Cytological methods for the detection, identification, and characterization of orchid viruses and their inclusion bodies

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Title:
Cytological methods for the detection, identification, and characterization of orchid viruses and their inclusion bodies
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vii, 83 leaves : ill. ; 28 cm.
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Ko, Nan-Jing, 1948-
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Subjects / Keywords:
Orchids -- Diseases and pests   ( lcsh )
Virus diseases of plants   ( lcsh )
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bibliography   ( marcgt )
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non-fiction   ( marcgt )

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Thesis:
Thesis (Ph. D.)--University of Florida, 1985.
Bibliography:
Includes bibliographical references (leaves 75-82).
Statement of Responsibility:
by Nan-Jing Ko.
General Note:
Typescript.
General Note:
Vita.

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Full Text







CYTOLOGICAL METHODS FOR THE DETECTION,
IDENTIFICATION, AND CHARACTERIZATION OF ORCHID
VIRUSES AND THEIR INCLUSION BODIES










By

NAN-JING KO


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY


UNIVERSITY OF FLORIDA


1985
























This dissertation is dedicated to my
mother, Min-Chu Ko.


J_ P %












ACKNOWLEDGMENTS


I wish to express my sincere gratitude to Drs. J. R.

Edwardson and F. W. Zettler whose guidance, encourage-

ment, and friendship made the fulfillment of my educa-

tional aspirations real.

Special thanks are due to Mr. R. G. Christie and to

Drs. E. Hiebert, D. E. Purcifull, H. C. Aldrich and T. J.

Sheehan for their guidance and helpful suggestions in the

preparation of this dissertation.

Appreciation is also extended to Mr. S. Christie, Mr.

W. Crawford, Ms. G. C. Wisler, Mr. C.-A. Chang, Ms. A. E.

Logan, Mr. M. S. Elliott and Ms. C. A. Baker for their

generous assistance during the course of this investigation.

I am also grateful to Dr. G. Erdos of the Microbiology and

Cell Science Department for technical assistance in immunogold

labelling and for supplying protein A-gold.

I am indebted to my family for their moral and financial

support and especially to my wife for her help and companion-

ship and to my brothers for taking care of my mother

throughout these years.

Appreciation is also expressed to the National Science

Council of the Republic of China and to the American Orchid

Society.


iii













TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS. . . . . iii

ABSTRACT. . . . . .. vi

CHAPTERS

1 INTRODUCTION. . . . . 1

2 USE OF LIGHT MICROSCOPY TO DETECT AND
DIAGNOSE ORCHID VIRUSES . . 5

Introduction . . . . 5
Materials and Methods . . . 6
Results . . . . 10
Odontoglossum Ringspot Virus. . 10
Cymbidium Mosaic Virus. . . ... 15
Cucumber Mosaic Virus . . 18
Bean Yellow Mosaic Virus. . . 22
Uncharacterized Bacilliform Virus . 25
Discussion . . . . 29

3 CONFIRMATORY STUDIES OF LIGHT MICROSCOPE
OBSERVATIONS. . . . .. 33

Introduction . . . .. 33
Materials and Methods . . ... 34
Leaf Dips . . . . 34
Thin Sectioning .. . . . 35
Immunofluorescence Microscopy . ... 36
Protein A-Gold Labelling. . ... 37
Results . . . . 38
VaMo Negative Staining . . ... 38
Thin Sectioning . . . 39
Immunofluorescence Microscopy . 43
Protein A-Gold Labelling. . ... 50
Discussion . . . .. 56

4 CONCLUSIONS .. . . . 62










Page

APPENDICES

I A SIMPLIFIED BIOASSAY TECHNIQUE FOR CYMBIDIUM
MOSAIC AND ODONTOGLOSSUM RINGSPOT VIRUSES 66

II AN EFFICIENT PROCEDURE FOR STAINING LARGE
NUMBERS OF ELECTRON MICROSCOPIC GRIDS . 70

III COMPARATIVE DIAGNOSES OF ORCHID VIRUSES BY
LIGHT MICROSCOPY, ELECTRON MICROSCOPY, AND
SEROLOGY. . . . . .. 73

LITERATURE CITED. . . . .. 75

BIOGRAPHICAL SKETCH . . . . 83








Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy


CYTOLOGICAL METHODS FOR THE DETECTION,
IDENTIFICATION, AND CHARACTERIZATION OF ORCHID
VIRUSES AND THEIR INCLUSION BODIES

By

Nan-Jing Ko


May 1985


Chairman: F. W. Zettler
Cochairman: J. R. Edwardson
Major Department: Plant Pathology

Light microscopy was explored for detecting and diagnosing

orchid viruses. Bioassay, serology and electron microscopy

have all been used for viral detection, but most studies have

involved only Cymbidium mosaic and Odontoglossum ringspot

viruses. The other orchid viruses are not well-characterized

and practical means for their diagnoses in orchids have not

been developed. In this investigation, light microscopic

techniques were developed and their diagnostic potential was

assessed. Calcomine orange-Luxol brilliant green (O/G)

combination and Azure A were used for selective staining of

proteinaceous inclusions and inclusions containing

nucleoprotein, respectively. Six viruses were studied,

Cymbidium mosaic (CyMV), Odontoglossum ringspot (ORSV), bean

yellow mosaic (BYMV), cucumber mosaic (CMV), and two

uncharacterized bacilliform viruses. Other techniques were

used to verify the light microscopic results. These included

vi








immunofluorescence microscopy, immunogold labelling, and

conventional electron microscopy (leaf dips and ultrathin

sectioning). The preparation of ultrathin sections was

facilitated considerably by the development of a device for

simultaneous staining of 10 or more grids.

