence and absence of virus) by grinding the leaves with equal volume of buffer

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solution (0.1 mol l-1 sodium phosphate) using a mortar and pestle. This filter-leaf sap is smeared on to the indicator plant which is predusted with 600 grade carborendum powder. If any virus is present in leaf sap, the indicator plant developed some characteristic symptoms. Some of the plant species used as indicator plant are: Chenopodium amaranticolour, V. Quino, Gomphrena glubosa, Nicotin tabacum, etc. (Klopmeyer, 2000). iii. Serological test: Among serological methods, ELISA (enzyme-linked Immunosorbent assay) is very popular to detect virus particles (Miller and Martin, 1988). It is more sensitive than bioassays, but it needs ELISA reader and enzymes. In ELISA test, antiserum against virus is collected from sheep or rabbit and purified. The primary antibody i.e., antiserum coated on to the microtitre (96-celled polystyrene plates) of ELISA reader to which add macerated plant extract of any test plant (suspected virus-infected plant) and incubate for some time. During incubation, if any virus is there, it will bind to the antiserum. After incubation, the microtitre well is thoroughly rinsed, to wash away the unbound particles. In next step, enzyme conjugated secondary antibody linked with alkaline phosphates is added to the well. During second incubation period, conjugated secondary body binds to any virus particles present in the well that was previously bound by the coating antiserum. So, plant extract (antigen) is sandwiched with antibodies (primary as well as secondary antibody; hence, it is popularly known as Sandwich ELISA test (Fig. 14.1). It is verified by adding an enzyme substrate and intensity of colour development in each well can signify the presence and relative concentration of plant virus.

Fig. 14.1. Sandwich ELISA test.

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Micrografting Initiating meristem cultures of some species, particularly woody perennial fruit crops, are very difficult. The shoot regenerated from meristem cultures of these plants quite often does not form roots. In such case, micrografting is undertaken. In the very beginning Morel and Martin (1952) micrografted the Dahlia meristem on to a virus-free stock (seedling) maintained and propagated in vitro. Micrografting had proved to be very useful for elimination of virus in fruit crops, especially apricot, wine grape, Eucalyptus, peach, etc.

14.10.4 Protocols 14.10.4.1 Virus Eradication in Potato Plants (Mellor and Stace-Smith, 1977, Razdan, 2005)

Reagent and other Equipment MS medium; growth regulators such as IAA/IBA, BAP/Kinetin. Equipment Growth chamber, binocular microscope. Plant material Potato tubers, virus-free stock (seedling) indicator plants such as Chenopodium amaranticolour and Gomphrena globosa. Procedure i. Collect single-eyed tuber peace and plant it in soil under normal environmental conditions. As soon as the first sprout is 15 cm tall, take a tip-cutting of 6-8 cm long. Remove the lower leaves and apply rooting hormone to the cut surface and plant the cutting in a 10 cm peat pot of sterilised soil. Cover it with glass beaker for 10 days. ii. After 3-4 weeks transfer the entire set-up to a growth chamber where the light is 3000-4000 lx for 16 hr a day and the air temperature about 36oC during the day and about 33oC at night. iii. Two weeks later cut the tip of the young plant to promote the growth of axillary shoots. iv. Give thermotherapy for 6 weeks and then collect one axillary shoot for bud excision, leaving at least two leaves on the plant to encourage further growth. v. From the excised shoot remove the lamina of leaves in such a way that the small basal portions of the petiole are left on the stem. Enfold the shoot in wet paper to prevent wilting. vi. Excise the meristem tip under binocular microscope. It is advisable to cut-off the meristems tip when the two youngest leaf primerdia are still left using a fine blade fixed in a suitable holder. vii. Transfer the excised meristem to suitable culture medium. viii. Store the cultures at about 23oC in a 16-hr light regime.

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ix. Transfer the regenerated rooted shoots to the soil. Maintain the plants under high humidity initially. In case, where shoots fail to develop root, their grafting may be performed. x. Perform the virus indexing before transplanting the plants in soil at regular intervals, Indicator plants to be used here are C. amaranti colour and G. globosa. 14.10.4.2 Virus-eradication in Ornamental Plants (Chawla, 2003)

Reagents and other equipment MS medium with growth regulators such as BAP, IAA/NAA, KIN, gibberllic acid, sterile distilled water. Dissecting stereo microscope and other paraphernalia required for plant tissue culture experiments. Plant material Chrysanthemum, Gerbera Procedure i. Collect seeds from the plant and raise the seedlings from seeds. ii. Maintain the sterile condition in the inoculation chamber by using the 70% alcohol and flame sterilisation. iii. Collect shoot apices from in vitro derived plantlets with sharp razor blade. iv. Remove the outer whorl of leaves of shoot apex with sharp blade. v. A piece of meristematic tissue (explant) is excised by making V-shaped cut just 0.3 to 0.5 mm below the tip of the shoot apex. The explant should have small portion of procambial tissue and it is immediately transplanted on MS agar medium culture tube. vi. MS medium is commonly used for this purpose. Medium is supplemented with auxins and cytokinins as they are useful for cellular multiplication. Gibberellins are necessary to ensure the elongation of the axis. Kinetin or BAP of 5×10-6, 5×10-7 M levels can be used, similar levels of NAA, IAA or IBA are employed here. GA3 should be at a level ranging from 10-7 to 10-8 M. vii. The culture tubes are kept in culture room maintaining under 16/8 h photoperiod at 3000 lx, 25±2oC. viii. When plantlets are formed after 4-6 weeks, take them out from the tubes, wash plantlets in running tap water and transfer them into pot conditions. ix. Keep the regenerants in small plastic pots with a sterile soil mix comprising sand: farmyard manure, soil in 1:1:1 proportion; see that soil is moist with water. Plants do not need to be watered for the first few days. x. Keep the plants in diffused light. Remove inverted beaker or plastic wrap so as to allow free exchange of air for 1 hr each day. After another week, increase gradually the period of exposure several hours per day. After 2-3

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weeks, remove the wrap and inverted beaker completely from the plants in order to adjust with natural environmental conditions. xi. Sap transmission test is performed on these regenerated plants and make sure that all are disease free plants.

14.11 Protoplast Fusion and Somatic Hybridisation In plant breeding programmes many desirable combinations of characters may not be transmitted through conventional methods of genetic manipulations. Alternate to this, Cocking (1979) has developed the technique, other than the sexual cycle, to get genetic recombinations. In this technique, fusion of two isolated somatic protoplasts is involved and this fusion product known as heterokaryon is subsequently developed into somatic hybrid. This phenomenon is known as somatic hybridisation (Fig. 14.2).

Fig. 14.2. Somatic hybridisation.

Through the fusion process, the nucleus and cytoplasm of both parents are mixed together and forms the heterokaryon. This results into the formation of various nucleo-cytoplasmic combinations of characters. Sometimes interaction between plastome (cytoplast carrying cytoplasmic characters) and nuclear genome contributes to the formation of cybrids (cytoplasmic hybrids). Cybrids, in contrast to conventional hybrids, possess a nuclear genome from only one parent but cytoplasmic genes from both parents. The process of protoplast fusion resulting in the development of ‘cybrids’ is known as ‘Cybridisation’ (Fig. 14.3). So, protoplast fusion is an important tool for the induction of genetic variability and combination of traits.

Fig. 14.3. Cytoplasmic hybridisation.

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The important steps involved in protoplast fusion are: (1) Isolation of protoplasts (2) Culture of protoplasts, (3) Protoplast fusion (4) Regeneration of somatic hybrids and cybrids.

14.11.1 Isolation and Purification of Protoplasts Sources of protoplasts (A) Leaves - The leaf is the best source of protoplasts because it allows isolation of a large number of relatively uniform cells. The following steps are involved in the isolation of leaf protoplasts: (a) sterilisation of leaves, (b) peel-off the epidermis, (c) incubation in the enzyme mixture, (d) isolation by filtration and centrifugation. (B) Callus cultures - Young callus cultures or actively growing callus cultures are also ideal materials for obtaining large quantities of protoplasts. Young callus is subcultured and it can be used after two weeks for protoplast isolation. (C) Cell suspension cultures - The cell suspension cultures are also excellent source materials for isolation of protoplasts. A high-density cell suspension is centrifuged. After removing the supernatant, cells are incubated in an enzyme mixture (cellulose + pectinase) in culture flask and liberated protoplasts are purified subsequently. Methods of isolation of protoplasts (A) Mechanical (non-enzymatic) method - Through this method cell walls are removed mechanically with sharp-edged knife. First, cells are kept in hypertonic solution for a few min. This brings about the plasmolysis in plant cells, resulting in the separation of protoplasts from cell walls. Next the cells walls are broken with the help of knife to release the protoplast from the cells. The isolated protoplasts are transferred to isotonic solution to prevent further damage. This technique has been employed for the isolation of protoplasts from highly vacuolated cells of storage tissues like onion bulbs, radish root, beetroot, etc. (B) Sequential enzymatic (two-step) - Take be et al. (1968) had developed the technique of isolation of protoplasts by using commercially available enzymes in two steps. In the first step, leaf segments are treated with pectinase (enzyme mixture A) and pectin is digested. In the second step, this tissue is treated with cellulase (enzyme mixture B) and cellulose is digested. In this way, whole cell wall is digested, and protoplasts are liberated. I step - enzyme mixture ‘A’ 0.5% pectinase (macerozyme + 0.3% potassium dextran sulphate in 13% mannitol at pH 5.8). II step - enzyme mixture ‘B’ - 2% cellulase in 13% mannitol at pH 5.4.

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(C) Mixed enzymatic (simultaneous/single step) method - In this method, leaf segments are treated with a mixture consisting of both macerozyme and cellulose enzymes. This can be completed in a single step. Enzyme mixture consists of both 0.5% macerozyme and 2% cellulase taken in 13% mannitol at pH 5.4. Leaf material is incubated in this mixture for 15-18 hr at 25°C. Protoplasts are isolated in a single step through this method. Isolation and purification of protoplast by mixed enzymatic method The protoplasts isolated as above may contain together with broken organelles and cell debris in the medium. So, purification of protoplasts is essential before proceeding for the protoplast fusion (Fig. 14.4).

Fig. 14.4. Flow chart of isolation and purification of protoplast by mixed enzymatic method.

Viability of Protoplasts Isolated protoplasts should be healthy and viable in order to undergo sustained division and regeneration. Protoplast viability could be tested by variety of methods including: (A) Presence of cytoplasmic streaming or cyclosis, (B) Presence of photosynthetic and respiratory activity. These tests indicate active metabolism in protoplasts. Besides these, protoplast viability is tested by using several dyes: (A) staining with fluorescein aseince (FDA), protoplasts fluoresce green/

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white; it should be examined within 5-15 min. after the FDA dissociates; (B) phenosafranin (0.1%) detects dead protoplasts, turn red whereas viable protoplasts remain unstained even after 2 hr in the stain solution. (C) Calcofluor white (CFW 0.1%) detects onset of cell wall regeneration around plasma membrane of viable protoplasts appears in the form of a ring of aseincence.

14.11.2 Culture of Protoplasts Since the isolated protoplasts are devoid of wall, these are very easy to undergo fusion in vitro. Protoplast fusion is employed to overcome the incompatibility barriers operating at interspecific and intergeneric levels. It is an important tool in somatic cell genetics and crop improvement programme. The first step in the protoplast culture is the development of cell wall around the plasma membrane of isolated protoplast. This is followed by induction of divisions in the protoplast-derived new cell giving rise to a small colony. By providing proper, nutritional requirement, cell colonies may be induced to grow callus continuously or to regenerate whole plant. Culture Media Generally, protoplast culture media contain 3-5% sucrose but in some systems (tobacco protoplast cultures) 1.5% sucrose content is required. Organic nitrogen in the form of CH (asein hydrolysate) and inorganic nitrogen NH4NO3 are supplementing the medium. Higher concentrations of iron, zinc and ammonia are used in these media. Various combinations of auxins (2, 4D, NAA, IAA) and cytokinins (Kinetin, zeatin, BAP) are used to induce cell wall formation and divisions in protoplasts. It has been observed that high auxin/ kinetin ratio is suitable and induce divisions in the protoplasts from actively growing cell cultures whereas high kinetin/auxin is for protoplasts derived from differentiated cells (leaf tissue, stem tissue, etc.). Protoplast Culture Techniques Various techniques have been employed for culturing of protoplasts. These are as follows: A. Bergman’s Cell Plating Technique For culturing of protoplasts in the solid agar medium, this technique may be followed. About 2 ml aliquots of isolated protoplasts of suitable density (103-105 cell ml-1) are mixed with equal volume of agar nutrient medium (temperature not to exceed 45° C). On solidification of agar, the culture plates are sealed and maintained in an inverted position and incubated in diffused light or in dark at 25°C. Here cell colonies are formed, and these can be conveniently observed under microscope. Plating efficiency is readily determined by the following formula: Plating efficiency =

Final no. of cell colonies Initial no. of cell units

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B. Culturing the Protoplasts at Low Densities Just like in cell cultures, the initial plating density of protoplasts has a profound effect on the plating efficiency. At high densities, such as 1× 104 to 1 × 105 protoplasts per 1 ml of medium, the cell colonies arising from individual protoplasts tend to grow into each other at an early stage in culture. This may lead to the formation of chimeral tissue if the protoplast population is genetically heterogeneous. In somatic hybridisation and mutagenic studies, cloning of individual cell/protoplast is essential (Fig. 14.5). This can be achieved by plating the protoplasts at very low densities (100-500 protoplasts ml-1). At low density, the development process of individual protoplast of heterokaryon (hybrid protoplast), can be easily monitored, thereby enabling the isolation and identification of hybrid colonies in the absence of a stringent selection system (Bhojwani and Razdan, 2005).

Fig. 14.5. Microchamber technique.

The nutritional components of the commonly used culture media are not suitable to induce cell division in protoplasts plated at low densities. A complex medium developed by Kao and Michayluk (1975) known as KM 8P, induces faster division in mesophyll protoplasts of alfalfa, pea, potato and tomato fusion products plated at low densities. The cultures in this medium are kept in darkness because KM 8P medium turns phytotoxic under strong light.

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14.11.3 Protoplast Fusion In this part of the chapter, various methods of protoplast fusion, and hybrid cell selections are given; also, practical applications of protoplast fusion and cybridisation technique have been discussed. 14.11.3.1 Methods for Protoplast Fusion

A. Spontaneous Fusion During enzymatic degradation of cell walls, some of the adjacent protoplasts may fuse together and form homokaryons (homokaryocytes) spontaneously. In this, plasmodesmatal connections in between the adjacent cells are enlarged and through these connections fusion of adjacent protoplasts takes place. However, the frequency of spontaneous fusion can be reduced by the sequential enzymatic method of protoplast isolation or exposure of cells to a strong plasmolyticum. B. Mechanical Fusion The giant protoplasts of Acetabularia have been fused mechanically by pushing together two protoplasts. This fusion does not need any fusioninducing agents. However, in this process protoplasts are likely to get injury. C. Chemical fusion Isolated protoplasts can be induced to undergo fusion with the help of fusogens (fusion inducing agents). In animals, Sendai virus is needed to induce fusion. In plants variety of fusogens are successfully employed to induce protoplast fusion, these are polyethylene glycol (PEG), NaNO3(Fig. 14.6) and high pH with high Ca++ ion concentration. (i) Polyethylene glycol (PEG) PEG has been used as fusogen in several species because of the reproducible high frequency of heterokaryon formation. About 0.6 ml of PEG solution (dissolve 1 g PEG mol. wt. 1500 in 2ml of 0.1 m glucose, 10 mM CaCl2 and 0.7 mM KH2 PO4) is added in drops to a protoplast pellet in the tube, tightly capped and incubate at room temperature for 40 min. Occasional shaking of culture tubes helps to bring the protoplasts in contact. This is followed by elution of PEG by the addition of 0.5 - 1 ml of protoplast culture medium in the tube after every 10 min. Wash the protoplast by centrifugation and resuspend in the same culture medium. Both molecular weight and the concentration of PEG are critical in inducing successful fusion. PEG less than 100 ml. wt. is not able to produce tight adhesions while that ranging upto 6,000 ml. wt. can be more effective in inducing fusion. Treatment with PEG in the presence of high pH/Ca++ is reported most effective in enhancing the fusion frequency and survivability of protoplasts.

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(ii) NaNO3 treatment Protoplast fusion induced by NaNO3 was first demonstrated by Power et al. (1970). Later, it was successfully employed in root tips of oat and maize seedlings. Isolated protoplasts are suspended in an aggregation mixture (5.5% sodium nitrate in 10% sucrose solution). This mixture induces the protoplast fusion on incubation (water bath maintained at 35°C). Finally, the mixture is replaced by liquid medium. The protoplasts in this culture medium are incubated again; this is repeated once or twice before plating the fused protoplasts on solid medium (Fig. 14.6). (iii) Calcium ions (Ca++) with high pH Isolated protoplasts of two different parents are incubated in a solution of 0.4 m mannitol containing 0.05 m CaCl2, with pH 10.5 (0.05 M glycine-NaOH buffer) and temperature at 37°C. Aggregation of protoplasts generally takes place at once and fusion occurs within 10 min. Many intraspecific and interspecific somatic hybrids have been produced using this method.

Fig. 14.6. NaNO3 treatment for production of Somatic hybrid of Nicotiana.

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(iv) Electrofusion Electric field can also be used for protoplast fusion, known as electrofusion. Culture tubes are placed in electric field in order to get fusion product. This method has been found to be simpler, quicker and more efficient than chemically induced fusion. More importantly, cells after electrofusion do not show any cytotoxic responses as generally found in PEG treatment of protoplasts. 14.11.3.2 Mechanism of Fusion

Protoplast fusion consists of three main phases: (i) Agglutination or adhesion - Two or more protoplasts come into close contact. This can be induced by a variety of treatments, e.g., PEG, high pH and high Ca++ ions; (ii) Plasma membrane fusion at localised sites - Membranes of protoplasts agglutinated by fusogen get fused at the point of adhesion, resulting into formation of cytoplasmic bridges between the protoplasts. Plant protoplasts carry a negative charge from -10 to 30 mv. Due to common charge plasma membranes of two agglutinated protoplasts do not come close enough to fuse. The high pH high Ca++ ions treatment has shown to neutralise the normal surface charge, so that agglutinated protoplasts come into close contact. High temperature also promotes membrane fusion due to perturbance of lipid molecules in plasmamembrane and fusion occurs due to intermingling of lipid molecules in membranes of agglutinated protoplasts. High molecular weight of PEG also promotes protoplast fusion. The PEG which is slightly negative in polarity can form hydrogen bonds with water, protein, carbohydrates, etc., which are positive in surface. When PEG molecule chain is large enough it acts as a molecular bridge between the surface of adjacent protoplast and adhesion occurs resulting into protoplast fusion. (iii) Heterokaryon formation - Fused protoplast increases its size forming spherical heterokaryon or homokaryon (see also Razdan, 2005 for details). 14.11.3.3 Identification and Selection of Hybrid Cells

After the fusion treatment, the protoplast population consists of a mixture of parental types homokaryones and heterokaryones. Since heterokaryones are potential source for regenerating the hybrids, efforts have been made to select them from the protoplast mixture. The following procedures have been employed for this purpose. A. Biochemical basis for selection Heterokaryones involving mesophyll protoplasts cannot be identified from the two parental types. Here biochemical markers are required allowing only the growth of heterokaryones in cultures or form somatic hybrid plants. (i) By using auxin-free medium: The auxin-free medium facilitates the hybrid Nicotiana glauca x N. longsdorfii. Parental protoplasts cannot

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grow in this auxin-free medium because these are auxin dependent to grow (Fig. 14.6). (ii) Drug sensitivity: Differential sensitivity of mesophyll protoplasts isolated from Petunia perodii and P. hybrida to the drug actinomycin D has been useful for hybrid selection (Razdan, 2005). Parental protoplasts are sensitive to the drug added to the MS medium. Whereas hybrid protoplasts are growing into callus which finally gives rise to somatic hybrid plant in the MS medium supplemented with this drug (Fig. 14.7). B. Visual Selection In most of the somatic hybridisation experiments, selection procedure involves fusion of chlorophyll-deficient (non-green) protoplast of one parent with green protoplast of other parent since this facilitates visual identification of heterokaryon at light microscopic level.

Fig. 14.7. Schematic representation of drug sensitivity test.

(i) Complementation selection coupled with differential media growth: Visual selection procedure coupled with complementary natural differences in the sensitivity of parental protoplasts to media constituents which enable only the hybrid cells to develop in cultures and regenerate plants, e.g., wild type protoplasts (mesophyll) of Petunia perodii fused with albino protoplasts isolated from cell suspension cultures of P. inflate and P. parviflora in separate experiments. In all these combinations, green perodii protoplasts got eliminated at the

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small colony stage, while the protoplasts of the other plant develop colourless callus. On the contrary, hybrid components proliferate into green calli, subsequently develop into somatic hybrid plant (Fig. 14.7). Similar procedures are followed in the selection of interspecific hybrids in Daucus, Datura,etc. In case of intergeneric hybridisation, e.g., hybrid between Atropa bellandnna and Datura innoxia, the parental protoplasts and heterokaryons are allowed to develop calli in cultures. The morphological differences in the resultant three types of calli permit the identification of hybrid tissue, which can then be selected out to regenerate somatic hybrid.

14.11.4 Cytoplasmic Hybridisation (Cbyridisation) In cytoplasmic hybridisation, fusion takes place between nucleus from one parent and cytoplasm of both parents, thus producing cytoplasmic hybrids, and called Cybrids. These cybrids can be obtained by using following methods: (i) fusion of normal protoplast from one parent with enucleated protoplast of other parent (Fig. 14.8); enucleated can be obtained by high speed centrifugation (20,000- 49-90 min) of protoplasts or by irradiate treatment, (ii) fusion of normal protoplast from one parent and protoplast containing non-viable nucleus from the other; this can be achieved by irradiation treatment, (iii) Selective elimination of one of the nuclei from the heterokaryon, and (iv) selective elimination of chromosomes of one parent at later stage after fusion. Cytoplasmic male sterility is obtained by cytoplasmic hybridisation. The identification of agronomically important traits regulated by the cell organelles of certain crop species, such as cytoplasmic male sterility (CMS) and cytoplasmic tolerance to the triazine herbicides (CD), as found in rapeseed, Brassica napus has made the production of cybrids commercially important. CMS is a valuable trait in breeding crops like rapeseed, since by preventing self-fertilisation, it can allow the economic production of single cross F1 hybrids. Besides this, incorporation CTT into rapeseed hybrids can allow the crop to grow in weed-infested areas. The weeds are eliminated by the application of triazine. In rapeseed, CTT is conferred by the chloroplast genome and CMS by the mitochondrion. In the procedure given in Fig. 14.8, mesophyll protoplast of CTT rapeseed ‘donor’ parent is fused with ‘recipient’ CMS rapeseed hypocotyl protoplast. When the fusion product undergoes organelle replication and random segregation it gives four types of plants. These are: (I) alloplasmic substitution cybrid which is male fertile and triazine tolerant A (mt-B/cp-B) but it possesses the nucleus of original CMS plant, (ii) reverse cybrid A (mt-B/cp-A) which is male fertile and triazine susceptible (due to opposite organelle combination, these are known as reverse hybrid) (iii) desired cybrid, A[mt-A(CMS)/cp-B(Cn)], which is cytoplasmic male sterile and triazine tolerant hybrid, and (iv) parental type A[mt-A(CMS)/cpA], is one which is male sterile and sensitive to triazine (Fig. 14.8)

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The technique of cybrid production has been exploited for obtaining the cytoplasmic male sterile Nicotiana and Petunia plants. Others like streptomycin resistance have also been transferred from Nicotiana tabaccumto N. sylvestris, using this technique.

Fig. 14.8. Cytoplasmic male sterility in Brassica napus (redrawn from Yarrow, 1999).

14.12 Anther and Pollen Culture Haploids are useful in crop improvement because they possess single copy of genes either dominant or recessive that can express even recessive mutations phenotypically. Large number of haploid plants have been produced by culturing an anther or pollen. Guha and Maheswari (1964, 1966) first reported pollen plantlets in anther cultures of Datura innoxia that caught the attention of plant breeders and geneticists all over the world. Induction of haploids by anther culture is known as Anther androgenesis or simple androgenesis. There are two types of obtaining the haploid plants, (a) Direct androgenesis in which pollen grain undergoes repeated cell division, differentiated into embryoids, that eventually giving raise pollen plantlet whereas, (b) indirect androgenesis in which callus formation takes place, later

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differentiated into embryoids, that subsequently giving rise pollen plantlets. The haploids plants are sterile, but they can be made fertile by diplodizing the chromosome number through colchicine treatment. Direct androgenesis e.g., Nicotiana, Datura, Atropa belladonna Indirect androgenesis e.g., Atropa, Arabidopsis, Asparagus Microspore has got four types of morphogenetic potentialities, these are: (1) to form a normal gametoplast (male gametotype), (2) to form a female gametophyte known as pollen embryosac; pollen embryo sacs were first observed by Nemac (1898) in Hyacinthus orientalis this is popularly known as Nemac phenomenon, (3) to form embryoids, e.g., Atropa, Datura, Nicotiana (4) to form callus tissue in Arabidopsis, Asparagus, etc. Anther culture For production for haploid plants, anthers rather than isolated pollen grains, are cultured because isolated pollen grains and their cultures have been successful in only a few cases. Anthers at right stage of pollen development should be excised from experimentally grown donor plant and inoculated onto a suitable nutrient medium. Different factors affecting the haploid production through anther culture are: (a) composition of culture medium, (b) stage of pollen development, (c) physiology of donor plant, (d) temperature and (e) genotype of experimental material. Culture medium Generally White, MS (Murashige and Skoog), Nitsch and Miller’s media are suitable for anther culture. These are supplemented with coconut milk and kinetic in various combinations. Embryoid formation from pollen has been observed on Nitsch’s medium alone but addition of cytokinetin (Kinetin or zeatin) that enhances good response (Misra, 2009). Sucrose at 2 per cent concentration has been found to be essential component in the medium. High concentration of sucrose seems to act as osmolyticum and tends to suppress callus formation and embryogenesis. Presence of iron in medium is equally important for pollen embryoid formation. Medium supplemented with activated charcoal (0.5-2 per cent) can stimulate androgenesis in some cases.

14.12.1 Ontogeny of Androgenic Haploids Microspore undergoes one unequal division, resulting into formation of one larger vegetative cell and smaller generating cell. Both cells combined or

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individually are participated in sporotypic type development. Repeated cell divisions take place and form the multicelled microscope. Later it gives rise to androgenesis haploid plant. During androgenic haploid development, three major phases are observed, these are: (a) Induction, (b) Early segmentation of microspore and (c) Later development. During induction period (from the time of anther culture to the first abnormal division) spreading about 6-12 days, there is complete degradation of gametophytic cytoplasm, and all cell organelles disappeared; ribosomes completely washed out. However, first population of ribosomes and other cell organelles reappeared afresh following the first sporophytic division of vegetative cell. That means, the vegetative cell programmed for gamtophytic development is now differentiated to take up sporophytic course of development. In the early segmentation of the microspore, four modes of in vitro androgenesis have been identified (Fig. 14.9). These are as follows:

Fig. 14.9. Early segmentation of the microspore and multicellular spore development.

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Pathway-I: Microspore undergoes equal division and both cells are distributed in sporophyte development, e.g., Datura innoxia. Pathway-II: Unequal division takes place, large vegetative cell contributes in formation of sporophyte, e.g., Hordeum vulgare, Nicotiana tobacum. Pathway-III: Unequal division; small generative cell is responsible for development of pollen plantlets, e.g., Hyoscyamus niger. Pathway-IV: Unequal division; both vegetative and genetic cells are contributed in pollen plantlet development, e.g., Datura innoxia, D. metel, etc. During later development, irrespective of the early pattern of divisions, the responsive pollen grain finally becomes multicelled one. This multicelled pollen gain later gives rise directly to embryoid (direct embryogenesis), subsequently transforms into plantlet. In the direct embryogenesis only one plantlet is formed, e.g., Atropa, Datura, Nicotiana, etc. However, in majority of plants, multicelled microspore further proliferates and it bursts open, releasing the callus. Now embryoids are differentiated on callus tissues. Numerous pollen plantlets are regenerated from the single mass of callus by media manipulation, e.g., Arabidopsis, Asparagus, Triticale, etc. (Fig. 14.10).

Fig. 14.10. Plantlet formation from multicelled pollen grain.

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14.12.2 Methods for Isolated Pollen Culture Pollen grains are collected at suitable stage of development from anthers. The isolated pollen gains are cultured by adopting following methods. i. Nurse Anthers Culture Technique: Sharp et al. (1972) developed this technique and raised the haploids plants from isolated pollen grains of Lycopersicon esculentum. In this technique, a healthy anther collected from some other plant; from Nicotiana glutinosa, Petunia hybrid has worked as nurse tissue. This nurse anther is taken in petridish; a filter paper disc is placed on nurse anther. Separately, a suspension of pollen grains from another anther of tomato was prepared in liquid medium (10 pollen per 0.5 ml of medium) and small aliquot of medium with 10 pollen grains is transferred to filter paper placed on nurse anther. Numerous clones are formed within a month. The clones obtained through this technique are uniformly haploid. ii. Anther Culture Technique: This technique is useful to raise haploids. In this technique, crushing of anther is avoided. Here whole anther excised from cold treated buds is floated on liquid medium (medium used for anther culture). After a few days, anther dehisces and discharging the batches of pollen grains including the induced pollen grains at various stages of development. All plants raised from the culturing of these pollen grains are uniformly haploids. To improve the efficiency of androgenesis, Wengel et al. (1975) and Rashid (1983) developed the technique, known as gradient centrifugation technique (Protocol given at the end of this section). In this technique, the embryogenic pollen grains capable of developing embryoids are separated from mixture of non-embryogenic pollen grains. Production of homozygous diploids Haploids were sterile ones since they contain only single set of chromosomes. They may grow normally up to the stage of flowering, but in the absence of homologous chromosomes, the meiosis is abnormal and consequently, viable gametes are not formed. Hence, it its essential to diploidise the chromosome complement of haploids to retain fertile homozygous diploids. In this context, the haploids have got immense importance in crop implement program. Diploidisation of haploids can be achieved through the classical method of colchicine treatment. In this method, colchicine of 1 g/L concentration can be used; young-pollen derived plants are immersed in colchicine solution for about 96 hrs and later transferred to culture medium to allow further growth. Alternatively, colchicine is added to pollen suspension culture medium; after 24 hrs of treatment, the haploids cells are transferred to fresh culture medium to allow further growth.

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Bulbosum method Hybridisation)

(Chromosome

elimination

through

Distant

Kasha and Kao (1970) demonstrated the Bulbosum method in which crossing between Hordeum vulgare (cultivated barley) (2n=14) and H. Bulbosum (wild barley) (2n=14) had involved. In this method, the female gamete of barley is fertilised in the H.bulbosum. During formation of embryo the chromosomes of H. bulbosum having the barley genome in the embryos are eliminated. Young embryos (immatured embryos) are excised and cultured to obtain haploid plantlets similar to that of H. vulgare in terms of morphology and chromosome complements.

14.12.3 Applications of Androgenic Haploids 1. The androgenic haploids are useful in crop improvement program. Mutations induced in haploids can be easily detected since haploids have only a single set of genes and hence there is no interference of dominant alleles. It is possible to develop haploids with desirable mutation/desirable characters, later their chromosome complements are diploidised to obtain fertile diploid plants with desirable characters. Several mutant cell lines have been developed through haploids, such as cell lines of resistant to environmental stresses, herbicides, viruses, drugs (antibiotics), etc. For example, salt resistant cell lines of Datura innoxia, nitrate reductase cell lines of Nicotiana tabecum, streptomycim resistant cell lines of Petunia hybrida and disease resistant cell lines of many crop plants have been isolated from pollen culture. 2. Haploids are useful in releasing new varieties through F1 double-haploid system. In haploid breeding technique, usually, only one cycle of meiotic recombination is involved. However, many agronomic traits (such as yield) are polygenetically controlled. One cycle of recombination is usually insufficient for the improvement of such quantitative traits since the linkage between polygenes will not release all potential variations available in a cross. To overcome this problem, alternative method of combining anther culture with sexual hybridisation among different genotypes of antherderived plants. The anthers of the hybrid progeny (F1) are excellent breeding material for raising pollen-derived homozygous plants (double haploids) in which complementary parental characteristics is combined in one generation (Fig. 14.11). This method shortens the breeding cycle, since in a normal hybridisation programme it takes several generations of backcrossing, or pedigree breeding to produce a stable homozygous line to release new variety (Luckett and Smithard, 1991).

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Fig. 14.11. Application of F1 double-haploid system in releasing new recombinants (Han, 1987).

3. Pollen embryos, being haploid, can be exploited for genetic transformation studies. Neuhaus et al. (1987) produced transgenic plants of Brassica napus by microprojectile bombardment of DNA into individual pollen embryos; haploid plants regenerated from transformed embryos on diploidisation expressed the recombinant transgene homozygously. 14.12.4 Protocols 14.12.4.1 Gradient Centrifugation Technique (Wenzel et al., 1975; Rashid and Reinert, 1981)

i. Collection of anthers of barley at proper stage of pollen development. ii. Prepare microspore suspension by gentle maceration. iii. Remove debris by repeated filtration and centrifugation and take the pollen suspension. iv. Pollen suspension is layered on solution containing 55% percol and 4% sucrose and again centrifuge it for 5 min. v. Androgenic, vacuolated pollen is formed a band at the top of the sucrose solution. vi. Band is pipetted, suspended in the washing medium (4% sucrose only) and centrifuge it; pellet at the bottom is deposited and supernatant discarded. vii. Take the pellet (rich in embryogenic pollen) and resuspend in same washing medium.

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viii. Now embryogenic pollen is cultured on pollen culture media (through anther nurse culture technique). ix. Embryogenic pollen is used as explant and from these androgenic haploids have been raised. 14.12.4.2 Raising of plantlets from isolated pollen (Nitsch, 1974; Razdan, 2005)

i. Collection of unopened flower buds, surface-sterilization with 2% sodium hypochlorite and wash thoroughly with sterile distilled water. ii. Place above anthers in 20 ml liquid medium and let anthers float for 2-3 days, and then squeeze out the microspores with a glass rod. iii. Filter the solution containing microspores and debris through nylon sieve (pore size 40 µm for Nicotiana, 25 µm for tomato and 100 µm for maize). iv. Centrifuge the suspension at 500-800 rpm for about 5 min. v. Discard the supernatant and resuspend the pellet containing pollen in fresh medium, again centrifuge it. Repeat iv and v steps twice. vi. Finally, mix the pollen with suitable culture medium at a density of 103104 grains ml-1. vii. Transfer the pollen suspension into the petri plate and see that the pollen grains should not be too deeply immersed in the medium. viii. Keep cultures in diffused light (500 lx) at 25oC temperature.

14.13 Micropropagation In general, plant propagation is of two types, asexual (multiplication through vegetative parts) and sexual (through zygotic seeds). Sexually propagated plants show the high amount of heterogeneity because of their seed progeny is not true-to-type unless they have been derived from inbred lines (Bhojwani and Razdan, 1985). On the other hand, asexual reproduction gives rise to plants that are identical to parental ones. Multiplication of genetically identical copies of any cultivar by asexual reproduction is known as clonal preparation. A population derived from a single individual by asexual reproduction is called clone. In plants like banana, fig, grape, chrysanthemum, vegetative multiplication is the only way because they produce little or no viable seeds. The widely used methods of cloning agricultural crops are cuttings of vegetative parts, layering, grafting and budding. The most of horticulturists have been adopting the asexual or vegetative reproduction for clonal multiplication of selected cultivars. The advantages of vegetative reproduction over sexual reproduction are as follows: i. A sexual multiplications are faster than the seed propagation in plants with a long-life cycle.

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ii. Bypassing the undesirable juvenile phase associated with seed-raised plants by propagating them vegetatively directly from adult plant materials. iii. Establishing gene banks by multiplying varieties among clonally propagated plants. Though the vegetative propagation is more advantageous, but it involves a lot of labour, time consuming and expensive approach. Tissue culture technology offers an alternative means of vegetative propagation. Clonal propagation through tissue culture means is called micro-propagation. It is a rapid method of clonal propagation which is cost-effective and time saving. By taking a small piece of plant material as explant and inoculate on suitable medium, it is possible to produce millions of plants within a short span of time. Micropropagation is an effective method of conservation of endangered and rare plant species. Use of tissue culture for micropropagation was initiated by Morel (1960) for orchid multiplication. Subsequently, it has been exploited by several horticulturists to develop and multiply various cultivars.

14.13.1 General Techniques of Micropropagation Micropropagation involves various stages. Murashige (1978a, b) proposed four distinct stages in Micropropagation of plants. Stages I-III are followed under in vitro, whereas stage IV is performed in a green-house conditions. Debergh and Maene (1981) suggested an additional stage ‘O’ for various micropropagating systems. All three stages are as follows (Razdan, 2009): Stage 0: Stage I:

Selection and maintenance of stock plants for culture initiation. Initiation and establishment of aseptic culture (explant isolation, surface sterilisation and washing, inoculation on suitable culture medium). Stage II: Multiplication of shoots or rapid somatic embryos formation using a defined culture medium. Stage III: Isolation and germination of somatic embryos, shoot function and rooting of regenerated shoots in vitro or in vivo. Stage IV: Transfer of plantlets to sterilised soil for hardening under greenhouse environment (in some cases rooting of in vitro-derived shoots may be performed at this stage by skipping stage III). Stage ‘O’ (Preparatory stage) It is the stage in which selection and maintenance of stock plants have been performed. Stock plants are grown under controlled conditions at relatively low humidity and good water facility, at least 3 months before the culture initiation. In woody and bulbous plants suitable temperature could help in breaking bud dormancy and provide more responsive plants. Systemic microbial contamination has to be checked in the stock plants.