Banded and crescent shaped inclusions were found in CyMV-

infected tissues stained with Azure A. Paracrystalline and

stacked-plate inclusions were formed in the ORSV-infected

tissues stained with either O/G combination or Azure A. In

CMV-infected tissues stained with Azure A, angular crystals

with or without clear centers were found. Cytoplasmic

cylindrical inclusions were observed in the BYMV-infected

tissues stained with the O/G combination. Nuclear inclusions

occurred in the bacilliform virus-infected tissues stained with

either the Azure A or O/G combination.

Direct fluorescence microscopy confirmed the presence of

viral-related agents in the CyMV-, ORSV- and CMV-infected

tissues. Vanadyl molybdate-phosphotungstate (VaMo) negative

staining was used for confirming the presence of CyMV and ORSV.

Protein A-gold labelling was applied to identify CyMV, ORSV,

BYMV and CMV in the thin sections of tissues embedded in LR

White resin.

Light microscopy was demonstrated to be a useful diagnostic

tool for orchid viruses. The diagnoses by light microscopy were

confirmed by VaMo negative staining, thin sectioning,

immunofluorescence microscopy and protein A-gold labelling.

vii















CHAPTER 1
INTRODUCTION

Orchidaceae is one of the largest and most diverse

families of plants. It includes 7-10% of all flowering plant

species (Dressler, 1981). Orchids are widely grown throughout

the world for sale as cut flowers or as potted plants. The

value of an individual mature plant can range from a few

dollars to several thousand dollars. In Florida, orchid

production has been estimated to have an annual sale value of

three million dollars (Anonymous, 1983), while the Federal

Republic of Germany imported $22.9 million of fresh-cut orchids

(Anonymous, 1978).

The value of individual orchids is reduced considerably by

plant pathogenic infections. Viruses are particularly

problematic in orchids wherever they are grown. The most

serious damage occurs when symptoms become evident on the

flowers. Twenty-two viruses have been reported in orchids

(Lawson and Horst, 1984). Cymbidium mosaic (CyMV) and

Odontoglossum ringspot (ORSV) viruses, the first orchid viruses

to be described, are the most prevalent. Many of the other

viruses of orchids have only recently been described and have

not been well-characterized. The prevalence of viruses in


-1-






-2-

orchids, particularly CyMV and ORSV, is presumably due to three

factors: i) some are readily transmitted by contaminated

cutting tools used during propagation and flower harvesting;

ii) the genetic diversity of the Orchidaceae results in

variable symptom expression and therefore symptoms are

unreliable as a means of diagnosis (Sheehan, 1980); and

iii) the viruses have been widely distributed through the

international exchange of individual plants.

Bioassay and electron microscopical and serological

techniques have been used previously for detecting and

diagnosing some orchid viruses (Lawson and Ali, 1975; Lawson

and Horst, 1984; Inouye, 1977; Lawson and Brannigan, in press;

Wisler et al., 1982). For CyMV, ORSV, and certain other

orchid viruses, these methods have proven to be reliable and

practical. Certain bioassay and serological techniques have

been modified specifically for orchids (Appendix I; Wisler et

al., 1982). However, such methods are not applicable for most

of the other orchid viruses listed by Lawson and Horst (1984).

This limitation is likely to be very significant to the orchid

industry, especially as some viruses have aerial vectors and

can spread rapidly in collections. Bioassays by manual

inoculation are not applicable to viruses such as the

Masdevallia isometric and Cypripedium filamentous viruses which

are apparently not transmissible by this means. Likewise,

serological techniques cannot be applied to any of the

rhabdoviruses for which antisera are not yet available.







-3-

Electron microscopy of negatively stained samples also has

limitations, particularly for isometric viruses such as

Cymbidium ringspot and/or cucumber mosaic viruses which have

similar particle morphologies. Electron microscopy of thin

sections is not also applicable for routine diagnosis due to

the time, labor, and expense involved in processing samples.

The purpose of this study was to explore the potential of

light microscopy as a method for detecting and diagnosing

orchid viruses. Immunofluorescence microscopy, protein A-gold

labelling, VaMo negative staining, thin sectioning,

immunodiffusion and slicing bioassay (Appendix I) techniques

were used to verify the results of light microscopic diagnoses.

Christie and Edwardson (1977) have described light

microscopic techniques as being very useful for diagnosing

plant viruses at the group level and, in some instances, at the

specific level. This system has also been shown to be useful

for diagnosis of viruses infecting certain crops, such as

peppers (Edwardson and Christie, 1979). The low cost of the

basic equipment and the relatively short time involved in

processing samples are some advantages of light microscopy.

However, certain limitations must be overcome before it can be

applied to crops such as orchids. Unlike many herbaceous

plants, such as peppers, most orchid leaves are thick and

leathery, and previously described techniques, such as

epidermal peels (Hiebert et al., 1984), are difficult to

obtain. For orchids, therefore, special modifications must be






-4-

developed. A second important problem to overcome is the lack

of background information about orchid viruses, most of which

are poorly characterized. Indeed, of the 22 viruses listed by

Lawson and Horst (1984), only 12 viruses have been assigned to

virus groups by the International Committee for the Taxonomy of

Viruses (ICTV) (Matthews, 1982). Finally, the resolution

limitations of light microscopy makes confirmation by other

means necessary regarding the characterization of some of the

inclusions observed. The lack of such information, especially

for orchid viruses, makes such studies a necessary contingency

for this type of investigation.













CHAPTER 2
USE OF LIGHT MICROSCOPY TO DETECT AND DIAGNOSE
ORCHID VIRUSES




Introduction


Odontoglossum ringspot (ORSV) and Cymbidium mosaic (CyMV)

are the best characterized orchid viruses, and several reliable

bioassay and electron microscopical and serological techniques

for their detection have already been developed (Appendix I,

Corbett, 1974; Inouye, 1977; Lawson and Ali, 1975; Lawson and

Brannigan, in press; Wisler et al., 1982). Since the discovery

of these two viruses, at least 20 additional orchid viruses have

been reported, some of which induce debilitating diseases

(Lawson and Horst, 1984; Lawson and Brannigan, in press). However,

only 12 of these viruses are sufficiently characterized to be

assigned to groups by the International Committee for the

Taxonomy of Viruses (Matthews, 1982). This deficiency of

information can be attributed in part to a lack of convenient

diagnostic techniques, which is especially true for those

viruses that are not manually transmissible and for which

antisera have not been prepared.

Light microscopy is useful for diagnosing viruses

representing 16 of the 28 recognized virus groups (Matthews,

-5-







-6-

1982) based on the inclusions they induce (Christie and

Edwardson, 1977; Edwardson and Christie, 1978; Hamilton et al.,

1981). This study explores the applicability of the light

microscope for diagnosing orchid viruses, as was done

previously for pepper viruses (Edwardson and Christie, 1979).

Electron microscopy of the inclusions found by light

microscopy was conducted to reveal the fine structure of the

inclusions.



Materials and Methods



Among the genera and viruses (in parentheses) studied in

this investigation were: Brassia (uncharacterized bacilliform

virus), Cattleya (ORSV, CyMV), Cymbidium (ORSV, CyMV,

uncharacterized bacilliform virus), Masdevallia (bean yellow

mosaic virus), Phalaenopsis (ORSV, CyMV, cucumber mosaic

virus), and Vanilla (CyMV). For each plant tested, an unopened

new leaf and the first and second mature leaves (with or

without evidence of symptoms) were used.

The Azure A and the calcomine orange/Luxol brilliant green

stain combinations (O/G) were used (Christie and Edwardson,

1977; Hiebert et al., 1984) for light microscopic examination

of viral inclusion bodies. The thick texture of most orchid

leaves precluded removal of epidermal strips for staining. To

overcome this problem, the following modifications were

employed: 1) For thin-leaved orchids (e.g., Phaius), leaf







-7-

pieces (ca. 0.5 x 3.0 cm) were cut, laid upon glass slides, and

gently rubbed with 320 grit sandpaper first and then 600 grit

sandpaper (cut into 1 x 4 cm pieces) to remove the cuticle and

permit stain penetration. Using this technique, it was

possible to remove enough overlying tissues to reveal the

epidermal layers below those being abraded. 2) For thicker-

leaved orchids such as Cattleya, tissues were sliced

paradermally or longitudinally with razor blades. Paradermal

sections were obtained by cutting leaves parallel to the leaf

surface (Fig. 1), while longitudinal sections were obtained by

inserting leaf tissues in either styrofoam or pith as a support

while they were being sectioned.

The thick-walled tissues infected with ORSV were stained

with the O/G combination at room temperature for 10 minutes and

then heated at 60 C for another 5 minutes. Leaves with thin-

walled cells were stained in the O/G combination at room

temperature for 15 minutes. In some instances, CyMV-infected

tissues were fixed in 5% glutaraldehyde (in 0.1 M sodium

phosphate, pH 7.0) for two hours and then rinsed with the same

buffer to preserve inclusion fine structure. For the

bacilliform viruses, paradermal strips were first treated with

either 5% Triton X-100 for 5 minutes or 1% Triton X-100 for 10

minutes followed by staining in Azure A at 60 C for 10 minutes.

Comparable tissues were also examined by electron

microscopy to confirm the light microscopic observations.

Tissues were fixed for thin sectioning in 5% glutaraldehyde,







-8-


Figure 1. Paradermal section (arrows) of Cattleya
showing the thickness of tissue to be cut
with a razor blade for staining.






-9-


postfixed in 1-2% Os4, dehydrated with acidified 2,2-

dimethoxypropane (Muller and Jacks, 1975), and embedded in

Spurr's low-viscosity medium (Spurr, 1969). Sections were made

with either a glass or a diamond knife and were stained with

potassium permanganate, uranyl acetate, and lead citrate (Ko

and Chen, 1982).