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Stage ’I’ (Initiation and establishment of aseptic cultures) During this stage, explants are collected from different sources, and these are disinfected under aseptic conditions; subsequently these are inoculated on suitable culture medium. In this stage, cultures are initiated from explants of several organs but shoot tips and root tips are mostly used for commercial micropropagation. Generally, sub-terminal and older segments can withstand the toxic effects of sterilising agents much better than the terminal cuttings. The physiological status of parental plant from which explant is collected has played vital role on the response of buds. Explants collected from actively growing shoots during early growing season generally give good results (Anderdon, 1980). Bulbs, corms, tubers and other organs should be subjected to the temperature and photoperiodic treatments required to break dormancy before excising the bud (Bhojwani and Razdan, 1985). It becomes necessary to take submillimeter shoot tips as explants when the objective is to produce a plant free of viruses from the infected plant (see section 14.10). The best explants for growing virus-free plants are nodal cuttings. For multiplication through adventitious bud formation with or without callusing, explants should be derived from root, stem, leaf or nucellus to form adventitious buds. Cultures are also initiated from leaf-base or scalebase explants which are highly meristematic in monocots. Immature zygotic embryos are suitable explants for propagation of woody species. Of course, in citrus and mango, nucellar cultures are highly embryogenic (Bojwani and Razdan, 1985). For rhizomatic plants such as strawberry and ornamental ferns, runner tips are taken as explant for their multiplication. Stage II (Multiplication of shoots) This is the most critical stage in which three approaches have been followed to achieve in vitro multiplication: i. Multiplication is achieved directly from explants excised from proliferating shoots (shoot meristem) and axillary buds. ii. Induction and multiplication of adventitious meristems through processes of organogenesis of somatic embryos directly on explants. iii. Multiplication of calli derived from organs, tissues, cells or protoplasts, and their subsequent differentiation either through somatic embryogenesis or organogenesis in serial subcultures. These callus-derived shoots are further multiplied. Multiplication by shoot tips and axillary buds: Each axillary bud has potential to give rise to axillary branching in vivo in the absence of apical dominance. Since the apical dominance is under the control of various growth regulators, it can be altered by media manipulation to induce the regeneration of axillary branching from axillary buds. Shoot tips cultured on

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a basal medium without any growth regulators typically develop into single seedling-like shoots with strong apical dominance. Here shoots with apical dominance can be derived by taking a single nodal explant (Fig. 14.12). On the contrary, when shoots of some explant materials are inoculated on to medium supplemented with cytokinins, such as BAP, 2 ip, zeatin; axillary shoots develop precociously which proliferate to form secondary and tertiary shoots. These cluster shoots can be further sub-divided into smaller clumps of shoots/individual shoot which, in turn, will give rise to similar cluster of shoots when subcultural on fresh medium. This subdivision process may be continued indefinitely, and it is possible to regenerate plants in the range of 0.1-3.0 x 106 within a year (Mantell et al., 1985). Many forest tree and orchid species are good enough to multiply by axillary buds.

Fig. 14.12. Single node method of plant propagation (redrawn from Pierik, 1989).

Multiplication by adventitious shoots/buds: Buds arise at any site other than normal axil region and are known as adventitious buds (or) adventitious shoots. The adventitious shoot regeneration is dependent on type of the explant used in clonal propagation. The structures like bulbs, corms, tubers, rhizomes, stems, internodes, leaf blades cotyledons, root elongation zone, etc. These organs can be used as a cutting in conventional clonal propagation. Bulbs and corms in Iris, Lilium, and Tulipa hybrids can proliferate the cluster of shoots. For bulbs with a strong apical dominance in Narcissus, Allium and Hyacinthus, it is necessary to destroy the main apex. Vegetative propagation through adventitious bud formation from root (blackberry, raspberry) and leaf (Begonia, Crasula, Peporomia) cuttings is

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standard horticultural practice (Bhojwani and Razdan, 1985). The rate of adventitious bud development can be improved by media manipulation. For example, in Begonia, buds normally originate only along the cut ends but in a medium supplemented with BAP the bud formation is so profuse that the entire cut surface is covered with shoot buds (Reuther and Bhandari, 1981). Under the influence of an appropriate combination of growth regulators, in vitro, adventitious buds can be induced on leaf and stem cuttings of even those species that are not propagated vegetatively. Adventitious embryos (also known as embryoids) raised in vitro through embryogenesis of explant under suitable media are good source of micropropagation of plant. Each embryoid isolated and inoculated on suitable medium promoting organogenesis, can regenerate the plant. For example, leaf pieces of coffee trees form embryoids directly when inoculate on a basal MS medium supplemented with high concentration of cytokinin (Razdan, 2005). Hence, embryoids are the best source for micropropagation of plants. Multiplication by callus culture: Explant inoculated on suitable medium, undergoes rapid divisions and gives rise to callus. Subculturing the callus and it can be induced to redifferentiate shoot buds or roots (Skoog and Miller, 1957) and somatic embryos. Somatic embryo formation and organogenic regeneration can be controlled by growth regulators. Induction of root/shoot from callus depends on auxin/cytokinin ratio. Stage ‘III’ At this stage, germination of somatic embryos and rooting of regenerated shoots in vitro is performed. Various factors are influencing at this stage, these are as follows: Physiological status of plant material: Explants collected from very young plant parts, have got good regenerative capacity. Seasons of the year are playing important role in regenerative capacity of tissue cultures, e.g., flowerstem explants of Tulipa should be excised during dry storage (dormant) phase and nodal explants of Dioscoreaalata excised from plants growing under long photoperiods (16 hr). Culture medium: Standard media have been used for stage I and II, whereas stage III, needs some modifications. Auxins and cytokinins play an important role in micropropagation. To induce adventitions root formation, auxins are added to medium whereas for shoot formation high concentration of cytokinins is required. Medium fortified with activated charcoal induces the root formation and detoxifies some harmful compound accumulated during in vitro cultures. The relative concentration of NH+4 and K+ influence the size and number of shoots and affect the induction of somatic embryogenesis. Culture environment: The in vitro derived plants need optimal light intensity, i.e., 100 lx. The quality of light is equally important in organogenic differentiation and growth of shoots. Blue light (467 nm) induces the bud formation in tobacco and further enhances number of shoots. Similarly,

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red light stimulates induction of flower bud and far-red light induces root production. Diurnal illumination of 16 hr day and 8 hr night may give satisfactory results in multiplication and proliferation of shoots. Optimum temperature 25oC is needed during incubation.

14.13.2 Acclimatisation of in Vitro - derived Plants to Soil Conditions Lastly stage IV involves the transfer of in vitro-derived plantlets to the sterilised soil for hardening under greenhouse environment. Generally in vitro-derived plantlets show some structural and physiological abnormalities, these are: (1) high humidity, poor development of cuticle, (2) they are not photosynthetically active because of immature development of mesophyll with larger air spaces (3) presence of open stomata causing the significant water loss during the first few hours of acclimatisation and (4) poor vascular connection, between shoot and root reduce to very poor water conduction. Therefore, these plantlets are transferred to potting mix consisting of peat, vermiculate soil, sand and inorganic nutrients and maintained the high humidity during initial stage of acclimatisation upto 15 days. Development of stomatal closure mechanic is important component of acclimatisation. The following measures should be taken to close the stomata as well as to reduce the humidity and extra deposition of epicuticular wax: i. Exposing the plantlets to CaCl2, ii. Medium supplemented with linolein, and iii. Opening the culture tubes and keeping inside the desiccator with CaSO4. In vitro-raised tree-legume plants (Leucaena leucocephala) show high survival rates, provided these should be given pretransplant treatment. Plants are transferred to screw-cap bottles containing sterilised quartz sand irrigated with inorganic nutrient solution containing Rhizobium strain (NGR 8). Initially, bottles are kept closed for two weeks and subsequently remove the caps to maintain plants under controlled conditions of light and temperature (25±2oC and 18 Wm-2) for another two weeks, later these are transferred to field conditions.

14.13.3 Industrial Applications of Micropropagation Commercial applications of micropropagation are given below: i. Production of pathogen-free plants; currently the most popular application of micropropagation is the mass clonal preparation of desirable genotypes. ii. Production of millions of plants by taking a small piece of tissue as explant within a limited time and space. iii. In vitro conservation and propagation of endangered and rare species.

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iv. Production of plants throughout the year irrespective of seasons, especially in herbaceous species. v. Vegetative propagation is extremely important in the case of herbaceous taxa where seed progeny yields 50% males and 50% females but plants of one of the sexes are more desirable commercially (Bhojwani and Razdan, 1985).

14.14 Experimental Protocols 14.14.1 In Vitro Multiplication of Dendrobium aphyllum (Rahamtulla, 2022) Micropropagation of D. aphyllum has been done by taking leaf segments as explants. Regeneration of plants is done through callus induction and PLB formation (Fig. 14.13).

Fig. 14.13. Flowchart of D. aphyllum plantlets regeneration from leaf explants.

I. Surface sterilisation of the leaf explants 1. Collect the young, fresh, healthy leaves from D. Aphyllum (Mother Plant).

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2. Wash the leaf with tap water to remove soil and dust particles deposited on the surface. 3. Transfer the washed leaf into a glass beaker containing tap water; add a few drops of liquid detergent – Tween 20. 4. Cover the beaker mouth with muslin cloth with the rubber band and keep under running tap water for 10 minutes to remove any waxy/oily deposition on the leaf surface. 5. Wash it twice with distilled water. 6. Transfer the leaf explant into a laminar airflow hood for further work to avoid contamination. 7. Wash the above leaf with sterile distilled water thrice, each washing should be for 3-4 minutes. 8. Treat it with 0.5% HgCl2 solution for 5-7 minutes. 9. After treating it with disinfectant, wash it with sterile distilled water thrice, each washing should be for 3-4 minutes. 10. Wash with 70% alcohol for 2-3 minutes and again wash it with sterile distilled water thrice. 11. Transfer the sterile leaf to a sterile Petri-plate. II. Callus induction 12. Cut the leaf into small pieces of about 0.5 × 0.5 cm with a sterile blade. 13. Inoculate the leaf segments (0.5 cm each in length and width) with the abaxial surface on MS basal medium (Murashige and Skoog, 1962). 14. Use the Plant growth regulators such as 2,4-D and BAP (0.5, 1.0, 1.5, 2.0 mg/L) alone and in different combinations in the medium. Some explants should be inoculated on MS medium without hormones as a control measure. 15. Later incubate the cultures at 25±2ºC in growth chambers under a 16/8 photoperiod. 16. Callus will start appearing within 2 weeks and good callus growth can be observed in 3-4 weeks. III. Callus subculturing and proliferation of PLBs 17. Subculture the callus induced from the leaf explants on the optimal medium. Callus can be sub-cultured after the 6th week on a fresh medium with the same composition. 18. Later, cut the 6-week-old callus sections grown on the optimal medium into approximately 0.3-0.5 cm diameter segments. 19. Inoculate the callus segments onto the culture medium with the same combinations of 2,4-D and BAP as done earlier. 20. After subculturing, the callus mass develops Protocorm-Like Bodies (PLBs) on MS medium without any growth regulators (The number of PLBs will increase when the appropriate amount of growth regulators is used).

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IV. Regeneration of plantlets from PLBs 21. Transfer the well-developed PLBs produced in the optimal medium to four different basal media such as MS medium, Mitra orchid medium, Vacin and Went orchid medium and Gamborg B5 medium without using growth regulators for the development of shoots and roots. 22. Do subculture on the same media for the development of the complete plantlets. V. Acclimatisation of in vitro derived plantlets (see also section 14.7) 23. Transfer the three-month-old in vitro-derived D. aphyllum plantlets to natural conditions for acclimatisation. 24. Clean the plantlets with tap water to remove the sticked agar, treat them with 0.2% Diathane M-45 and transplant them into small pots. 25. Use different materials such as coco peat, charcoal, and red brick pieces as potting media separately and in combinations. 26. Arrange the potted plantlets into groups and place them on a wooden table and maintain the temperature ranging between 22°C to 25°C, illumination between 1200-1400 lux, and relative humidity of 80-90%. 27. Spray the water daily, and foliar sprays of NPK (1:1:1) weekly once to the potted plantlets. Result: D. aphyllum leaf explants inoculated on MS medium supplemented with 1.5 mg/L 2,4-D and 1.0 mg/L BAP, produced a green compact callus with a diameter of 1.92 cm (Fig. 14.14A). The combination of 1.5 mg/L 2,4-D and 1.0 mg/L BAP is the most favourable medium for callus induction (Rahamtulla and Khasim, 2022). After subculturing on the optimal medium, calli initially exhibited 3 different appearances: light green, yellow and light brown. The light green calli grew rapidly, consisting compact mass of isodiametric granules, and produced 1.2 PLBs on an average per 0.5 cm diameter of callus on MS medium without growth regulators. A low concentration of 2,4-D (0.5 gm/L) and a high concentration of BAP (1.5 gm/L) favoured the PLB formation. Different combinations of 2,4-D and BAP generated a greater number of PLBs in comparison to their individual effects. Media supplemented with 0.5 mg/L 2,4-D and 1.5 mg/L BAP produced 7.6 PLBs on average from the callus segment and the highest percent (76%) of PLB formation had observed (Fig. 14.14B). Further subculturing of these PLBs regenerated complete plantlets. After 6 weeks about 4.2 plantlets were formed from each PLB in MS basal medium (Fig. 14.14C). The highest shoot length (7.4 cm) and a greater number of leaves (3.8) on average were observed in the plantlets grown in MS basal medium (Fig. 14.14D, E). The plantlets with well-formed leaves and roots got established within 20 days of transplantation in the pots. The combination of charcoal and brick pieces supported the maximum survival frequency (100%) and growth of roots and leaves (Fig. 14.14F).

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Fig. 14.14. A. Green compact callus on MS medium + 1.5 mg/L 2;4-D + 0.5 mg/L BAP; B. The appearance of 7-8 PLBs from callus segments on MS medium + 0.5 mg/L 2;4-D +1.5 mg/L BAP; C. Regeneration response of PLBs on MS medium; D. Plantlets regenerated from single PLB subcultured on the MS basal medium; E. Plantlet growth on MS medium; F. Plantlets in the combination of redbrick and charcoal pieces.

14.14.2 Micropropagation of Grammatophyllum Speciosum Blume Sopalun et al. (2010) studied micropropagation of G. speciosum Blume. Protocorm-like bodies (PLBs) were induced from shoot tips.The highest frequency of PLBs (93%) were observed on explants incubated on half-

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strength ½ Murashige and Skoog (MS) liquid medium containing 2% (w/v) sucrose without any plant growth regulators (PGRs). Tests with different carbon sources compared to sucrose revealed that maltose promoted the highest relative growth of PLBs (7-fold increase); while, trehalose and sucrose yielded 5-fold and 4-fold increases, respectively. In ½ MS liquid medium, addition of 15 mg/l of chitosan promoted a 7-fold increase in PLB growth; while, 25 mg/l promoted a4-fold increase. However, the relative growth rate in solid culture was significantly lower in liquid culture. In addition, Chitosan supplementation in solid medium promoted shoot formation but not rooting. Plantlet regeneration was induced using a combination of NAA and BA supplementation in ½ MS solid medium with optimum induction shoot and root formation at 2.0mg/lNAA and1.0 mg/l BA. Using this protocol, approximately 8 months was required to obtain a hundred plantlets from one shoot tip. The plantlets showed no changes in ploidy when tested by flow cytometry. Fig.14.15 and 14.16 showed the establishment of G. Speciosum Blume micropropagation and transfer plantlets to the saranhouse, respectively. Summary of the micropropagation of G. speciosum is given Fig. 14.17.

Fig. 14.15. Establishment of Grammatophyllum speciosum Blume micropropagation; A: A shoot tip was excised under a stereo microscope; B: PLBs were induced in ½MS liquid medium (2 months after culture); C: PLBs were subcultured for multiplication (1 month after culture); D: PLBs were cultured on ½MS solid medium supplemented with 0.05% (w/v) of activated charcoal, 2.0 mg/l NAA and 1.0 mg/l BA; E and F: Shoots and roots were induced, respectively (3 months after culture). Bar = 1 mm.

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Fig. 14.16. Transfer of G. speciosum Blume plantlets to the saranhouse. A: Plantlets (6.0-8.0 cm long) were obtained after 3 months of culture; B: Plantlets were removed from the bottle and washed with tap water; C: Plantlets were transplanted to pots filled with small pieces of coconut husk; D: Plants grown 6 months in the saranhouse; E: Plants grown 1 year in the saranhouse; and F: Plants grown 2 years in the saranhouse. Bar = 1 cm (A to D).

Fig. 14.17. Flow chart of micropropagation of Grammatophyllum speciosum Blume.

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14.14.3 Micropropagation of Vanda Coerulea Griff. ex Lindl. Wasiksiri et al. (2010) studied the effects of 6 different liquid media including, modified Vacin and Went medium (Vacin and Went, 1949), Murashige and Skoog medium (Murashige and Skoog, 1962), Schenk and Hildebrandt medium (Schenk and Hildebrandt, 1972), half strength MS (½MS), half strength SH (½SH) and half strength KNO3 in SH (½KNO3SH) which contained 100 ml/l coconut water and without sucrose on multiple shoot formation from lateral buds of Vanda coeruleaGriff. ex Lindl. After 8 weeks of culture, the survival of lateral buds (100%) was observed on ½SH and ½KNO3SH but the protocorm-like structure was found only in ½ KNO3SH. For increasing multiple shoots or protocorm-like bodies (PLBs), lateral buds were cultured

Fig. 14.18. Micropropagation of a lateral bud of Vanda coerulea Griff. ex Lindl. in ½ KNO3SH liquid medium after 8 weeks of culture in medium. A: without PGR, B: 0.1 mg/l TDZ, C: 0.5 mg/l TDZ, D: 1 mg/l TDZ, E: 1.5 mg/l TDZ, F: 2 mg/l TDZ, G: 1 mg/l BAP, H: 2 mg/l BAP, I: 0.1 mg/l NAA and 1 mg/l BAP, J: 0.5 mg/l NAA and 1 mg/l BAP, K: 1 mg/l NAA and 1 mg/l BAP, L: 0.1 mg/l NAA and 2 mg/l BAP, and M: 0.5 mg/l NAA and 2 mg/l BAP. Bar = 0.5 cm.

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Fig. 14.19. Effects of organic additives and combinations with concentrations of sucrose and organic additives on embryogenesis on PLBs of Vanda coerulea Griff. ex Lindl. under culture in ½KNO3SH contained 100 ml coconut water and 1 g/l activated charcoal after 8 weeks of culture. A: ½KNO3SH agar medium without organic additives and sucrose. B: ½KNO3SH agar medium supplemented with 10 g/l sucrose. C: ½KNO3SH agar medium supplemented with 20 g/l sucrose. D: ½KNO3SH agar medium in combination with 50 g/l banana without sucrose. E: ½KNO3SH agar medium in combination with 50 g/l banana and supplemented with 10 g/l sucrose. F: ½KNO3SH agar medium in combination with 50 g/l banana and supplemented with 20 g/l sucrose. G: ½KNO3SH agar medium in combination with 50 g/l potato without sucrose. H: ½KNO3SH agar medium in combination with 50 g/l potato and supplemented with 10 g/l sucrose. I: ½KNO3SH agar medium in combination with 50 g/l potato and supplemented with 20 g/l sucrose. J: ½KNO3SH agar medium in combination with 50 g/l banana and 50 g/l potato without sucrose. K: ½KNO3SH agar medium in combination with 50 g/l banana and 50 g/l potato and supplemented with 10 g/l sucrose, and L:½KNO3SH agar medium in combination with 50 g/l banana and 50 g/l potato and supplemented with 20 g/l sucrose. Bar = 1 cm.

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on ½SH and ½KNO3SH media supplemented with different concentrations of plant growth regulators (TDZ, BAP, and NAA). The highest fresh weight of PLBs (total fresh weight of 1.4 g) was observed on ½KNO3SH medium supplemented with 1 mg/l BAP and 0.1 mg/l NAA (Fig. 14.18). Development of PLBs was established by organic additives (activated charcoal, potatoes, bananas, and sucrose). The results revealed that PLBs which cultured on ½KNO3SH supplemented with 1 g/l activated charcoal, 50 g/l potatoes, 50 g/l bananas, and 20 g/l sucrose produced the highest shoots (434 shoots/lateral bud). These new shoots were able to develop into plantlets. New shoots were cultured on ½KNO3SH supplemented with 1 g/l activated charcoal, 50 g/l bananas, and 20 g/l sucrose. After 16 weeks of culture, the best growth of plantlets was found (Fig. 14.19). Summary of the micropropagation of V. coerulea is given Fig. 14.20.

Fig. 14.20. Flow chart of micropropagation of Vanda coerulea Griff. ex Lindl.

15 Chromatography Chromatography is the most powerful technique to separate various components from a mixture of chemical substances based on their physicochemical properties. The term ‘Chromatography’ was coined by Russian botanist Tswett in 1906 (Greek words Chroma meaning ‘Colour’ and graphs to write’). In chromatography, the sample is applied at one end of a porous support that holds stationary phase (solid or liquid) and the mobile phase (liquid or gas) on another end is allowed to run over it. The various components in the sample get separated due to differences in their position or distribution behaviour between the stationary and mobile places. The partition or distribution coefficient (Kd) is defined as the ratio of concentration of compound in two phases (A and B) at equilibrium. If, both the phases are liquids, the formula is as follows: Kd =

Concentration of compound in solvent 'A' Concentration of compound in solvent 'B'

Separation of various compounds in the mixture takes place based on their distribution coefficient. In other words, the compounds in the sample that have greater affinity for stationary phase; the movement of such compounds is retarded since they tend to spend more time in that phase. On the other hand, the compounds with relatively higher affinity for mobile phase travel at a much faster rate as these are moved along with following mobile phase. This leads to separation of various compounds in the sample. Chromatographic techniques may broadly be classified on the basis of nature of stationary phase. These include: i. Paper chromatography: In this system, a filter paper sheet is used as a support for stationary phase. ii. Thin layer chromatography (TLC): A glass plate, plastic sheet or a piece of metal foil can be used as support for stationary phase that has been applied in the form of a thin layer in these materials. iii. Column chromatography: In this, stationary phase is packed into a tubular glass, polypropylene or metal columns. The various chromatographic techniques have also been classified based on the forces or interacting phenomenon between the solute molecules and the stationary phase. These are as follows:

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Partition chromatography Adsorption chromatography Affinity chromatography Ion-exchange chromatography Molecular sieve chromatography

The other modern instrument known as High Performance Liquid Chromatography (HPLC) is highly expensive. The classical techniques such as paper chromatography and thin layer chromatography can be set up in ordinary laboratory without much expenditure. The separation, identification and quantification (semi) of amino acids using paper chromatography is given below. The same method can be employed to separate other smaller molecules such as sugars, organic acids etc., by changing the mobile phase and spraying (detection) reagents (Table 15.1).

15.1 Paper Chromatography Depending upon the direction of flow of mobile phase the systems have been used in our laboratories. These are ascending and descending paper chromatography. In the ascending type, the solvent is kept at the base of the tank. Whatman filter paper is immersed in the solvent. The sample is applied at the edge of paper and see that the sample spot does not get dipped into the solvent; it should remain just above the surface of the solvent (Fig. 15.1). The solvent ascends (moves up) the paper by capillary action and the separation of compounds takes place according to the differences in their partition coefficients. The most polar substances will be at the bottom with respect to the tank whereas the least polar will be on the top end of the tank (Fig. 15.1). Ascending technique is relatively slow process.

Fig. 15.1. Ascending paper chromatography. (Redrawn from Sawhney & Singh, 2001).

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In the descending method, the tank containing solvent (mobile phase) is fitted at the top of the chamber. The filter paper is hung in such a way that the side where the sample has been spotted is dipped in the tank. The solvent travels down the paper under the force of gravity (Fig. 15.1). In this technique most polar substances will be on the top with respect to tank whereas least polar ones will be at the bottom.

15.2 Thin Layer (planar) Chromatography (TLC) Thin Layer Chromatography (TLC) is very useful in research laboratories to separate, identify and characterise various compounds. A variety of molecules such as sugars, amino acids, organic acids, lipids etc., are separated by this technique. The great advantage of this technique is within a short time, about 30 min or more than 90 min, separation is achieved. The principle of TLC is similar to that of column chromatography (adsorption chromatography). In the adsorption process, the solute competes with the solvent for the surface sites of the adsorbent. Depending on the distribution coefficients, the compounds have been separated on the surface of the adsorbent. The adsorbent normally contains a binding agent such as calcium sulphate which facilitates the holding of the adsorbent to the glass plate.

15.2.1 Preparative TLC Instead of thin layer, layer of adsorbent is coated and a greater amount of sample can be applied onto the plate. After the separation, the area of separated compound is scraped off, eluted with a suitable solvent and recovered in a relatively pure form. This modified TLC is known as preparative TLC. In a preparative method, a large amount of sample is applied in the form of streak rather than a spot. The compounds are separated as a series of bands and these may be scraped off and eluted with a suitable solvent. Extraction i. For extraction of amino acids, sugars, phenols, phenolic acids and plant acids follow the procedure given for amino acid extraction in the section Paper chromatography. ii. Neutral lipids: Macerate the flesh leaf tissue in 20 volume of isopropyl alcohol (cold), the re-extract with chloroform-methanol (2:1). Seed tissue is extracted directly with chloroform and ethanol (2:1) mixture. The lipids in cereals can be extracted with chloroform, ethanol, water 200:95:5 mixture. Store the extracts at 5oC in the presence of antioxidants (Butylated hydroxytoluene BHT 0.005%).

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Table 15.1. Identification of compounds by their colour* Absorbent

Solvent system

Mode of Identification

Colour

Amino acids

Silica gel G

(i)Ethanol and water (70:30) (ii)Butan-1-ol acetic and water (80:20:20)

Spray with 0.1% ninhydrin in acetone and heat the plates for 5 min at 100-110oC

Pink or purple

Mono and disaccharides

Kieselghur G. (sodium acetate)

(i)Ethyl acetate and propan-1-ol (65:35)

Spray with 2% diphenylamine (in water) saturated with n-butanol (or) in n-butanol and methanol (1:1) containing 5% trichloroacetic acid. Heat at 100oC for 10-15 min.

Aldohexoses give brown spots; aldopentoses give purple spots.

Neutral lipids (alkanes, triglycerides, diglycerides and monoglycerides

Silica gel G

Petroleum ether, diethyl ether and acetone (90:10:1) (or) Isopropyl ether acetic acid (24:1)

(a) Spray with ethanolic 0.2% 2,7 dichlorofluorescein, observe under UV light

Spots fluoresces light green.

Plat acids

Silica gel G

Ethanol, water and 25% ammonium hydroxide (78:9,5:12,5)

Spray with 0.1% 2,6 dichlorophenol indophenols in ethanol (brief heating enhances the colour)

(b) Spray with 0.5% ethanolic rhodomine B

Yellow or blue violet spots on a pink background.

Pink spots on sky blue background (oxalic acid gives rocket-like spot).

(Contd.)

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Compound

Table 15.1. (Contd.) Phenols

Silica gel G

Acetic acid and chloroform (1:9) (or) Ethyl acetate and Benzene (9:1)

Spray with vanillin HCl

Orcinol and 2-methyl resorcinol given blueish-pink colour; 4-methyl resorcinol and resorcinol give red colour.

Purine and Pyrimidine bases

Silica gel G

Ethanol acetic acid and water (81:5:14)

Spray with 0.25% mercuric acetate and 0.05% diphenyl carbazone

White spots on violet background

*(adopted from Sadasivan and Manikam, 2005)

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15.3 Column Chromatography In column chromatography, separation is achieved by passage of sample through vertically fixed tubular glass or polypropylene column which is loaded with an appropriate chromatography media. These columns with a porous sintered plate fixed at their base, are commercially available. Alternatively, a simple glass burette with a plug of glass wool at the base can be used as column. At the base, there is a small capillary tubing through which eluent from the column flows into test tubes from which fractions are collected. Chromatographic media (stationary phase) in the column are water insoluble, porous and resolution of sample components occurs depending upon the size and shape of the stationary phase, accordingly flow rate depends. Large size and coarse particles have higher flow rate and give poor resolution, while finer particles with large surface to volume ratio have slower flow rate but greater resolution efficiency. Generally, for better resolution, the smaller particles of 200-400 mesh have been employed. For fractionation and separation of components, the sample is loaded on the top of the column and eluted with an appropriate buffer. The effluent (eluent) emerging from the base of the column is collected in the form of fractions of fixed volume of fixed time in individual test tubes by using either an automatic fraction collector or manually. The collected fractions are analysed for the presence of desired substances or compounds. Various types of column chromatography have been classified based on the nature of interaction between solutes and the stationary phase. These are given below:

15.3.1 Absorption Chromatography In adsorption chromatography, the sample compounds (solute) are hold on to the surface of a solid adsorbent, having specific absorption sites, via weak forces such as van der Waal’s forces and hydrogen bonding. Different compounds bind with varying strengths and hence can selectively be desorbed. For good resolution, selection of good adsorbent and eluent (or mobile phase) is very important. Some of the commonly used absorbants are charcoal, silica, alumina, etc. Eluent influences quality of separation since polarity of mobile phase influences the adsorption considerably. In general, the polar solvents are preferred for the substances having polar or hydrophilic groups and non-polar solvents for substances with hydrophobic or non-polar groups. For example, alcoholic solvents for OH group containing substances; acetone or ether for substances with carbonyl groups; and hydrocarbons such as toluene or hexane for non-polar substances.

15.3.2 Affinity Chromatography Purification of sample by affinity chromatography is different from all other forms of chromatography that this technique does not make use of the

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differences in physical properties (such as solubility, adsorption, molecular weight and ionic properties) of the molecule to be separated. Rather it exploits one of the unique and fundamental properties of biopolymers, i.e., the specificity of their interaction with other biomolecules. In other words, affinity chromatography is a type of adsorption chromatography in which substance to be isolated is specifically and reversibly bound to a complementary binding substance (ligand) immobilised on an insoluble chromatographic bed material (matrix). The unbound substances will be washed away while bounded substance subsequently recovered by displacement from the ligand either by specific (affinity) elution or by non-specific (change in pH or ion concentration) elution (Fig. 15.2). Some of the macromolecules separated by employing specific ligand in affinity chromatography are given in the Table 15.2.

Fig. 15.2. Schematic representation of principle of Affinity chromatography. Table 15.2. Specific ligands commonly used in Affinity chromatography Ligands

Macromolecules/cells

Bicocytin Tryptophan Benzamidine Heparin Poly(U) or poly dT Lysine Concanavaline A Insulin, Concanavalin A

Avidin Α-chymotrypsin Thrombin Interferon Poly(A) messenger RNA Ribosomal RNA Glycoproteins, glycolipids Fat cells

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The commonly used matrices are cross-linked dextrans (e.g.,sephacryl), agarose (sepharose), polyacrylomide gel (biogel P), polystyrene, Cellulose, Porous glass and silica. Spaces arm: It facilitates effective binding to macromolecules (proteins) to the ligand and matrix. The length of the spaces arm is critical. If it is too short, the arm is ineffective and the ligand fails to bind the substance, if it is too long, non-specific effects become pronounced and reduce the selectivity of the separation. So the optimum length of the spaces arm is generally 6-10 c-atoms or their equivalent. The chemical nature of the spaces arm (hydrophobic or hydrophilic) is also critical for the success of separation. The approaches have been followed for the coupling of the legend. In first one, spaces arm is first linked to the matrix followed by the coupling of the ligand, whereas second approach involves the binding of the spaces arm to the ligand which is then linked to the matrix. The first approach is more convenient than the second one.

15.4 High Pressure Liquid Chromatography (or) High Performance Liquid Chromatography (HPLC) High Pressure Liquid Chromatography or High Performance Liquid Chromatography (HPLC) is an advanced column chromatography which has been used to separate various compounds from the mixture of various chemical compounds. It helps in identify, quantify and purify the individual compounds from the mixture of sample. In HPLC, the principle involves that some compounds of the sample take longer than others to pass through the chromatography column. The time taken by the molecule depends on affinity of the molecule with mobile phase (liquid or gas) and stationary phase (solid or liquid). Those compounds with greater affinity with the stationary phase take longer time to pass through the column (longer retention time). HPLC made of pump to pass a pressurised solvent containing the sample mixture through a column filled with a solid absorbent material. Each component in the sample interacts slightly different with the adsorbent, causing different flow rates for the different components and leading to the separation of compounds as they flow out of the column. In this technique, the sample is forced by a liquid at high pressure (the mobile phase) through a column which is packed with a stationary phase composed of irregularly or spherically shaped particles (a porous monolithic layer or a porous membrane). HPLC is distinguished from traditional (low pressure) liquid chromatography; in HPC, operational pressure is significantly higher (50-350 bar), while ordinary liquid chromatography typically depends on the force of the gravity to pass the mobile phase through the column. As the small amount

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of sample to be separated in the analytical HPLC, typical column dimensions are 2.1-4.6 mm diameter, and 30-250 mm length. Moreover, HPLC columns are made with small adsorbent particles of about 2-50 µm particle size. Hence, HPLC gives superior resolving power to separate various compounds from the mixture of compounds. HPLC typically consists of a pumping system, simple injector, a chromatographic column of 5-25 cm length, stationary and mobile phases, detector and data collecting device (Fig. 15.2). The pump delivers the desired flow (50 MP pressure) and composition of the mobile phase through the column. The injector introduces a narrow stream of variable and reproducible sample volume onto the top of the analytical column. The optimum level of the analyte is maintained in the analytical column at an acceptable pressure drop. The detector generates a signal proportional to the amount of sample component emerging from the column, hence allowing for quantitative analysis of the sample components. A digital microprocessor and user software control the instrument and provide data analysis. HPCL has been used for several purposes, such as separation of compounds from the biological samples, detecting vitamin D, levels in blood serum, separation of various synthetic chemicals, during the production process of pharmaceutical and biological products etc.

15.4.1 Normal Phase Liquid Chromatography (NPLC) In Normal Phase Liquid Chromatography (NPLC), stationary phase is more polar (hydrophilic, e.g., silica) than the mobile phase (toluene). The polar molecules which have greater affinity with hydrophobic phase would pass slowly through column than the non-polar molecules (Fig. 15.3).

Fig. 15.3. Normal Phase Liquid Chromatography.

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15.4.2 Reverse Phase Liquid Chromatography (RPLC) In Reverse Phase Liquid Chromatography (RPLC), stationary phase is hydrophobic (non-polar, octadecylsilyl-C18), whereas mobile phase is more hydrophilic (polar). In this, non-polar molecules would move slowly through the column when compared with the polar molecules (Fig. 15.4).

Fig. 15.4. Reverse Phase Liquid Chromatography

Selecting the mobile phase (solvent) is one of the most important steps when performing HPLC and is selected based on polarity. Solvent polarity relates to the ability of the components to partition into the phase. The relative polarity of various solvents is given below (Table 15.3). Table 15.3. Relative polarity of mobile phase (solvents) Solvent Cyclohexane 1-chlorobutane Carbon tetrachloride Toluene Tetrahydrofuran (THF) Ethanol Methanol Acetonitrate Ethylene glycol Water

Relative polarity 0.04 1.0 1.6 2.4 4.0 4.3 5.1 5.8 6.9 10.2

In the HPLC analysis of a mixture, the components will separate based on their retention times. This will produce a chromatogram. Either the peak height or peak area can be used to estimate the concentration.

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These values are proportional to the concentration and the peaks are sharp, and the flow rate is carefully produced. A calibration curve is prepared by plotting either peak height or peak area as a function of compound concentration.

15.4.3 Ion Exchange HPLC In Ion Exchange HPLC, partition of ions takes place between a polar liquid phase and a stationary phase with ion exchange sites. The ion exchange sites are typically immobilised in small beads of resin that are formed by crosslinked polymer. Bonded phase columns in which the ion exchanger is bonded to small particles of silica are also available. Cations are separated on cation exchange resins which contain negatively charged functional groups such as SO3 and –COO--. Anions are separated on anion exchange resins which contain positively charged functional groups such as CH2N+(CH3)3. Resin affinity increases with increasing charge density.

15.5 Adsorption Chromatography In Adsorption chromatography, the stationary phase is a solid of polar nature such as hydrated silica or alumina. The mobile phase and solvent mixture are in completion for active adsorption on the stationary phase particles. Hence, more strongly adsorbed components are retained longer than weakly adsorbed components. As more polar compounds adsorb on a polar surface to a greater extent than the less polar compounds, retention in column is related to sample extract polarity.

15.6 Gas Chromatography/Mass Spectrometry (GC/MS) The different unknown samples can be separated by high resolution capillary Gas Chromatography (GC) coupled with an ion tap detector (ITD). The ITD is a variation of a quadrupole mass spectrometer (MS) and is designed to function specifically as a GC detector. Separation of sample into its various compounds depends on solubility differences of the sample vapour in a liquid (the stationary phase). The stationary phase is coated in a thin layer on solid particles (solid support) of large surface area and then packed uniformally into a column. A constant flow of the carrier gas passes through the column and transports solute molecules in the gas phase. The column has oven inside for precise temperature control. ITD records the total number of ions entering the mass analyser from the column. The chromatogram produced is called the total ion chromatogram. Each point in chromatogram is a mass spectrum. The basic principle in MS, it utilises the nature of ions. By accelerating an ion (anatom or molecule with an electric charge) to a certain speed, and

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passing it through a magnetic field, the path of the ion will be deflected by magnetic field. The amount of deflection depends on the intensity of the magnetic field and the mass number of the ion. Each compound can be identified by comparing its retention time (RT) to that of standard. RT is defined as ‘the length of time compound remains present in the column’. RT depends on the column temperature, its length, type of stationary phase and the carrier gas velocity. Furthermore, ITD provides the structure of compound. Therefore, compounds can be detected not only by their retention time but also their mass spectral data.

15.7 Protocols 15.7.1 Paper Chromatography (Sadasivan and Manikam, 2005) Materials and Reagents i. ii. iii. iv. v. vi.

vii.

viii.

ix. x.

Chromatography chamber Whatman No.1 filter paper Hair-dryer Automixer Micropipette Mobile phase (Solvent system) – Take n-butanol, glacial acetic acid and water in the ratio of 4:1:5 in a separating funnel, mix them thoroughly and stand to equilibrate for 30 min. Drain off the lower aqueous phase into beaker and place it inside to saturate the chromatography chamber. Upper organic phase is retained and use it for developing chromatogram. Extraction of sample – With mortar and pestle grind a known quantity of the sample material in 10-fold volume of 70% ethanol. Shake the contents thoroughly at 55oC for 30 min. Centrifuge it at 10,000 rpm for 10 min. Collect the supernatant. Repeat the extraction of the pellet at 55oC at least twice. Collect supernatant and shake vigorously (for leaf extracts, treat with equal volume of petroleum ether 40-60oC). Discard the petroleum ether layer containing chlorophyll. Evaporate the ethanol fraction to dryness under vacuum by using evaporator at 40-45oC. Dissolve the residue in a known volume of absolute ethanol or water for analysis. Dissolve different individual amino acids in a distilled water at a concentration of 1 mg/ml. Use very dilute (0.05 N) HCl to dissolve the free amino acids tyrosine and phenylalanine. Dissolve tryptophase is very dilute 0.05 N NaPH. Ninhydrin reagent – Dissolve 100 mg ninhydrin in 100 ml acetone. Elution mixture – Prepare 1% copper sulphate solution. Mix ethanol and copper sulphate in the ratio 80:20.