The identities of ORSV, CyMV, bean yellow mosaic virus

(BYMV), and cucumber mosaic virus (CMV) were confirmed in tests

using antisera to the respective viruses. Expressed sap was

tested against antisera to the respective viruses in sodium

dodecyl sulfate (SDS) double radial immunodiffusion tests as

described previously (Purcifull and Batchelor, 1977; Zettler et

al., 1978; Kuwite and Purcifull, 1982; Wisler et al., 1982).

The immunodiffusion medium used contained 0.5% SDS, 1% NaN3,

0.8% Noble agar. Slicing bioassays were used to identify the

CyMV and ORSV on the indicator hosts, Cassia occidentalis and

Gomphrena globosa, respectively (Appendix I). Conventional

bioassay techniques (Lawson and Brannigan, in press) were used

for CMV and BYMV. Cucumber mosaic virus infections were confirmed

by inoculation of several hosts, including cucumber (Cucumis

sativus 'Marketer'), tomato (Lycopersicon esculentum),

Nicotiana benthamiana, N. x edwardsonii, N. glutinosa, N.

tabacum Xanthi nc., cowpea (Vigna unguiculata), and V. radiata.

Bean yellow mosaic virus infections were confirmed by






-10-

inoculation of peas (Pisum sativum 'Alaska', 'Ranger', and

'Little Marvel') and N. benthamiana.

Freshly sectioned orchid tissues were exposed to antisera

(refer to chapter 3) conjugated with tetramethyl rhodamine

isothiocyanate (TRITC), and were examined for virus-induced

inclusions with a Nikon Fluophot fluorescence microscope

(Hiebert et al., 1984).



Results

Odontoglossum Ringspot Virus

Crystalline stacked plate and paracrystalline inclusions

typical of ORSV and some other tobamoviruses (Christie and

Edwardson, 1977; Edwardson and Zettler, in press) were readily

observed by light microscopy in epidermal and mesophyll tissues

of infected plants (Figs. 2 and 3). These inclusions are

aggregations of virus particles (Figs. 4 and 5), as determined

by electron microscopy of thin sections.

Thick-walled tissues required prestaining at room

temperature for 10 minutes followed by heating at 600C for 5

minutes to achieve stain penetration. Treatment with the O/G

combination without heat resulted in poor staining of

inclusions (Fig. 6). Tissues were stained with Azure A for 15-

20 minutes at 60 0C to detect the nucleic acid in inclusions.

These inclusions were stained reddish-violet (Figs. 8 and 9),

but they did not stain with Azure A at room temperature (Fig.

7). Heat treatment (60C for 1-2 minutes) has also been shown

to be necessary for staining crystalline, paracrystalline,

angled-layered-aggregate and fibrous mass inclusions of other







-11-


Figure 2.




Figure 3.


Light micrograph of paracrystals (P) and
polar view of stacked-plate inclusions (SI)
of ORSV-infected Cattleya leaf cells
stained with O/G combination. N: Nucleus.
Bar = 2 Am.
Light micrograph showing side view of
stacked-plate inclusion (SI) in ORSV-
infected Cymbidium leaf cells stained with
O/G combination. Bar = 5 .um.







-12-


* P


4-


Electron micrograph of a portion of
paracrystal induced by ORSV in Cattleya
leaf mesophyll cell. Bar = 400 nm.
Electron micrograph of a portion of
stacked-plate inclusions induced by ORSX
in Cattleya leaf mesophyll cell.
Bar = 300 nm.


Figure 4.


Figure 5.






-13-


Figure 6.



Figure 7.


Light micrograph of poorly stained
paracrystalline inclusions (P) induced by ORSV
in Cattleya leaf cells. Tissue stained with
O/G without heat. Bar = 3 Aim.
Light micrograph of ORSV-infected Cattleya
leaf cells stained with Azure A without heat
showing poorly stained inclusions. P:
Paracrystal, SI: Stacked-plate inclusion. Bar
= 5 Am.







-14-


1V


Figure 8.



Figure 9.


Light micrograph of paracrystal (P) induced by
ORSV in Cattleya leaf cells stained with Azure
A heated at 60 C for 20 min. N: Nucleus.
Bar = 3 Aum.
Light micrograph of stacked-plate inclusion
(SI) (polar view) induced by ORSV in Cattleya
leaf cell, stained with Azure A heated at 60 C
for 20 min. N: Nucleus. Bar = 5 Aum.





-15-


tobamoviruses when using Azure A (Christie and Edwardson,

1977).

In fluorescence microscopy, the ORSV inclusions fluoresced

when treated with TRITC-conjugated ORSV antiserum (refer to

chapter 3). However, they did not fluoresce when either TRITC-

conjugated CyMV antiserum or normal serum was used. Expressed

sap from ORSV-infected plants used in this study reacted

homologously in SDS immunodiffusion tests with ORSV antiserum.

In the bioassay, ORSV infection caused chlorotic spots on the

leaves of Gomphrena globosa as described by Lawson and

Brannigan (in press).

Cymbidium Mosaic Virus

Reddish-violet inclusions typical of other potexviruses

were found in Azure A-stained epidermal and mesophyll cells of

plants infected with CyMV (Christie and Edwardson, 1977). In

glutaraldehyde-fixed tissues, some of these inclusions were

banded (Fig. 10). Crescent- or tear-shaped inclusions were

also readily found in the Azure A-stained tissues (Fig. 11).

Thin sections of banded-body inclusions consisted of large

aggregates of tiered (Fig. 12) or whorled, and intertwined

virus particles (Fig. 13) like those described previously for

CyMV and other potexviruses (Chen et al., 1983; Christie and

Edwardson, 1977; Hammond and Hull, 1983; Hanchey et al., 1975;

Hiruki et al., 1980; Lawson and Hearon, 1974; Purcifull and

Edwardson, 1981).

CyMV-induced inclusions fluoresced when exposed to

TRITC-labelled CyMV antiserum (refer to chapter 3), but not






-16-


-y *
S


I


Figure 10.


Figure 11.


Light micrograph of banded-body inclusion (I)
induced by CyMV in Cymbidium leaf cell fixed
by 5% glutaraldehyde. CW: Cell Wall, N:
Nucleus. Bar = 3 um.
Light micrograph of non-banded, crescent-
shaped inclusion (NI) induced by CyMV in
Cymbidium leaf cell fixed by 5%
glutaraldehyde. N: Nucleus. Bar = 2 Am.







-17-


4 r


I_ 'C'


Figure 12.



Figure 13.


Electron micrograph of a portion of a
banded-body inclusion (side view)
induced by CyMV in a Cymbidium leaf
cell. CW: Cell Wall. Bar = 420 nm.
Electron micrograph of banded-body
inclusions (polar view) induced by CyMV
in a Cymbidium leaf cell. CW: Cell
Wall. Bar = 360 nm.






-18-


when labelled ORSV or normal serum was used. Reactions of

identity were obtained between antigens in expressed sap and

antigens of a known CyMV isolate when tested against CyMV

antiserum in SDS immunodiffusion tests. In the bioassay, CyMV

infection caused dark brown spots on the cotyledons and

leaflets of Cassia occidentalis as described by Lawson and

Brannigan (in press).

Cucumber Mosaic Virus

Color breaks on the flowers (Fig. 14) and chlorosis on the

leaves were found in the Phalaenopsis orchid infected with CMV

(Fig. 15). Angular shaped, densely stained inclusions of

variable sizes (Fig. 16) were seen in the tissues of this

plant. In some instances, the inclusions had clear internal

areas, similar to those noted by others for CMV and other

cucumoviruses (Christie and Edwardson, 1977). The same type of

inclusion was also found in the other systematically infected hosts,

such as N. x edwardsonii (Fig. 17). Inclusions were much more

abundant in mesophyll than epidermal cells of the orchid flower

or leaf tissue. They also occurred much more abundantly in the

newly emerged immature leaves. Thin sections revealed these

inclusions to consist of massive crystalline arrays of densely

packed virus particles (Fig. 18) (Christie and Edwardson,

1977; Russo and Martelli, 1973). Such inclusions fluoresced

when exposed to TRITC-labelled CMV antiserum (refer to chapter

3). Expressed sap of CMV-infected tissues reacted homologously

in immunodiffusion tests with CMV antiserum produced by Kuwite







-19-


Figure 14. Color breaking of Phalaenopsis flowers
caused by CMV infection.
Figure 15. Chlorotic Phalaenopsis leaf infected with
CMV.








-20-


Figure 16.



Figure 17.


a) and b) Light micrograph of CMV induced
crystalline inclusions (C) in Phalaenopsis
leaf cells stained with Azure A. N: Nucleus.
Bar = 5 uAm.
Light micrograph of CMV induced crystalline
inclusions (C) in N. x edwardsonii leaf cells
stained with Azure A in low magnification.
X: Xylem tissue. Bar = 24 um.







-21-


Figure 18. Electron micrograph of a portion of a
CMV induced crystalline inclusion in
Phalaenopsis petal cell. Bar = 350 nm.
Inset shows higher magnification of virus
particles. Bar = 120 nm.






-22-


and Purcifull (1982). It also reacted specifically with CMV

antiserum in the undecorated immunosorbent electron microscopy

test. This virus was mechanically transmitted to cucumber,

tomato, N. benthamiana, N. x edwardsonii, N. glutinosa, N.

tabacum Xanthi nc., cowpea, and V. radiata. It caused systemic

infection in the first six hosts and brown-colored lesions in

the inoculated leaves of the last two hosts.

Bean Yellow Mosaic Virus

Cytoplasmic inclusions (Fig. 19), typical of potyviruses

and stable in 5% Triton X-100, were not frequently found in

epidermal and mesophyll tissues of Masdevallia stained in 0/G.

These inclusions, which are proteinaceous and do not contain

nucleic acid, failed to stain with Azure A, as would be

expected (Christie and Edwardson, 1977). Thin sections

revealed subdivision II cylindrical inclusions (Fig. 21) as

described previously for this virus (Edwardson, 1974). Dense

bodies such as those described for bean yellow mosaic virus

(Zettler and Abo El-Nil, 1977) were rarely found in these

sections. Cytoplasmic inclusions were readily detected in the

epidermal strips of N. benthamiana, 'Alaska' and 'Ranger' peas

(Fig. 20). In thin sections of these tissues, numerous dense

bodies and cylindrical inclusions (Fig. 22), which are

characteristic ofbean yellow mosaic virus (Edwardson, 1974),

were seen. 'Little Marvel' pea plants did not become infected.

This is consistent with the reported insusceptibility of this

cultivar to BYMV (Zettler and Abo El-Nil, 1977).