Chromatography 321

Procedure i. Cut the chromatography sheet carefully to the size 40x24 cm. Draw a line with pencil across the sheet about 5 cm away from one end. Mark a number of points at intervals of 3 cm. ii. By using micropipette apply 25 µl of each amino acid as a separate small spot-on filter paper. iii. Also spot different known aliquots of sample extract. iv. After spotting, set the chamber for ascending or descending type of chromatograph (see previous section). v. Add the organic (phase) solvent to the tank and close the chamber airtight. Develop the chromatogram, preferably overnight, till the solvent moves almost to the other end. vi. Mark the solvent front and dry the chromatogram in a fume chamber. vii. Spray the chromatogram with ninhydrin. Dry the filter paper at room temperate for 5 min and followed by at 100oC in an oven for 2-3 min. Amino acids appear as purple spots; hydroxyproline and proline give yellow-coloured spots. Mark all the spot and calculate their retardation factor, Rf values by the following formula: Rf =

Distance moved by the solute from the origin (cm) Distance (cm) moved by the solvent from the origin

Comparing the Rf values with that of the authentic amino acids that are co-chromatographed, amino acids present in sample is identified. i. For quantitative estimation, cut each spot from the chromatogram and also cut into several small bits and transfer them to the bottom of the test tube. Add 3 ml of elution mixture, shake the tubes vigorously for 15 min. Decant the liquid and again elute the pieces with another 2 ml of elution mixture. Repeat this step till unit bits become colourless. Combine and clear the elute by centrifuging at 10,000 rpm for 10 min. take the readings of intensity of purple colour at 570 nm in a colorimeter, compare this with leucine (50 µg) run as standard.

15.7.2 Separation and Identification of Sugars by Adsorption Thin Layer Chromatography (Sawhney and Singh, 2010) Sugars are separated on the basis of differential adsorption onto silica gel. The sugars that have higher affinity for stationary phase are adsorbed more strictly and hence, they migrate slowly when mobile phase moves over them. Whereas, these sugars having lower affinity for stationary phase are absorbed very weakly and, they move more rapidly on the mobile phase. These separated sugars clearly appear when the plate is sprayed with aniline diphenylamine.

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Reagents and materials Silica gel (G) Solvent system: Prepare a mixture of ethyl acetate: isoprepanol: water: pyridine (26:14:7:2 v/v) Standard sugar solution: Prepare 1% standard sugar solutions such as glucose, ribose, lactose etc., in 10% iso-propanol (v/v). For mixture in which sugars have to be identified, mix the sugar solution in equal proportions. Aniline diphenylamine reagent: Mix 5 volumes of 1% aniline and 5 volumes of 1% diphenylamine in acetone with 1 volume of 8.5% phosphoric acid. Glass plates (20 × 20 cm) Spreader TLC chromatographic tank Micropipettes Sprayer Hair drier Oven maintained at 105oC Procedure i. Tape clean and neat dried glass plates (20 x 20 cm) on a flat plastic tray side by side without any gap. ii. Prepare the slurry of stationary phase (silica gel G) without clumps in the water or in a suitable buffer. iii. With help of spreader, spread the uniform layer of 250 µm thickness by making it from one end of the tray to its other end. iv. Activate the glass plates by keeping them at 105oC for 30 min. Allow the plates to cool in a desiccator before use. v. Gently put marks in a straight line with a help of a pin at a distance of 2 cm from one edge of the plate. The adjacent marks should be about 15-20 cm apart from each other. Extreme care should be taken that silica plate should not get scratched during this process. vi. Carefully apply the solution of individual standard sugars and the mixture of alcoholic extract of the sample on the separate marked spots. vii. Gently put marks or draw a line 1 cm from the opposite edge. viii. Place the glass plates in chromatographic tank that has already equilibrated with the solvent taking care that the base line on which samples have been applied should not dip into the solvent. ix. Now close the chromatographic tank with air-tight lid and allow the solvent to ascend along the silica plate by capillary action till the solvent reaches the marked line of the upper side of plate. Run the chromatogram for 90 min. x. Take out the plates from the tank and allow them to dry at room temperature. xi. For determining the location of sugars on the TLS plates, spray them

Chromatography 323

with freshly prepared aniline-diphenylamine reagent and ensure that silica gel should not blow off during spraying. xii. Later keep the plates in hot air oven at 100oC for 10 min. Bluish spots on the white background appeared that indicate the presence of sugars at that region of the plate. xiii. Measure the distance from the base line to the cooled spot and calculate the Rf value of each sugar; and identify the sugar or sample by comparing the Rf value of the standard sugar. We shall observe bluish spots at different locations on the silica plates after spraying with aniline-diphenylamine reagent. By calculating Rf value, we can identify the type of the sugar present in the plant sample. In the experiment, the size of the sample applied on the silica plate should be as small as possible. If large volume of the sample has to be spotted, then it should be done in small aliquots with an intermittent drying. Overloading is generally avoided in the adsorption thin layer chromatography.

15.7.3 Column Chromatography and HPLC Analysis [Analysis of Phenolic Compounds from the Solvent Extract of Salvinia Molesta Mitchell (Giri and Jothi, 2018)] In this protocol, separation of phytocompounds in the ethyl acetate extract of Salvinia molesta by column chromatography and determination of phenolic compounds present in the active fractions by HPLC are taken-up. Chemicals and Phenolic standards Hexane, ethyl acetate, ethanol, methanol, acetone, vanillin-H2SO4spray, acetonitrile, phosphoric acid, acetic acid, chromanorm water, gallic acid, catechol, benzoic acid, resorcinol, ascorbic acid, vanillin, quercetin, silica gel and sea sand were procured from Hi Media Pvt. Ltd., Mumbai, India. Plant materials Clean plant materials of S. molesta under running tap water and shade drying at room temperature for three weeks. The dried materials are pulverised into fine powder, filter through a sieve (mesh no.40) and stored airtight bottles. Procedure i. Prepare plant extracts using four solvents, namely hexane, ethyl acetate, ethanol and methanol; sequential extraction has been carried out starting from low polarity to high polarity. ii. Powdered plant material of 50 grams is soaked in 200 ml of hexanes in a stoppered container and placed on orbital shaker at 120 rpm for 72 h at room temperature. iii. Later, filter the mixes through Whatman No.1 filter paper and their concentrate under reduced pressure using a rotary evaporator.

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The same procedure (steps i, ii, iii) is followed for the other three solvents. The extraction process is carried out in triplicate with each solvent. The dried crude extract is collected and stored in amber vials and placed in refrigerator at 4oC. Column chromatographic fractionation of ethyl acetate extract Ethyl Acetate Extract (EAE) is subjected to Silica gel column chromatography for the isolation of phytoconstituents. A vertical glass column (40 mm width × 60 mm length) made of borosilicate material is used for the fractionation: iv. Rinse the column with acetone and let it dry for some time. v. Place a piece of glass wool at the bottom of the column with the help of glass root. vi. Add sea sand (50-70 particle size) to the top of the glass wool to 1 cm height. Sand particles are rinsed down with the solvent. vii. Then pour hexane into the column up to 3/4th level by closing stopcock. viii. Silica gel 200 g (60-120 mesh size) is used as packing material. Prepare silica slurry with hexane; this silica slurry is poured from the top of the column up to 2/3rd level with simultaneous draining of the solvent to aid proper packing of the column. ix. Then add sea sand to the top of the silica slurry to 1 cm height and sand particles are rinsed down with solvent. x. Mix 20 g of EAE with minimum quantity of hexane and pouring down from the top of the column along the side and rinse down with the solvent. xi. Add sea sand to the top of the extract to 1 cm height. See that solvent level up to 6 cm above the extract should be maintained in order to prevent the drying of the column. xii. Follow the gradient elution method to separate fractions from EAE using solvents from low polarity to high polarity (from hexane to methanol) in varying ratio. Flow rate is adjusted to 5 ml/min and 40 ml solvent collected for each fraction. TLC of fraction xiii. Collect fractions separately and subjected to TLC (20x20 cm aluminium sheets coated with silica gel 60 F254) to detect the pressure of phytochemicals. xiv. Now spray TLC plates with vanillin-conc. H2SO4 spray (15 g of vanillin in 250 ml of ethanol + 2.5 ml of conc. H2SO4) and dried at 100oC in hotair area for 20-30 min. The Rf value of each spot is calculated. xv. Fractions with the same Rf values are mixed together and condensed to dryness with a rotary evaporator; the dry weight of these combined fractions is calculated. HPLC analysis The condensed fractions and EAE are further analysed by HPLC for the presence of antioxidants phenolic compounds. This can be performed by two

Chromatography 325

methods using two different mobile phases selected on the basis of varying gradation of solvent systems in specific retention times and elute detections. In this protocol, analysis of all samples was performed using Shimadzu LC-10 AT VP, Luna 54 C18 reverse-phase analytical column (250 × 4.6 mm) with binary gradient mode, SPD-M10A VP photo diode array detector (PDA); injection volume 20 µl, total flow 1 ml/min, column oven temperature 25oC and detection wavelength 280 nm; 55 mg of EAE and each fraction dissolved in 3 ml of methanol for the analysis. Solvents to be used for mobile phases are previously filtered through Millipore and degassed prior to use. Quercetin, ascorbic acid, benzoic acid, gallic acid, vanillin, resorcinol and catechol are used as standard solutions for the quantification of phenolic compounds. Method A HPLC analyses of ascorbic acid, benzoic acid, gallic acid, vanillin, resorcinol and catechol are performed by method A. Gradient elution of two solvents is used for the quantification of these compounds; solvent A: Acetontrile and solvent B: phosphoric acid 0.1% in water. Gradient elution program begins with 92% of solvent B and it is held at this concentration for 0-35 min; then followed by 78% of solvent B for the next 35-45 min. Total run time is 45 min only. Method B This method is used for HPLC analysis of quercetin. Gradient elution of two solvents is used for the quantification of quercetin; solvent ‘A’ with methanol, solvent ‘B’ 4% acetic acid. Gradient elution program begins with 100% of solvent ‘B’ held for 0-4 mins, then followed by 50% of solvent ‘B’ for 4-10 min. Then it is reduced to 20% of solvent ‘B’ for the next 10-20 min. And further increased to 50% of solvent ‘B’ for the next 20-22 min. Total run time is 20 min. Results: HPLC analysis indicates the presence of phenolic compounds, viz., ascorbic acid, quercetin, gallic acid, resorcinol, catechol, vanillin and benzoic acid with specific retention times.

15.7.4 GC-MS Screening of Solvent Extracts of Cymbidium Aloifolium (L.) Sw. (Orchidaceae) (Rampilla and Khasim, 2020) Equipment and its conditions Agilent GC-MS (Model-5975C inert MSD with Triple-Axis Detector, USA) equipped with an HP-5 MS fused capillary column (30 m × 0.25 mm i.d., film thickness 0.25 µm) has been employed. While operating the equipment following chromatographic conditions are employed: Helium as the carrier gas, the flow rate of 1 ml/min; and the injector is operated at 200oC and column oven temperature is programmed as a rate of 10oC/min injection

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mode. Following Mass Spectroscopy conditions are used: ionisation voltage of 70 eV; ion source temperature of 250oC; interface temperature of 250oC; the mass range of 50-600 mass units. Soxhlet apparatus Vacuum rotary evaporation Reagents & materials n-hexane, methanol What man no. 41 filter paper Collection of plant material and sample preparation i. The fast green leaves of Cymbidium aloifolium growing on Borassus flaballifer host plant are collected from their natural habitat and packed in polyethylene bags. ii. The plant samples are shade dried and pound into fine powder and stored in air-tight polythene bags unit use. iii. The dried leaves powder (150g) is extracted in methanol at room temperature by using soxhlet extractor for 12-18 h. iv. The crude methanol extract is prepared and evaporated by vacuum rotary evaporation under reduced pressure. v. The crude extract is diluted with water and extracted with n-hexane. The extract is filtered using What man no.41 filter paper to obtain particlefree extract. The residue is re-extracted twice with solvent to obtain final extract. vi. The sample of 2 µl is injected into the GC-MS instrument for phytochemical analysis. vii. Interpretation of GC-MS data is carried out using the National Institute of Standard and Technology (NIST) database library 2.0 version which has more than 62,000 patterns. The spectrum of the unknown component is compared with the specimen of the known one stored in NIST library. Result: Based on retention time and peak area (Table 15.4), some of compounds such as n-hexadecanoic acid (Fig. 15.5a), 9, 12-Octadecadienoic acid (Z,Z) (Fig. 15.5b), 9,12,15-octadecatrienoic acid (Z,Z,Z), Octadecatrienoic acid (Fig. 15.6a) and phytol (Fig. 15.6b) have been recorded. Table 15.4. Compounds recorded in n-hexane extract of C. aloifolium Compound n-Hexadecanoic acid 9, 12-Octadecadienoic acid (Z,Z) 9,12,15-Ocxtadecatrienoic acid (Z,Z,Z) Octadecanoic acid Phytol

Retention time

Peak area %

50.409 52.803 52.872 53.110 52.538

22.6956 28.5815 38.3769 5.7647 4.5813

Chromatography 327

Fig. 15.5a. n-hexadecanoic acid; b. 9, 12-Octadecadienoic acid (Z,Z) (Courtesy: Rampilla and Khasim, 2020).

Fig. 15.6a. Octadecatrienoic acid; b. Phytol(Courtesy: Rampilla and Khasim, 2020).

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The identified various bioactive compounds have therapeutic properties that can be useful for the treatment of various diseases. These compounds reported from this investigation have some phylogenetic significance too.

16 Phytochemical Methods Secondary metabolites including primary metabolite products possess biological activity and are distributed across the plant kingdom. These chemical compounds are referred to as phytochemicals. They provide health benefits for humans, and these are classified under macro and micronutrients (Haster and Blumberg, 1999). Further, these phytochemicals act in plant defense against pathogen attacks and diseases (Mathai, 2000). Nearly 4,000 phytochemicals have been reported and studied for their protective function, and physical and chemical characterization (Meagher and Thomson, 1999).

16.1 Basic Equipment for Phytochemical Studies i. Separatory funnel Generally, the separatory funnel is used in liquid-liquid extractions to separate (partition) the components of a mixture into two immiscible solvent phases of different densities. ii. Rotary evaporator It is used to remove solvents efficiently and gently from samples by evaporation. The purpose of distillation is to separate a given mixture into its components based on their respective volatilities, through the process of evaporation and condensation (liquid-gas-liquid). The main purpose of the rotary evaporator is as follows: i. to concentrate non-volatile components in a mixture ii. to extract the volatile aroma and flavour molecules from mixtures gently and at low temperatures iii. Reflux Reflux is a distillation technique involving the condensation of vapours and the return of this condensate to the system from which it originated. It is used to supply energy to reactions over a long period. iv. Soxhlet extraction When a compound of low solubility needs to be extracted from a solid mixture a Soxhlet extraction can be carried out. The technique places specialized glassware in between a flask and a condenser. The refluxing solvent repeatedly washes the solid extracting the desired compound into the flask.

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A Soxhlet Extractor has three main sections such as a percolator (boiler and reflux) which circulates the solvent, a thimble (usually made of thick filter paper) which retains the solid to be laved, and a siphon mechanism, which periodically empties the thimble.

16.2 Steps Involved in Plant Collection Various steps are involved in plant sample collection. These are explained in the proceeding sections (Fig. 16.1).

16.2.1 The Plant Material Collection and Identification Plants that are useful to mankind, the pharmaceutical industry, and taxonomic importance, should be taken up for phytochemical analysis. Ideally, fresh plant materials should be collected and plunged into boiling alcohol immediately after their collection (Harborne, 2013). Alternatively, plants may be shade dried at room temperature before extraction. Once thoroughly dried, these are kept stored under safe conditions for quite a long period. In fact, for the analysis of alkaloids, flavonoids, quinones and terpenoids, herbarium plant tissues (dating back many years) have been used (Harborne, 2013). In this context, Sanford and Heinz (1971) observed that the myristicin content of nutmeg Myristica fragrans fruits increased slowly on storage, while the more volatile β-Pinene content decreased over time. On the contrary, flavonoids, and alkaloids in the herbarium specimens are remarkedly stable over time; therefore, the leaf sample of Strychnos nux-vomica originally collected in 1675 still contained 1-2% by weight of alkaloid (Philipson, 1982). For phytochemical studies, botanical identification of plants is needed, and it should be authenticated by plant taxonomists. Regional floras are referred to and helpful in the identification of plants. For further purposes, it is a common practice to deposit the herbarium specimen in the recognized herbaria for future reference (see chapter 9).

16.2.2 Powdering After complete drying of moisture, the plant samples are to be powdered. There are different types of powdering, they include the following: i. Grinding can be done by milling in an electric grinder or by a spice mill or it can also be done in mortar or pestle. ii. Grinding increases the efficiency of the extraction due to the increased surface area of the plants. The decrease in the surface area can lead to the dense packing of the material. iii. Milling the plants into a fine powder is always ideal but if they are too fine this affects the solvent’s flow and produces more heat which could degrade some thermolabile compounds.

Phytochemical Methods 331

Fig. 16.1. Flowchart of plant collection, identification, and solvent extraction.

16.3 Choice of Solvent The solvent that is being used for the extraction process is very important in determining the biologically active phytochemicals from the plants. These solvents must be less toxic, easy to evaporate in less heat, should preserve the compounds in them, and should not dissociate them. The various solvents commonly used for extraction include: i. Water: It is a universal solvent; plant extracts with anti-microbial activities are usually extracted with water. But the organic solvents give consistent results in anti-microbial activities when compared to water. Water-soluble compounds in the extract cannot give significant results. ii. Alcohol: The alcoholic extracts of plants show more activity than aqueous extracts due to the presence of higher amounts of polyphenols. This is because of the higher cell wall and seed degradation by the alcohols that release the polyphenols which will be degraded in the case of aqueous extracts. But ethanol is more microbicidal than water. More bioactive compounds are extracted in 70% ethanol than pure ethanol. Ethanol is also found easier to extract intracellular ingredients from plant materials. Polar solvents like methanol, ethanol, and their aqueous mixtures are used for the extraction of phenolic compounds. The addition of water to alcohol will improve the rate of extraction. Methanol is more polar, but it is unsuitable for extraction due to its cytotoxic nature (Balamurugan et al., 2019). iii. Acetone: Acetone dissolves many hydrophilic and lipophilic compounds from the plants, and it is miscible with water. It is low toxic and volatile and it is used for extracting antimicrobial activities. Extracting tannins and other phenolic compounds is done with acetone. They are also used to extract saponins.

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iv. Chloroform: Terpenoid lactones are obtained from barks by extraction with chloroform. Tannins and Terpenoids are treated with less polar solvents. v. Ether: They are used for the extraction of coumarins and fatty acids. The solvent used for the extraction of medicinal plants is also known as the menstruum. The choice of solvent depends on the type of plant, the part of the plant to be extracted, the nature of the bioactive compounds, and the availability of solvent. In general, polar solvents such as water, methanol, and ethanol are used in the extraction of the polar compound, whereas nonpolar solvents such as hexane and dichloromethane are used in the extraction of nonpolar compounds. During liquid-liquid extraction, the conventional way is to select two miscible solvents such as water-dichloromethane, water-ether, and water-hexane. In all the combinations, water is present because of its high polarity and miscibility with an organic solvent. The compound to be extracted using liquid-liquid extraction should be soluble in organic solvents but not in water to ease the separation. Furthermore, the solvent used in extraction is classified according to their polarity, from n-hexane which is the least polar to water the most polar (Table 16.1). During fractionation, the selected solvent is added according to the order of increasing polarity, starting from n-hexane, the least polar to water with the highest polarity. If you like to select five solvents during fractionation, the usual practice is to choose two solvents with low polarity (n-hexane, chloroform), two with medium polarity (dichloromethane, n-butanol), and one with the highest polarity (water). Table 16.1. Various solvents of extractions arranged according to the order of increasing polarity S.No.

Solvents

Polarity

1.

n-Hexane

0.009

2.

Petroleum ether

0.117

3.

Diethyl ether

0.117

4.

Ethyl acetate

0.228

5.

Chloroform

0.259

6.

Dichloromethane

0.309

7.

Acetone

0.355

8.

n-Butanol

0.586

9.

Ethanol

0.654

10.

Methanol

0.762

11.

Water

1.000

Phytochemical Methods 333

16.4 Extraction Methods Extraction involves the separation of the medicinally active constituents of plant tissues from the active or inert component by using solvent(s) and by using one of the standard extraction procedures. The products obtained from plants are relatively impure liquids, semisolids, or powders, intended only for oral or external use. Methods of extraction can be divided into cold methods and hot methods.

16.4.1 Cold Extraction Methods It is the process of extracting a substance from a mixture via a cold solvent. The procedure is carried out at room temperature (15-25 ºC). i. Maceration This simple widely used procedure involves leaving the pulverized plant to soak in a suitable solvent in a closed container. Simple maceration is performed at room temperature by mixing the ground plant powder with the solvent (plant powder solvent ratio: 1:5 or 1:10) and leaving the mixture for several days with occasional shaking or stirring. This method is best suitable for thermolabile compounds. The main disadvantage of maceration is that the process can be quite time-consuming, taking from a few hours up to several weeks. There are three types of maceration, simple (single maceration), double maceration and triple maceration (Agrawal and Paridhavi, 2012). In double and triple maceration, the menstruum is equally divided into two and three parts respectively and used. The purpose is to obtain a more extractable yield. ii. Percolation Percolation (from Lat. percōlāre, to filter) concerns the movement and filtering of fluids through porous materials. The powdered plant material is soaked initially in a solvent. In a percolator, the additional amount of solvent is then poured on top of the plant material and allowed to percolate slowly (dropwise) out of the bottom of the percolator. If necessary, the same material can be re-extracted with a second solvent to yield more extract.

16.4.2 Hot Extraction Methods i. Infusion Infusion is applicable for soft drugs only and drugs containing watersoluble constituents (Agrawal and Paridhavi, 2012). The drug is mixed with water. It is allowed to stand for 30 minutes. Shaking or stirring is done if required. Drug + H2O

Filtrate

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Here heat is not applied, and it is filtered. The filtrate is called an infusion. The infusion should be used within 24 hours. Sometimes 20-25% ethanol is used as a preservative and stored for quite a long period, e.g., compound infusion of gentian and chirata (Agrawal and Paridhavi, 2012). ii. Decoction A decoction is applicable only for water-soluble and heat-stable drugs obtained from hard and woody materials. The term decoction is derived from the Latin ‘decoquere’ (meaning to boil down), de = from; coquere = to cook. A decoction is a method of extraction by boiling, dissolved chemicals, from hard plant material, which may include stems, roots, bark, and rhizomes on a source of heat or direct flame then agitating until the active constituents will be dissolved in the solvent. Here the solvent used depends on the active constituent and source of heat. Drug + H2O

heating for 15-20 minutes

filtrate.

The filtrate is called decoction. It should be consumed within 24 hours. iii. Digestion Digestion is a modified form of maceration. Here heat is applied for macerated material. It is suitable for heat-stable substances only. iv. Continuous hot extraction methods Reflux condenser: Plant material is immersed in a solvent in a roundbottomed flask connected to a condenser. The solvent is heated until it reaches its boiling point. As the vapour is condensed, the solvent is recycled into the flask. Soxhlet apparatus: The plant powder is placed in a cellulose thimble in an extraction chamber, which is placed on top of a collecting flask beneath a reflux condenser. A suitable solvent is added to the flask, and the setup is heated under reflux. When a certain level of condensed solvent has accumulated in the thimble, it is siphoned into the flask beneath. The main advantage of Soxhlet extraction is that it is a continuous process for the extraction of active constituents decomposed by direct heat.

16.4.3 Aqueous Alcoholic Extraction by Fermentation Some medicinal preparations of Ayurveda (like asava and arista) adopt the technique of fermentation for extracting the active principles. The extraction procedure involves soaking the crude drug, in the form of either a powder or a decoction (kashaya), for a specified period, during which it undergoes fermentation and generates alcohol in situ; this facilitates the extraction of the active constituents contained in the plant material. The alcohol thus generated also serves as a preservative. If the fermentation is to be carried out in an earthen vessel, it should not be new: water should first be boiled in the vessel. In large-scale manufacture, wooden vats, porcelain jars, or metal vessels are used in place of earthen vessels. Some examples of such preparations are

Phytochemical Methods 335

karpurasava, kanakasava, and dasmularista. In Ayurveda, this method is not yet standardized but, with the extraordinarily high degree of advancement in fermentation technology, it should not be difficult to standardize this technique of extraction to produce herbal drug extracts.

16.4.4 Counter-current Extraction In counter-current extraction (CCE), wet raw material is pulverized using toothed disc disintegrators to produce a fine slurry. In this process, the material to be extracted is moved in one direction (generally in the form of a fine slurry) within a cylindrical extractor where it comes in contact with the extraction solvent. The further the starting material moves, the more concentrated the extract becomes. Complete extraction is thus possible when the quantities of solvent and material and their flow rates are optimized. The process is highly efficient, requiring little time and posing no risk from high temperatures. Finally, a sufficiently concentrated extract comes out at one end of the extractor while the mark (practically free of visible solvent) falls out from the other end (Handa et al., 2008).

16.4.5 Ultrasound Extraction (Sonication) The procedure involves the use of ultrasound with frequencies ranging from 20 kHz to 2000 kHz; this increases the permeability of cell walls and produces cavitation. Although the process is useful in some cases, like the extraction of Rauwolfia root, its large-scale application is limited due to the higher costs. One disadvantage of the procedure is the occasional but known deleterious effect of ultrasound energy (more than 20 kHz) on the active constituents of medicinal plants through the formation of free radicals and consequently undesirable changes in the drug molecules.

16.4.6 Supercritical Fluid Extraction (SFE) of Natural Products Supercritical fluid extraction (SFE) on an analytical and processing scale is quite widespread in the food industry for the extraction of fats and oils from seeds, foodstuffs, and other materials. It is an alternative method to reduce the use of organic solvents and to yield more extract. The factors to consider include temperature, pressure, sample volume, analyte collection, modifier (cosolvent) addition, flow and pressure control, and restrictors. Generally, cylindrical extraction vessels are used for SFE, and their performance is good beyond any doubt. In this method carbon dioxide (CO2) is used as extracting fluid. In addition to its favourable physical properties, carbon dioxide is inexpensive, safe, and abundant. But while carbon dioxide is the preferred fluid for SFE, it possesses several polarity limitations. Solvent polarity is important when extracting polar solutes and when strong analyte-matrix

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interactions are present. Organic solvents are frequently added to the carbon dioxide-extracting fluid to alleviate the polarity limitations. Of late, instead of carbon dioxide, argon is being used because it is inexpensive and more inert. Typically, supercritical carbon dioxide at pressures of at least 250 bar and a temperature of 20 and 40ºC conditions suitable for the extraction of oil from oil seeds of Helianthus annus and Brassica napus (Agrawal and Paridhavi, 2012).

16.4.7 Phytonics Process A new solvent based on hydrofluorocarbon-134a and a new technology to optimize its remarkable properties in the extraction of plant materials offer significant environmental advantages and health and safety benefits over traditional processes to produce high-quality natural fragrant oils, flavours, and biological extracts. Advanced Phytonics Limited (Manchester, UK) has developed this patented technology termed the “Phytonics process” (Handa et al., 2008). The products mostly extracted by this process are fragrant components of essential oils and biological or phytopharmacological extracts which can be used directly without further physical or chemical treatment. The properties of the new generation of fluorocarbon solvents have been applied to the extraction of plant materials. The core of the solvent is 1,1,2,2-Tetrafluoroethane, better known as hydrofluorocarbon-134a (HFC134a). This product was developed as a replacement for chlorofluorocarbons. The boiling point of this solvent is -25° C. It is not flammable or toxic. Unlike chlorofluorocarbons, it does not deplete the ozone layer. It has a vapour pressure of 5.6 bar at ambient temperature. By using modified solvents with HFC-134a, the process can be made highly selective in extracting a specific class of phytoconstituents. Similarly, other modified solvents can be used to extract a broader spectrum of components. The biological products made by this process have extremely low residual solvents. The residuals are invariably less than 20 parts per billion and are frequently below levels of detection. These solvents are neither acidic nor alkaline and, therefore, have only minimal potential reaction effects on the plant constituents. The processing plant is totally sealed so that the solvents are continually recycled and fully recovered at the end of each production cycle. The only utility needed to operate these systems is electricity and, even then, they do not consume much energy. There is no scope for the escape of the solvents. Even if some solvents do escape, they contain no chlorine and therefore pose no threat to the ozone layer. The waste biomass from these plants is dry and “eco-friendly” to handle (Handa et al., 2008).

16.5 Preparation and Analysis of Plant Extracts After selecting a suitable solvent, the below-given steps should be followed to extract chemical compounds:

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1. Grind the plant material into a fine powder using a mixer or mortar and pestle. 2. Place the powdered plant material in a container that should be sealed. 3. Add the solvent to the container until the plant material is completely submerged. 4. Seal the container and allow it to sit for 24-48 hours, shaking it occasionally. 5. After the extraction time has elapsed, open the container and filter the plant material from the solvent using a muslin cloth or filter paper. 6. Collect the filtered solvent in another container, the crude extract. 7. Further refine the extract by evaporating the solvent using a rotary evaporator or some other method.

16.5.1 Qualitative Phytochemical Screening of Crude Extracts The crude extracts obtained were used for the qualitative analysis of primary (carbohydrates and proteins) and secondary metabolites (alkaloids, flavonoids, phenols, saponins, steroids, tannins, and terpenoids). The preliminary phytochemical screening was performed by following the standard protocols formulated by Harborne (2013), Trease & Evans (1989, 1996) and Sofowara (1993). 16.5.1.1 Test for Carbohydrates

A. Molisch’s Test: i. Take 2 ml of the plant extract in a clean test tube. ii. Add 2-3 drops of Molisch reagent slowly. iii. Now add concentrated sulphuric acid along the sides of the test tube. iv. The acid forms a layer at the bottom. v. Note the junction of the two layers. Result: Formation of the violet ring indicates the presence of carbohydrates. B. Benedict’s Test: i. Take the plant extract to be tested in a clean test tube. ii. Add 5 ml of Benedict’s reagent to it. iii. Boil the solution for about 2 minutes. iv. Cool the solution and observe the solution. Result: Formation of a green, red or yellow precipitate indicates the presence of reducing sugars. 16.5.1.2 Test for Proteins

A. Biuret Test: i. Take a cleaned and dried test tube. ii. Add the plant extract into the test tubes.

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iii. Add 2 ml of Sodium hydroxide and 5 to 6 drops of copper sulphate solution to it. iv. Shake the test tube gently to mix the ingredients thoroughly and allow the mixture to stand for 4-5 minutes. Result: The appearance of bluish-violet colour indicates the presence of protein. B. Xanthoproteic Test: i. Take a cleaned and dried test tube. ii. Add the plant extract into the test tubes. iii. Add a few drops of concentrated Sulphuric acid and shake the test tube. iv. Heat the test tube gently on a bunsen burner. Result: The formation of a yellow precipitate indicates the presence of protein. 16.5.1.3 Test for Alkaloids (Dragendorff’s Test)

i. Take a cleaned, dried test tube and add 0.5 gm of plant extract. ii. Later add 5ml of 1% HCl, boil the mixture and filter it. iii. To the above filtrate add 2-3 drops of Dragendorff’s reagent. Result: The formation of a reddish-brown precipitate indicates the presence of alkaloids. 16.5.1.4 Test for Flavonoids (1% Aluminium Chloride (AlCl3) Test)

i. Take a cleaned, dried test tube and add a small quantity of plant extract. ii. Add a few drops of 1% AlCl3 to the plant extract. Result: The appearance of yellow colour indicates the presence of flavonoids. 16.5.1.5 Test for Phenols (10% Ferric Chloride (FeCl3) Test)

i. Take a cleaned, dried test tube and add a small quantity of plant extract. ii. Later add a few drops of 10% aqueous FeCl3 solution. Result: The appearance of blue or green colour indicates the presence of phenols. 16.5.1.6 Test for Saponins (Frothing Test)

i. Take 2 ml of the plant extract in a clean test tube. ii. Add about 5 ml of distilled water and mix well. Result: The formation of persistent froth indicates the presence of saponins.

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16.5.1.7 Test for Steroids (Liebermann-Burchard Test)

i. Take 1 ml of the plant extract in a clean test tube and dissolve it in 1 ml of chloroform and acetic anhydride. ii. Add a few drops of concentrated H2SO4 from the sides of the test tube. Result: The appearance of a brownish-green ring at the interface indicates the presence of steroids. 16.5.1.8 Test for Tannins (Iodine Test)

i. Take 2 ml of the plant extract in a clean test tube. ii. Add a few drops of Iodine solution. Result: The appearance of faint bluish colour indicates the presence of tannins. 16.5.1.9 Test for Terpenoids (Salkowski Test)

i. Take a few drops of the plant extract in a clean test tube and add 2 ml of chloroform. ii. Add concentrated H2SO4 to form a lower layer. Result: The appearance of reddish-brown colour at the interface denotes the presence of terpenoids.

16.6 Case Studies 16.6.1 Quantitative Analysis of Secondary Metabolites in Dendrobium aphyllum (Roxb.) C.E.C. Fischer (Rahamtulla, 2022) i. Estimation of alkaloids (Harborne, 2013) About 5 g of the powdered plant sample is transferred to a 250 ml beaker, and 200 ml of 10% acetic acid in ethanol was added and covered, allowed to stand for 4 hours. Later the sample is filtered, and the filtrate is brought to ¼th of its original volume in a water bath. Further, a few drops of conc. NH4OH is added to the extract until the precipitation was complete. The precipitate formed in the solution is washed with dilute NH4OH and the residue is transferred to petri plates, dried and weight is recorded. The weight is expressed in terms of gram % of alkaloids on a dry weight basis. ii. Determination of total phenols (Kaur and Kapoor, 2002) The total phenolic content of plant extracts is measured by the FolinCiocalteau reagent method with minor modifications (Kaur and Kapoor, 2002). About 100 µL of plant extract (1mg/mL) is added to 900 µL of methanol and 1 mL of Folin-Ciocalteau reagent. Later 1 mL of 20%

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(w/v) of Na2CO3 solution is added to the above mixture. The mixture is incubated at room temperature for 30 min. in dark. Afterward, the absorbance is measured at 765 nm. The total phenolic content is expressed in terms of gallic acid equivalent (µg/mg of extract). iii. Determination of total flavonoids (Liu et al., 2007) The total flavonoid content of plant extracts is assessed by the Aluminium chloride method with slight modification (Liu et al., 2007). About 500 µL of plant extract (1mg/mL) is added to 0.5 mL of methanol and 1 mL of 5% (w/v) sodium nitrite solution. Subsequently, about 1 mL of 10% (w/v) Aluminium chloride solution was added to the above mixture and shaken well. Later 100 µL of 1 M NaOH solution is added to the above mixture, and its absorbance is measured at 510 nm. The total flavonoid content is expressed in terms of (µg/mg of extract) quercetin equivalent. Results: The total alkaloid content of different plant parts of D. aphyllum is determined quantitatively (Table 16.2). Methanol leaf extract showed the highest alkaloid content (1.232 mg/gm dry weight), followed by root and stem extracts. Whereas phenolic content is high in methanol root extract (327.10 µg/mg of gallic acid equivalent) followed by leaf and stem extracts. In the case of flavonoid content is high in the methanol root extract (65.06 µg/mg of quercetin equivalent (QE)), followed by methanol leaf and stem. Table 16.2. Quantitative estimation of alkaloids, total phenols and total flavonoids in ethyl acetate and methanol extracts of different parts of D. aphyllum Solvents used

Ethyl acetate

Methanol

Plant parts Used

Total alkaloid content (mg/g) dry weight

Total phenolic content (µg/mg of gallic acid equivalent)

Total flavonoid content (µg/mg of quercetin equivalent)

Leaf

0.890

133.43

4.67

Stem

0.342

244.54

16.71

Root

0.702

180.17

11.14

Leaf

1.232

293.87

44.56

Stem

0.942

286.60

14.92

Root

1.004

327.10

65.06

Apart from the above methods, various separation techniques such as Column Chromatography, Gas Chromatography, and HPLC have been employed for the separation of various compounds (see chapter 15). Further, spectral data obtained from Ultraviolet spectroscopy, Infrared absorption spectroscopy, and Mass spectroscopy are useful for the identification of these chemical compounds (see chapter 7).

17 Cytological and Vital Staining Techniques The smear or squash technique is the most suitable for cytological studies. This technique has a great advantage over the sectioning method as the entire process is rapid and more suitable for critical observations. In properly prepared smears and squashes, one can make studies on separated single cells; besides this, cells that are released undergo much enlargement in volume, affording wider space for the chromosomes to become scattered. The only disadvantage of this method, when specially applied to somatic chromosomes, is that the individual cells being released from one another shift from their original site and the original topography is altered.

17.1 Cytological Techniques The terms ‘smears’ and ‘squashes’ are often loosely used as one process but strictly speaking, these are two different processes (Sharma and Sharma, 1972). In smears, the cells are directly spread on the slide without fixation or without pre-treatment to secure cell separation. Pollen mother cells from anthers are the most convenient for making smears. In squashes, on the other hand, special treatments are needed to dissolve the middle lamellae, so that cells are freed from the compact mass of tissue. This treatment may be carried out after fixation or even after staining. After passing through the required steps, softened bulk material or tissue is neatly squashed on the slide by applying pressure or tapping with a needle over the cover slip. The mitotic behaviour of chromosomes can be studied by squashing the root tips.