-23-


191


Figure 19.



Figure 20.


Light micrograph of large group of
cylindrical inclusions (CI) induced by BYMV
in cytoplasm of a Masdevallia leaf cell
stained with O/G. N: Nucleus. Bar = 5 .m.
Light micrograph of large group of
cylindrical inclusions (CI) induced by BYMV
in a pea leaf cell stained with O/G. N:
Nucleus. Bar = 5,um.





-24-


c ^v:^ef


Figure 21.



Figure 22.


Electron micrograph of BYMV-infected
Masdevallia leaf cell showing portions
of subdivision II cylindrical inclusions.
L: Laminated aggregates, P: Pinwheel
arms, M: Mitochondrion. Bar = 340 nm.
Electron micrograph of BYMV-infected pea
leaf cell showing subdivision II
cylindrical inclusions and dense bodies
(D). L: Laminated aggregates, P:
Pinwheel arms, V: Virus particles.
Bar = 710 nm.






-25-


Uncharacterized Bacilliform Viruses

These viruses caused striking symptoms in their hosts.

Chlorotic and necrotic fleck lesions occurred on Cymbidium

leaves (Fig. 23), while necrotic and ringspot lesions occurred

on Brassia leaves (Fig. 24). Nuclear, but not cytoplasmic

inclusions were observed in mesophyll and phloem tissues of

plants infected with bacilliform viruses. Those in Brassia

were relatively large and solitary (Fig. 25b), whereas those

found in the Cymbidium specimens were smaller, and usually

there was more than one inclusion within each nucleus (Fig.

26). The O/G stain combination readily stained these

inclusions (Fig. 27). With Azure A, however, penetration was

facilitated considerably by pretreatment of tissues in 1%

Triton X-100 for 10 minutes and heating at 60 0C for 10 minutes

during staining. Without these treatments, the nuclear

inclusions did not stain well (Fig. 25a). Thin sections for

both isolates revealed rhabdovirus particles. These nuclear

inclusions are composed of virus particles and granular

materials (Fig. 28). Particles ca. 45 nm in diameter occurred

in aggregates within the nucleus and in the cytoplasm. Those

in the cytoplasm were sometimes arranged in a "spokewheel-like"

fashion, like those described previously for Dendrobium leaf

spot, orchid fleck, and Phalaenopsis chlorotic spot viruses

(Lesemann and Begtrup, 1971; Petzold, 1971; Lesemann and







-26-


Figure 23.


Figure 24.


Chlorotic fleck and necrotic lesions of
Cymbidium leaves associated with a
rhabdovirus infection.
Necrotic and ringspot symptoms of
Brassia leaves associated with a
rhabdovirus infection.





-27-


Figure 25.






Figure 26.


Light micrograph of nuclear inclusions
(arrows) induced by a rhabdovirus in Brassia
cell stained with Azure A. (a) Poorly
stained by conventional method. (b)
Inclusion well stained after treatment with
Triton X-100 and heating to 600C while
staining. No: Nucleolus. Bar = 5 Am.
Light micrograph of nuclear inclusions
(arrows) induced by a rhabdovirus in a
Cymbidium leaf cell stained with Azure A.
Bar = 5 Am.











0


'C


N

4
a
*

I

Im


Figure 27.

Figure 28.


Light micrograph of nuclear inclusions
(arrows) induced by a rhabdovirus in Brassia
leaf cells stained with O/G. Bar = 10 ,m.
Electron micrograph of a portion of a nuclear
inclusion consisting of virus particles and
granular materials in a rhabdovirus-infected
Brassia leaf cell. V: Virus particles. Bar
= 460 nm. Inset showing virus aggregations.
Bar = 460 nm.


-28-


27






-29-


Doraiswamy, 1975; Chang et al., 1976). Dispersed virus

particles and spokewheel-like virus aggregations were not

detected by light microscopy.

Discussion



Infected orchid tissues examined by light microscopy

contained inclusions characteristic of tobamo-, potex-, cucumo-,

rhabdo-, and potyviruses (Table I). The diagnoses based on

inclusions were confirmed by electron microscopy and, for ORSV,

CyMV, and CMV, by fluorescence microscopy using TRITC-

conjugated antisera and immunodiffusion tests. Thus, like

other plant viruses (Christie and Edwardson, 1977), orchid

viruses can be reliably detected by light microscopy. While

other techniques (i.e., serology, bioassay, electron microscopy)

have been used previously for diagnosing orchid viruses

(Lawson and Ali, 1975; Lawson and Horst, 1984; Lawson and

Brannigan, in press; Corbett, 1974; Wisler and Zettler, in press),

this is the first systematic study using light microscopy for

this purpose. Potential difficulties posed by the exceptional

thickness of most orchid leaves in processing samples can be

readily overcome by using sandpaper to abrade the leaf surface

or by sectioning leaves. The sandpaper technique also proved

useful for working with geminiviruses of legumes (Ko and

Christie, unpublished). In some instances, it was necessary to

apply mild heating and/or Triton X-100 to assure stain

penetration, but neither of these modifications induced







-30-


TABLE I. SUMMARY OF STAINING PROCEDURES AND
INCLUSION TYPES OF ORCHID VIRUSES


VIRUS STAINING TYPE OF INCLUSION
PROCEDURE
CYTOPLASMIC NUCLEAR


ORSV O/G, Azure A Paracrystal
Stacked plate

CyMV Azure A Banded, crescent-
and tear-shaped
inclusion

CMV Azure A Crystal

BYMV O/G Cylindrical
inclusion

Rhabdovirus Azure A, Nuclear
O/G inclusion






-31-


significant problems in processing tissues. Glutaraldehyde

fixation is necessary to preserve the configuration and banding

of the inclusions in CyMV-infected tissues.

Viruses infecting orchids other than those included in

this study have been described, such as Cymbidium ringspot, a

tombusvirus (Hollings and Stone, 1977; Hollings et al., 1977;

Matthews, 1982). Distinctive inclusions induced by this virus

have been described (Russo and Martelli, 1981; Martelli and

Russo, 1981). Light microscopy should also be useful in

detecting this virus as well. Dendrobium vein necrosis virus,

whose properties are similar to those of closteroviruses,

should form fibrous and banded-body inclusions in phloem tissue

(Lesemann, 1977; Matthews, 1982). These types of inclusion can

be detected by light microscopy (Christie and Edwardson,

1977).

The specific identities of the rhabdoviruses infecting

Brassia and Cymbidium described herein have not been

established. The particle dimensions and spokewheel-like

configurations are similar to those described for the third

subgroup of plant-infecting rhabdoviruses (Matthews, 1982), but

their relationship to one another and to the viruses examined

in this study remain to be determined. The cylindrical

inclusions of bean yellow mosaic virus noted in this study

conform to those belonging to "subdivision II" (Edwardson,

1974). However, BYMV cylindrical inclusions and dense bodies

occurred rarely in Masdevallia tissues, indicating that they






-32-

could be undetected in studies confined to thin sections. This

may be attributed to relatively low virus titers in the orchid

host.

The use of light microscopy has special advantages for

diagnosing orchid viruses. Many orchid viruses, such as the

rhabdoviruses, are poorly characterized, and using alternative

techniques for their diagnosis is currently either impractical

or unreliable. Light microscopy can be used to detect multiple

infections. For example, a Phalaenopsis, which was received

for diagnosis, was determined to be triply infected with ORSV,

CMV, and a potyvirus. The presence of one virus does not

interfere with other viruses with regard to types of inclusions

induced. These observations have also been noted for multiple

infections of other crops (Edwardson and Christie, 1979; Russo

and Martelli, 1973).

Light microscopy proved to be a reliable method for diagnosis

of orchid viruses when compared with electron microscopy of

negatively stained exudates (Lawson and Brannigan, in press) and

immunodiffusion techniques (Wisler et al., 1982). Fifteen of 30

samples (Appendix III) gave the same results with all three

techniques. Nine of these were infected with CyMV, ORSV or both

viruses; the other six were negative. Infections of either a

rhabdovirus or a potyvirus were detected by light microscopy alone

in 11 of these samples. In 2 samples, ORSV was detected by

serology, but not light microscopy. Additional tests indicated

this discrepancy was probably due to a sampling error.










CHAPTER 3
CONFIRMATORY STUDIES OF
LIGHT MICROSCOPIC OBSERVATIONS


Introduction



Light microscopy has proved to be very useful in 1)

viral diagnosis and classification, 2) the selection of

tissues for ultrastructural studies, 3) monitoring host

tissues for virus infections, and 4) monitoring viral

inclusion purification (Christie and Edwardson, 1977;

Hiebert et al., 1984). The large field of view, the

chemically selective stains, and the speed and ease of

tissue preparation and examination are some of the

advantages of light microscopy over electron microscopy in

studying viruses. In the second chapter, the use of light

microscopy for the diagnosis of orchid viruses has been

described. However, the resolution of light microscopy is

limited. In recent years, the demands for comparative

light and electron microscopic studies of individual cells

or specific tissue structures has led to the development

of various procedures which permit examination of the same

tissue area by both types of instrument (Rossi et al.,

1972; Nilsen et al., 1982; Perrie and Webb, 1982). In

this study, modified electron microscopic methods are used

to reveal the fine structure of viral inclusions whose

-33-






-34-

detection by light microscopy were described in the first

chapter. In addition, immunofluorescence microscopy and

immunogold labelling were used in revealing the specific

protein composition of viral inclusions.



Materials and Methods

Leaf Dips

For CyMV and ORSV, vanadyl molybdate-phosphotungstate

(VaMo) was used for negative staining (Boothroyd and

Israel, 1980). The VaMo staining solution consisted of 1

part vanadyl molybdate (a mixture of 1 part of 1% vanadyl

sulphate and 4 parts of 1% ammonium heptamolybdate), 3

parts of 2% sodium phosphotungstate and 4 parts of 0.025%

bacitracin. One to 3 cuts were made into virus-infected

tissue (ca. 3 x 4 mm) in the freshly prepared staining

solution. A drop of this suspension consisting of

diffusing plant sap and staining solution was transferred

to a grid such that the droplet covered about one-third of

the grid. After 30 seconds the excess liquid was removed

using a piece of filter paper. The grid was then examined

with an electron microscope. The droplet edge was located

on the grid at low magnification and then enlarged to

locate virus particles accumulated there. The

phosphotungstic acid (PTA) staining was processed in the

same manner as VaMo staining.