17.1.1 Smears Smear method is a very quick and simple method for studying mitosis and meiosis. In this method, loosened cells are uniformly spread on the chemically cleaned slide without causing any injury to them. No adhesive is required for affixing the cells. The cells may be killed and fixed, dehydrated and stained properly and finally made permanent. The smear method is limited to those cells which are not firmly united to one another by middle lamellae. Microsporocytes of higher plants are

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of this sort, and they can be easily separated with this method. However, microsporocytes of some plants (e.g., Acacia and some orchids, in these pollinia are present) and testis cells of many animals cannot be satisfactorily smeared but some other methods should be employed for these specimens. Before smear preparation, microslides are chemically cleaned in the following solution: Potassium dichromate Conc. sulphuric acid Distilled water

20 g 100 ml 100 ml

Dissolve potassium dichromate in distilled water and add sulphuric acid slowly. Cool the mixture as heat is produced during the process. Clean the slides in this mixture and later wash thoroughly in running tap water. Immerse the slides for sometime in strong alcohol containing some ammonia, then running tap water and dry them with muslin cloth. 17.1.1.1 Preparation of Smears from Anthers (Microsporangium)

In normal course, anthers are taken out from the flower bud at least 15 days before the shedding of pollen grains. The anthers of Tradescantia may be fixed during mid-day to show good meiosis. Two ways of preparing smears are given below (Johansen, 1940): (i) Take out the anther from the flower bud and place it at one end of a chemically cleaned microslide and cut it transversely into small pieces with the help of a razor blade. Put a scalpel flat over the anther and quickly draw the scalpel lengthwise, applying the pressure gently to get a thin film of microsporocytes. (ii) Another method is to place pieces of anther at the centre of the slide. Hold another slide crosswise over the first, exert gentle pressure to crush the anther pieces and spread the microsporocytes by drawing it back and forth. Smears prepared by the above methods are now killed and fixed in suitable fixative. Besides Radolph’s modified Navashin’s fluid, the following fluid is also suitable for chromosome studies: 10% aqueous chromic acid 2% osmic acid in 2% chromic acid 10% aqueous acetic acid Distilled water To the above mixture add 1% saponin.

1 ml 7.5 ml 10 ml 41.5 ml

17.1.1.2 Making Permanent Mounts

i. Take a petri dish (about 16 cm in diameter), place two slender glass rods

Cytological and Vital Staining Techniques 343

ii. iii. iv. v. vi. vii. viii. ix. x. xi.

(cut into proper length) in the petri dish parallel to each other and pour the killing solution into the dish, just sufficient to immerse the rods. Invert and place the slide over the glass rod in such a way that the smeared surface should come in contact with the killing fluid. Keep it for 10 minutes or more by covering another petridish. Later take out the slide and wash in slow running water for 15 mins. Immerse the smear in freshly prepared 1% aqueous solution of methyl violet 2B (do not use boiled solution) or any other suitable stain for 15 mins. Rinse the slide in water for 30 minutes. Differentiate in 70 and 95% alcohol (containing 0.5% picric acid) for 10 secs in each grade. Dip the slide in 95% alcohol containing three drops of ammonia for 15 secs. Completely dehydrate in absolute alcohol for 10 seconds. Completely differentiate in clove oil for 10-15 seconds keeping the slide moving back and forth. Wash in xylene containing few drops of absolute alcohol. Transfer to pure xylene for 2 hrs and mount in Canada balsam.

Belling’s Iron-acetocarmine method Johansen (1940) has modified one of the Belling’s formulae for the preparation of iron-acetocarmine: Carmine dye Glacial acetic acid Aqueous ferric acetate Distilled water

1g 90 ml few drops 110 ml

Add 110 ml distilled water to 90 ml glacial acetic acid and heat the solution upto boiling point. Now stop boiling and add 1 g carmine dye, cool to ice temperature and decant. Add a few drops of aqueous ferric acetate to the mixture till the colour appears wine-red (take care not to add more ferric acetate which causes precipitation). There are several ways of using iron-acetocarmine for different materials. Some of the methods are described below (refer to Johansen, 1940): Method I i. Place the fixed anther in a drop of acetocarmine on a clean slide. ii. Remove excess fluid with blotting paper. iii. Put fresh drop of stain over the material and squeeze out the microsporocytes by applying slight pressure; remove anther walls and other debris. iv. Put the cover slip and remove excess stain with blotting paper. v. Seal the coverslip with molten wax.

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vi. Leave the preparation for few days to facilitate the penetration of stain into microsporocytes. Method II i. Follow above steps (i) to (iii) as in section 10.1.1. Carnoy’s solution may be used as fixative here (see section 3.1.2.2.D for Carnoy’s preparation). ii. Put a drop of acetocarmine on the smear and place a cover slip on it. iii. Seal the cover slip with molten wax. Method III i. Keep anthers in the vial containing 1:3 glacial acetic acid and absolute alcohol mixture for 12 hrs or some weeks to improve the staining ability of chromosomes. ii. Transfer the anther to clean slide and put a drop of acetocarmine over it. iii. Squeeze out the microsporocytes by applying pressure gently, remove anther wall and other debris. iv. Gently heat the slide over the flame for a second so that the spread cells flatten. Do not allow to boil. v. Transfer the slide to a jar containing equal parts of acetic acid and absolute alcohol for 5 mins. vi. Later pass through the following mixtures keeping for a minute in each: (a) 1:3 acetic acid and absolute alcohol (b) 1:9 acetic acid and absolute alcohol (c) 1:1 absolute alcohol and xylene vii. Mount with thin balsam (diluted with xylene). 17.1.1.2 Preparation of Smears from Root Tips

The root tips are obtained from germinated seeds. Seeds are allowed to germinate in wet blotting paper discs or sand. The sand is thoroughly washed to make it free from any debris and seeds are sown in this sand. For teaching purpose, the seeds of Medicago falcata (ver. Methi) are suitable because they germinate soon, and their chromosomes are bigger in size. The young root tips are cut during morning around 7-9 a.m. During this time, the cells are actively dividing mitotically. The onion root tips are also suitable for teaching purpose. Root tips of onion (Allium cepa) are obtained by placing a bulb at the mouth of a glass jar full of water, so that the basal part of the onion bulb is always in contact with water. It is placed in dark to produce root tips for 3-4 days. Root tips are cut during morning hours. Several cytological fixatives are being used but Radolph’s modified Navashin’s fluid (CRAF) has been found to be better for fixing root tips (for CRAF see section 3.1.2.2C). Some of the methods for both temporary and permanent preparation of smears are described below:

Cytological and Vital Staining Techniques 345

Method I (temporary preparation) i. Take out root-tip, thoroughly wash to remove fixative and cut into small pieces, about 0.5 mm long. ii. Place the root tip with two drops of acetocarmine on microslide and tease it lengthwise with the help of needles. iii. Heat the slide over the flame for a second and repeat the process 4-6 times to allow the cells to spread and flatten. Take care that it should not be boiled. iv. Put a cover slip and wrap around the slide covering the coverslip with blotting paper. Now apply pressure gently with finger over the blotting paper to separate the cells. Take care to see that neither the cover slip is broken nor does it move sideways. v. Remove the blotting paper and seal the cover slip with nail polish or .paraffin wax. Method II (Warmke’s 1935 temporary preparation method) i. Fix the root tips in 1:3 acetic acid and absolute alcohol for 12 hrs to several weeks to improve the stainability of chromosomes. ii. Take out the root tip from the fixative and place it in a mixture containing equal parts of 95% alcohol and concentrated HCl for 5-10 mins. This serves to dissolve pectin substances of middle lamellae. iii. Transfer the root tip to Carnoy’s fluid (with chloroform) for 5 mins or more (the tissues made soft by acid treatment, will become hard again). iv. Put the root tip with a drop of acetocarmine on a clean slide and press it gently with the help of scalpel to separate the cells. v. Place the cover slip over the material. vi. Heat the slide over the flame, each time not more than a second. vii. Seal the edge of cover slip with nail polish or paraffin wax. Method III (see also McClintock’s permanent preparation method, Johansen, 1940) Follow steps (i)-(vi) of above Warmke’s method for preparation of smear: i. Keeps the slide in a dish containing 10% aqueous acetic acid. ii. Remove the cover slip carefully with the help of forceps when it gets slightly raised after dipping in acetic acid. iii. Transfer the slide and cover slip carefully to the jar containing equal parts of acetic acid and absolute alcohol for 5 mins. iv. Pass through the following mixtures keeping for a minute in each: (a) 1:3 acetic acid and absolute alcohol (b) 1:9 acetic acid and nine parts absolute alcohol (c) 1:1 absolute alcohol and xylene v. Mount it with thin balsam (dilute with xylene) covering with same cover slip over the smear.

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Method IV (Hillary’s permanent preparation method, Johansen, 1940) Preparation of Leuco-basic fuchsin Basic fuchsin 1g Boiling distilled water 200 ml N/I HCl 30 ml Potassium metabisulphite 3g + Norit or activated charcoal 0.5 g Dissolve 1 g basic fuchsin in 200 ml boiling distilled water. Shake well and cool it at 50°C. Then filter, add 30 ml N/I HCl, 3 g potassium metabisulphite, shake well and leave for 24 hrs in dark and finally add 0.5 activated charcoal and shake well for a minute. Filter quickly through filter paper and keep the colourless stain in dark for future use.

Procedure i. Fixed root tips should be washed thoroughly in water. ii. Stain with leuco-basic fuchsine for an hour or more. iii. Dehydrate through dioxan giving three changes. iv. Take the root tip on a clean slide and put a drop of balsam (diluted with dioxan). v. Tease the root tip lengthwise with the help of needle. vi. Add more balsam if necessary, put a cover slip and gently press it with the finger to allow the cells to spread. This method may also be used for preparing anther smears. 17.1.1.3 Smears and squash preparations for meiosis and mitosis studies in plants

Rapidity of fixation and handling of the material is the principle advantage of smear and squash preparation over hand/microtome sections. The smears will not only save time but increase the efficiency of the worker to examine single layer of cells in intact condition. Material: The best material for laboratory studies of smear and squash preparation is Allium cepa and A. Sativus as mentioned earlier. However, for meiotic studies Phlox drummondii and Lathyrus species are also preferable. Squash Preparation: This method is useful to study the chromosomes and proved to be simple and best available method. Squash method was first used by Schneider and adapted for chromosome work by Belling in 1921. A pre-treatment by certain chemicals like para-dichlorobenzene, alphabromonaphthalene, etc., is given to the material to get good preparations with all separated chromosomes. Fixation: Fixation of the material to be studied is an important stage because, it helps in reducing the staining of the cytoplasm and thus giving

Cytological and Vital Staining Techniques 347

a clear contrast of chromosomes. Even though different workers adopted different fixatives, 1:3 Acetic acid: Alcohol has been giving interesting results. Storage of the material: The material may be stored in 70% alcohol at 0-40 upto 60 days. But later, the stain will suffer. Staining: 1% acetocarmine is generally used for studying meiosis in PMCs while Fuelgen’s stain is suitable for studying the mitotic chromosomes. The method for Fuelgen’s staining is as follows: i. ii. iii. iv. v.

Cut the tip of actively growing roots. Pre-treat them with 0.025% colchicine for 3 hours. Wash and dry them. Then transfer the roots to 1:3 acetic-alcohol. After 24 hrs of fixation wash the root tips and hydrolyse them with 1N HCl at 600 for 8-10 min. vi. Again, wash them dry and transfer the root tips to Basic Fuschin (Fuelgen’s stain) and keep them for 45 mins in it, in dark. vii. Wash them and put in distilled water. viii. Now the root tips (only the meristematic zone) are stained and ready for squash preparation. Spreading: The material is to be taken over a slide (i.e., anther for meiosis and root tip in case of mitosis). Crush the material with the help of a brass rod in a drop of acetocarmine stain and remove the debris with the help of forceps and put a cover slip over it. Gentle heating over a spirit lamp flattens the cells, sticks them to the slide and coverslip, and spread the chromosomes. The degree of spreading is proportional to the pressure of the coverslip partly to the amount of stain fixative used and also the pressure applied on the coverslip. Unless all the debris is removed, the possibility of well spread chromosomes becomes difficult. Gentle finger-tip pressure on the cover slip under the filter paper folds, is useful for further improvement of the preparation. It is always advisable to use appropriate amount of stain. Mordanting: A trace of ferric chloride may be added to the fixative or carmine stain is suggested to get clear preparation. The mordant helps in softening and makes the material plastic thereby facilitates easy flattening. Storage of slides: In many cases duration of 4 days improves the quality of preparations. The edges of the coverslips should be sealed with rubber solutions or by bee-wax. They should be kept in refrigerator at 40oC. Permanent method: Different researchers have provided several strategies for making the slide permanent, however no single type/method for making permanent slides is best for all materials. However, in general practice the following method is used: i. Remove the rubber solution put over the edge of the cover slip. ii. Invert the slide in a dish of 1:3 acetic acid-alcohol. iii. Now take the slide and pass it through series of mixture of acetic acid:

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alcohol in the ratio of 1:3, 1:6, 1:9 and twice through absolute alcohol in series. Then let the alcohol evaporate itself. Pass the slide over a spirit lamp to ensure equal spreading of mounting medium. Similarly, pass the cover slip and finally mount in euparol taken on a slide. The slide must be kept in oven at 600oC for 10-12 days.

17.1.2 Squashes In squash preparation, the first step is the softening of the tissue. The different schedules can be divided into two categories: these are: (a) softening performed prior to staining, and (b) softening, clearing and staining performed in the same fluid. In the first category, various chemical agents have been employed by different authors including the use of enzymes. Chemical agents: The widely used chemical agent for softening the tissue is diluted hydrochloric acid. In Feulgen staining, this step is essential for securing Schiff s reaction for aldehydes. In addition to liberating aldehydes of sugar, normal hydrochloric acid at 58°C serves two important purposes, namely dissolution of middle lamella, thus helping in cell separation and clearing of the cytoplasm. These two properties of hydrochloric acid have been employed in other schedules also after fixation. If dilute hydrochloric acid (10%) is used, for best results, the treatment should be carried out in slightly warm temperature such as 58-60°C for 4-5 mins until softening, later acid is washed off thoroughly either in 45% acetic acid solution or tap water before staining. Softening and maceration of the tissue can also be achieved during fixation by using the mixture consisting of equal parts of 95% alcohol and concentrated hydrochloric acid as fixative. No warming is needed and within 5 mins of treatment the tissue becomes fixed and softened. If necessary, hardening of tissue may be carried out by keeping it in Carnoy’s fluid for 10 mins. For hard materials, alcohol and hydrochloric acid mixed in the 3:1 ratio is more effective. Though there are some disadvantages, trichloroacetic acid can also be employed for cytoplasm clearing and softening of the tissues (for detail discussion refer to Sharma and Sharma, 1972). Chromic acid is also employed for maceration of tissues, but it is time consuming and may take at least 24 hrs to soften the tissue. Since a strong concentration of chromic acid may injure chromosome parts, prolonged treatment with a diluted chromic acid (1%) is generally preferred (Sharma and Sharma, 1972). This procedure is generally not recommended for the study of chromosomal morphology.

Cytological and Vital Staining Techniques 349

In addition to acids, other reagents such as hydrogen peroxide and sodium hydroxide are also employed for softening of the tissues. Hydrogen peroxide is used with a trace of sodium or lithium carbonate added and the middle lamella is initially attacked. Satisfactory dissolution of the middle lamella has been obtained by using the mixture consisting of equal proportion of saturated aqueous ammonium oxalate solution and hydrogen peroxide. Ford (cited in Darlington and La Cour (1960) has recommended it for meristematic tissue fixed in osmic acid. However, for comparatively stiff materials, this reagent is not suitable for softening. Alkali treatment, prior to squashing, was used by Tandler (cited in Sharma and Sharma, 1972) in plants. Enzyme treatments: The most reliable method for softening and clearing without causing injury to cellular parts is enzyme treatment (see also chapter 14 for isolation of protoplast). The use of pectinase for dissolution of middle lamella has been applied by McKay and Clarke (cited in Sharma and Shanna, 1972). Later 5% pectinase (macerozyme) solution in 1% aqueous peptone has been employed for this purpose. But this procedure is time consuming and takes 2-5 hrs of treatment. Sharma and Sharma (1972) noted that a 2% aqueous solution of pectinase, if applied for half an hour at 37°C, results in considerable softening of plant materials. In addition to the above-mentioned methods, there are a number of schedules in which softening is carried out together with staining. In some cases, the tissue is heated after fixation, over a flame for a few seconds in a mixture of the acidic dyes and hydrochloric acid. The commonly used acetic solutions of dyes are acetic-orcein, acetic-lacmoid and acetic-carmine (go through the next part of the chapter). 17.1.2.1 Other protocols

A. Enzyme digestion method for root-tip cytology (Lattier et al., 2017) In this enzyme digestion method, the root tips of gymnosperms and angiosperms were taken and, these were prefixed, postfixed and finally enzyme digestion had carried out (Lattier et al., 2017). This technique reduces the handling of the roots allowing for long-duration enzyme digestion.

Plant material Root tips are collected from various plants grown in glasshouse; these plants are Ribes sanguineum Purch, Quercus robur L., Thuja occidentalis L., Cercidiphyllum japonicum Siebold & Zucc., Acer tartaricum subsp. ginnala (Maxim.) Wesm. (collected field location) and Hibiscus syriacus. Reagents 8-Hydroxy quinoline 2Mm Cycloheximide 0.24 Mm

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Carboy’s fluid (6 parts 95% alcohol: 3 parts chloroform: 1 part glacial acetic acid; by volume Ethanol 70% (storage solution) Enzyme mixture (0.5% cellolase from Trichoderma reesei + 0.5% cytohlicase from Helix pomata + 0.5%pectolyase from Aspergillus japonicas) in a sodium citrate buffer at pH 4.5.

Procedure i. Root-tips are collected in the micro-centrifuge tubes and suspended in a solution of 2mM 8-hydroxyquinoline and 0.24 mAQWM cyclohexamide. The root tips are kept in the dark at room temperature for 2-5h before a cold period in a refrigerator at 4°C for 2.5 h. ii. After 5h in prefixative reagent, then the material is prefixed in Carnoy’s fluid overnight at room temperature. Later, the material is transferred to storage solution of 70% etanol and stored in refrigerator at 4°C. iii. Then root squash is prepared in following way: The roots are transferred to a small beaker containing sterilised water for 5 minutes, swirling occasionally. This step is repeated twice for a triple rinse of each root a total of 15 minutes. Then, root is taken on the clean slide and root tips excised under the dissection microscope. The remaining material is discarded and excess water is removed with a single-ply, low-lint tissue (VWR International, Rednor, PA) leaving only a small droplet encompassing the root tip to maintain the hydration. iv. A ring of ultraviolet region is placed around the root tip and overlapping a waxed paper pull tab. A region is set in an ultraviolet crosslinker, and enzyme digestion solution is added to well to encompass the root tip. Enzyme digestion is carried out in an oven at 37°C with the slide in a petridish atop a small weighing dish and moist filter paper. v. After the digestion, a pull tab is used to remove the enzyme digestion well, and the root tips are rinsed with a droplet of water before being wicked dry. vi. A single drop of modified carbol fuchsin stain (Kao, 1975) is added, and a cover-glass is lowered on the root-tip with half of a double-sided razor blade. vii. The slide is covered with a bibulous paper, and the root is squashed applying the gentle pressure from a pencil eraser. viii. The prepared microslide may be stored in the Petri-dish and observed under microscope. Results: Ploidy levels including diploids, triploids, and tetraploids with chromosome number ranging from 2n = 16 to 2n = 80 are observed. R. Sanguineum is confirmed as diploid, 2n = 16 whereas Q. robur is as a diploid, 2n = 24. Then T. occidentalis is confirmed as diploid, 2n = 22 while C. japonicum as diploid with 38 chromosomes. A. Tartaricum subspecies granula is confirmed as triploid (3x = 39) resulting from an interploid cross in

Cytological and Vital Staining Techniques 351

the isolated block (wild location), and H, syriacus is confirmed as tetraploid (4xx = 80). The range of haploid 2c genome sizes is spanned from 15.4 to 24.71. This protocol is useful for plant biologist working with taxa that exhibit a wide range of genome size and ploidy levels. B. Oxyquinoline-Acetic-Orcein Squash Method for Root-tips (Tjio and Levan, 1950; Radford et al., 1974)

Preparation of Reagents i. Stock solution of 8-hydroxyquinoline: Dissolve 0.3 g 8-hydroxyquinoline in one litre of distilled water. Store in refrigerator and use for several weeks. ii. Aceto-orcein: Dissolve 2.2 g of orcein in 100 ml glacial acetic acid. Gently boil the glacial acetic acid to which add slowly the orcein; later cool it and filter. For use, dilute the stain upto 45% strength acetic acid by adding distilled water. iii. HCl-orcein: HCl and aceto-orcein stain in 1:9 proportion is prepared. Procedure i. Root-tips are collected from germinating seeds and put in vials containing the aqueous oxyquinoline solution. ii. Place the vials in a cool room for 4 hrs. Specimens like Pinus with large, long chromosomes require 24-30 hrs pre-treatment. iii. Later place the root-tips in watch glass to which add few drops of HClorcein (1:9) solution. Heat gently over the spirit lamp. The sample is to be heated 3 or 4 successive times at intervals of a minute. Take care as heat hastens the macerating effect of the HCl. Leave the specimen in this solution for 20-30 mins. iv. Keep the root-tips in aceto-orcein stain (45% solution) for 20-30 mins; for small chromosomes, much longer period (12-16 hrs) of staining is needed. v. Stained root-tips are taken on the clean slide and broken into small pieces (not more than 1-2 mm in length) with the help of scalpel. Place the small pieces of tissue on another clean slide and cover it with glass immediately. Now gently tap the material with blunt instrument (handle of forceps) to tease and spread the material uniformly. Later invert the slide on blotting paper. Apply pressure slightly over the material with the ball of the thumb. The pressure applying here varies with the type of the material. The important point to be noted is that orcein preparation should not be heated. vi. Slides are kept in a refrigerator overnight without sealing. Later, these may be sealed with paraffin or with finger nail polish. They may be made permanent by the McClintock method or Conger dry ice method (Conger and Fairchild, 1953).

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C. Alcoholic Hydrochloric Acid-Carmine (Snow’s stain) for chromosomes in squash Preparations (Snow, 1963; Radford et al., 1974)

Staining Solution Carmine Conc. HCl Distilled water 85% ethanol

4g 1 ml 15 ml 95 ml

Add carmine and conc. HCl to distilled water. Mix well and boil gently for 10 mins while stirring. Cool the solution to which add 85% ethanol and filter. Store in brown bottles.

Procedure i. Fix the material (root tips or buds) in 3:1 ethyl alcohol and glacial acetic acid or in 3:1:1 ethyl alcohol, chloroform and glacial acetic acid. ii. Wash the fixative by giving several changes in 70% ethanol, one hour in each change. iii. Leave the materials in stain for 24 hrs or more. To hasten the staining process, place capped vials in oven at 50-60°C for 24-48 hrs. iv. Rinse the materials with water or 70% alcohol and prepare squashes substituting 45% acetic acid in place of aceto-carmine. v. Dissection of buds and maceration of the tissue, if needed, should be performed in 45% acetic acid. vi. For permanent slides, mount in equal pans of Hoyer’s mounting medium (30 g of gum arabic be soaked in 50 cc of distilled water about 24 hrs, add 200 cc chloral hydrate and let the solution stand until the whole material dissolves and finally add glycerine), and 40% acetic acid and squash. D. Feulgen Technique The Feulgen stain, or leuco-basic fuchsin stains DNA specifically and is used for cytophotometric determination of DNA in nuclei or chromosomes. The basic principle involved in this technique is the liberation of the aldehyde groups of DNA by hydrolysis, a chemical reaction binding the free aldehyde groups to the leuco-basic fuchsin, followed by the removal of unbound stain from the chromosomes. The following steps are involved in the Feulgen squash technique performed in somatic tissues in Nicotiana.

Procedure i. Fix the tissue in Carnoy’s fluid (six parts 95% alcohol, three parts chloroform, one part glacial acetic acid) or 3:1 absolute alcohol and glacial acetic acid for 2-24 hrs.

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ii. Later transfer the tissue in 70% ethanol and squash or store in refrigerator. iii. Hydrolyse the tissue in 10% conc. HCl for 10 mins at 60°C. iv. Later rinse in several changes of distilled water. v. Now the tissue is stained in leuco-basic fuchsin for 10-15 mins. vi. Later unbound stain is removed by rinsing three times with SO2 water followed by distilled water. vii. Transfer to 40% acetic acid. viii. Squash in 45% acetic acid over the slide. ix. Cover with cover glass. E. Staining with Lacto-Propiono-Dimethylsulfoxide Carmine (Bonga and Venkateswaran, 1976) It is observed that a combination of dimethylsulfoxide and carmine stains the tissues intensely when compared to aceto-carmine (Bonga and Venkateswaran, 1976). But it also causes some swelling of chromosomes. This undesired swelling can be counteracted by adding lactic acid and propionic acid to the DMSO-carmine. The staining procedure is as follows: (i) Prepare the Lacto-propiono-dimethylsulfoxide carmine (LPDC) stain solution: Carmine (alum lake) Dimethylsulfoxide (DMSO)

2g 150 ml

The above two are added and mixture is shaken for 30 mins at room temperature. Excess carmine is removed by filtration. Now this filtrate is known as DMSO-carmine. To the DMSO carmine add the following reagents: Lactic acid Propionic acid

25 ml 25 ml

The above mixture is now referred to as LPDC and stored at room temperature. (ii) Root tips and callus tissue are fixed in 3:1 alcohol-acetic acid (iii) Later tissues are stained in 1 ml of LPDC in tightly stoppered screw cap vials in an oven at 90°C for 1 hr and then squashed under a coverglass in a drop of DMSO on a microslide. The LPDC staining is more superior than aceto-carmine staining. Cell separation is much better in squashes of tissue stained in LPDC than the squashes of tissue stained in aceto-carmine. Chromosomes, nuclei and nucleoli are stained more intensely and vacuoles and ctyoplasmic strands are clearly visible with LPDC staining.

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F. Use of Cellophane for Permanent Preparation of Squashes (Breugel, 1980) Root tips are fixed in ethyl alcohol: acetic acid (3:1) for 1 hr. Later macerate them in 1N HCl for 15 mins at 60°C. Rinse in water briefly and stain by Feulgen method (see Protocol C).

Later Procedure i. Place small pieces of root tips in a drop of 45% acetic acid on microslide. ii. Presoak the 1 cm square of cellophane sheet, type 300 P (uncoated), density 1.4 available as cellophane 300 pT 50/50 (uncoated) in 45% aqueous acetic acid for few mins. iii. Spread cellophane over tissue and cover with blotting paper. iv. Apply the thumb pressure gently to spread the cells; later rinse in 96% ethyl alcohol for 1 or 2 mins. v. Lift the cellophane sheet and rinse in fresh 100% ethanol. The squashed tissue sticks to the cellophane sheet. vi. Mount the wet sheet upside down in a drop of Euparal and put the coverslip. vii. Citric acid pre-treatment may be given for animal tissues. G. A Quick Squash Method for Permanent Cytological Preparation (Lavania, 1981) i. Place a drop of Haupt’s adhesive and rub it gently with finger tip on the micro slide or coverslip. ii. Wash the other side of the slide with water for a few seconds. iii. By placing the coverglass squash the material gently, either fresh or after maceration, in a drop of 45% acetic acid. iv. Invert the squashed slide over a glass rod in a petri dish containing ethanol and apply gentle pressure to detach the coverslips. The squashed material remains stuck to the slide. Separation in ethanol will dehydrate the cells and will also harden them for better adhesion. v. Air dry the preparation after a fresh rinse in ethanol and mount in Euparal, DPX or any other suitable mounting medium. H. Technique for Preparation of Synoptonemal Complexes (SCs) from solanaceous plants (Stack, 1982) Lycopersicon esculentum (2n = 24) and Solanum tuberosum were used by Stack (1982).

Stains a) Dissolve 1 g phosphotungstic acid (PTA) in 25 ml of water; filter the solution and mix with 75 ml of 95% ethanol. b) Prepare 50% (w/w) silver nitrate solution just before use.

Cytological and Vital Staining Techniques 355

Medium 0.1 ml liquid medium contains 0.8 M sorbitol (Sigma), 1% polyvinyl pyrrolidone (PVP, Ave. MW 10,000 Sigma), 0.6 mM KH2PO4, 1.0 mM CaCl2, 1.6 mM MgCl2, 0.1 mM 1,4 Piperazine diethanesulfonic acid (PIPES) (acid, Sigma), and 0.3% potassium dextran sulphate (Ave. MW 8000, Calbiochem), pH adjusted to 5.0 with 0.1 M KOH. Procedure i. Excise and place the anthers containing primary microsporocytes showing leptotene through pachytene stages in cavity microslides (1.5 × 1.5 mm depressions). ii. Put 0.1 ml of above medium in the depression. Gently squeeze the anther contents and remove anther walls. Now add 1 mg β-glucuronidase from Helix pamata (No. G0751, Sigma). Β-glucuronidase should be desalted on a G25 sephadex column and lycophilised. Briefly stir the digestion medium to dissolve the enzyme. iii. Transfer the slide to a petri dish containing wet paper, towel and cover while digestion proceeds at room temperature for 20-40 minutes. iv. After incubation, the cell suspension is drawn into a micropipette with a tip approximately 0.2 mm in outside diameter. Cell masses are broken into individual cells in this process. v. Place approximately 5µl of cell suspension on microslide which is previously coated with Falcon plastic (to obtain 1% plastic solution 1 g of plastic from a Falcon plastic petridish should be dissolved in 100 ml of chloroform). New slides are wiped with a paper towel, dipped in plastic solution and dried in a vertical position. vi. Gently cover the cell suspension with an 18 × 18 mm cover-glass that is siliconised with ‘Prosil-28’ (Clay Adams) to make it hydrophobic. Now gently place the 16 g weight on the cover-glass before placing the 20 × 30 mm piece of thin, absorbent tissue (Kimwipe Kimberly Clark) against the left edge of the cover-glass. A capillary attraction is established by placing a drop of water on the edge of the paper immediately adjacent to the cover-glass. After 15-30 secs when as much of the solute as possible has been drawn from beneath the cover-glass, 2.5 µl of distilled water is placed on the right edge of the cover-glass which is pulled under the cover-glass, displacing most of the solution while cells remained trapped. vii. Now observe the slide under phase contrast microscope to determine whether there are numerous burst cells and spreads of synaptic complexes (SCs) on the slide. viii. Remove the cover-glass by the dry ice method. The hydrophobic coverglass tends to separate and leave the cells on the hydrophilic plastic surface.

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Fixation ix. Air-dry the slides briefly and fix in a freshly prepared 4% formaldehyde buffered with borate buffer (pH 8.2) for 10 mins (solution should be ice cold). x. Rinse the slides in 4% photoflo 200 buffered with borate buffer (pH 8.2) for 10-15 seconds and air-dry briefly. Staining and Light microscopy xi. Dip the slides in PTA solution for 10 mins. Rinse briefly in 95% ethanol and air dry. xii. Perform the silver staining by following either the ammoniacal silver (Ag-As) technique of Goodpasture and Bloom (1975) or the silver nitrate incubation (Ag-I) technique of Bloom and Goodpasture (1976). xiii. The 50% (w/w) silver nitrate solution should be prepared immediately before use. xiv. Observe PTA-stained SCs under phase contrast optics. Silver stained SCs can be viewed very well with bright field optics and are more suitable for photography: Electron Microscopy i. Locate the good chromosome spreads with light microscope and record their coordinates. Add water from either end of coverglass till a thick film is formed underneath the coverglass. Immerse the slide in petri dish containing water; the coverglass is floated free on the surface of water. Drain-off the water and coverglass, and air-dry the slide. ii. Relocate the good SCs spread with LM and cover with 50-mesh copper grid. Grids are made tacky, by dipping them in a solution prepared by dissolving adhesive from l-inch of cellophane tape in 10 ml of ethylene dichloride; allow the grids to dry. iii. While monitoring the operation at a magnification of 100 × under a compound microscope, the slightly adhesive grids are moved into exact positions using an eye lash mounted on an applicator stick. The plastic is scored around each grid with a steel dissecting needle to leave a margin of several millimetres. A drop of water is placed on the outer edge of the score mark, and the plastic is teased up with a dissecting needle. By gradually adding water the plastic is usually separated from the glass with the grids in place. iv. Float the freed sheet of plastic onto the surface of the bowl of water. Cover the grids and plastic with a piece of glassine weighing paper to which the plastic adheres. Now lift the paper, plastic and grids from the water and air-dry. v. Place the grids on a glass slide and observe under bright field or phase contrast microscope, and record the grid squares with chromosome spreads.

Cytological and Vital Staining Techniques 357

vi. Finally, grids are transferred to AEI EM6B electron microscope for observation and photography. Results: Two parallel lateral elements are present in each SC. In tomato, SCs in distal segment of chromosomes are more darkly stained than SCs in proximal segments. PTA stains kinetochroes, lateral elements, central elements of SCs. Ovoid structures about 100 nm in their longest dimensions are observed lying on the central element of the SCs. These are probably recombination nodules on pachytene chromosomes. This technique offers two-dimensional visualisation of complete sets of SCs by both light and electron microscopes and it is a powerful tool for genome analysis by allowing (1) determination of relative and absolute lengths of SCs and chromosome arm ratios at pachytene, (2) analysis of complex patterns of synapsis, and (3) the location of what are probably recombination nodules and the length of SCs. I. A Modified Fluorescence-plus-Giemsa Technique for Differential Staining of Sister Chromatid (Andersson, 1985) Roots of cuttings of Tradescantia paludosa (clone ‘V’ and clone ‘O’) were used in this technique by Andersson (1985). To develop roots, the cuttings are immersed in tap water for 5-6 days (in a green house-temperature cycle: 13½ hrs, 25°C; 10½ hrs, 18°C). After this, cuttings with roots of 10-20 mm length are transferred to an incubator where they are kept in the dark at 20°C.

Procedure i. Expose roots of 20-40 µm length to the aqueous solution of 100 µM 5-bromodeoxyuridine (BrdUrd), 0.5 µM 5-fluorodeoxyuridine (FdUrd). This is done in glass tubes containing 40 ml of solution. ii. Roots which have been exposed to BrdUrd solution for 16 hrs, are transferred to aqueous solution of 100 µM thymidine (dThd) and 5 µM uridine, and kept there for 18 hrs. iii. Later treat the roots with 0.5% colchicine for 3 hrs. iv. Fixation: Fix the material in cold 3:1 methanol, acetic acid for 1-2 days. The tubes containing the BrdUrd and dThd solutions should be covered with black tape to protect the roots from light. v. After fixation, rinse the roots in the same slightly acidic buffer (0.01 M citric acid C6H8O7. H2O - sodium citrate Na3C6H5O7. 2H20, pH 4.7) in which maceration is going to perform. vi. Maceration: Macerate the roots either with a 1% pectinase solution (Sigma, from Aspergillus niger) or with a mixture of 0.5% pectinase and 0.5% cellulase (Sigma, from Aspergillus niger) for 2 hrs at 27°C. The maceration is terminated by transferring the roots to distilled water at room temperature. Half an hour later, squashing is performed.

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vii. Squash preparation: Squash in 45% acetic acid on slides coated with a mixture of 10% gelatin and chrome alum (Reidel-De Haen, Seelze, Hannover). viii. Remove coverslips on dry ice and pass through 100, 85, 70, 50 and 30% ethyl alcohol and distilled water. ix. Later place 200 µl of RNase (Sigma, Ribonuclease A from bovine Pancreas) solution [1 mg dissolved in 10 ml 0.5 × SSC (saline sodium citrate) (0.075 M NaCl + 0.0075 M Na3C6H5O7. 2H2O)] on each preparation and put the cover-slip, incubate for 50 minutes at 27°C. x. Fluorochome staining: After a brief rinse in 0.5 × SSC, stain the preparation in a solution of 1 mg-Hoechst 33258-pentahydrate in 2000 ml phosphate buffered saline (Oxford Limited, England) for 20 minutes, later briefly rinse in 0.5 × SSC. xi. Place the slides in 0.5 × SSC in Coplin jars and immediately expose for 40 mins at a distance of 15 cm to light from an Osram 300 W UlatraVitalux sun lamp. xii. The irradiation should be followed by a treatment with 0.5 × SSC for one minute at 55°C. xiii. Giemsa staining: Transfer the slides to 0.017 M phosphate buffer (pH 6.8) and keep them for 5-10 mins. Later stain in a 3% Giemsa (Gurr R66) for 4 mins in the same buffer. xiv. Rinse in running tap water and air dry. Dip the dry preparation in xylene and mount in Canada balsam. Results: The concentration of FdUrd is necessary to give rise to a good sister chromatid differentiation varying between 5 × 10-9 and 1 × 10-7 M in the Vicia faba. Twice the concentration of pectinase is needed to get good separation of the meristematic cells. To get rid of background staining caused by cytoplasmic ribonucleic acid, the preparations are treated with RNase. For practical reasons, the slides are treated for 50 mins with RNase, but the time of exposure does not seem to be critical. J. A Haematoxylin Staining Procedure for Maize Pollen Grain Chromosomes (Kindiger and Beekett, 1985) Preparation of Haematoxylin stain Solution A: Dissolve 2 g of haematoxylin (Fisher, Lot 701073) in 100 ml 50% propionic acid. Allow it to ripen for one week and store it in a stoppered brown bottle without refrigeration. Solution B: Dissolve 0.5 g ferric ammonium sulfate [FeNH4(SO4)2] in 100 ml 50% propionic acid. This solution keeps indefinitely in stoppered brown bottle without refrigeration. Mix equal volumes of solutions A and B. The mixture, which turns dark brown, is ready for use immediately. The stain remains good for about two weeks.

Cytological and Vital Staining Techniques 359

Collection and Preparation of Pollen grains Collect maize tassels when about 3 cm of the tassel becomes visible above the leaf whorl. This will usually give many male gametophytes at the 1st and 2nd mitotic divisions. Material that is to be retained for longer periods should be placed in a 14:1 mixture of 70% ethyl alcohol: formaldehyde and stored in the refrigerator. Pre-fixation: It is not necessary, but it can be performed before staining so as to provide greater definitions of mitotic chromosomes. Dissect out anthers and pre-fix for 30-45 mins in a solution containing 100 ml tap water, four drops of monobromonaphthalene and one drop of dimethyl sulfoxide (DMSO). Monobromonaphthalene inhibits the spindle at metaphase allowing observation of shortened, well defined chromosomes. Fixation: Later anthers are fixed in Carnoy’s fluid (3:1 95% ethanol acetic acid) or in 3:1 ethanol-chloroform for 30 minutes. Staining Procedure i. Dissect out anthers and place in a drop of 45% acetic acid on a slide. ii. Add a crystal of chloral hydrate; macerate to liberate pollen grains and to dissolve the chloral hydrate. Remove debris. iii. Add a drop of stain and mix on the slide. For fixed material, wait for a minute before applying the coverslip. iv. Then apply the coverslip, blot of excess stain and apply gentle pressure on it. v. Gently heat the slide and then cool. Upon cooling, chromosomes and nuclei will darken and cytoplasm will clear. vi. Continue heating and cooling until the desired, contrast is obtained. vii. To make the slide permanent, seal the edges of the coverslip with permount and allow it to dry. Haematoxylin procedure is faster, gives more consistent staining, and provides superior contrast between nucleus and cytoplasm when compared to Belling’s iron acetocaramine stain. Slides cannot be kept indefinitely, as the stain begins to fade after about three months. Satisfactory staining of fixed material is obtained if the pollen is allowed to soak in the stain longer before heating.

General Slide preparation for male meiotic studies i. Flower buds of appropriate size are fixed in freshly prepared Carnoy’s fluid (1:3 acetic acid: ethanol) for 4-5 days at room temperature. ii. After fixation, the flower buds are stored in 70% ethanol at 4°C. iii. Sepals and petals are removed from the selected flower buds and the anthers are taken; and then anthers are squashed in a drop of 2% acetocarmine stain. iv. In some cases, ferric chloride solution is used as a mordant and the stained cells are washed by adding drops of 45% acetic acid.

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v. The prepared slide was covered using a cover slip and was slightly warmed by passing over a flame. vi. Finally, the prepared slide is wrapped in a fold of blotting paper to remove access of stain leaving only a thin film of fluid in between slide and cover-slip. vii. The prepared slide is observed under a microscope. K. Efficient preparation of Plant Chromosomes for SEM Study (Martin et al., 1994) Root tips of 3-day old seedlings of Barley (cv. ‘Jgri’), barley telotrisomic lines and wheat-barley telosomic addition lines are taken.

Procedure i. Synchronise the meristematic tissue either by pre-treating the root tips in ice water for 24 hrs or by following Pan et al. (1993) and Busch et al. (1994). ii. Fix in 3:1 ethanol-acetic acid (v/v) and store it at 20oC. Digestion i. Chop and digest the root-tips in a solution of 2.5% cellulase ‘Onozuka’ R (Sigma chemicals, Munich), 2.5% Pectolyase Y-23 (Kikkoman, Japan) in 75 mM KCl (pH 4.0) at 30oC for 50-60 mins. ii. Filter the suspension through nylon cloth (100 mm). iii. Treat with hypotonic solution of 75 mM KCl at room temperature for 8 mins. iv. Spin down the protoplasts at 80g for 7 mins. v. Resuspend in ethanol-acetic acid (3:1) and spin again at 80g for 7 mins. This step should be performed four times. vi. Finally resuspend the pellet in an appropriate volume of fixative and drop on ice-cold cleared slides. vii. Spreading of cells and subsequent steps will be monitored in a ‘ZEISS Axiophot’ light microscope. viii. Add a drop of 45% acetic acid before completely drying out the specimen. ix. Cover the preparation with a coverslip, gently squash and subsequently freeze on dry ice. x. Remove the coverslips and transfer the slides immediately to the fixative containing 2.5% glutaraldehyde in 75 mM Cacodylate buffer and 2mM MgCl2 (pH 7.4). xi. Osmium impregnation, dehydration, critical point drying, and sputter coating is accomplished following Wanner et al. (1991). xii. Observe the chromosomes under a Hitachis 4100 field emission scanning electron microscope. xiii. Microdissect the chromosome with glass rods using a Zeiss inverted microscope.

Cytological and Vital Staining Techniques 361

Results: This technique offers the production of relatively high numbers of chromosome spreads that can be viewed at high resolution showing structural details below 10 nm. This technique is not only restricted to metaphase chromosomes, but also to the chromosomes of all stages of the cell cycle.

17.1.3 Chromosome Banding Technique Traditionally, chromosome identification has been done based on their morphological characteristics such as relative length, arm ratio and presence or absence of secondary constrictions. Besides this, nowadays chromosome banding technique has become popular and a useful tool in the identification and characterisation of chromosomes within the complement. In banding technique, various chromosome segments are selectively stained with an appropriate fluorescent agent (fluorochrome) that binds selectively to bases in chromosomal DNA. For example, quinacrine mustard (QM) fluoresces in alternate light and dark bands when viewed under ultraviolet light. Like fingerprints these fluorescent banding patterns are unique for each chromosome. These bands are generally consistent for a taxon except for minor variations. There is a general agreement that the technique, which involves denaturation of DNA followed by slow renaturation, permits identification of constitutive heterochromatin, because it mainly consists of repetitive DNA. Various fluorochromes have been employed and they show different banding patterns. Acridine orange (AO) produces variable banding in plants while ethridium bromide gives reverse bands. These bands are made visible through low and high intensity regions under the fluorescent microscope or as differentially stained areas under the light microscope. Heteromorphy for chromosome bands has also been demonstrated in many cases, so that the possibility of using bands as ‘markers’ has been established. Bands have also been detected after treatment of chromosomes with restriction enzymes (RE) and these bands are described as RE bands (Gupta, 1997). The dye is usually applied to the chromosome preparation (either squashed or air dried) and then mounted in a suitable medium, and observed under fluorescent microscope. All these dyes show a pronounced specificity for nucleic acids e.g., QM reacts with DNA by alkylation of certain purine bases, and AO, proflavine and acriflavine bear positive charges that allow them to interact with the DNA-phosphate backbone. It is further stabilised by the intercalation of the flat acridine rings between nucleic acid base pairs (Sharma, 1976). AO is also used to study the strandedness of nucleic acids, particularly DNA in chromatin and chromosomes. After staining, AO molecules are dispersed within the helix of a double-standard DNA combining as a monomer without intermolecular interactions and fluorescene green, absorbing light at 500 nm. On the other hand, stacks of AO molecules are formed combining as polymers on single-stranded DNA or RNA which fluoresces red, absorbing light at 463 nm.

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Methods to demonstrate the Giernsa-stained banding patterns of mammalian chromosomes have been described by many scientists using basic principle of in vitro hybridisation of DNA. The earlier methods depended on alkaline denaturation followed by renaturation in an adequate buffer. Later schedules for studying the G-banding are simpler, based on assumed differential denaturation only. The agents for producing bands are alkali used in different temperatures, proteolytic enzyme and protein denaturing substance, including detergents, usually aniomic ones. The process of differential processing used in these simplified G-band techniques seems to depend on the temperature, pH, ionic strength, divalent cation content and proteolytic enzyme content of the medium. According to Chromosome Band Nomenclature (adopted at the Paris Conference in 1971), the following types of banding in human chromosomes have been recognised:

Q-band: by quinacrine staining and fluorescence. G-band: by staining with Giemsa and related stains after appropriate pretreatment. R-band or banding reverse to Q-band: by staining with Giemsa after heating to 87°C. C-band: for constitutive heterochromatin, demonstrated by the denaturation-re-association technique. E-band: Produced by enzymatic digestion, as classified by Lejeune (1973 quoted in Sharma, 1976), show swollen and shrunken regions corresponding to the dark and faint regions of G-banding. The same nomenclature can be applied to banding patterns observed in other eukaryotic chromosomes. Other banding patterns include:

CT-bands: The centromeric and telomeric segments show bands after treatment in barium hydroxide, incubation and staining in ‘Stains All’ (4, 5, 4’, 5’-dibenzo-3, 3’-diethyl-9-methyl thiacarbocyanine bromide) in mammals. N-bands: at nucleolus-organising regions, possibly due to acidic proteins in mammals. O-bands: by orcein staining, mainly for plant chromosomes; both intercalary and centromeric heterochromatin show bands. Comings in 1973 (quoted in Sharma, 1976) has found that centromeric heterochromatin appears in the C-bands and the intercalary heterochromatin in the G-bands. The former is rich in highly repetitive satellite DNA while the latter is with non-repetitive poorly methylated DNA. Protein-DNA interactions are also important factors in formation of chromosome bands. He observes that in C-banding the selective extraction of non-band DNA plays a major role but the intensity of Giemsa staining is enhanced by DNA protein interactions.

Cytological and Vital Staining Techniques 363

In both G and R bands, base composition may be involved but DNA-protein interactions seem to be the major role. In Q-banding, negative or pale staining regions are due to the presence of non-histone proteins binding in the small groove of DNA and preventing the intercalative binding of quinacrine. A similar mechanism may be responsible for G-banding (Comings, 1974). Neither C- nor G-banding appears to depend on the differential denaturation of DNA. Various attempts have been made to induce chromosome bands by differential digestion of the chromosomal DNA by nucleases. By using restriction endonucleases, it is to localise highly repetitive DNAs in human metaphase chromosomes. A different approach is undertaken by Schweiger (1977), who produced R-bands by DNase digestion of chromosomes after protecting the G-C rich DNA sequences with the DNA-binding antibiotic chromomycin A3. 17.1.3.1 Protocols

A. Methyl Green-Pyronin Dye to Stain C-banded Chromosomes (Puertas et al., 1983) Fix the root tips of rye, Secale cereale L. in acetic: alcohol (1:3) before squashing in one drop of 45% acetic acid. Dehydrate the material in absolute alcohol for 24 hrs.

Preparation of Stains a. Giemsa: Mix 3 ml of Gurr’s R66 Giemsa in 100 ml of 1/15 M phosphate buffer (pH 7.0). b. Methylgreen-pyronin: Dissolve 2 g of methyl green in 100 ml of distilled water; solution is repeatedly extracted with chloroform to remove traces of methyl violet formed when methyl green is exposed to air. Pyronin 2% solution (in distilled water) is also washed with chloroform. Stock solutions can be stored in the refrigerator. At the time of use, 7.5 ml of 2% methyl green, 12.5 ml of 2% pyronin, 2.5 ml of 95% ethanol, 80 ml of distilled water and 0.5 g of phenol are mixed, adjusted at pH 4.5 with IN HCl.

Treatment before Staining i. Hydrolyse the material in IN HCl at 60°C for 12 mins. ii. Incubation is performed in bovine pancreas ribonuclease (one mg ribonuclease is dissolved in 50 ml of 0.05 M acetate buffer at pH 5; the solution is heated to 80°C for 10 mins to eliminate possible DNase activity) at 37°C for 1 hr. iii. C-banding-Routine method (Giraldez et al., 1979). a. Hydrolysis is accompanied by treating the roots with 0.2 N HCl at 60°C for 3 mins.

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b. Immerse in saturated solution of Ba(OH) at room temperature for 12 mins. c. Incubate in 2 × SSC at 60°C for 1 hr. iv. As treatment (iii) but without hydrolysis with 0.2 N HCl. v. As treatment (iv) but with incubation in ribonuclease as in treatment (ii).

Staining Procedure a. Giemsa staining: In treatment (ii), (iii) and (iv) control slides are stained with Giemsa, cleared in xylene and mounted in DPX. b. Methyl green-pyronin staining: Preparations after treatments (i) to (iv) are immersed in methyl green-pyronin for 2-5 mins. Later rinse in tertiary butyl alcohol, clear with toluene and mount in DPX. To check the ability of pyronin to stain proteins, ovalbumin and bovine albumin are subjected to electrophoresis using the two different 12% starchgel electrophoretic systems given below: a. 0.15 M Tris-citric acid, pH 7.75, as the gel buffer and 0.3 M NaOH boric acid, pH 8.6, as the electrode buffer. b. 0.043 M Tris-citric acid, pH 7.0 as the electrode buffer and 0.006 M histidine, pH 7.0 as the gel buffer. Gels are stained with pyronin prepared as above. Slides stained with methyl green-pyronin show red bands. Results: Giemsa stained slides are obtained best by the use of treatment: (staining procedure a) Methyl green-pyronin staining gives good results with treatment; (staining procedure b) in this clear blue nucleus with red heterochromatic blocks are observed; mitotic cells show blue euchromatic portions of chromosomes with red heterochromatic bands. Untreated and methyl green stained preparations produce blue-green nuclei and with pyronin alone produce red nuclei, nucleoli and proteins subject to electrophoresis but do not show any affinity for pyronin. B. Heterochromatin patterns in Crotalaria L. (Gayatri et al., 1993) Dry seeds of four species of Crotalaria L. viz. C. juncea Linn., C. Laburnifolia Linn., C. verrucosa Linn., and C. trifoliastrum Wild. were used in this study.

Procedure i. Roots collected from germinated seeds are pretreated with cold water at 4°C for 24 hrs and fixed in 3:1 alcohol: acetic acid for 24 hrs. ii. Later roots are washed and subjected to hot hydrolysis in 1N HCl for 10 mins at 60°C. iii. After hot hydrolysis, wash again for 15 mins. iv. Stain the root tips with pararosanilline chloride for 45-60 mins.

Cytological and Vital Staining Techniques 365

v. Later root tips are squashed in 0.5% acetocarmine, put the coverslip and seal it with paraffin wax. Gayatri et al. (1993) studied the variation with reference to the total number of bands, band size and position in between the chromosomes within a complement as well as between the four species; in C. laburnifolia and C. trifoliastrum and reported nearly equal amount of heterochromatin which was more than that of the other two species. The quantity of heterochromatin present in an arm of the chromosome is calculated as follows: The quantity of heterochromatin in each arm of the chromosome is calculated as follows: The quantity of heterochromatin in each arm ___________________________________________ × 100 Total quantity of heterochromatin in haploid genome C. Giemsa G-banding in Root Tips of Allium (Peffley and Vries, 1993) Root tips of Allium fistulosum, A. Cepa ‘Jumbo’ and their interspecific cross were taken for Giemsa G-banding studies.

Procedure i. Pretreat the root tips in a saturated solution of l-bromonaphthalene at 4°C for 2 hrs. ii. Later fix the material in fresh 96% ethanol, glacial acetic acid (3:1) at 4°C for 30 mins. iii. Some chemically fixed roots are squashed in 45% acetic acid and then frozen in liquid nitrogen, later coverslips are removed. Other roots are kept in the fixative for 2 hrs and washed with distilled water. iv. Incubate the tissue in a mixture of cellulase (1.5%, w/v) and pectinase (15%, w/v) in citrate buffer at pH 4.8 for 1 hr at 37°C to soften the tissue. v. Rinse in distilled water. vi. Spread root tips on a glass slide in a drop of 60% acetic acid with the help of a pair of needles. vii. Air dry the slides. Staining Method i. Immerse the slides in 0.01 M NaHC03, pH 9.0-9.1, at 27-30°C for 1-5 mins. ii. Stain in 2 or 25% Giemsa, in pH 6.8 buffer solution (from Gurr’s buffer tablets). iii. Rinse the slides in distilled water twice. iv. Air dry the slides. v. Observe under light microscope to evaluate the banding pattern in chromosomes. If no bands are visible, the slides are rinsed briefly in 96%

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ethanol: glacial acetic acid (3:1) fixative to partly remove the Giemsa stain and follow the staining method of Drewry (1982) given below: vi. Place the slides again in 0.01 M NaHCO3, pH 9.0-9.1.

Staining Method of Drewry (1982) i. Keep the above-prepared slides at 45-60°C for 1-12 hrs. ii. Immerse the slides in trypsin solution [Gibco, trypsin-EDTA (10x) 5.0 ml, Hank’s BSS 4.5 ml, H2O 40.5 ml] at room temperature for 1-40 mins. iii. Stain in 2 or 25% Giemsa (Gurr) in pH 7.9 buffer (Gurr’s buffer tablets adjusted with NaHCO3) for 3-6 mins. iv. Rinse in buffer solution, pH 6.8 (Gurr’s buffer tablets) and in distilled water twice. v. Mount in DPX (Gurr). Result: In all treatments, prophase and prometaphase chromosomes of Allium show rows of numerous dots and bands of different sizes extending along the length of the chromosome: i. C-bands were located exclusively at telomers and interstitial C-bands at centromeric region. ii. The number of bands visible along the entire length of chromosomes in metaphase is reduced to chromosomes in late prophase. iii. G-bands stain more intensely without trypsinisation and with increasing trypsin incubation chromosomes become more dispersed and obscure the banding pattern to a large extent. G-banding visualised in the chromosomes of Allium using Giemsa stain is the first demonstration of its kind.

17.2 Vital Staining Vital dyes are used to study living tissues. According to Barbusa and Peters (1971), vital staining was developed by Grew in 1682. The extensive literature on vital staining is available in German. Detailed discussion on this aspect is outside the scope of this book. Stadelmann and Kinzel (1972) published the English review. A general opinion among biologists is that vital stains do not affect the structure and functioning of the cells being stained. However, this needs thorough investigation. Indeed, Barbusa and Peters (1971) concluded from their studies that vital dyes not only may be toxic to the target organism but may also cause a number of pathological symptoms in the organism, some so subtle that they may be overlooked. One must be careful while interpreting the results of vital staining with plant cells. For example, neutral red is commonly used by botanists. Accumulation of this dye by vacuoles is usually considered to indicate cell viability but O’Brien and McCully (1981) found that it is capricious. This

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supports Bailley’s warning (1930) that vital staining of vacuoles in plant cells is due to tannins precipitating the dye, but not due to the uptake of dye through tonoplast. In this regard, Boyer (1963) has shown that tannin-filled vacuoles in pine needles stain strongly with neutral red. Stefano et al. (2017) have studied plant vacuoles using various fluorescent probes including fluorescent-tagged proteins. For example, AFVY-RFP, a soluble marker based on the fusion of a manomeric red fluorescent protein (RFP) to the tetrapeptide AFVY, a C-terminal sorting signal of soluble proteins to the vacuole (Hunter et al., 2007) (Fig. 17.1). The figure clearly shows the vacuolar lumen in the abaxial epidermal cells at 9 days after the germination (DAG) of seeds of Arabidopsis thaliana (17.1A). The VAC-YFP is another fluorescent-tagged tonoplast marker based on the fusion of the C-terminus of a v-TIP protein (Nelson et al., 2007) to the yellow fluorescent protein (YFP) (Fig. 17.1B). The figure shows the tonoplast and transvacuolar strands in abaxial epidermal cells expressing the VAC-YFP at 12 DAG (Fig.17.1 B).

Fig. 17.1. Staining of vacuoles with fluorescent proteins. (A) Confocal images of Arabidopsis abaxial epidermal cells (9 DAG) stably expressing AFVY-RFP, which marks the vacuolar lumen indicated by an asterisk. (B) VAC-YFP (12 DAG) in abaxial epidermal cells. Arrow points to tonoplast, arrowhead indicates bulbs, and empty arrowhead indicates transvacuolar-strand. Bar = 10 μm (Courtesy: Stefano et al., 2017).

17.2.1 Protocols A. Staining of plant cell vacuoles (Stefano et al., 2017) The plant vacuoles are extremely important for growth and development, and influence the vital cellular functions such as photosynthesis, respiration and transpiration. One of the important roles of vacuole is

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to regulate the cell turgor in response to different stimuli. Therefore, the morphology, dynamics and physiology of plant vacuole are fundamentally important for the advancement of plant science, particularly plant cell biology (Stefano).

Plant materials and chemicals Arabidopsis thaliana seedlings at 3 to 9 days after germination (DAG) Agrobacterium cultures Lipophilic styryl dye, N-(3-triethylammonium propyle)-4-(pdiethylaminopropyl- hexatrienyl) pyridinium dibromide (FM4-64) AFVY-RFP---a soluble fluorescent marker based on the fusion of a monomeric red fluorescent protein (RFP) to the tetrapeptide AFVY, a C-terminal sorting signal of soluble proteins to the vacuolar (Hunter et al., 2007). VAC-YFP, a tonoplast marker based on the fusion of the C-terminus of a v-TIP protein to the yellow fluorescent protein (YFP) (Nelson et al., 2007). Plant growth i. Prepare the petriplates containing Linsmaier and Skoog (LS) medium. The seeds of A. thaliana are incubated on LS medium at pH 5.7 and incubate in a growth chamber at 22°C temperature, 55% humidity for 16 hours of light and 8 hours of dark cycle. ii. Keep the plates vertically to image root seedlings or position horizontally to take picture of the abaxial pavemental cells in cotyledon or leaf. Stable or transient expression of vacuolar reporters in Arabidopsis lines AFVY and VAC transgenic lines: Grow the Arabidopsis AFVY-RFP stable transformant lines as mentioned above and below and similarly VAC-YFP also. For transient expression of vacuolar markers of thaliana, the following materials are required: i. Prepare plates with LS medium to stratify the seeds; one litre of medium consists of 1.18 g LS, 1% sucrose and 0.8% agar and then autoclave the medium for 25 minutes. ii. Prepare the co-cultivation medium with LS medium, acetosyringone 100 µm, silvet L-77 OF 0.005% and incubate agrobacterium along with seedling. iii. Prepare the washing solution using MgCL2 of 10 µM and acetosyringone 100 µM and rinse the seedlings co-cultivated with agrobacterium.

Procedure AVFY-RFP and VAC-YFP samples for confocal microscopic studies a. Stable expression of genetically-encoded vacuolar markers in A.thaliana:

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i. Sterilise the seeds expressing the transgene (AFVY-RFP or VAC-YFP) as follows: Take some seeds in eppendorf tube, then add 1 ml of ethanol 70%, invert the tube, centrifuge at low speed and discard the solution. Then add 1 ml of 20% bleach solution, invert the tube few times, centrifuge briefly and discard the solution. Wash the seeds thrice with distilled water and inoculate on LS medium and verbalise at 4°C for 48 hours. ii. Later transfer the petriplates to growth chamber maintaining 16 hours light and 8 hours dark cycle at 22°C. iii. Conduct the confocal microscopic analyses on seedlings starting from 3 DAG until desired to follow the vacuolar marker (up to 12DAG). b. Transient expression of genetically-encoded vacuolar markers in A.thaliana i. Sterilise the seeds as previously described and these sterilised seeds are inoculated on LS medium. ii. Transfer the inoculated plates to the growth chamber maintaining 16 hours light and 8 hours dark at 22°C. iii. Two days after germination, prepare the co-cultivation medium. iv. Three days after germination, inoculate the Agrobacterium carrying the binary vector with the vacuolar marker of interest, into LS with appropriate antibiotics, then shake at 200 rpm, at 28°C until the optical density read at 600 nm reach a value which is greater than 1.5. v. Centrifuge the agrobacterium culture at 5000 rpm for 3 minutes, then discard the supernatant. vi. Resuspend the agrobacterium pellet in 100 mL of wash solution by vortex. Repeat this step once again. vii. Again resuspend the pellet in 100 mL of wash medium. viii. Add agrobacterium culture to 100 mL of cocultivation medium to reach a final optical density of 0.5, then pour the medium in the petriplate. ix. Incubate the plates in the dark for 24 hours. x. Open the plates, remove the medium, and wash atleast twice with 10 mL of cocultivation medium xi. Seal the plate with micropore 3M and incubate in standard growth conditions. xii. After 48-96 hours after transformation, cotyledons or root tissue can be observed under the confocal microscope [for YFP, 514 nm (excitation) and 560-600 nm (emission; for RFP, 560 nm (excitation) and 600-650 nm (emission). To have an image of a good fluorescent signal of the vacuole or tonoplast, it is necessary to observe the sample after 48 hours because the signal may be higher in the endoplasmic reticulum before this time.

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B. The use of Fluorescein Diacetate and Phenosafranine for determining viability of cultured plant cells (Widholm, 1972) Composition of Tobacco medium (in mg/L) NH4NO3, 1650; KNO3, 1900; CaCl2.2H20, 440; MgSO4.7H20, 370; KH2PO4, 170; H3BO3, 6.2; MnSO4.4H2O, 22.3; ZnSO4.4H2O, 8.6; Kl 0.83; Na2MoO4.2H20, 0.25; CuSO4.5H2O, 0.025; COCl2.6H2O, 0.025; NaFe EDTA, 33.

Fluorescein diacetate (FDA) FDA, 0.01 % is prepared by diluting a 5 mg/ml acetone solution (stored in freezer) with the tobacco medium. This diluted solution is useful for few hours. Procedure i. Grow the culture of tobacco pith (Nicotiana tabacum L. cv. Xanthi and Wisconsin 38), Tomato stem (Lycopersicon esculentum Mill.), garden carrot root (Daucus carota L.), germinating rice seed (Oryza sativa L., Blue Bonnett 50) and soybean stem (Glycine max Merr. cv. Wayne) as described by Widholm (1971). ii. Pipette the cells of one gram into 100 ml fresh liquid medium and take fresh weights after filtration on Miracloth. iii. Test the dyes (0.1% prepared in tobacco medium) by mixing a drop of the dye solution with a single drop of a cell suspension on a glass slide. iv. Later, put the coverslip and observe under phase contrast, after 5 and 30 min with a Zeiss microscope. The fluorescein and fluorescein diacetate treated cells are viewed with the same microscope utilising fluorescence technique. An Osram HBO mercury-vapour lamp with exciter filter BG12 is used in combination with barrier filter 47. The other compounds tested by Widholm are phenosafranine, neutral red (Mann Research Laboratories), 2,3,5-triphenyl tetrazolium-Cl, acid fuchsin, acridine orange, crystal violet, helianthin, indigo carmine, neutral red iodide, orange G, rhodamine B, rhodamine 6G, Sudan III (Allied chemical), isatin, methylene blue (Eastman Organic Chemicals), aniline blue, methyl green, safranine O (Fisher Scientific Co.), brom phenol blue, brom phenol red, Janus Green B (Matheson Coleman and Bell), malachite green, nile blue A (Hartman-Leddon Co.), methyl red (General Chemical Co.) and brom thymol blue. Results: FDA stains live cells whereas the others like phenosafranin, brom phenol blue, methylene blue, stain only dead cells. FDA probably stains live plant cells in the same way that it does with animal cells (Rotman and Papermaster, 1966) and pollen (Heslop-Harrison and Heslop-Harrison, 1970). The non-polar molecule enters the live cells where esterage cleaves off the acetate residues leaving fluorescein which then accumulates. The fluorescein molecule can fluoresce. Taking cyclosis and plasmolysis as the main criteria,

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cells at various stages of culture growth were also examined for their viability by Widholm using fluorescein diacetate. C. Vital staining for root hairs in perennial grasses (Marianne et al., 1990) Roots of Paspalurn notatum, P. vaginatum, Stenotaphrurn secundaturn. Eremochloa ophiuroides, Zoysia japonica and Cynodon dactylon were taken by Marianne et al. (1990). Excise 10 cm long adventitious roots with intact primary and secondary branches and store in 20 ml glass scientillation vials containing 17 ml of 0.05 N phosphate buffer (pH 7.0), 4°C for 18 hrs. Place control vial in boiling water for 10 mins, later cool and store at 4°C.

Stains Used Prepare phenosafranin (PS), neutral red (NR) and methylene blue (MB) at a concentration of 0.1 g/L, Evans blue (EB) at 0.5 g/L and congo red (CR) at 1.0 g/L in 0.05 N phosphate buffer at pH 7.0. Procedure i. Stain the root hairs of each grass in the above-mentioned stains for 4-24 hrs. ii. Washing is performed in multiple baths of water and 0.05 M phosphate buffer (pH 7.0) for 45 mins to 72 hrs. iii. Observe under the light microscope and compare the colours developed in different materials with reference to control. Evans blue (EB) is found to be the best stain in terms of precision and least variability to show the quantity, size and viability of root hairs. D. Pollen fertility with PHR stain (Ramanjini Gowda et al., 1991) Pollen of Dolichos, ash gourd, sweet potato, Bougainvillea and Hibiscus were taken.

Preparation of P.H. Ramanjini (PHR) stain Collect fully ripe purple-blue coloured fruits of Cocculus hirsutum. Crush these fruits and filter through muslin cloth. Add 20% phenol (v/v) to extract before using as stain. PHR stain may be modified by adding water (60% v/v) and lactic acid (20% v/v) to it. Procedure i. Take the pollen dust onto a micro slide and put 1-2 drops of stain. ii. Mix-up properly the pollen dust and stir with stainless steel needle. iii. Mount the coverglass and examine after 5 minutes. Result: The viable pollen grains appear dark to red in colour whereas nonviable ones do not take up stain. The intensity of the staining is dependent on the thickness of the pollen grain walls.

18 Electron Microscopic Studies Basically, the biological specimen preparation for electron microscopic (EM) studies is similar to that of light microscopy. However, some modifications are made to meet the specific requirements. Plant materials should be chemically killed and fixed (preserved) and followed by infiltration and embedding. Later, cut into ultrathin sections on ultra-microtomes and stain them to bring out contrasts in tissues and cell organelles.

18.1 Specimen Preparation for EM Studies Sample preparation includes fixation, infiltration, embedding, sectioning and staining for EM studies (Table 18.1). Table 18.1. Characteristic difference in sample preparation between electron microscopy and light microscopy Electron microscopy

Light microscopy

Fixation

Glutaraldehyde, Osmium tetroxide

FAA, formaldehyde, acrolein

Embedding

Monomeric epoxy resins

Paraffin wax

Sectioning

Glass knife or diamond knife Steel knife

Thickness of sections

0.05-0.1 µm

Staining

Heavy metals like uranyl acetate, lead citrate

2-10 µm and more thickness Dyes

18.1.1 Fixation The primary objective of the killing and fixation is to arrest the growth processes and preserve the material in some fixative reagent with minimum damage to the tissues. Several fixatives are currently employed such as osmium tetroxide, certain aldehydes, acrolein, potassium permanganate, etc.

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Osmium tetroxide (OsO4) is widely used as a fixative for EM studies. Its penetration is slow, but it reacts rapidly with proteins, unsaturated fatty acids and phospholipids. Now-a-days, it is common for Transmission Electron Microscopy (TEM) that the material is prefixed in glutaraldehyde and postfixed in OsO4. Glutaraldehyde: It is a non-coagulant, and its structural formula is CHO(CH2)3-CHO (Sabatini et al., 1963). A solution of 4% glutaraldehyde in phosphate buffer (pH 7.0) is prepared. Specimen is prefixed in this solution for 2 hrs to overnight. If cytoplasm is abnormally electron dense or tissue shrunken, fix the material for 2 hrs at 0-4°C and then at room temperature. Osmium tetroxide (OsO4): Specimen is post-fixed in 2% OsO4 prepared in phosphate buffer at pH 6.8 (O’Brien and McCully, 1981) for 1-2 hrs. Under post-fixed condition, OsO4 gives positive electron contrast to membranes and preserves unsaturated lipids that would otherwise be extracted during dehydration and embedding (O’Brien and McCully, 1981). OsO4 penetrates fresh tissues slowly but after glutaraldehyde treatment, it appears to penetrate more rapidly.

18.1.2 Embedding Generally, harder embedding medium (harder than paraffin wax) is used for EM studies. After dehydration, the specimen is infiltrated and embedded in epoxy resins like Epon, Araldite and Maraglas. Extreme care is necessary while handling these chemicals since they are carcinogenic and cause skin reactions. The materials are embedded in epoxy resin which is then polymerised to form a hard block. This medium allows smooth cutting of the specimen without damage to tissue on ultra-microtome. Epon embedding mixture (Luft, 1961): Generally embedding is carried out in gelatin capsules of various sizes. Also, there are special moulds available for the embedding of materials. Epon mixture is prepared by mixing solution-A and solution -B in varying proportions according to the degree of hardness required (Table 18.2). Table 18.2. Preparation of epoxy embedding mixture No. 1 2 3 4

Hardness of final block

Solution -A (ml)

Soft Standard Medium Hard

80 60 50 30

Solution-B (ml) 20 40 50 70

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Epon

Solution-A Mix 25 ml Epikote 812 and 50 ml of dodecenyl succinic anhydride (DDSA) Solution-B Take 50 ml Epikote 812 and mix slowly with 50 ml of nadic methyl anhydride (NMA)

During mixing of solutions A and B, 2% benzyl-dimethylamine is added to speed up the process of polymerisation. Gelatin capsule is filled with embedding medium and the specimen to be embedded is kept into the capsule. Allow the specimen to sink slowly through the viscous fluid. The polymerisation is performed by keeping this preparation in an oven at 60°C for 24 to 48 hrs. Araldite mixture (Mercer and Birbeck, 1966): Araldite mixture consists of the following ingredients: Dodecenyl succinic anhydride Araldite M 2,4,6-trimethylaminomethyl phenol (DMP-30) Dibutylphthalate

20 ml 20 ml 0.6 ml 2.0 ml

The mixture is heated to 50°C and after thorough mixing, this can be used as described above for Epon. The other embedding media such as Vestopal W (Ryter and Kellenberger, 1958) and Maraglas epoxy resin (Desai and Desai, 1980) are also employed.

18.1.3 Ultramicrotomy Since electrons have low penetrating power, ultrathin sections of 60-100 nm thickness are needed to observe under TEM. This can be achieved by employing ultra-microtome fitted-with glass or diamond knife. If high quality sections are needed, diamond knife should be used which is available with water trough attached. Water trough is prepared by wrapping a short strip of scotch insulating tape (no. 33) around the back of the knife and sealing it with paraffin wax (Fig. 18.1). After trimming, the blocks are fixed on to a microtome and the sections are taken. Sections float onto the surface of water present in the trough and remain together in the form of a ribbon (see Fig. 18.1). The thickness of the sections cut with ultramicrotome will be judged with the help of colours generated in the ribbon by reflection of light. Grey and pale gold to light silver-coloured sections having thickness of about less than 10 nm and 80 nm respectively is

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useful for ultrastructural studies. Then the floating sections are collected from the trough onto a small circular copper grid (Fig. 18.2; for preparation of grids see Desai and Desai, 1980).

Fig. 18.1. Glass knife with floating sections in the trough prepared from the adhesive (Redrawn from Desai and Desai, 1980).

Fig. 18.2. Copper mesh grid.

18.1.4 Contrast Enhancement In light microscopy, various staining techniques are employed largely to improve the specimen contrast with respect to the background. Many staining

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reactions rely on colour contrast to achieve the contrast image of the specimen. But colour contrast is not seen with transmission electron microscope since the image is viewed either on a fluorescent screen or recorded photographically in black and white. The contrast in the TEM image of a specimen arises largely due to the scattering of the electron beam by its interaction with atomic nuclei in the specimen (Agar et al., 1974; Hayat, 1975). The number of scattering increases with the thickness of any element encountered and with its atomic number (i.e., mass thickness). Thus, the scattered intensity for any given solid angle is proportional to the mass thickness of the specimen (O’Brien and McCully, 1981). Chemically, biological materials are mainly composed of elements like carbon, nitrogen, oxygen and hydrogen. These elements have low electron scattering power, hence the poor quality of the image of the specimen is seen. Various techniques are being developed to improve the quality of the TEM image of the specimen [e.g., lead staining of sections by Lever (1960), lead citrate staining by Venable and Coggeshall (1965), etc.]. 18.1.4.1 Staining

Some fixatives like osmium tetroxide (OsO4) and potassium permanganate (KMnO4) have got staining ability. The materials fixed in these fixatives are further stained to increase the contrast. Staining can be done before dehydration, during dehydration or after sectioning. Two general types of staining techniques are currently employed for TEM studies: (a) Negative staining and (b) Positive staining. (a) Negative staining: This technique is widely applied to study cell fractions, viruses and other particulates. Negative staining provides a lighter image against the electron-dense background (Fig. 18.3). This technique was extensively reviewed by Haschemeyer and Meyrs (1972). Hall (1955) was the first person to employ this technique to study the structure of the bushy stunt virus. In the negative staining technique, particles (biological material) are taken in a dilute solution (0.1-2%) of the salt of a heavy metal and placed on a support film covering an EM grid. Excess fluid is removed, allowing the film to solidify by evaporation. The metal salt (acting as a stain) encases the particles on the support film and penetrates any irregularities to which it has access. Now the negatively stained particles appear as light areas or holes due to their poor electron-scattering capacity relative to the encasing metal salt (for details see O’Brien and McCully, 1981). Various metal salts viz. sodium or potassium phosphotungstate, uranyl acetate, uranyl formate and ammonium molybdate are currently used in this technique. Negative staining has been widely used in routine diagnosis of plant viruses (Desai and Desai, 1980).

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Fig. 18.3. Negatively stained specimen (Redrawn from Haschemeyer and Meyers, 1972).

(b) Positive staining: Positive staining makes electron-transparent particles visible against a relatively transparent background. This technique is more useful for ultrathin sections. Stains containing lead e.g., lead tartarate, lead citrate, etc., give more satisfactory results. Many researchers prefer to stain first with 2% uranyl acetate and then with lead salts. In some cases, Potassium permanganate (KMnO4) has been used to study the lignification of the xylem (Zhou et al., 2015). Layering structure (alternating narrow and broad layers of several wall thickening) of sclerenchyma fibres and xylem vessels in Miscanthus sinensis is clearly seen when the sections are stained with KMnO4 (Figs. 18.4A, B; 18.5A, B).

Fig. 18.4. TEM images showing layering structure of sclerenchyma fibre (Sf) in Miscanthus sinensis cv. internode tissue, stained with 1% w/v KMnO4. (A) Sf adjacent to xylem; (B) Sf adjacent to phloem. Ccml, cell corner middle lamella; Cml, compound middle lamella; Mxv, metaxylem vessel; Par, parenchyma; Sw, secondary wall (Adopted from Zhou et al., 2015).

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Fig. 18.5. TEM images showing layering structure of M. sinensis internode tissue. (A) Metaxylem vessel (Mxv); (B) protoxylem vessel (Pxv) (Adopted from Zhou et al., 2015).

18.1.4.2 Shadow Casting

Shadow casting is widely used as means of contrast enhancement for electron microscopy. This technique involves the deposition of an electron-dense material at an angle over a specimen. For this purpose, various heavy metals viz. chromium, nickel, platinum, uranium, gold or their alloys are evaporated in the specially designed apparatus. In this process, the specimen acquires a shadow on the uncoated side (Fig. 18.6). This technique provides the topographical details of the specimen.

Fig. 18.6. Deposition of metal evaporated onto a specimen in Shadow-casting technique.

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Shadow casting was widely applied to study the cellulose orientation in extracted walls (Preston, 1974; Roland, 1978) and to demonstrate the substructure of scales and flagellae of marine algae (Manton, 1966). 18.1.4.3 Freeze Etching

In freeze etching, the specimen soaked in 20% glycerine is frozen (-100oC) and fractured with a cool knife. The exposed surface is then etched, causing sublimation of the ice block. To give contrast to the etched surface, shadow casting with heavy metal is performed. Later carbon replica is prepared from this preparation. The shadowed specimen is destroyed by chemical treatment and the carbon replica is examined under EM. This technique is useful to study cell walls and membrane structures.

18.1.5 Protocols for Transmission Electron Microscopic (TEM) Studies 18.1.5.1 Sections Staining with Lead Hydroxide (Lever, 1960)

The specimen is fixed in OsO4 and embedded in methacrylate or araldite. Preparation of stain: Dissolve 1 gr of lead hydroxide in 100 ml of distilled water. Later boil the solution, cool and filter it. To filtrate (faintly clouded), add 2 N Potassium hydroxide drop by drop until it becomes clear; the mixture contains some lead in plumbite form (salt formation). Staining Procedure i. Float the grids face down over a few drops of staining solution for 5 mins in a covered watch glass. ii. Remove the excess stain and wash with distilled water. iii. Agitate each grid in 1% KOH for 3-4 secs and immediately wash with distilled water. iv. Grids are blotted with filter paper, dried and examined in TEM. Note: Lead impregnation of the specimen appears to be reduced when treatment in alkali is prolonged. 18.1.5.2 Staining with Lead Citrate (Venable and Coggeshall, 1965)

Preparation of Stain Dissolve 0.01-0.04 gr of lead citrate in 10 ml of distilled water in a centrifuge tube to which add 0.1 ml of 10 N Sodium hydroxide and shake it vigorously till lead citrate dissolves. Staining Procedure i. Stain grids in a small quantity of lead citrate for 1-5 mins. ii. Wash in distilled water, twenty dips in each of the two or three vessels.

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iii. Air-dry on filter paper. Double staining with aqueous uranyl acetate followed by lead citrate can also be performed. For different tissues, trials should be made with various concentrations of lead citrate as well as time intervals (Lane and Europe, 1965). 18.1.5.3 Removal of Uranyl Acetate Precipitation using Oxalic Acid (Avery and Ellis, 1978)

Material is fixed in 3% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.2) and post-fixed in 1% OsO4. Staining: Specimens are divided into three groups. i. Enbloc staining with 0.5% uranyl acetate in 95% ethanol for 1 hr. ii. Enbloc staining with 2% aqueous uranyl acetate for 8-10 hrs. iii. Without enbloc staining. Dehydration: Dehydrated in ethanol, embedded in Spurr’s (1969) low viscosity plastic. Removal of precipitate from sections: Silver gold or gold sections are processed on Marinozzi rings (Marinozzi, 1961) and treated with 1-15% aqueous oxalic acid, 15% oxalic acid in 5% methanol, 5% sodium thiosulphate or 1% EDTA. Washed with respective solvent and post-stained with 5% uranyl acetate in 50% methanol followed by lead citrate (Reynolds, 1963). Examine before and after oxalic acid treatment without post-staining. Result i. Methanolic oxalic acid 15% for 30 mins removes precipitate from stained sections. ii. Aqueous oxalic acid 1-15% for 30-60 mins at 40°C destained the tissue. iii. Complete removal of precipitate and staining is done with 1% aqueous oxalic acid for 30 secs. iv. Sodium thiosulphate does not remove the precipitate. v. EDTA 1% (pH 12.5) for 30-60 mins does not remove precipitate. For removal of precipitates from stained sections refer to O’Brien and McCully, 1981. 18.1.5.4 Fixatives used for Transmission Electron Microscopy

A. Caulfield’s osmium tetroxide-sucrose (Caulfield, 1957) Veronal-acetate-HCl buffer 2 ml 8 ml 2% OSO4 Distilled water 4 ml 0.1 N HCl 2 ml Crystalline sucrose

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B. Buffered Potassium Permanganate (Luft, 1956) Buffer and 0.1 N HCl stock solutions are prepared as given in Appendix. Fixative 1.2% KMnO4 solution Veronal-acetate-HCl buffer

1 part 1 part

The pH is adjusted to 7.4-7.6 with 0.1 N HCl. The specimen is fixed for 15 mins-12 hrs at OoC. Later specimen is rinsed for several mins in cold 25% ethanol and brought down to room temperature in fresh 25% ethanol. Then the specimen is dehydrated through graded ethanol. C. Phosphate-buffered osmium tetroxide (Millonig, 1961, 1962) Phosphate buffer 50 ml OsO4 0.5 gr The final fixative is prepared as follows: Stock buffer solution 5 ml 2% OsO4 10 ml The stock solution has a molarity of about 0.2 and pH 7.4. The pH can be adjusted with HCl. The final concentration of OsO4 in this solution is 1.33%. D. Potassium Permanganate-Potassium bichromate (Kawakami, 1961) i. Fix specimen in 0.5% potassium permanganate, containing 1% calcium chloride neutralised with calcium carbonate for 10-30 mins. ii. Wash in 1% calcium chloride neutralised with calcium carbonate. iii. Post-fix with 0.5% potassium bichromate neutralised with potassium hydroxide to pH 7.2 and containing 1% calcium chloride for 10-30 mins. iv. Wash again with neutral 1% calcium chloride. v. Dehydrate with ethanol series. vi. After staining, sections may be immersed in 1% lead acetate adjusted to pH 4.2 with 1% acetic acid for 10 mins. E. Glutaraldehyde-Osmium tetroxide mixture (Trump and Bulger, 1966) 50% aqueous solution of glutaraldehyde 1 part 4% aqueous solution of OsO4 2 parts 0.1 M collidine buffer 5 parts F. Glutaraldehyde-Potassium bichromate-Osmium Tetroxide (Sugihara et al., 1966) Potassium bichromate mixture (pH 7.2) Potassium bichromate (5%) 16 ml 2.5 N KOH 2 ml Distilled water 2 ml Sorensen’s phosphate buffer 20 ml

Electron Microscopic Studies 383

i. ii. iii. iv. v. vi.

Procedure Fix the specimen in 6.25% glutaraldehyde buffered with 0.075 M NaCacodylate (pH 7.2) for 30-60 mins. Wash the specimen thrice in cold 0.33 M sucrose solution. Later fix the specimen in potassium bichromate mixture for 30 mins. Wash again thrice in the sucrose solution. Post-fix in 1 to 2% osmium tetroxide buffered with veronal-acetate (pH 7.5). Post-stain the sections by immersion in uranyl acetate followed by lead acetate.

G. Glutaraldehyde-acrolein-Potassium permanganate (Hayat, 1968a, b) Glutaraldehyde-acrolein mixture is a rapid general preservative and potassium permanganate enhances contrast. It is a good fixative for algae, fungi and tissues of higher plants. It preserves well the cytoplasmic details including lipoprotein membranes. Procedure i. Fix specimen in a mixture (1:1) of glutaraldehyde (3%) and acrolein (3%) buffered with 0.2 M sodium cacodylate (pH 7.2) at 4°C for 1 to 2 hrs. ii. Later wash the specimen in the same buffer. iii. Post-fix in aqueous 6% potassium permanganate at 4°C for 20 mins. iv. Rinse the specimen twice in the same buffer. v. Post-stain the sections by immersion in lead citrate for 5 mins. H. Glutaraldehyde-acrolein-paraformaldehyde-osmium tetroxide 10% glutaraldehyde 6 ml 10% acrolein 3 ml 6% paraformaldehyde 5 ml 0.2 M buffer 5 ml Distilled water 1 ml

18.2 Electron Tomography in Plant Cell Studies Electron tomography (ET) protocols are mainly based on imaging a biological specimen, particularly a plant cell, at different tilt angles by transmission electron microscopy (TEM). ET can be applied to both plastic-embedded and cryo samples (frozen samples). In the advancement in TEM, direct electron detection, automated image collection and image processing algorithms allow for 2-7 nm scale axial resolution, in tomographic reconstructions of cells and organelles (Otegui and Pennington, 2018). Conventional ETM can resolve cellular macromolecules at a resolution of 1-2 nm. However, it generates 2D projections and requires specimen fixation

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and processing, which sometimes causes articrafts and unwanted changes to the cell organelles.

Fig. 18.7. Electron tomographic reconstruction of aleurone and starchy endosperm Cells at 22 DAP. (A) Tomographic slice (4.3 nm thick) of a starchy endosperm cell containing multiple protein bodies (asterisks) within the ER. (A’) The tomographic model is derived from the tomogram shown in (A); (B) Tomographic slice (4.3 nm thick) of an aleurone cell. PSVs with large inclusions (asterisks) and LB which is the predominant storage compartment in these cells. (B’) The tomographic model is derived from the tomogram shown in (B). Here only PSVs, PrVC, and a G are shown. This is a serial tomogram reconstructed from four serial 250-nm-thick serial sections. (C) and (C’) Tomographic slice and tomographic model of a PSV. In this, a large inclusion (asterisk), IM and a GL are seen. CW, cell wall; ER, endoplasmic reticulum; G, Golgi stack; GL, globoid; IM, intravacuolar membranes; LB, lipid bodies; PSV, protein storage vacuole; PrVC, prevacuolar compartments. Bar = 100 nm. (Reprinted from Reyes et al., 2011, The Plant Cell; with copyright permission from Oxford University Press).

Electron Microscopic Studies 385

Electron tomography (ET) approaches are instead based on collecting images of an individual sample at different angles and combining the individual 2D projections onto a 3D reconstruction of tomogram via either Fourier or real space methods (Brooks and Chiro, 1975; Ercius et al., 2015). ET of Plastic-embedded samples and Cryo-ET of vitrified material have allowed cell biologists to image macromolecular complexes and organelles in their native, 3D cellular context with an axial resolution of 2-7 nm or even higher when combined with image analysis approaches such as sub-tomogram averaging (Schur et al., 2014; Castano-Diez et al., 2017). In combination with other molecular techniques, electron tomography is a powerful technique to study the fundamentals of cellular processes. ET of plastic sections and a few Cryo-ET studies of plant cells and algal cells have been published and reporting the variety of cellular structures and processes, such as preprophase band assembly (Mineyuki, 2014; Takeuchi et al., 2016; Karahara et al., 2009), phragmoplast organisation (Austin et al., 2005) and organelle dynamics during the cell cycle (Segui-Simarro, 2006). Further, Reyes et al. (2011) studied the zein-rich protein inclusions in maize endosperm cells. In this study, a combination of molecular approaches, in vivo imaging of fluorescent proteins and structural analysis by electron tomography have been adopted to reveal the synthesis and transport of storage proteins in aleurone cells. Semi thick sections (250 nm) from Epon-embedded endosperm samples at 14 and 22 days after pollination (DAP) have been taken. They found the protein bodies enclosed in a continuous ER network in the starchy endosperm (Fig. 18.7 A, A’). The architecture of the endomembrane system in aleurone cells is strikingly different with protein storage vacuole (PSV) and lipid bodies occupying most of the cellular volume (Fig. 18.7 B, B’). In addition to the zein-rich protein inclusions, the aleurone PSVs contained one or more globoids (crystals of phytic acid salts) and a large system of intravacuolar membrane (Fig. 18.7 C, C’).

18.3 Scanning Electron Microscopic (SEM) Studies The preparation of biological material for EM depends on the nature of the sample itself. In fact, all the techniques currently available are an attempt to gain optimum structural and cellular preservation with minimum shrinkage and distortion. Robards (1978) defines three basic criteria for the successful preservation of biological samples mechanical stability, complete removal or immobilisation of water and effective conduction of absorbed electrons to earth. There are few samples that meet these criteria for a short period of time without any processing at all (Baker and Parsons, 1971; Ledbetter, 1976). However, in general, some sample preparation is needed since few samples are able to withstand vacuum desiccation. Unless the water is stabilised or removed before keeping the specimen in the vacuum of SEM, it will boil and

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lead to deformation and cellular collapse (Boyde and Wood, 1969; Robards, 1978; Hayat, 1974). The sample preparation for SEM varies from the simple technique of water stabilisation to the more complex process of water removal by freeze drying, critical point drying, and resin etch techniques (O’Brien and McCully, 1981). The simplest preparation technique is to stabilise the water by quickly freezing the sample in Freon cooled by liquid nitrogen, and then transferring the sample to a modified microscope stage which can maintain the temperature at -100°C while it is being viewed under vacuum (Heywood, 1969; Trougton and Donaldson, 1972; Troughton and Sampson, 1973). Freezing of the specimen must meet Robards’ (1978) three basic criteria: rigidity of the sample (therefore no mechanical vibration), immobilisation of water (no surface tension stresses) and moderately effective earth created via the solidified water (for a detailed discussion refer to O’Brien and McCully, 1981).

18.3.1 Sample Preparation for SEM The surface of an object to be studied by SEM should meet some of the important characteristics such as it must be free from foreign particles or cell debris, stable when put under a high vacuum, remain stable after exposure to electrons, and should develop as few surface charges as possible, and it must emit enough secondary electrons. Biological specimens need special attention because most of them are soft, full of extracellular fluids, and poor conductors. There are several preparative techniques depending on the nature of the tissue and the type of studies to be carried out. Preparation of specimens for SEM studies involves the following steps. Specimen selection Specimen size depends upon (a) stage capacity (x-y movement) of the SEM, (b) type of fixation and (c) drying process used. It should be the smallest size possible, to reduce the preparation artifacts. Tissue culture, single cells or microorganisms can be processed in suspension on attached to a suitable base (coverslips) before going through preparative procedures. As extracellular products such as mucus, blood, other body fluids and tissue fragments may obscure the surface to be examined, they must be removed by gentle washing with cold isotonic buffer. Microorganisms and cell suspensions can be separated from their liquid environment by centrifugation. Sterilisation of specimen Apart from hard objects such as teeth, bone and wood, all other biological materials must be stabilised to prevent from undergoing structural changes, which is done by chemical fixation or physical fixation (cryo-fixation). the

Electron Microscopic Studies 387

fixative and the fixation protocol are same as in the case of TEM specimen preparation. Washing and dehydration To remove the unreacted fixative, the specimen is washed in buffer (which was used for preparing the fixative), especially after secondary fixation with OsO4. The later may be reduced by dehydrating agents, causing precipitates to form on the specimen thus obscuring the surface morphology. Chemically fixed objects must be dehydrated before drying. The purpose of dehydration is to remove water before drying. The most commonly dehydrating are ethanol and acetone. To avoid initial osmotic damage to specimen, starting solvent (acetone or ethanol) should be 30% or still lower. Then specimens are dehydrated using increasing concentration of the dehydration agent up to dry absolute acetone stage. After dehydration, specimens are dried and there are different approaches of drying such as air drying and critical point drying. Air drying is the simplest method of drying, but it creates maximum distortion. It causes flattening of specimen surface because of compressive forces of surface tension of liquid gas interface, whereas critical point drying is the commonly used techniques to dry biological samples. This sample is transferred from an organic dehydration medium (acetone) to drying medium (liquid CO2 or freon 13) in a chamber that is cooled and put under pressure. Surface coating of specimen When the dehydrating agent (acetone) has been completely removed and impregnated with the metal (gold) that coats the surface of the specimen. Many metals, when heated in vacuum, evaporate readily into a mono-atomic state. The high temperature required to start evaporation of the materials can be achieved by resistive heating. The coating involves the following steps: The specimen is mounted on rotating specimen stage to get even coating; thin wire of metal to be evaporated is wrapped around tungsten filament; then the chamber is evacuated; the tungsten filament is heated by supplying voltage; finally, metal will start evaporating and a thin layer will deposit on a rotating specimen. Sometimes it causes uneven coating; whereas sputter coating allows uniform coating on the specimen even on the parts that are not directly facing the metal to be evaporated. Hence it is the most acceptable technique. In this technique, specimens are placed on a cooled base plate which functions as an anode. The other one, which is the metal (e.g., gold) to be evaporated, is positioned above the specimen. Then the chamber, in which the cathode and anode are located is evacuated. Inert gas such as argon is flooded into the chamber and is again evacuated. The negative high voltage applied to the cathode. This result in glow discharge and argon gas molecules gets ionised. The positively charged ions move towards the cathode and strikes on it, there by releasing atoms. The dislodged metal atom due to numerous collisions

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with the gas molecules get deflected several times and hit the specimens at different angles. Thus, a uniform thin layer is deposited on the specimens. The thickness of coat depends up on the current applied, anode and cathode distance and sputtering time. With Balzer SCD 020 sputter coater, using gold as the target (cathode), a coating of about 35 nm thickness can be obtained under the conditions, such as an electrical current of 21.5 mA, pressure 0.05 mbar, argon gas, cathode to anode distance about 30 nM and sputtering time one minute. After the metal coating, specimen is ready for observation under SEM.

18.3.2 Protocols for SEM Studies 18.3.2.1 General Protocol of Specimen Preparation for SEM

Sample size for SEM studies should be 2-3 mm. Steps to be followed are given below: i. Primary fixation is performed in 2.5% glutaraldehyde for 2-6 hours at 4°C. ii. Washing is due with 0.1 M phosphate buffer (3 washings), each washing of 15 minutes at 4°C. iii. Post-fixation (optional) is carried out in 1% OsO4 for 2 hours at 4°C. iv. Final washing is with 0.1 M phosphate buffer for three changes each of 15 minutes at 4°C. v. Dehydration is performed in various grades of acetone (30%, 50%, 70%, 80%, 90%, 95%, 100%), 15 minutes in each grade under 4°C temperature. vi. Critical point drying – Critical point drying with liquid CO2 at its critical point, i.e., 31.5°C at 100 psi (pound per square inch). vii. Mounting of samples should be done on to the aluminium stubs. Sputter coating is preferred; using silver or gold, coat the samples with 20-30 nm thick film. Now it is ready for observing under SEM. viii. Image interpretation and magnification. Magnification on EM images Organelles are measured in micrometres (µm) and nanometre (nm) 1 mm = 1000 µm 1 µm = 1000 nm 1 nm = 10 A° The true size of objects in micrograph can be calculated by using simple and useful thumb rule, i.e., there are 1000 µm in one mm, therefore,1 µm object will appear to be 1mm in size when the magnification is ×1000. To put a scale mark representing 1 µm on micrograph, simply draw a line as many mm long as there are thousands in magnification. For example, Magnification × 50000, i.e., 1 µm is magnified 50000 times; since 1mm equals 1000 µm, 1 µm equals to 50 mm or 5 cm.

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i.e.,

Precise measurement of objects in the micrograph The original size of the organelle seen from ultra micrograph can be measured either in micrometre or nanometre by using following calculation: Size of micrograph (µm) × 1000 Magnification

= True size in µm

or Size of micrograph (nm) × 1000000 Magnification

= True size in nm

Calculating the total magnification of a picture from Bar magnification e.g.,

2 µm 1 µm Since 1 mm 1 µm

= = = =

10 cm 5 cm = 50 mm 1000 µm 50000 µm (50000 x)

ix. Precautions while working in the EM lab Some of the following precautions should be taken while working in the EM laboratory. These are as follows: a. Always use disposable hand gloves while handling the EM chemicals. b. The glassware should be kept clean and dry before starting the tissue processing protocols. c. If the skin comes into contact with any EM chemical, it should be cleansed with water. d. Care should be taken while weighing uranium salts; uranyl acetate should be weighed in a room free from the air current. e. Always use detergent to clean hands if you touch spur or other plastics. f. Use fume hood while dealing with OsO4. g. Never dispose the used EM chemicals into wash basin. Collect them separately into bottle or boxes and dispose into a pit or incinerator.

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18.3.2.2 Partial Embedment in Epoxy Resins (Chambers and Hamilton, 1973)

i. Fix the material in 5% glutaraldehyde. ii. Dehydrate in acetone series of 25, 50, 75, 95 and 100%. Two changes are given in 100% acetone. iii. Add Spurr’s resin, mix for 4-5 hrs. Spurr’s resin has got the following ingredients: ERL-4206 (Vinyl cyclohexene dioxide) D.E.R. 736 (Diglycidyl ether of polypropylene glycol) NSA (Nonenyl succinic anhydride) S-l accelerator

10.0 gr 6.0 gr 26.0 gr 0.4 gr

iv. After 36-48 hrs infiltration in pure resin mix, cut tissues to expose surfaces which are being studied under SEM. v. Place in oven at 70°C for 1 hr. vi. Take out the material and rinse with acetone. vii. Again, put it in the oven in acetone for 5 mins. viii. Then remove from the oven and rinse vigorously with acetone till tissue has a mat like appearance when viewed with a dissecting microscope. ix. Place tissue on filter paper and polymerise at 70°C. x. Mount on stubs and coat with gold. This method gives good preservation of wall ingrowths of transfer cells and cells maintain turgidity and life-like appearance (O’Brien and McCully, 1981). 18.3.2.3 New Imprinting Material for SEM (Dwivedi et al., 1991)

Seeds of Cicer arietinum and Sansevieria roxburghiana are taken. Procedure i. Dissolve thermocol material in chloroform to obtain 5% solution (w/v). ii. Clean and moisture the surface of the seed with a cotton swab soaked in chloroform. iii. Spread the solution on the surface with a glass dropper and allow it to dry for 5 mins. iv. After drying, the film on the surface is peeled off and mount (imprint surface should be kept above) on specimen stub coated with double sided adhesive tape. v. Stubs are then sputter-coated with approximately 200 A° thick coating of gold, vi. Samples are scanned with JEOL JSM 35 C SEM at 15 KV and photographed.

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Negative replica provides reliable information, and this can be compared with that obtained through customary methods. 18.3.2.4 New Drying Agent for SEM Study of Pollen (Chissoe et al., 1994)

Sample Preparation i. Pollen grains of Avena are collected from herbarium sheets and freezer storage. ii. Pollen stored in test tubes are acetolysed, sieved through fine wire mesh (to remove undigested plant fragments), and placed on sucrose pads (for removing finer particles). iii. Dehydrate in graded ethanol series, 5 mins in each grade. iv. Give three changes in absolute alcohol, 5 mins each. v. Later two changes of 100% hexamethyldisilazane (HMDS) are given. vi. Pollen is centrifuged and it is pipetted on to copper specimen mounts (JEOL-TEM SCAN 100-CX 2.4 × 5.6 mm). vii. Sprinkle the pollen onto adhesive-coated specimen mounts after HMDS has evaporated. (Specimen mounts are coated with thermoplastic adhesive “Tempfix”). viii. In some test Tempfix is sputter-coated with gold for 4 mins prior to pollen application. ix. All samples are sputter-coated with gold for 4-6 mins and observed under JEOL-880 SEM fitted with lanthanum hexaboride gun. Pollen grains are almost distortion-free and retain original shape with HMDS treatment. 18.3.2.5 Chromosomal Study with SEM (Martin et al., 1994)

Root-tips of three-day old seedlings of Barley (cv. ‘Igri’), barley telotrisomic lines and wheat-barley telosomic addition lines are taken. Synchronise the meristematic tissue either by pretreating in ice water for 24 hrs or by following Pan et al. (1993) and Busch et al. (1994). Later fix the material in 3:1 ethanolacetic acid and store it at 20 ºC until preparation of specimens. Digestion i. Cut and digest the root-tips in a solution of 2.5% cellulase, 2.5% pectinase in 75 mM KCl (pH 4.0) for 50-60 mins at 30°C. ii. Filter the solution through nylon mesh (100 µm), iii. Treat (hypotonic) with 75 mM KCl for 8 mins at room temperature. iv. Centrifuge the suspension for 7 mins at 80 g. v. Resuspend the protoplasts in 3:1 ethanol-acetic acid and pellet again for 7 mins at 80 g, perform centrifugation four times. vi. Finally resuspend the pellet in an appropriate volume of fixative and take a drop on ice-cold, clean slide.

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vii. Spread protoplasts on the slide by observing under ‘ZEISS Axiophot’ light microscope. viii. Add a drop of 45% acetic acid before completely drying off the specimen. ix. Cover the preparation with a coverslip, gently squash and subsequently freeze on dry ice. x. Remove the coverslips and transfer the slides to a fixative containing 2.5% glutaraldehyde in 75 mM cocalodylate buffer and 2 mM MgCl2 (pH 7.4). xi. Osmium impregnation, dehydration, critical point drying, and sputter coating are performed (see section 18.3.1). xii. Observe chromosomes under Hitachis 4100 field emission scanning electron microscope. xiii. Microdissect the chromosome with glass rods using a ZEISS inverted microscope. Results: This procedure gives good results for chromosome studies. This also allows to observe plant chromosomes of all stages of life cycle. 18.3.2.6 Sequential SEM Observations of the Reproductive Meristem by Non-invasive SEM Technique (Hemandez, 2001)

i. Expose the floral meristem of sunflower (Hlianthus annuus L) CV. Dekalb G100 (Dekalb Argentina S.A.) at a floral stage 5 (Mart and Palrner, 1981), i.e., 32-34 days after seedling emerges by removing the last formed apical leaves and involucral bracts. ii. Sequential replicas of a single receptacle surface are obtained during floret formation. To make a primary mould, exposed meristem surface is covered daily with a polyvinyl impression material SDS Kerr’s extrude wash (Sybron Dental Specialities, West Collins Orange, CA, USA). iii. After 10 mins of polymerisation, remove the primary mould with tweezers. iv. To protect the apex from desiccation, the receptacle meristem is immediately covered with a cap made from wet tissue paper and aluminium foil. v. Invert the mould and affix to the microslide. vi. To prepare the cast, fill the moulds completely with Spurr’s resin (hard mixture) and place in an oven at 70°C for 72 hrs. vii. After hardening, mount the cast on aluminium SEM stubs and sputter coated (Green and Linstead, 1990; Hemandez and Green, 1993). viii. Observe each replica in a JEOL JSM35CF scanning electron microscope at 10 KV and photograph.

Electron Microscopic Studies 393

18.3.2.7 Orchid Seed Study with SEM (Khasim et al., 2015; Ramudu et al., 2020)

Reagents Glutaraldehyde 2.5%; 0.2 M cacodylate buffer (pH 7.2); graded ethyl alcohol: acetone series Procedure i. Seeds of orchid species such as Calanthe triplicata, Cymbidium aloifolium, C.giganteum and Malaxis densiflora were collected and fixed in 2.5 glutaraldehyde prepared in 0.2 M cacodylate buffer (pH 7.2), kept in room temperature for two hours.

Fig. 18.8. SEM photographs of orchid seeds under high magnification. (A) Part of the testa in Malaxis densiflora; (B) Testa cells with transverse walls in Calanthe triplicata; (C) A pore between testa cells in Cymbidium aloifolium; (D) Part of the testa with pores in Cymbidium giganteum.

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ii. Later seeds samples were dehydrated in graded ethyl alcohol, acetone series. iii. Seeds were dried at critical point dryers. iv. After dehydration, these dehydrated seeds were mounted on to copper stubs and gold coated for five minutes. v. Now the processed specimens were examined and photographed on a HITACHI 53000 N model SEM. Results SEM photographs clearly showed the testa cell wall thickening in orchid seeds; Malaxis densiflora and Calanthe triplicata (showed the transverse testa cell wall thickenings whereas Cymbidium aloifolium and C. giganteum with longitudinally oriented cell wall thickenings (Fig. 18.8 A, B, C, D). Various shapes of orchid seeds such as filamentous, rope-like, spindle shaped and fusiform shape were also studied with the SEM (Ramudu et al., 2020).

19 Autoradiography

Basic Principle The technique of Autoradiography involves the incorporation of radioactive substance into a cell, and subsequent detection of that material using a photographic emulsion. The primary source of radioactive substance used in cell biology is an organic molecule containing tritium, the radio-active form of hydrogen. Radioactive carbon, phosphorous and iodine are occasionally employed. Tritiated thymidine (3H-thymidine) is often used to study the synthesis and localisation of DNA. Thymidine is a soluble base which is specific to DNA. Upon fixing the cells for histological observations, the DNA molecules (with their incorporated radioactive thymidine) are precipitated or crosslinked as permanent parts of the cell. Un-incorporated thymidine is removed from the cell, as it remains soluble and is disposed off in the tissue washing procedures. When the tissues are sectioned and taken on to a glass slide, they contain radioactive nuclei; here nuclei should be ‘S’ phase of cell cycle during the exposure of cells to 3H-thymidine. Later photographic emulsion is applied directly over the section, so that it would expose to radioactive substance. Developed slides are now examined under microscope. Process of autoradiography is outlined in Fig. 19.1 (See also Alberts et al., 1994). If one wants to localise the newly synthesised DNA in the regenerating cells of rat liver, here the cells are actively dividing and, therefore, DNA synthesis occurs; the first step in this technique is to supply the precursor, i.e., thymidine which is rapidly taken up by the cells and it is specific for DNA. Thymidine is labelled with tritium (H3) and it is injected in the form of H3thymidine into the cells. Now cells are allowed for some time to synthesise new DNA. The tissue is then fixed, embedded and sectioned. Slides (or grids) with sections containing new synthesised H3-DNA are then coated with a thin layer of very-fine-grain photographic emulsion. This step should be performed in dark since emulsion is light sensitive. The coated slides are kept in dark for a period ranging from days to weeks. Photographic emulsions are composed of silver halide crystals which are reduced to metallic

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Fig. 19.1. Autoradiography (Adopted from Alberts et al., 1994).

silver by the energy from light in photography. In autoradiography, the energy is from the radioactive disintegrations of the isotopic H3 in the DNA. Because H3 is a weak energy emitter (see Table 19.1), its radioactive disintegrations travel only 1 µm and will reduce silver grains present just above the site of H3-DNA. Following the exposure period, the emulsioncoated slides are incubated in photographic developer (in the dark only). Table 19.1. Isotopes commonly used to study the Biological Materials Isotope H3 14C 32P

Maximum Energy (Mev) 0.018 0.155 1.71

Half-Life (Years) 12.3 5570.0 0.04

Autoradiography 397

This substance essentially supplies electrons to the tiny reduced silver grains, and they become precipitated with more silver. These precipitated silver grains are clearly visible under microscope. Following fixation to stop these reactions, the slides are observed under light microscope. Silver grains, 0.2-1.0 µm in diameter, appear over the region where the newly synthesised H3-DNA is located (nucleus). In the same way, RNA(H3-uridine as precursor) and proteins (H3-amino acids as precursor) can be localised in the cell auto radiographically.

19.1 Protocols 19.1.1 General Protocol for Autoradiography Reagents and other materials Alcohol-acetic acid fixative Xylol, permount 3H-thymidine Onin sets, Jars Materials for feulgen reaction Paraffin embedding Rotary microtome Kodak nuclear track emulsion Kodak D19 Developer Water bath at 42oC Kodak fixer Glass slide, cover slips Compound microscope Procedure i. Take onion on the small jar/beaker filled with water and allow it to develop roots for 4-7 days. ii. Then transfer the root tips to another beaker containing the 10 µ/ml ^3H-thymidine solution. Root tips are be remained in the radioactive solution for 1 hr at room temperature. iii. After the radioactive treatment, rinse root tips by dipping several times, in water taken in another beaker and allow them to remain in the beaker for four hours. iv. Cut root tips into pieces and fix in alcohol-acetic acid (3:1) overnight. v. Take out root pieces and wash them thoroughly for several minutes and put them in 1N HCl at 60oC for 12 minutes. vi. Dehydrate the root tips in 70%, 90% and 100% alcohol for 20 minutes in each grade, embedded in paraffin wax and section at 10-15 microns in rotary microtome.

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vii. Take sections in the microslide and deparaffinise in alcohol: Xylene series (3:1, 1:1, 1:3 – alcohol: xylene). viii. Stain the sections with Feulgen reaction. ix. Rehydrate the sections briefly in 70, 90 and 100% alcohol for two minutes each. x. In a dark room, melt some amount of liquid auto-radiographic emulsion on a water bath at 42°C. xi. Place two slides back-to-back and dip slowly into the moulted emulsion. Later take out, allow to drain out excess emulsion and keep the slides in light-proof container and allow them to dry in a vertical position. xii. After drying, place the slides in an opaque slide box containing drierite. Wrap in aluminium foil and place in a refrigerator. xiii. The slides should be kept in the refrigerator until a proper exposure has been made. This may vary from 5 days to two weeks. After one week, a trial slide is taken out and develop the autoradiogram in following method. Examine the slide under bright field microscope. A correct exposure is determined by the appearance of black silver graining over the cells. If the trial slide is correct, then, remaining slides should be processed immediately. xiv. Develop the autoradiogram in the dark as follows: a. b. c. d. e. f.

Develop in Kodak D-19 developer at 20°C for 3 minutes. Wash with distilled water for 16 seconds. Fix with Kodak Fixer for 3 minutes. Wash in running tap water for 15 minutes. Dehydrate by placing in 95%, 100% alcohol for 3 minutes each. Clear in two changes of xylol for 3 minutes each.

xv. Mount the slide with permount by placing the coverslip. Result: Slides are observed under the microscope. Those nuclei exposed to radioactive substance are readily identified. The number of silver grains can be counted to give a quantitative measure of ^3H-thymidine incorporation, that shows DNA synthesis very clearly in the cell. It is also possible to count the cells that are undergoing cell division.

19.1.2 Liquid Emulsion Method (Fieq, 1959; CF Jensen, 1962) i. Use freeze-dried, freeze-substituted or chemically fixed tissue in 1:3 acetic acid: alcohol. ii. Cut the material containing H3 at about 5 µm thickness and if it contains C14 at about 5-10 µm. The tissue may also be stained with Feulgen and squashed. iii. Mount the section on slides pre-coated with gelatin-chrome alum adhesive (adhesive is prepared by dissolving 5 gr gelatin in 1000 ml warm distilled water and add 0.5 gr of chromium potassium sulphate).

Autoradiography 399

iv.

v.

vi. vii. viii.

ix.

x. xi. xii.

xiii. xiv. xv. xvi.

Deparaffinise with xylene or toluene. Rinse in absolute alcohol and airdry. Fill a large beaker (1000 ml) with water at 50°C to serve as a water bath. In another beaker (25 ml) take distilled water about one third the amount of emulsion to be used. The third small beaker tied with fine cheese cloth at the top is taken. Carry all these beakers including glass rod and tissue paper into the darkroom. Bring the emulsion (K5, or K5-Ilford in gel form; life 2-3 months) maintained at 4°C to room temperature. In the dark room (use a Wratten safelight lamp series to illuminate the dark room), open the emulsion container and with the help of glass rod put emulsion in a small beaker containing water. Keeping the bottom of this beaker in the warm water of a large beaker, stir the emulsion and water with glass rod until it forms a rich cream. Carefully pass the emulsion through the cheese cloth covering the second beaker to remove the air bubbles. Remove the cheesecloth from the beaker and wipe out the glass rod with tissue paper. Now pour a small amount of emulsion on the centre of the slide to form a pool roughly of 20 mm. Carefully and quickly spread the emulsion with glass rod. Take care that the tissue should not be rubbed. Place the slide on a glass plate in a box about 2 × 2 × 2 ft size fitted with light-tight door, to prevent any light to enter, and with trays containing calcium chloride. Emulsion will reach full sensitivity only when it is dried (3-5 hrs). Slides are kept in the dark for 2-4 days. After proper exposure time, develop the emulsion in a mixture containing 1.125 gr of amidol (diaminophenol), 4.5 gr anhydrous sodium sulfite, 2 ml 10% solution of KBr and 250 ml water. Develop in the dark for 20 mins at 14°C or less. A stop bath of 1% acetic acid may be used to halt the process of developing the emulsion. Fix in sodium thiosulfate (hypo) at 1/3 saturation for 30-45 minutes or until the emulsion is clear. The hypo solution is maintained at 14°C. Wash in running tap water for about 1 hr. Water temperature should not be more than 14°C during washing since the thick emulsion slows diffusion of chemicals used in developing. Higher temperature causes bubbling in emulsion that leads to ruining of autoradiograph. Place the slides in 70% alcohol for 30 mins. This expands the gelatin, that facilitates the staining of the tissue. Stain with methyl green-pyronin or Azure B for 5-30 mins depending on the type of tissue. Differentiate in water or 70% alcohol. The emulsion should lose most of the dye in this step. Air-dry and mount with any permanent mounting medium.

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19.1.3 Stripping-film Method (Doniach and Pelc, 1950; C.F. Jensen, 1962) i. Freeze-dried, or freeze-substituted or chemically fixed (1:3 aceticacid: alcohol) material is taken. FAA is also a good fixative for autoradiography. ii. Deparaffinise and rehydrate in water. iii. Use Kodak autoradiographic AR10 or V1042 stripping film. This film consists of a glass plate, a layer of gelatin and a layer of 4 µ thick emulsion with grain size of 0.2 µ. The film is stored in a refrigerated heavy metal or lead container. iv. Illuminated by a very dim red light. Cut out the emulsion of an area of about 1½ square in the dark room with a razor blade. Handle the film with extreme care at this step because stretching will cause the formation of latent images. v. Float the film, emulsion-side down, on distilled water at 21-24°C. Allow it for a few minutes. vi. Slip a slide, tissue upward, into the water below the floating film and lift the film out on top of the sections. vii. Now dry the slide by using an electric drier. Care should be taken to prevent dust from blowing and avoid exposure of the film to possible sparking of the drier. The slide is kept in a brown plastic box wrapped with black paper. This box is placed in a desiccator which is refrigerated. viii. Kodak D-19B developer is diluted with distilled water (1:2) and filtered. ix. Develop for 5 minutes at 18°C. x. Fix by placing the slide directly in Kodak acid fixer at 18°C until it becomes clear. xi. Wash in running tap water for 30 minutes at 18°C. xii. Dry at room temperature under dust-free conditions. xiii. Stain in haematoxylin or Unna’s stain. Finally, the slide is dehydrated and mounted in Canada balsam and observe under the microscope.

19.1.4 Restripping Method (Andresen et al., 1952, C.F. Jensen, 1962) Follow steps (i)-(iii) as described above. Mark the slide with India in to help you later in orientation of the restripped emulsion; (iv) Cover the tissue with nylon film. Preparation of nylon film: Dissolve Dupont nylon (type FM 6901) in warm amyl alcohol. Allow drop of this solution to fall on dust-free water surface (film should be thin enough). On a circular wire frame the film is lifted and allowed to dry in a desiccator. This film is placed on the slide to cover the tissue. Dip a glass rod with smooth round tip in the same nylon solution and press on the film in a circle around the sections. Care should be taken not

Autoradiography 401

to enter the air bubbles at this step. The nylon film will break away when the circle is completed. Apply additional nylon solution around the edge to make it watertight. i. Place the stripping film over the slide and the nylon film as described in above protocol and handle the slide in the same way until developing. ii. After exposure, re-strip the film from the sections by cutting the emulsion at the edge of the slide and gently pulling it away from the section. When the film is completely freed from the slide, grip it with a plastic clamp. iii. Now develop the film with clamp for 5 minutes at 18°C in D 19B developer. iv. Transfer the developed film to acid fixing bath at 18°C. v. Wash the film in distilled water for 15 mins giving two changes. vi. After washing, free the film from the clamp and mount it emulsion side up on a slide with the help of brush. vii. After emulsion is dried, mount it with Canada balsam. viii. Carefully remove the nylon film from the tissue and stain it. The tissue may also be used for making another radiograph.

19.2 Radiography of Botanical Material by Means of Low-energy X-rays (O’Brien and McCully, 1981) Relatively low-energy X-rays (soft X-rays in the range of 1-30 KV) are employed here to study the plant structure. Fresh material or herbarium specimens are radiographed with this simple technique. Procedure i. Cover the X-ray film or other suitable photographic emulsion with a thin layer of black plastic to protect it from light. ii. Put the plant specimen directly upon the black plastic. iii. Now expose to X-rays; the optimum combination of energy and time of exposure varies, and it should be standardised on a trial-and-error basis. iv. Develop the film photographically and it can be examined either as a negative or as a positive after contact printing. The method has got potential importance in studying the leaf venation and distribution of calcium oxalate crystals in leaf, and useful in taxonomic studies (O’Brien and McCully, 1981).

20 Cryopreservation Development for Germplasm Storage Plant Cryopreservation (Cryo is derived from the Greek word; ‘Cryos’ means cold or freezing and preservation mean storage) is the preservation of plant species in the form of plants outside natural habitats. (ex situ conservation), including DNA, seeds, pollen, buds, dormant buds, embryonic axes, zygotic, and somatic embryos at ultra-low temperatures (−130°C) but generally prefer to store the species using liquid nitrogen at −196°C, which is the temperature level that arrests biological activities, including all biochemical reactions of cells and no genetic changes during storage (Engelmann, 2004). Therefore, it is a reliable method for long-term storage of plant materials, although in some cases, such as storing seeds at −20°C, is more appropriate and less expensive than storage in liquid nitrogen and buds can be stored at −4 to −80°C, but storage cannot last long. Therefore, cold storage is another option for longterm storage. But stored living materials should be checked periodically.

20.1 Principles of Cryopreservation Plants that grow from warm to cold can withstand cold weather down to below 0°C. When the cold season passed, spring replaced it. Plants will begin to sprout new branches and leaves. Plants can withstand extreme cold weather because outside the cells contain more water than inside the cells, then ice crystals form outside the cells first. The water will move sufficiently out of the cells due to lower water potential than the outside until there is only a little water inside the cells which do not form ice crystals inside the cells when exposed to very low temperatures at −30°C. Prof. Dr. Akira Sakai (Sakai, 1960) conducted an experiment about 60 years ago using small branches of temperate trees, such as the willow and mulberry by reducing the temperature of those branches to −30°C before being immersed in liquid nitrogen or liquid helium for 1 year. After removing those branches from liquid nitrogen, they were still alive. This experiment is the first experiment by experimenting with pieces of plants in the cold regions that cause water loss before being immersed in liquid nitrogen. Later, there were many experiments and various methods that were developed to be

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carried out easily, conveniently, quickly, and high efficiently by increasing survival and low cost. However, the same principles are still used by reducing the amount of water inside the cells adequately by various methods, after that take plant materials to store at very low temperatures. The cells will be vitrified instead of ice crystals which have sharp edges to damage the cells and cell membranes. In terms of structure and properties, choosing a semi-permeable membrane will cause the cells to finally die. Success in preserving plant varieties in super cold conditions based on the cooling rate and the concentration of chemicals (cryoprotectants) preventing ultra-low temperature in order not to make the solution inside the cells to be ice crystals, but to be glass transition (Panis et al., 2005). Plant materials to be preserved must be in the suitable growth stage, type of cell groups, and the size that can be dehydrated the cells sufficiently and efficiently. Table 20.1 shows the plant materials used for preserving plant species in super cool conditions and the advantages and disadvantages of preservation. Table 20.1. Plant materials used for preserving varieties in super cool conditions Plant materials

Groups

Advantages

Disadvantages

Seeds

Small seeds, seeds that are resistant to dry and extreme conditions, cannot be asexual propagated.

Easy to preserve and are orthodox seeds (tolerate to dry conditions).

Cannot be used with large seeds that do not tolerate cold and dry conditions (recalcitrant seeds) and asexual propagation.

Pollen

Many groups

Easy for different genera and helpful in plant breeding.

Can store only half of the total number of chromosomes

Dormant buds

Trees that grow in the cold weather.

The buds are rest and ready to be kept in the collection. Requires less time and labour. No need for tissue culture laboratories.

Resistance to cold, depends on seasons and genotypes. Requires more storage space than other methods. Must have expertise in grafting or budding after storage.

Cryopreservation Development for Germplasm Storage 405 In vitro shoot tips

Many groups

Can collect the tip all year round. Easy to handle both anatomy and physiology.

No techniques have been developed for all plants. Must have laboratories and skilled workers.

Embryonic cultures

Many groups

In general. callus is easy to preserve.

Some plants do not produce somatic embryos. Techniques are not yet widely used for preserving plant species.

Embryonic axes

Plants which seeds are not resistant to dehydration and have large seeds (recalcitrant seeds).

Easy to remove and proceed.

Must do tissue culture to get the plantlets after storage.

Zygotic embryos

Plants which seeds are not resistant to dehydration and have large seeds (recalcitrant seeds).

Removing seed coats may increase survival,

Time consuming and quite difficult techniques.

Preservation of plant materials in cold conditions need to understand the basic mechanism of this principle which are: 1. Slow cooling (slow freezing) and 2. Vitrification.

Slow cooling This principle is used in the initial stages of preserving plant varieties in super cool conditions. Plants can endure the cold weather in the fall and very cold, lower than the freezing point in the winter, then sprouting new branches and leaves in the spring. When the weather starts to be cold, the water that is in the solution outside the cells which has more water than inside the cells will begin to form ice crystals. The remaining solution that does not form ice crystals will have a higher concentration. Therefore, to maintain the balance of water inside and outside the cells, water from inside the cells will move out from the protoplasm according to the water potential from higher to the outside which has lower water potential. Due to the presence of water outside the cells more than inside the cells, the cells will be dehydrated and wither. The concentration of the solution inside the cells will gradually increase and at the same time the temperature will slowly decrease. Until the end, the volume of cells that are not frozen will have very little water, and outside the membrane,

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the ice crystals will reach a temperature of −40°C, which is enough to store in liquid nitrogen (−196°C). The solution in the cells will have very high viscosity that does not form ice crystals but form glass-liked (vitrification) instead (Fig. 20.1). Therefore, the preservation of plant varieties in an extremely cold state used the principle of slow cooling. It is the method used in the beginning to keep the dormant buds of trees in the cold region by collecting plant materials from the trees in the winter through cold weather using slow cooling method, cultured at very low temperatures, such as 0°C for a while or immersed in cryoprotectants, such as glycerol and sucrose at low concentration for 1020 minutes, then use programmable freezing controller (Fig. 20.2) which is expensive, complicated, difficult to use, high cost, and takes many hours to work to lower the temperature at a rate of 0.5-2°C per minute, depending on crop type and growth stage until −40°C. Then, plant materials will be stored in liquid nitrogen. This method was applied to non-organised tissues, such as cell suspension and callus. After removing plant materials from liquid nitrogen, the temperature must be slowly increased the same as changing from the winter to the spring. Since the solution inside the cells still has low water content, if the temperature rises quickly, it may affect the metabolism of cells.

Fig. 20.1. The water inside the cell drops to a point where the solution inside the cell is very concentrated when exposed to extremely low temperatures, such as below 0°C, the water inside the cells changes from liquid to glass, which is called non-crystalline or amorphous phase or vitrification, the water molecules will form ice crystals or solid state. As a result, the amount of water in the solution outside the cell decreases, so the water potential outside the cell is lower than inside the cells. Water then moves from the inside of the cells to the outside of the cell enough to not cause the formation of ice crystals inside the cells. The rate of temperature reduction must be appropriate in order not to injure various organs inside the cell and the cell membrane. The cell is still alive.

Cryopreservation Development for Germplasm Storage 407

Fig. 20.2. Programmable freezing controller.

Vitrification The glass transition is a process of physical changes from liquid to noncrystalline, amorphous, and a glass state (Fahy et al., 1984). Cells can be preserved in non-ice crystals which do not damage cell membrane (Benson, 2008) (Fig. 20.3). Changing the state of the cell to a glass state requires two changes: (1) The rate of temperature reduction is very fast and (2) The solution within the cell is highly concentrated (Sakai, 2000). Normally, when plant materials in a cryotube are put into liquid nitrogen, the temperature drops quickly (about 60°C/sec). Later, there is a development to reduce the temperature faster by dripping the anti-coolant solution containing the plant materials onto the aluminium foil, called the droplet method, which can reduce the temperature to around 4,000°C per minute (Panis et al., 2005; Schafer-Menuhr et al., 1996) and cryo-plate, which will reduce approximately 5,000°C per minute (Yamamoto et al., 2011). That means lowering the temperature at a fast rate and warming it at a fast rate after removing liquid nitrogen will increase the survival of cells. The Vitrification method was developed by Sakai et al. (1990) and applied to various plants in the tropics, sub-tropic, and temperate. The method is based on the creation of glass by immersing the plants in a highly concentrated anti-cold chemical, such as PVS2 solution at temperatures of 25°C and 0°C (Matsumoto et al., 1994) to extract water from the cells without damaging the tissue. The moisture content of cells must be about 20-30% (Matsumoto et at., 1995) before being stored in liquid nitrogen and form the condition of glass inside and outside the cells (Figs. 20.3, 20.4, and 20.5). Vitrification method is an easy, convenient, less time consuming, and low cost, requires no expensive and complicated equipment (a programable freezing controller). After that, plant materials are removed from liquid nitrogen, then do rapid warming to prevent the transition from a glass state to ice crystals but skip a liquid state. Therefore, it will not damage the cell condition and cryopreserved cells are still alive.

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Fig. 20.3. Characteristics and structure of water molecules in a liquid state (A, D), an ice crystal (B, E), and a glass state (C, F). Cell Membranes are destroyed when the water inside the cells is in an ice crystal state (E).(Image courtesy of Dr. Bart Panis).

Fig. 20.4. Vitrification method using concentrated chemicals to prevent cold conditions. The most widely used solution is PVS2 (Plant Vitrification Solution Formula 2), developed by Prof. Dr. Akira Sakai contains 30% (w/v) glycerol,15% (w/v) ethylene glycol, 15% (w/v) dimethyl sulfoxide (DMSO), and 0.4 M sucrose. Glycerol and DMSO will move into the cells. Ethylene glycol will not move into the cells but will make the water which has higher water potential from inside the cells moves out of the cells. Therefore, preserved plant materials in liquid nitrogen are in a glass state, both inside and outside the cells.

Cryopreservation Development for Germplasm Storage 409

Before storage, cells grow normally.

Intracellular ice crystals after storage in liquid nitrogen without exposure to PVS2 solution, cells die.

Cells after exposure to PVS2 solution and be stored in liquid nitrogen, cells survive. Fig. 20.5. Cryopreservation of jackfruit embryonic axes using the vitrification method (Thammasiri, 1999).

Figure 20.6 describes the concentration of solution inside the cells. As the solution rises, the temperature of glass transition will decrease and the area where ice crystals form will be reduced. Therefore, increasing the concentration of solution inside the cells will result in reducing the occurrence of ice crystals inside the cells, leading to higher survival. However, the cells must be resistant to dry conditions at 20-30% moisture content inside the cells. Plant tissue containing a lot of water are not tolerant to dry conditions, except pollen, seeds, and somatic embryos of plants that are tolerant to dry conditions (orthodox species). Further, Fig. 20.6 clearly explains the slow cooling mechanism that allows plants to be preserved in a super cold condition and still survive. From the autumn to the winter, water begins to form ice crystals when the temperature is below 0°C in external cells first because there is more water content than inside the cells, then the water from inside the cells will be more and gradually move out from the cells causing the water inside the cells to decrease. The solution inside the cells gets more concentrated. The area of the ice crystals will

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Fig. 20.6. The relationship between temperature and concentration of solution to liquid, solid (ice) and glass phases (Tm = Melting point temperature; Th = Homogeneous nucleation temperature; Tg = Glass transition temperature) (Image courtesy of Dr. Bart Panis).

gradually decrease as the concentration of the solution increases. Therefore, the chance of the formation of ice crystals within the cells will be less. It takes some time from liquid to ice crystal state; therefore, if the temperature keeps getting lower, and the concentration of solution increases steadily to the interception point of Th and Tg, the glass state will be formed. The formation of ice crystals will occur very little or not at all. The important thing is to naturally reduce the temperature slowly. If decreasing too quickly, the concentration of solution inside the cells remains low because water does not move out of the cells quickly, resulting in ice crystal formation. Since ice crystals form from −40°C or reduce the temperature rapidly will cause ice

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crystals at low temperature. In summary, the plants must be cooled slowly so that the amount of water inside the cells is very low to about −40°C and then can be stored in liquid nitrogen and the cells are still alive. On the other hand, preservation of plant materials by vitrification is the opposite of slow cooling by exposure to highly concentrated solution, such as PVS2 solution for suitable time and reducing the temperature rapidly to have about 20-30% water content inside the cells. In order not to be toxic to cells and the reduction of exposure time, some chemicals with a low molecular weight, commonly use DMSO and glycerol will rapidly increase the concentration of solution inside the cells and help to eliminate free radicals and stabilise cell membranes. Another part of the chemicals due to their high molecular weight will be outside cell membrane, such as sucrose which helps to move water out of the cells and to stabilise cell membrane condition. Therefore, freezing of anti-coolant substances, such as PVS2 solutions, causes the solution inside the cells to rapidly increase concentration by the chemicals moving in and the water moving out according to the water potential. The amount of water in the cells should be 20-30% to be stored in liquid nitrogen. Therefore, when the concentration of solution increases rapidly, the temperature reduction must also be fast. Due to the high concentration of solution, the formation of ice crystals is even lower and rapid temperature reduction will reduce the time of exposure to plants, thus decreasing the toxicity. For example, the Droplet method reduces the temperature at around 4,000°C per minute and Cryo-plate method decreases the temperature at around 4,500°C per minute. From liquid to ice crystal state, it takes time for the formation of ice crystals. Therefore, quickly lowering the temperature would be able to overcome the ice crystal state to be glass state and increasing survival. Evidence of glass state must be studied from the physical process. The general method used is measuring the latent heat released. In summary, slow cooling uses pre-treatment chemicals to withstand low temperature conditions that have low concentration and slowly reduce the temperature, while vitrification uses highly concentrated solution and rapid cooling rate. The two methods have the same goal to prevent ice crystal formation within the cells to get high survivals after being taken out of the storage in a super cold condition.

20.2 Steps in Plant Cryopreservation There are 7 main steps in plant cryopreservation. Some plants may not require every step (skipping steps 1, 2, 5, or 6) in preservation depending on plant species and cryopreservation methods. The whole protocol is as follows: i. Preculture The cells will be adjusted to low temperature using 0.3 M sucrose (1-3 days) for (a) adding sucrose and, (b) adding ABA (abscisic acid) to stabilise cell membrane under dry condition.

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Fig. 20.7. The cells of wasabi plants immersed in preculture chemicals, a solution of 0.4 M sucrose for 20 minutes. There is a noticeable loss of water in some cells (Image courtesy of Prof. Dr. Akira Sakai).

ii. Loading solution Using chemicals to protect osmotic pressure, such as 2 M glycerol + 0.4 M sucrose (20 minutes) for dehydrating the cells to reduce osmotic pressure and help to withstand highly concentrated solution, such as PVS2 solution and adjust to dry and vitrification conditions.

Fig. 20.8. Immersion of the cells of Wasabi into a highly concentrated PVS2 solution. The water inside the cells is reduced sufficiently to prevent ice crystal formation inside the cells when exposed to low temperatures. Instead, the glass condition will be formed and not harm the cells when stored in liquid nitrogen. In this figure, we can see that the size of the cytoplasm. dramatically decreases when compared with immersion in preculture solution (Image courtesy of Prof. Dr. Akira Sakai).

iii. Dehydration Nowadays, scientists combine dehydration chemicals at a suitable rate for each plant. These chemicals are called plant vitrification

Cryopreservation Development for Germplasm Storage 413

solutions (PVS) which have many formulas (Table 20.2). The formula used in the present 4 formula is PVS1 (Usagami et al., 1989), PVS2 (Sakai et al., 1990), PVS3 (Nishi gawa et al., 1993), and PVS4 (Sakai et al., 2000). Highly concentrated solution, such as PVS2 solution at 25oC or 0oC (for suitable time) is used for dehydrating the cells sufficiently. The chemical composition in each formula is shown in Table 20.2. Table 20.2. Chemical composition of PVS solution of 4 formulae Chemical (g / l)

PVS1

PVS2

PVS3

PVS4

Glycerol

220.0

300.0

500.0

350.0

Ethylene Glycol

150.0

150.0

-

200.0

Propylene Glycol

150.0

-

-

-

Dimethyl sulfoxides

70.0

150.0

-

-

Sucrose

-

136.9

500.0

205.0

Sorbitol

91.1

-

-

-

These chemicals have different functions during the cold and warm process. Sugar helps stabilise cell membrane and proteins. DMSO will capture free radicals, increase the absorption of membrane cells and protect the cell structure during the cold condition. Glycerol helps stabilise cell membrane. But the work of these chemicals is not yet fully understood: iv. Storage in liquid nitrogen All the cryopreserved cells will be vitrified from −115oC to −196oC (in liquid nitrogen). v. Rapid warming Warm the cryopreserved cells rapidly at 35-40oC in water bath to change from glass state to liquid state. vi. Unloading solution Move the cells to concentrated 1.2 M sucrose for 20 minutes to reduce osmotic pressure and not let water move into the cells too fast. This step is for acclimation before culture. vii. Culture on suitable medium Culture the cells on suitable medium for plantlet development. Then, transfer the plantlets to the greenhouse, applying acclimatisation for 2 weeks.

20.3 Methods in Plant Cryopreservation Preservation of plant materials in cold conditions need to understand the basic mechanism of this principle which can be done in two ways: 1. Slow cooling or slow freezing (cell dehydration due to cold temperature) and 2. Vitrification (glass formation). In slow cooling, dormant bud and slow freezing methods are concerned. In vitrification, cells are dehydrated by exposure to highly concentrated chemicals or air drying before freezing, followed by rapid warming to avoid intracellular ice formation. Six different vitrification-based methods are: 1. vitrification, 2. encapsulation-dehydration, 3. encapsulation-

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vitrification, 4. droplet-vitrification, 5. vitrification cryo-plate method (V cryo-plate), and 6. dehydration cryo-plate (D cryo-plate). Up till now, there are eight methods chronologically as follows:

20.3.1 Dormant Bud Method This is the first plant cryopreservation method that dormant buds were collected during winter when buds were dehydrated at −10 to −30oC before storage in liquid nitrogen (−196oC). This method was successful in willows (Salix koriyanagi) and poplar (Populus sieboldii) (Sakai, 1960). Later, dormant bud method was studied in apple (Sakai and Nishiyama, 19781; Forsline et al., 1998; Towill and Bonnart, 2003), cherry (Towill and Forsline, 1999), blueberry (Jenderek and Reed, 2017), etc.

Dormant Buds Fig. 20.9. Diagram showing Dormant Bud method.

The protocol for preserving dormant buds is shown in Fig. 20.9. This method works well with cold perennials, such as apples, willows, mulberry, conifers, etc., and is another option for plant cryopreservation.

20.3.2 Slow Freezing Method Slow freezing was the standard method in the early time (Panis and Lambardi, 2005), but need a programmable freezing controller to reduce the temperature at the rate of 0.5-2°C/min depending on the plant species and the growth stage until about −40°C, then store in liquid nitrogen. The protocol takes many hours to complete and requires expensive tools. This method was popular during 1980-1990 to store undeveloped plant tissues, such as suspended cells and calli of various plants.

Cryopreservation Development for Germplasm Storage 415

Slow Freezing Fig. 20.10. Diagram showing the Slow Freezing method.

The protocol for Slow freezing method is shown in Fig. 20.10. Slow freezing method is used to preserve plant species in extremely cold conditions compared to other methods. Due to complications, it takes a long time and must use a programmable freezing controller but may work better in some plants in China at the National Gene bank in Beijing of the 82 plants collected, only 3 mulberry plants were kept dormant by slow freezing method and had a survival rate of 90% (Zhang et al., 2014).

20.3.3 Vitrification Method Vitrification is a method developed to reduce the temperature and to warm rapidly, all parts of cells and tissues in a glass state. Vitrification method involves treatment of explants with plant vitrification solution (PVS), such as PVS1, PVS2, PVS3, and PVS4 to induce dehydration of explants during cooling and warming to avoid intracellular ice-crystal formation (Uragami et al., 1989; Sakai et al., 1990). The key to successful cryopreservation by vitrification method is to prevent injury by optimising exposure time to PVS for dehydration (Niino, 2007) because over exposure time to PVS may result in cell injury and intracellular ice formation during cooling. The optimum exposure time to PVS depends on explant size and species specific. The suitable dehydration duration was related to the sample size, the composition, and loading solution (Chen and Wang, 2002). Sakai et al. (1990) succeeded to cryopreserve nucellar cells of naval orange using PVS2 solution, after that many plants were experimented with success.

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Vitrification Fig. 20.11. Diagram showing Vitrification method.

The protocol for the Vitrification method is shown in Fig. 20.11. Vitrification is a method developed with short processing time, low cost, and less skilled work. It gives high survival, but it needs accurate time because of a short protocol time. In addition, the use of chemicals to dehydrate cells may be toxic to cryopreserved plant materials.

20.3.4 Encapsulation-dehydration Method Encapsulation-dehydration method is developed from artificial seed production that explants are encapsulated in alginate beads, precultured with high sucrose, desiccated by air-drying in a laminar air-flow cabinet or with silica gel, and then plunged into liquid nitrogen (Sakai et al., 2000; Matsumoto and Sakai, 1995). The advantages of this method are easy for manipulation of encapsulated explants (Hirai, 1998), preventing direct contact of toxic chemicals with plant materials, and non-toxic cryoprotectants are applied to protect during dehydration (Niino and Sakai, 1992). However, this method is the longer dehydration procedure than vitrification method (Thammasiri, 2000). Fabre and Dereuddre (1990) succeeded to cryopreserve tomato apical meristems by using this method. There are many plant species to be cryopreserved by encapsulation-dehydration. Jitsopakul et al. (2008a) successfully cryopreserved protocorms of Vanda coerulea by encapsulation-dehydration in combination with a loading solution (Fig. 20.12). Protocorms were selected 70 days after sowing seeds, harvested from 7-month-old fruits. After encapsulation in an alginate matrix composed of 2% Na-alginate, 2 M glycerol plus 0.4 M sucrose (loading solution), the protocorms were precultured in modified VW liquid medium (Vacin and Went, 1949) supplemented with 0.7 M sucrose on a shaker (100 rpm) at 25±3oC for 20 h. Encapsulated protocorms were then dehydrated

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in a sterile air-flow in a laminar air-flow cabinet at 25±3oC for 1-10 h, and then directly plunged into liquid nitrogen for 1 d. After thawing at 49oC for 2 min, cryopreserved beads were cultured on modified VW agar medium for regrowth. The highest regrowth of 40% was observed with cryopreserved bead with 35% water content for 8 h dehydration. No morphological variation was detected between non-cryopreserved and cryopreserved plantlets, and ploidy level was unchanged because of cryopreservation.

Fig. 20.12. Regrowth of Vanda coerulea protocorms after cryopreservation by encapsulation-dehydration in combination with loading solution. A: Precultured beads after sterile air-flow dehydration for 10 h. B: Cryopreserved protocorms after 20 days of regrowth. C: 3 months of culture on modified VW agar medium showing shoot growth. D: Plantlets after 8 months of culture on modified VW agar medium (left: non-cryopreserved, right: cryopreserved plantlet). E: Plantlets derived from cryopreserved protocorms after 5 months and F: 15 months of culture in the greenhouse. Bar for A-C = 1 mm, for D = 0.5 cm, and for E and F = 1 cm. Source: Jitsopakul et al., 2008a (with permission from Cryo Letters Journal).

The protocol for the Encapsulation-dehydration method is shown in Fig. 20.13. No highly concentrated chemicals used for dehydration, but air blow is used instead; therefore, it is not toxic to plant materials and environment. Encapsulation helps to reduce the damage from moving plant materials and air blow, as well as taking longer time (20-30 minutes more) for operation which help flexible operation. In addition, there is no unloading solution after rapid warming since the water from outside the cells will not move in fast due to the presence of calcium alginate encapsulation and in a dry condition including without highly concentrated chemical solution. The disadvantages of this method are long operation of 2-3 days, medium or low survival depending on adjusted protocols, and high sucrose concentration after dehydration which some plant species cannot tolerate.

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Encapsulation-Dehydration Fig. 20.13. Diagram showing Encapsulation-Dehydration method.

20.3.5 Encapsulation-vitrification Method Encapsulation-vitrification method is a combination of encapsulationdehydration method and vitrification method. The explants are encapsulated in alginate bead, and then subjected to dehydration by highly concentrated plant vitrification solutions, such as PVS1, PVS2, PVS3, and PVS4. This method gives higher survival than encapsulation-dehydration (Hirai and Sakai, 1992; Sakai et al., 2008; Matsumoto, 2017).

Encapsulation-Vitrification Fig. 20.14. Diagram showing Encapsulation-vitrification method.

The protocol for Encapsulation-vitrification method is shown in Fig. 20.14. Encapsulation-vitrification method is the method that is applied from Encapsulation-dehydration. Survival is higher than the Encapsulation-

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dehydration method and close to or slightly lower compared with the vitrification method.

20.3.6 Droplet-vitrification Method This recent method is the fast freezing from small drops of plant vitrification solution (PVS) on aluminium foil. Droplet-vitrification method is a combination of droplet-freezing and solution-based vitrification, then placed on aluminium foil strip in droplet of vitrification solution and then frozen by rapidly immersion in liquid nitrogen (Sakai and Englemann, 2007). Rapid warming was done by dipping the aluminium foil strips in unloading solution without using a water bath (Kim et al., 2006). During the cooling and warming procedures, rapid heat transfer is needed to avoid freezing injury (Agrawal et al., 2004; Kim et al., 2006; Yoon et al., 2006). Aluminium foil has an efficient thermal conductivity, resulting in quick and uniform heat distribution among tissue Halmagyi et al., 2005; Kim et al., 2006; Yoon et al., 2006). A high warming rate was employed to avoid recrystallisation of intracellular ice or addition cell dehydration by extracellular ice (Kim et al., 2006). The first success of droplet-vitrification method was studied in potatoes (Pennycooke and Towill, 2000). Later, successes in papaya (Ashmore et al., 2001), prunes (De Boucaud et al., 2002), jam (Leunufna and Keller, 2003), chrysanthemum (Halmagyi et al., 2004), banana (Panis et al., 2005), rose (Halmagyi and Pinker, 2006), Bletilla striata (Jitsopakul et al., 2008b), Grammatophyllumspe ciosum(Sopalun et al., 2010), Vanda coerulea (Jitsopakul et al., 2008a), Sugar cane (Barraco et al., 2011), Vanilla orchid (Gonzalez-Arnao et al., 2009; Hernandez-Ramirez et al., 2014), Oil Palm (Gantaitet al., 2015), Grapes (Pathirana et al., 2015), Orange (Volk et al., 2012), etc. Jitsopakul et al. (2008b) studied droplet-vitrification method for cryopreservation of Bletilla striata mature seeds (0 day after sowing), zygotic embryos (3 days after sowing) and protocorms (6, 9, and 12 days after sowing). Mature seeds were surface-sterilised and sown on solidified New Dogashima (ND) medium supplemented with 3% sucrose and cultured under illumination provided at an intensity of 62.0 µmol. m-2.s-1 for 16 h/d at 25°C for preparation of zygotic embryos and protocorms. Mature seeds, zygotic embryos, and 6-day-old protocorms were precultured in liquid ND medium supplemented with 0.3 M sucrose for 3 h on a shaker (110 rpm) and then dehydrated with 2 M glycerol and 0.4 M sucrose in ND liquid medium (loading solution) for 15 min, followed by exposure to PVS2 solution for 60 min at 25°C. Then, plant materials were soaked in liquid nitrogen by droplet-vitrification method, and then were cultured on solidified ND medium supplemented with 3% sucrose, germination rates of cryopreserved mature seeds, cryopreserved zygotic embryo, and survival rate of cryopreserved 6-day-old protocorms were 93%, 91%, and 84%, respectively. Cryopreserved 9-day-old protocorms gave the highest survival rate of 66% when precultured with 0.5 M sucrose for 3 h on a shaker, dehydrated with loading solution

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Fig. 20.15. Mature seeds of Bletilla striata. A: Development of mature seeds sown on solidified ND medium supplemented with 3% sucrose under illumination provided at an intensity of 62 µM m-2.s-1 for 16 h/d at 25°C for 3 days, B: 3 days, C: 6 days, D: 9 days, E: 12 days, and F: Twenty droplets of PVS2 solution (2 µl) with protocorms, placed on sterilised aluminium foil strip (7 × 20 mm2). Bar: A-E = 0.5 mm, F = 1 cm. Source: Jitsopakul et al., 2008b (with permission from Cryo Letters Journal).

for 15 min, followed by exposure to PVS2 solution for 40 min at 25°C and cultured on solidified ND medium supplemented with 480 mg/l ammonium nitrate and 3% sucrose. No survival was observed in cryopreserved 12-oldday protocorms. Fig. 20.15 A-F showed the development of mature seeds. Mature seeds developed into zygotic embryos at 3 days after sowing (Fig. 20.15B). Protocorms formed green spots on zygotic embryos at 6 days of sowing (Fig. 20.15C). Protocorms formed apical meristems at 9 days of sowing (Fig. 20.15D) and formed primary leaves at 12 days of sowing (Fig. 20.15E).

Droplet-Vitrification Fig. 20.16. Diagram showing Droplet-vitrification method.

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The protocol for Droplet-vitrification method is shown in Fig. 20.16. Droplet-vitrification method is a relatively new method which is improved from Vitrification. The explants are dehydrated by using just one drop of highly concentrated plant vitrification solution and place on an aluminium foil which contributes to heat. It is convenient to move 10-12 explants on the aluminium foil at the same time. High survival is obtained from rapid cooling and warming from aluminium foil and rapid warming is skipped from this protocol.

20.3.7 V Cryo-plate Method A new method for preserving plant species in cold conditions was developed 11 years ago by Yamamoto et al. (2011) is the V cryo-plate method. It is the combination of encapsulation-vitrification and droplet-vitrification methods to make artificial seeds to attach well (high quality) on aluminium sheets (cryo-plate) size 7 mm × 37 mm × 0.5 mm, which has 10-12 holes (hole size 1.5 mm × 0.75 mm) in which the aluminium sheet is thicker than the aluminium foil (aluminium foil) used with the droplet-vitrification method. The thermal conductivity of the aluminium sheet is better than the aluminium foil which has a cooling rate of approximately 4,000°C/min and an increase of temperature of around 3,000°C/min; while aluminium sheets have a temperature reduction of around 5,000°C/min and an increase of temperature around 4,500 °C/min. Therefore, V cryo-plate method gives higher survival than droplet-vitrification method because the cooling and warming rates are faster. The advantages of the V cryo-plate method are: 1. The cooling and warming rates are very fast. Therefore, providing high survival. 2. The protocol is easy and convenient because the plant materials are attached to the aluminium sheet throughout the operation, not in a suspended state in a super cool solution, such as PVS2 and PVS3. Using this method does not need special skills. Jitsopakul et al. (2021) studied orchid pollinia cryopreservation using the V cryo-plate and apply it to some orchid species for breeding. Pollinia of Rhynchostylis gigantea (L.) Ridl. were collected from completely open flowers in the morning and then were placed on aluminium cryo-plates embedded in alginate gel, then immersed in loading solution containing 2 M glycerol and 0.4 M sucrose for 15 min at room temperature (29°C), and then dehydrated with PVS2 solution for 0-60 min at 29°C. The cryo-plates with pollinia were directly plunged into liquid nitrogen for 40 min, and rapidly warmed in 1 M sucrose for 15 min. The cryopreserved and non-cryopreserved pollinia were used for hand-pollinating flowers of the same species for producing hybrids. The results showed that the viability of non-cryopreserved and cryopreserved pollinia dehydrated with PVS2 was 100%. The highest fruit set after pollinating flowers with cryopreserved pollinia dehydrated with PVS2 for 40 min was also up to 100%. The protocol for cryopreservation of

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Fig. 20.17. Cryopreservation of R. gigantea pollinia using V cryo-plate method for hand-pollination with R. gigantea var. harrisoniana flowers. (a) R. gigantea flowers and (b) pollinia. (c) Pollinia were placed on aluminium cryo-plates embedded in 3% sodium alginate solution, then soaked in 100 mM CaCl2 and PVS2, respectively. (d) Cryo-plates with pollinia were transferred to uncapped 2 mL cryotubes held on cryo-canes and then directly plunged into liquid nitrogen. (e) Non-cryopreserved and cryopreserved pollinia staining red were considered viable, while the unstained pollinia were considered non-viable. (f) R. gigantea var. harrisoniana flowers. (g-h) Capsule set after hand-pollination of cryopreserved pollinia of R. gigantea with R.gigantea var. harrisoniana flowers. (i) Capsule and seeds of R. gigantea var. harrisoniana x R.gigantea. (j-k) Seeds of R. gigantea var. harrisoniana x R. gigantea germinated and developed into protocorms and then plantlets on modified VW agar medium. (l) Hybrid plantlets grown in the greenhouse at Rajamangala University of Technology Isan, Surin Campus, Surin province, Thailand for one year. Source: Jitsopakul et al., 2021 (with permission from Cryo Letters Journal).

R. gigantea (L.) Ridl. pollinia using V cryo-plate method was successfully applied for the cryopreservation of nine Thai orchid species (Fig. 20.17). The exposure time to PVS2 affected the pollinia viability (range 40-100%; average 93%) and capsule set (range 20-100%; average 78%) of the nine species. The

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successful capsule set and seed production after pollination with cryopreserved pollinia in all orchid hybrids were observed. Seed germinated into protocorms and developed to plantlets cultured on modified Vacin and Went (1949) agar medium. Cryopreservation of freshly collected orchid pollinia using V cryoplate method is an efficient tool for the long-term storage of plant germplasm and for orchid breeding.

V cryo-plate Fig. 20.18. Diagram of V cryo-plate method.

The protocol for V cryo-plate method is shown in Fig. 20.18. The V cryo-plate method is developed from a combination of Encapsulationvitrification and Droplet-vitrification methods. The advantages of the V cryoplate method are: 1. The cooling and warming rates are very fast. Therefore, providing high survival. 2. The protocol is easy and convenient because the plant materials are attached to the aluminium sheet throughout the operation, not in a suspended state in a super cool solution, such as PVS2 and PVS3. Using this method does not need special skills.

20.3.8 D Cryo-plate Method Niino et al. (2013) developed the D cryo-plate method, which is a combination of encapsulation-dehydration with the V cryo-plate method (Niino et al., 2014). As some plants may be affected using PVS2 solutions, there is low survival. Therefore, to avoid this damage, cells were dried by blowing air from a laminar air-flow cabinet. In addition, the cryopreserved plant materials can be larger than those using the V cryo-plate method because there is no problem of dehydration and the toxicity of the PVS2 solution after prolonged immersion. Both V cryo-plate and D cryo-plate methods have the same protocol except

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for the cell dehydration by air-drying, silica gel, or drying beads, instead of using concentrated chemical solutions, such as PVS2 solution. Later, Cordova II and Thammasiri (2016) developed D cryo-plate method using silica gel and drying beads for dehydrating cells. In this experiment, protocorms were placed in the preculture solution consisting of 0.7 M sucrose on a shaker (110 rpm) at 25+3°C for 1 d. After that, protocorms were placed one by one in the wells which filled before with the alginate solution containing 2% (w/v) sodium alginate in calcium-free ½MS basal medium with 0.4 M sucrose. The cryo-plates were hardened for 20 min by slowly dispensing the calcium chloride solution containing 0.1 M calcium chloride in ½MS basal medium with 0.4 M sucrose. Then the cryo-plates were surface dried using sterile filter paper, placed in Petri dishes containing silica gel or drying beads in a laminar air-flow cabinet (Fig. 20.19). Cryo-plates were dehydrated for 5 h until 25% moisture content was achieved. Dehydrated cryo-plates were placed in 2 ml cryotubes and plunged directly into liquid nitrogen for 1 d.

Fig. 20.19. Cryo-plate method dehydrated with silica gel or drying beads. A: Protocorm development, B: Preculture of protocorms in ½MS liquid medium with 0.7 M sucrose for 1 d, C: Pour the alginate solution containing 2% (w/v) sodium alginate in calcium-free ½MS basal medium with 0.4 M sucrose in the wells, D: Place the precultured protocorms in the wells one by one, E: Pour the calcium chloride solution containing 0.1 M calcium chloride in ½MS basal medium with 0.4 M sucrose, F: Dehydration with 50 g silica gel, G: Dehydration with 30 g drying beads, H: Put each cryo-plate in a 2 ml cryotube, I: Plunge 2 ml cryotubes into liquid nitrogen for 1 d, J: Warming in 1.2 M sucrose solution for 20 minutes, K: Plate on ½MS agar medium, L: Regrowth, and M: Regrowth after 60 days. Source: Cordora II and Thammasiri, 2016 (with permission from Cryo Letters Journal).

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Cryo-plates were removed from cryotubes and warmed in unloading solution (1.2 M sucrose solution) for 20 min. Protocorms were then removed from the cryo-plate and placed on ½MS agar medium for regrowth. Growth conditions were conducted using 16 h light at 25+3°C. For effect of the cryoplate method, regrowth of control treatments dehydrated using silica gel was observed to be 90%. Regrowth of control treatments dehydrated using drying beads was observed to be 92.1%. In all other treatments, regrowth was observed to be 73.8% using silica gel for dehydration. Regrowth for all other treatments dehydrated using drying beads was observed to be 76.5%. Regrowth was observed at the 2nd week of transfer to ½MS media. Dehydration using silica gel or drying beads did not significantly affect regrowth rate. Protocorms dehydrated using silica gel or drying beads developed into normal plantlets. The protocol for D cryo-plate method is shown in Fig. 20.20. The development of the V cryo-plate method, which is a combination of Encapsulation-vitrification and Droplet-vitrification method and followed by the D cryo-plate method which is a combination of Encapsulationdehydration with V cryo-plate method by using an aluminium sheet that conducts heat well as a vehicle to carry 10-12 plants at the same time throughout every step. Therefore, making the operation convenient, fast, efficient and with high survival. Since it has a cooling rate of 5,000°C/min and a warming rate of 4,500°C/min.

D cryo-plate Fig. 20.20. Diagram showing D cryo-plate method.

21 Microbiological Techniques Microorganisms live together in mixed populations in nature. It is possible to isolate a particular type of organism by using the enrichment culture technique, first described by Winogradsky (quoted in Tauro et al., 1993).This method enables the growth of a particular type of organism, which is achieved by employing suitable growth substances. For example, to isolate the sulphur oxidising bacteria from the soil, the medium is supplemented with sulphur, which only enables the growth of sulphur bacteria. Subsequently,the isolation of pure culture is possible by employing one of the purification techniques described below. Similarly, a medium incorporated with sodiumazide facilitates the growth of lactic bacteria whereas it inhibits others. The isolation of fungi from a mixed population of fungi and bacteria is made possible by incorporating the antibacterial substance in the medium. Thus, a variety of methods are available by which a selected group of organisms can be isolated very easily.

21.1 Isolation Methods 21.1.1 Serial Dilution Technique The serial dilution technique was originally developed by Lister (quoted from Tauro et al., 1993).In this technique, the mixed culture sample is serially diluted in a liquid medium so that the final concentration will contain one or none of the microorganisms. Growth in the last tube of the dilution series is presumed to be from a single cell. This method is useful for the isolation and purification of bacteria and yeasts.

21.1.2 Pour Plate Technique The pour plate technique involves plating an aliquot of appropriately diluted culture suspension onto molten agar medium at 40-45°C and pouring it into sterile petri dishes. Upon solidification, the plates are incubated. Pour plates exhibit both surface and subsurface growth. Though it is not an ideal method to isolate pure cultures of all types of organisms, it is suitable to isolate and purify the cultures of aerobic bacteria. These bacteria can be further purified by serial dilution technique or by the repeated streak plate technique.

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21.1.3 Streaking Technique In the streaking technique, the petri plate is divided into four sectors; a small aliquot of different dilutions is streaked on the surface of the solid medium with the help of an inoculation loop in these four sectors (Fig. 21.1). Upon incubation of the streak plate, pure colonies are developed, each arising from a single mother cell. Mother cell divides in exponential numbers, resulting in billions of daughter cells and forming a pure colony. Remember,a colony is a pure colony if it does not overlap any other colony (Beishir, 1996).

Fig. 21.1. Streak plate technique (Redrawn from Beishir, 1996).

21.1.4 Isolation of Anaerobic Bacteria Anaerobic bacteria are isolated either by pour plating technique using a medium containing a reducing agent like thio glycolate and incubating under anaerobic conditions or by roll-tube technique. In the roll-tube technique, an aliquot of appropriate dilution is added to the molten, reduced agar medium, and the tube is rolled in such a way that a thin layer of medium is formed on the inner walls of the tube. The tubes are sealed with butyl stoppers and anaerobic conditions are maintained by providing oxygen-free CO2 or nitrogen. Tubes are dried during a prolonged incubation period which is necessary because of the slow growth of obligately anaerobic bacteria.

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21.2 Media A medium may be synthetic (chemically well-defined) containing chemically known compounds in known quantities or it may be non-synthetic containing unknown organic substances such as yeast extract, potato extract, coconut milk, beef extract, tomato juice, etc. Generally, a medium should contain carbon sources (sucrose or glucose), inorganic salts, and in some cases vitamins, amino acids, and other growthpromoting substances for the satisfactory growth of microorganisms. The recipe for the medium depends on the microorganism to be grown and some of the commonly used media for the growth of microorganisms are mentioned in the appendix (see also Sullia and Shantharam, 1997). A liquid medium to which agar is not added is known as broth. For the growth of aerobic microorganisms in liquid culture, a proper supply of oxygen is necessary. For providing proper aeration, liquid culture flasks are continuously agitated on the rotary shaker, and it provides uniform growth of microorganisms. Cultures on a large scale are prepared in special devices called fermentors where sterile air is kept continuously bubbling through the medium. To obtain a solid medium, a solidifying agent like agar (obtained from algal weeds Gelidium, Gracilaria, etc.) should be added to the broth. Agar contains complex carbohydrates generally not utilised by microorganisms and has no nutritive value. Agar gel has some advantages over other gelling agents, they are: (i) it will not react with any other constituents of the medium, (ii) it will not be degraded by any other enzymes used in the medium, and (iii) it melts only at 100°C and upon solidification at 40°C forms solid agar medium which is hard and transparent in the petri dishes (when they are stored upside down). Sterilisation of media Media sterilisation can be achieved either by autoclaving or by filtration through membrane filters. Culture media in glass containers sealed with cotton plugs or aluminium foils should be autoclaved at 15 pounds per square inch (Psi) and 121oC for 15 mins. The pressure should not exceed 20 Psi as higher pressure may lead to the decomposition of carbohydrates and other constituents of the medium. Autoclave time depends on the volume of the liquid to be sterilised (Table 21.1). Table 21.1. Minimum time required for autoclaving nutrient media Volume (ml)

*Sterilisation time (min)

1-200

15

200-1000

20

1000-2000

40

*At 15 Psi and 121°C.

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Some thermolabile substances like vitamins, amino acids, hormones, natural plant, and animal extracts, etc., may decompose during auto claving. These substances require filter sterilisation. Seitz Filter: Seitz filters are asbestos discs about 1 mm thick. The asbestos disc is fitted into a metal container supported by a metal grid and screwed tightly. An ehrlenmeyer flask with a side arm is fitted to the apparatus. The whole setup is autoclaved for 5 mins at 15 Psi and 121°C. After cooling the apparatus, the liquid medium is filtered through a filter by applying suction. After filtration, the Seitz filters are discarded. The disadvantage of this method is that some compounds of the media may get absorbed into the filter disc. Sintered Glass Filters: These filters are made of finely ground glass fused sufficiently to leave small pores. These filters are useful for chemically defined media. Less adsorption of chemical compounds takes place in these filters than in the asbestos discs. Membrane Filters: These are very thin membranous filter discs made up of cellulose acetate (millipore filters) with a pore size of 0.43-0.47 µm. The membrane filter allows large volumes of liquid to pass through rapidly under suction. These filters do not cause the adsorption of chemical compounds and are used as disposable items. These are widely used among other bacterial filters. Filter sterilised solutions are then added to the autoclaved substances and a completely sterilised medium is prepared. This can be used for culturing microorganisms.

21.3 Maintenance and Preservation of Cultures It is necessary to maintain and preserve the pure cultures of microorganisms for further research. Some of the simple methods for the maintenance and preservation of microbial cultures are given below (see also Tauro et al.,1993).

21.3.1 Transfer to Fresh Media Microbial cultures are maintained continuously by periodic transfer onto a fresh medium in the culture tubes. The frequency of transfer may be varied with the organism, e.g., a culture of E. coli needs to be transferred at monthly intervals. To keep the cultures viable, it is necessary to use the appropriate growth medium and to maintain proper storage temperature. Many heterotrophic bacteria can be maintained on agar medium by transferring to fresh medium after every 20-30 days.

21.3.2 Lyophilisation In this method, cell suspensions are taken in the culture vials and these are frozen by immersing in a mixture of dry ice and acetone or liquid nitrogen.

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The culture vials are evacuated and dried under a vacuum, sealed, and stored at low temperatures. By this method, cultures are preserved for a longer period without any change in the organism. However, it has been reported that the loss of vacuum during storage leads to the inactivation of cultures.

21.3.3 Overlaying with Mineral Oil Many bacteria and fungi are well preserved by covering the fresh growth in agar slants with sterile mineral oil. The mineral oil is placed at the tip of the slanted surface. Care must be taken to cover the entire surface of the culture with the mineral oil. In this method, cell viability is very high when compared to frequent transfer and storage at low temperatures. These cultures are stored at 0-5°C. With this method, microorganisms may be preserved satisfactorily for more than 5-20 years.

21.3.4 Storage in Sterile Soil This method is widely employed for preserving spore-forming bacteria and fungi. Spore suspensions are added to sterile soil (sterilised at 15 Psi and 121°C for 2-3 hrs at intervals of 1-2 days) and the mixture is dried at room temperature and stored in the refrigerator. Bacterial cultures maintained by this method remain viable for 70-80 years.

21.3.5 Storage in Silica Gel The microorganisms such as bacteria and yeasts can be stored in silica gel at low temperatures for about 1-2 years. In this method, finely powdered, heat sterilised, and cooled silica powder is mixed with a thick suspension (paste) of cells and stored at low temperatures. The basic principle involved in this technique is quick desiccation at low temperatures which allows the cells to remain viable for longer periods.

21.4 Preparation and Staining of Specimens Generally, living microorganisms are directly examined under a light microscope. Often, they must be fixed and stained to increase visibility and to study specific morphological features. Sections are generally used to study the parasitic microorganisms in plant and animal tissues.

21.4.1 Fixation The primary aim of fixation is to terminate the life processes of the cell as well as preserve external and internal structures of the cell with minimum alteration when compared to the living state. In other words, the microorganism is killed and attached firmly to the micro slide.

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There are two different types of fixation: (1) Heat-fixation of bacterial smears by gently flame heating the air-dried film of bacteria. This adequately preserves the overall morphology of bacteria. (2) Chemical fixation preserves the cellular organelle intact and protects the overall morphology of bacteria. Chemical fixatives penetrate the cells and react with cellular components, usually proteins and lipids, to render them inactive, insoluble, and immobile. Common chemical fixatives are ethanol, mercuric chloride, acetic acid, formaldehyde, glutaraldehyde, etc. (see also Chapter 3).

21.4.2 Dyes and Simple Staining The dyes used to stain microorganisms have two common features: (1) they have chromophore groups with conjugated double bonds that give the dye its colour, and (2) they can bind with cells by ionic, covalent, or hydrophobic bonding, e.g., a positively charged dye binds to negatively charged molecules on the cell. Ionisable dyes may be divided into two general classes based on the nature of their charged group (see also Chapter 4).They are as follows: Basic dyes (Cationic dyes) - These dyes have positively charged groups and are generally sold as chloride salts. Basic dyes bind to negatively charged molecules in the specimen such as nucleic acids and many proteins. Since the surfaces of bacterial cells are negatively charged, basic dyes are very commonly used in bacteriology. Some of the cationic dyes are safranin, crystal violet, basic fuchsin, methylene blue, malachite green,etc. Acid dyes (anionic dyes) - These dyes possess negatively charged groups such as carboxyls (-COOH) and phenolic hydroxyls (-OH). Acid dyes bind to positively charged cell structures. Some of the acid dyes are fast green FCF, eosin, aniline blue, rose bengal, acid fuchsin, etc. The pH may alter the staining effectiveness since the nature and degree of the charge on cell components change with pH. So anionic dyes stain best under acidic conditions when proteins and other molecules carry positive charges; basic dyes are most effective at higher pHs. In most dyes, ionic interactions are probably the most common means of attachment. Dyes can bind through covalent bonds and their solubility characteristics decide the affinity with the specimen. For example, DNA can be stained by the Feulgen method in which Schiff’s reagent is covalently attached to its deoxyribose sugars after HCl treatment. Sudan III (Sudan Black) selectively stains lipids because it is lipid soluble, but it will not dissolve in aqueous portions of the cell. Simple stains are also referred to as monochrome stains because only one dye is employed for staining the microbes. Basic dyes like crystal violet, methylene blue, and carbolfuchsin are frequently used to determine the size, shape, and arrangement of bacteria.

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21.4.2.1 Simple Staining Method for Bacteria in Milk by using Breed’s Methylene Blue

Breed’s Methylene Blue Take 0.3 g methylene blue chloride and add 30 ml ethanol. To this mixture add 100 ml of 2% phenol in water and mix properly. This mixture is directly used for staining the bacteria. Procedure 1. Take the clear microslide on which place 0.01 ml of milk sample and spread it with a needle. Dry the smear with gentle heat. 2. Immerse the slide in xylene or chloroform to remove the fat. 3. Fix the smear with 95% alcohol for 2 mins. 4. Stain the smear with Breed’s methylene blue for 2 mins. 5. Wash the smear with 90% alcohol till the smear appears faintly blue, air dry, and examine under oil-immersion objective. Result: Bacteria appear dark blue against a light blue background. 21.4.2.2 Demonstration of Anthrax bacilli by using Polychrome Methylene Blue

Preparation of Polychrome Methylene Blue Solution Take 100 ml of Loeffler’s methylene blue solution in a brown bottle and allow it to ripen for more than 10 months. Shake the solution intermittently during this period. The above process is hastened by adding 1% K2CO3 to the solution. The slow oxidation of methylene blue forms a violet compound that gives polychrome properties to this stain. In addition to simple staining, polychrome methylene blue solution is also used for Mc Fadyean’s reaction in which anthrax bacilli appear blue against purple granular background (Desai and Desai, 1980).

21.4.3 Protocols 21.4.3.1 Differential Staining of Bacteria

The staining procedure which differentiates two groups of bacteria known as gram-negative and gram-positive is termed a differential staining technique. The basic principle involved in this technique is due to differences in chemical and physical properties of the bacterial cell wall. They react differentially with the staining reagents. There are two important differential stains viz., Gram stain and Acid-fast stain. Gram Stain The Gram stain developed by the Danish physician Christian Gram in 1884 is the most widely employed staining technique in bacterial staining. It has great

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taxonomic significance and has importance in the identification of an unknown prokaryotic organism. For eukaryotic cells, this stain is not significant since most of them are gram-negative except yeasts and a few moulds which are gram-positive. In general, gram-negative bacteria are more widely distributed than gram-positive and their morphological features are given in Table 21.2. Based on the Gram stain technique, bacteria are divided into two groups: (i) Gram-positive, those that retain primary dye such as crystal violet and appear purple in colour, (ii) Gram-negative, which lose the primary dye on destaining with alcohol and take the colour of counterstain like safranin or basic fuchsin and they appear red in colour. Preparation of Reagents (i) Gram’s crystal violet. It consists of two solutions: Solution A Crystal violet 2.0 g Ethanol, 95% 20.0 ml Dissolve the crystal violet in the ethanol: Solution B Ammonium oxalate 0.8 g Distilled water 80.0 ml Dissolve the ammonium oxalate in the distilled water. Now mix solutions A and B and shake well, properly mixed. (ii) Gram’s iodine Iodine Potassium iodide Distilled water

10 g 2.0 g 300.0 ml

Grind both iodine and potassium iodide together by using a mortar and pestle to which add water and mix thoroughly. Store the solution in a tightly closed bottle. (iii) Safranin

Safranin Ethanol, 95% Distilled water

0.25 g 10.0 ml 100.0 ml

Dissolve the safranin in the ethanol and mix thoroughly. Add the distilled water and mix well. Later filter the solution and use it directly. (iv) Acetone alcohol Ethanol, 95% Acetone Mix the above two thoroughly.

70.0 ml 30.0 ml

Microbiological Techniques 435

Procedure i. Prepare and fix the bacterial smear on the clean micro slide. ii. Stain the smear with a basic dye, crystal violet for 30 secs. iii. Wash the slide carefully with distilled water from a wash bottle. (Do not squirt water directly onto the smear). iv. Now add Gram’s iodine to the smear and allow it to stand for 10 secs. v. Wash off the iodine carefully with distilled water. vi. Destain the smear with 95% ethyl alcohol for about 10-20 secs. This is a critical step. The degree of alcohol decolourising depends on the thickness of the smear. vii. Wash gently with distilled water. viii. Add basic stain safranin for 30 secs. ix. Wash with distilled water and blot the slide dry with absorbent paper and examine under the oil immersion objective. Table 21.2. Gram reaction and morphology of some common bacterial genera (Adopted from Desai and Desai, 1980) Gram reactiona

Morphological characters

Genera

Positive

Rods in chain, non-sporulating

Lactobacillus

Rods either in the chain or single sporulating

Bacillus

Besides the above characteristics, spores bigger than the cells

Clostridium

Bigger rods, often forming obtuse angle or Mycobacteriab small bundle of parallel cells

Negative

Cocci in bunch

Staphylococcus

Cocci in chain

Streptococcus

Cocci, ovoid or lanceolate, occurring in pair

Diplococcus

Cocci, sporulating

Sporosarcina

Rods either in single or short chain

Azotobacter, Corynebacterium Escherichia Rhizobium Proteus, Pseudomonas, Salmonella, Serratia, Shigella

Rods ovoid

Francisella Pasteurella Yersinia (Contd.)

436

Plant Techniques: Theory and Practice Table 21.2. (Contd.)

Gram reactiona

Morphological characters

Genera

Rods, curved Cocci in pair Cocci in single

Vibrio Neisseria Acidaminococcus Megasphaera Veillonella

Spiralc

Borrelia Leptospira Spirillum Treponema

a

Gram reaction of various genera listed above are based on the results of the test with 24 hrs old cultures. b The Gram reaction is debatable. c The staining of these organisms requires a special staining technique.

Result: Gram-positive appear purple by retaining crystal violet and gramnegative red by taking counterstain. Biological reaction: The primary stain, crystal violet, and Gram’s iodine form a complex called CV-1. Gram’s iodine acts as a mordant in the formation of this complex. This complex is taken up by the peptidoglycan layer in the bacterial cell wall (Fig. 21.2).

Fig. 21.2. Cell walls of Gram-positive and Gram-negative bacteria.

There is no universally accepted explanation for Gram’s reaction. The most widely accepted theory is based on the difference in permeability of the cell wall. Gram-negative cells have got a greater lipid content, in their walls than Gram-positive. Lipids are soluble in acetone and alcohol which are used

Microbiological Techniques 437

as a decolouriser in Gram staining. Removal of lipids by the decolouriser is thought to increase the porosity of cell walls and cause the rapid decolourisation of Gram-negative bacteria. Another interpretation is that gram-positive cell walls consist of many layers of peptidoglycan (Fig. 21.2; see Table 21.3); the CV-I is trapped in the cell wall following alcohol treatment which causes a decrease in the diameter of the pores in the cell wall peptidoglycan. In gramnegative bacteria, the peptidoglycan layer is much thinner and there is less cross-linkage than in the walls of gram-positive bacteria. Because of the large pores left in the gram-negative bacteria, the CV-I gets extracted upon alcohol treatment. Table 21.3. Some characteristic differences between gram-positive and gram-negative bacteria Character

Gram-positive

Gram-negative

1. Gram’s staining

Purple or dark violet

2. Cell wall composition 3. Peptidoglycan in cell wall 4. Teichoic acid in cell wall 5. Sensitivity to penicillin 6. Nutritional requirements

Low in lipid (1-4%) Many layers in thickness Present More Generally complex, only few species autotrophic More resistant

Red (the colour of the counterstain) High in lipid (11-20%) Very thin Absent Less Relatively simple, many species autotrophic Less resistant

7. Resistant to physical disruption Examples

Lactobacillus Clostridium Staphylococcus Streptococcus Diplococcus

Corynebacterium Salmonella Shigella, Proteus Pseudomonas Escherichia Azotobacter Rhizobium Pasteurella Vibrio, Spirillum

21.4.3.2 Modified Staining Technique for Gram-positive and Gramnegative Bacteria (Garvey et al., 1986)

Solution Crystal violet--dissolve 1.0 g crystal violet in 100 ml distilled water. Sodium bicarbonate--dissolve 2.5 g sodium bicarbonate in 100 ml distilled water. Gram’s iodine--dissolve l.0 g. iodine and 2.0 g. potassium iodide in 300 ml distilled water. Do not heat the solution because iodine sublimes.

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Plant Techniques: Theory and Practice

Cello solve (ethylene glycol monoethyl ether) Carbol new fuchsin stock solution Solution A – dissolve 1 g new fuchsin in 5.0 ml ethanol Solution B – mix 5.0 ml phenol in 90.0 ml distilled water Combine solutions A and B and filter before use. Working carbol new fuchsin is prepared by diluting 25.0 ml stock carbol new fuchsin solution with 75.0 ml distilled water. Tartrazine solution--dissolve 0.01 g tartrazine or acid yellow (C.I. 19140, Matheson, Coleman and Bell) in 100 ml distilled water to which 2.0 ml acetic acid has been added. Histoclear Staining procedure i. Placental tissues injected with a broth culture of Staphylococcus aureus,Clostridium perfringens and Escherichia coli and various autopsy and surgical sections were fixed in 10% neutral buffered formalin, cleared in Histoclear, embedded in paraplast plus and cut at 4 µm. ii. Deparaffinise and hydrate to distilled water. iii. Keep sections in working crystal violet solution (combine 1 ml 1%crystal violet and 0.2 ml 2.5% sodium bicarbonate just before use) for 1 minute. iv. Wash with tap water to remove excess crystal violet. v. Then keep in Gram’s iodine for 1 minute. vi. Wash with tap water to remove excess iodine. vii. Differentiate sections in cello-solve until obvious extraction of blue colour stops, then leave in the differentiator for 2 minutes. viii. Rinse in absolute alcohol. ix. Keep sections in working carbol new fuchsin solution for 90 seconds. x. Rinse briefly in tap water to remove excess carbol new fuchsin. xi. Counterstain in 0.01% tartrazine in 2% acetic acid solution for 15 seconds. xii. Dehydrate within 15 secs, clear, and mount. Result: Gram-positive bacteria and some fungi appear blue; gram-negative bacteria and nuclei appear red; background appears yellow. In this procedure, the inclusion of sodium bicarbonate in the crystal violetstain increases the resistance of the gram-positive bacteria to decolorisation. The brightness of gram-negative bacteria is maintained through the reduction of background staining brought about by acetic acid in the tartrazine counterstain. 21.4.3.3 Acid-faststaining

Acid-fast staining is also a differential staining employed to diagnose Mycobacterium tuberculosis and M. leprae which causes tuberculosis and leprosy respectively.

Microbiological Techniques 439

Preparation of reagents Ziehl-Neelsens’ Carbolfuchsin Basic fuchsin Ethanol, 95% Phenol crystals Distilled water

0.3 g 10.0 ml 5.0 g 95.0 ml

Dissolve basic fuchsin in the ethanol. In a separate container, dissolve phenol crystals in water. Mix these two solutions together thoroughly. Acid-alcohol decolouriser Conc. HCl (37%) Ethanol, 95%

3.0 ml 97.0 ml

Add conc. HCl to the ethanol and mix well. Loeffler’s methylene blue Methylene blue chloride Ethanol, 95% Distilled water

0.3 g 30.0 ml 100.00 ml

Dissolve methylene blue in the ethanol, to which add distilled water. Later filter and use it. Procedure i. ii. iii. iv. v. vi. vii.

Prepare and fix the bacterial smear onto a clean microslide. Stain the smear with carbolfuchsin for 5 minutes and heat gently. Gently wash the slide with distilled water. Now apply the decolouriser onto the slide for 1 minute. Wash carefully with distilled water. Counterstain the smear with methylene blue for 1 minute. Wash with distilled water and blot dry. Observe the acid-fast slide under oil immersion.

Result: Acid-fast bacteria appear red whereas non-acid-fast cells appear blue by accepting the counter stain. Biological reaction: The cell walls of acid-fast organisms contain a wax-like lipid called mycolic acid which makes the cell wall impermeable to most of dyes. Carbolfuchsin is easily soluble in carbolic acid (phenol) solution. Carbolfuchsin mixed with carbolic acid easily penetrated the cell wall. Here carbolic acid acts as a chemical intensifier for penetration. Heat is also applied to penetrate the stain into the cell wall. Once the cell wall is stained with dye, it resists decolourisation when washed with an acid-alcohol decolouriser.

440

Plant Techniques: Theory and Practice

21.4.3.4 Negative Staining

The bacterial capsule is composed of polysaccharides which are water soluble and non-ionic. Since the bacterial capsule is non-ionic, it cannot be stained in the usual manner. Several techniques have been developed that allow the background to be stained and leave the capsule unstained. This is known as negative staining. By using India ink, the background looks dark, and the capsule appears like a clear ring around the bacterial cell (Fig. 21.3).

Fig. 21.3. Gins’ method of capsule stain.

Modified Gins’ Capsule Stain using the Blood Smear Method (Beishir, 1996) In the modified Gins’ capsule stain, both negative and positive stains are employed. This can be observed under the oil immersion objective and magnified 1000 times. i. By using inoculating loop add Klebsiella pneumoniae to equal amounts of India ink and water and take it on one end of the clean slide. ii. With the help of the second slide spread the mixture on the slide. iii. Allow the smear to air-dry. iv. Now stain the smear with crystal violet for a minute. v. Drain and rinse the smear. vi. Allow to air-dry and observe under the microscope. Result: Indian ink stains the background dark leaving the capsule unstained and clear (negative stain) with the small purple cell body in the centre of the ring (positive staining by crystal violet, see Fig. 21.3). 21.4.3.5 Staining of Vesicular Arbuscular Mycorrhizae (VAM)

There are three major types of mycorrhizae such as ectomycorrhiza (ectotrophic mycorrhizae), endomycorrhizae and ectendomycorrhizae (a form of mycorrhizae in which hyphae may penetrate the plant cells).

Microbiological Techniques 441

The endomycorrhizae caused by the aseptate hyphae, particularly the members of Endogonales have often been referred to as Vesicular Arbuscular Mycorrhizae (VAM) because these fungi form dichotomously branched haustoria with the host cell called arbuscules. Apart from that, the fungal hyphae also bear a swollen thick structure called vesicles. These are extremely common and widespread in the plant kingdom (Aneja, 2003). Standard mounting media for determining VAM are water, lactophenol or lactoglycerine. Staining the spores with lactophenol cotton blue is useful for revealing the hyphal characteristics and for interpreting some types of the apparent surface texture of the spores. Chemical reagents and other materials

• • • • • • • • • •

VAM infected root 18% Lactophenol FAA 10% KOH 95% Ethyl alcohol Chlorazol black E Chloral hydrate Glycerine Basic fuchsin Mounting fluid [chloral hydrate 20 g, gum Arabic 20 g, glycerine 20 g, Glucose syrup 3 ml, basic fuchsin 10 drops (0.3 g., 10 ml 95% ethyl alcohol), 35 ml water] • Autoclave-resistant jars • Water bath at 90 ºC • Clean microslides Procedure i. Wash VAM-infected roots thoroughly and fix them overnight in FAA. ii. After that roots are washed with tap water several times to remove FAA. iii. Now roots are transferred to KOH solution taken in autoclave-resistant jars. iv. Clear the samples by autoclaving them at 15 psi for 15 minutes. v. Rinse the samples several times with tap water. vi. Wash with deionised water. vii. Stain roots with staining solution (equal volumes of 80% lactic acid, glycerine and distilled water with 0.1% chlorazol black E) for 1 hour or longer at 90ºC. viii. Now stained roots are kept in glycerine for destaining. ix. Finally mount the roots on a clean micro slide using mounting media. Observe the stained VAM fungi under the bright field microscope for their structures. Here, the staining solution is to be prepared several hours

442

Plant Techniques: Theory and Practice

before use and it is kept undisturbed to allow undissolved particles to settle down later the particles are discarded. 21.4.3.6 Lactophenol Cotton Blue Mounting of Fungi

Lactophenol cotton blue stains the fungal cytoplasm and gives a light blue background, against which the walls of the hyphal can readily be seen. The stain contains four constituents, such as phenol, lactic acid, cotton blue and glycerine. Phenol serves as a fungicide; lactic acid acts as a clearing agent; cotton blue stains the cytoplasm of the fungus, and finally, glycerine provides the semi-permanent preparation. Chemical reagents and other materials

• • • • • • • •

A young culture of Penicillium or Aspergillus (5-7 days old) Lactophenol cotton blue in a dropper bottle Microslides and coverslips Mounted needles (2) Spirit lamp 70% alcohol Nail polish (or) paraffin wax Compound microscope

Procedure i. Put a drop of lactophenol cotton blue on a clean microslide. ii. Transfer a small tuft of the fungus (containing spores and spore-bearing structures) onto the slide using a flamed, cooled needle. iii. Gently tease the sample using the needles. iv. Mix gently the stain with mould structures. v. Place a coverslip over the preparation (see that to avoid air bubbles trapping in the stain). vi. To keep the slides for several months, apply a thin layer of nail polish around the edge of the coverslip and let the preparation dry overnight. Observations and Results Observe the slide under a compound microscope with high-power lenses. The fungal cytoplasm is seen as a lightly stained blue region forming a layer inside the unstained cell wall of hyphae, conidiophores, phialides, conidia, that is surrounded by light blue background on the slide. 21.4.3.7 Antimicrobial Activity

Plants and other natural sources can provide a huge range of complex and structurally diverse compounds. Recently, many researchers have focused on the investigation of plant and microbial extracts, essential oils, pure secondary metabolites and new synthesised molecules as potential antimicrobial agents. A variety of laboratory methods can be used to evaluate or screen the in vitro

Microbiological Techniques 443

antimicrobial activity of an extract or a pure compound. The most known and basic methods are the disk-diffusion and agar well methods. Disk Diffusion Method (Tendencia, 2004) This method is based on the principle that an antibiotic-impregnated disk, placed on agar previously inoculated with the test bacterium, pick-up moisture, and the antibiotic diffuses radially outward through the agar medium producing an antibiotic concentration gradient. The concentration of the antibiotic at the edge of the disk is high and gradually diminishes as the distance from the disk increases to a point where it is no longer inhibitory for the organism, which then grows freely. A clear zone or ring is formed around an antibiotic disk after incubation if the agent inhibits bacterial growth. The disk diffusion method is performed using Mueller-Hinton Agar (MHA), which is the best medium for routine susceptibility tests because it has good reproducibility, low in sulphonamide, trimethoprim, and tetracycline inhibitors, and gives satisfactory growth of most bacterial pathogens. The inoculum for the disk diffusion method is prepared using a suitable broth such as tryptic soy broth. This medium is prepared according to the manufacturer’s instructions, dispensed in tubes at 4-5 ml and sterilised. A sterile 0.9% salt solution may also be used. Media are supplemented with 1-2% sodium chloride (NaCl) is intended for marine organisms. I. Preparation of agar medium i. Prepare MHA from the dehydrated medium according to the manufacturer’s instructions. Media should be prepared using distilled water or deionised water. ii. Heat with frequent agitation and boil to dissolve the medium completely. Sterilise by autoclaving at 121°C for 15 min. iii. Check the pH of each preparation after it is sterilised, which should be between 7.2 and 7.4 at room temperature. This is done by macerating a small amount of medium in a little distilled water or by allowing a little amount of medium to gel around a pH meter electrode. iv. Cool the agar medium to 40-50°C. Pour the agar into a sterile glass or plastic petri dish on a flat surface to a uniform depth of 4 mm. v. Allow to solidify. vi. Prior to use, dry plates at 30-37°C in an incubator, with lids partly ajar, for not more than 30 minutes or until excess surface moisture has evaporated. Media must be moist but free of water droplets on the surface. The presence of water droplets may result in swarming bacterial growth, which could give inaccurate results. They are also easily contaminated. Storage If plates are not to be immediately used, they may be stored in the refrigerator inside airtight plastic bags at 2-8°C for up to 4 weeks.

444

Plant Techniques: Theory and Practice

Unpoured media may be stored in airtight screw-capped bottles under the conditions specified by the manufacturer. Control Before use, check the ability of the agar to support the growth of control strains (listed in the Introduction) by streaking bacterial cultures on the agar medium. It is also advisable to check the ability of each batch of media to support the growth of a representative member of the species to be tested. II. Inoculum Preparation i. From a pure bacterial culture (not more than 48 hours, old except for slow-growing organisms), take four or five colonies with a wire loop. ii. Transfer colonies to 5 ml of Trypticase soy broth or 0.9% saline. iii. Incubate the broth at 30°C or at an optimum growth temperature until it achieves or exceeds the turbidity of 0.5 Mac Farland standard (prepared by adding 0.5 ml of 0.048 M BaCl2 to 99.5 ml of 0.36 NH2 SO4; commercially available). iv. Compare the turbidity of the test bacterial suspension with that of 0.5 Mac Farland (vigorously shaken before use) against a white background with a contrasting black line under adequate light. v. Reduce turbidity by adding sterile saline or broth. III. Inoculation of plates i. Dip a sterile cotton swab into the standardised bacterial suspension. ii. Remove excess inoculum by lightly pressing the swab against the tube wall at a level above that of the liquid. iii. Inoculate the agar by streaking with the swab containing the inoculum. iv. Rotate the plate by 60° and repeat the rubbing procedure. Repeat two times. This will ensure an even distribution of the inoculum. v. Allow the surface of the medium to dry for 3-5 minutes but not longer than 15 minutes to allow for the absorption of excess moisture. IV. Antimicrobial disks i. The number of antimicrobial agents to be tested should be limited. To make the test practical and relevant, include only one representative of each group of related drugs; those indicated for veterinary use to control or prevent disease, and those that can be useful for epidemiological or research purposes. Use antibiotic disks purchased from a reputable manufacturer. The disk diameter is approximately 6 mm. Disks should be properly stored in a tightly sealed container with a desiccant at 2-8°C. Expired disks should not be used. Application: i. Using sterile forceps or a disk dispenser, place the antibiotic disk on the surface of the inoculated and dried plate.

Microbiological Techniques 445

ii. Immediately press it down lightly with the instrument to ensure complete contact between the disk and the agar surface. Do not move a disk once it has come into contact with the agar surface since some diffusion of the drug occurs instantaneously. iii. Position disks such that the minimum center-center distance is 24 mm and no closer than 10 to 15 mm from the edge of the petri dish. A maximum of six disks may be placed in a 9-cm petri dish and 12 disks on a 150 mm plate. Reduce the number of disks applied per plate if overlapping zones of inhibition are encountered. Control Include one plate inoculated with a control strain (Table 21.4) for every set of plates and incubate together. Incubation i. Incubate plates in an inverted position at 30°C or at an optimum growth temperature. ii. Observe for the zone of inhibition after 16 to 18 hours. Slow growing organisms may require longer incubation period. Reading and measurement of zones of inhibition i. The zone of inhibition is the point at which no growth is visible to the unaided eye. ii. Record the presence of individual colonies (arrow) within zones of inhibition. iii. Record occurrence of fuzzy zones (arrow). In measuring the zone diameter, the fuzzy portion of the zone should be ignored as much as possible. The zone limit is the inner limit of the zone of normal growth. iv. Read and record the diameter of the zones of inhibition using a ruler graduated to 0.5 mm. v. Round up the zone measurement to the nearest millimeter. Interpretation of results Compare the diameter of the zone of inhibition of the test isolates with those in the chart of interpretative standards for veterinary pathogens. Report result as follows: Resistant (R), Intermediate (I) or Susceptible (S). Example Disk used: Chloramphenicol, 30 µg (C-30) Zone of inhibition: 16 mm Result/interpretation Intermediate is based on the zone diameter interpretative chart (Table 21.5) Susceptibility test results using agents other than those listed in the chart are interpreted based on the presence or absence of a definite zone of inhibition and is considered only as qualitative until such time as interpretative zones have been established.

Disk

Escherichia coli

Staphylococcus aureus

Pseudomonas aeruginosa

Streptococcus pneumoniaea

Content

ATCC 25922

ATCC 25923

ATCC 27853

ATCC 49619

30 µg

19-26

20-26

18-26

-

20/10 µg

18-24

28-36

-

-

Ampicillin

10 µg

16-22

27-35

-

30-36

Cefazolin

30 µg

21-27

29-35

-

-

Cefoxitin

30 µg

23-29

23-2

Cephalothin

30 µg

15-21

29-37

-

Chloramphenicol 30 µg

21-27

19-26

-

26-32

Clindamycin

2 µg

-

24-3

-

19-25

Erythromycin

15 µg

-

22-30

-

25-30

Gentamicin

10 µg

19-26

19-27

16-21

-

Imipenem

10 µg

26-32

-

20-28

-

Kanamycin

30 µg

17-25

19-26

-

-

Oxacillin

1 µg

-

18-24

-

17

< 14

Gentamicin*

10 µg

> 15

13-14

< 12

Kanamycin*

30 µg

>18

14-17

< 13

Spectinomycin

100 µg

> 14

11-13

< 10

Staphylococci

20/10 µg

> 20

-

< 19

Other organisms

20/10 µg

>18

14-17

< 13

Pseudomonas aeruginosa

75/10 µg

> 15

-

< 14

Gram(-)enteric organisms

75/10 µg

>20

15-19

< 14

Enterobacteriaceae

10 µg

> 17

14-16

< 13

Staphylococci

10 µg

> 29

-

< 28

Amoxicillinclavulanic acid*

Ticarcillin-clavulanic acid*

Ampicillin*

Enterococci

10 µg

>17

-

< 16

Streptococci (not S. pneumoniae)

10 µg

> 26

19-25

< 18

1 µg

> 13

11-12

< 10

10 units

> 29

-

< 28

Oxacillin* Staphylococci Penicillin* Staphylococci

10 units

> 15

-

< 14

1 µg oxacillin

> 20

-

-

10 units

> 28

20-27

< 19

Pseudomonas aeruginosa

75 µg

> 15

-

< 14

Gram (-) enteric organisms

75 µg

> 20

15-19

< 14

10 units/30 µg

> 18

15-17

< 14

Imipenem*

10 µg

> 16

14-15

< 13

Cephalothin*

30 µg

> 18

15-17

< 14

Enterococci S. pneumoniae Streptococci (not S. pneumoniae) Ticarcillin*

Penicillin-novobiocin

Microbiological Techniques 449 Cefazolin*

30 µg

> 18

15-17

< 14

Ceftiofur

30 µg

> 21

18-20

< 17

Enrofloxacin (canine/feline)

5 µg

> 23

-

Enrofloxacin (chickens/turkeys)

5 µg

> 23

17-22

< 16

Enrofloxacin (bovine)

5 µg

> 21

17-20

< 16

Difloxacin

10 µg

> 21

18-20

< 17

Orbifloxacin

10 µg

> 28

-

Clindamycin

2 µg

> 21

15-20

< 14

Pirlimycin

2 µg

> 13

-

< 12

Streptococci

15 µg

> 21

16-20

< 15

Organisms other than Streptococci

15 µg

> 23

14-22

< 13

Tilmicosin (Bovine)

15 µg

> 14

11-13

< 10

Tilmicosin (Swine)

15 µg

> 11

Streptococci (not S. pneumoniae)

30 µg

> 21

18-20

< 17

S. pneumoniae

30µg

> 21

-

< 20

Organisms other than Streptococci

30 µg

> 18

13-17

< 12

Florfenicol

30 µg

> 19

15-18

< 14

Tiamulin

30 µg

>9

-

19

16-18

< 15

Organisms other than S. pneumoniae

1.25/23.75 µg

> 16

11-15

< 10

Streptococcus pneumoniae

5

> 19

17-18

< 16

Organisms other than Streptococci

5

> 20

17-19

< 16

250 or 300

> 17

13-16

< 12

30

> 23

19-22

< 18

17-22

18-22

< 16

< 17

Erythromycin*

< 10

Chloramphenicol*

Trimethoprimsulfamethoxazole*

Rifampin*

Sulfisoxazole* Tetracycline* Streptococci

(Contd.)

450

Plant Techniques: Theory and Practice Table 21.5. (Contd.)

Antimicrobial Agent

Disk Content

Zone Diameter (mm) S

I

F

R

Vancomycin* Enterococci

30

> 17

15-16

< 14

Streptococci

30

> 17

-

-

Other gram-positive organisms

30

> 12

10-11