-35-


Thin Sectioning

After staining with either the O/G combination or

Azure A (see Chapter 2), ORSV-infected tissues containing

inclusions were rinsed with 50% ethanol to remove Euparal

and then fixed with 5% glutaraldehyde and postfixed with

1-2% osmium tetroxide. Tissues were then dehydrated with

acidified 2,2-dimethoxypropane as described by Muller and

Jacks (1975), and embedded in Spurr's low-viscosity medium

(Spurr, 1969). Sections were made with either a glass or

a diamond knife and stained with potassium permanganate,

uranyl acetate, and lead citrate (Ko and Chen, 1982).

The O/G and Azure A stains were used to select tissues

infected with ORSV, CyMV, CMV, BYMV and rhabdoviruses to

be processed for embedding. Semithin (0.5-1.0 um)

sections were cut with a glass knife mounted on a Sorvall

Porter-Blum MT2-B ultramicrotome. Sections were then

placed on a glass microscope slide and stained with 1%

toluidine blue (w/v dissolved in 1% sodium borate

solution) for 10-30 seconds on a hot plate at 55-60 C.

The semithin sections were then examined by light

microscopy. When inclusion-containing areas were located,

ultrathin sections were made from the same block still

mounted on the microtome. Resulting ultrathin sections

were stained in a conventional manner as described above

and examined with an electron microscope.







-36-

Immunofluorescence Microscopy

Non-fixed ORSV-infected Cattleya, CMV-infected

Phalaenopsis and fixed (5% glutaraldehyde) CyMV-infected

Cymbidium tissues were used. The CMV antiserum used was

obtained from D. E. Purcifull (Kuwite and Purcifull, 1982)

and the antiserum against ORSV and CyMV from G. C. Wisler

(Wisler et al., 1982). The immunoglobulins (IgG) of CMV,

CyMV and ORSV antisera were purified by the use of protein

A-Sepharose CL-4B (Pharmacia Fine Chemicals, Sweden) as

described by Miller and Stone (1978). IgG was conjugated

with TRITC by dialysis (Hiebert et al., 1984). Healthy

tissue extracts were made by triturating tissues with 10

volumes (w/v) of 20 mM sodium phosphate buffered saline at

pH 7.4 (PBS). The extracts were then filtered using

Whatman No. 1 filter paper. The staining solution was

made by mixing 27 Ail TRITC-conjugated antiserum, 27 ul

healthy tissue extract, and 6 ul dimethylsulphoxide (DMSO)

in saline (Herbert et al., 1982). This solution was

incubated for 30 minutes before tissue staining. Tissue

sections prepared as described in Chapter 2 were floated on

the above solution for another 30 minutes. After incubation,

tissue sections were blotted with filter paper, washed

three times with 20 mM PBS and incubated for another 15

minutes on top of a large drop (about 200 ul) of PBS. All

the procedures were carried out in a moist chamber at room

temperature (ca. 25 C). Tissue sections were picked up






-37-

with the aid of a wooden applicator stick, blotted with

filter paper and transferred to an aqueous, non-

fluorescing medium (Aqua-mount, Lerner Laboratories, New

Haven, Connecticut 06513) placed upon a glass microscope

slide. A cover slip was then placed over the mounting

medium and the tissues were observed with a Nikon (Nippon

Kogaku K. K., Tokyo, Japan) Fluophot microscope with epi-

illumination capabilities. The interference excitation

filter and barrier filter were used. Observations were

recorded with an automatic camera using Kodak Ektachrome

400 film (Eastman Kodak Co., Rochester, New York). TRITC-

conjugated normal serum and heterologous antisera were

used for staining infected tissues as controls. Healthy

tissues were also studied as controls.



Protein A-Gold Labelling

A solution of protein A-gold (containing colloidal

gold particles 15 nm in diameter) was passed through a 0.2 um

millipore filter before use. The tissues studied were as

follows: CMV-infected Phalaenopsis, BYMV-infected pea,

ORSV-infected Cattleya, CyMV-infected Cymbidium and

healthy Cattleya and Cymbidium. These tissues were fixed

with 5% glutaraldehyde and dehydrated through an ethanol

series (25, 50, 75, 95, 100%), and embedded in LR White

resin, medium grade (Polaron Equipment Limited,

Hertfordshire, England). Polymerization of LR White resin






-38-

was done at 55-60 0C for 12 hours. Ultrathin sections were

placed upon Formvar carbon-coated copper grids and exposed

to 10% ovalbumin solution for 10 minutes. The excess

liquid was then blotted from the grids with filter paper.

Following this, grids were floated on a drop of PBS-

diluted antiserum (1/500-1/2000) for 30 minutes. Grids were

then rinsed 3 times with 20 mM PBS buffer for 10 minutes

each and then incubated with protein A-gold for 30

minutes. Finally, grids were rinsed with PBS buffer for

20 minutes, followed by a 10 minute rinsing in distilled

water. The sections were poststained with 2% uranyl

acetate and 0.2% lead citrate and examined by electron

microscopy. Tissues incubated with normal serum were used

as controls for all samples studied.





Results



VaMo Negative Staining

The VaMo negative staining method was very useful for

detecting CyMV and ORSV particles in orchid leaf and

flower tissues. With this method, most virus particles

accumulate at the edge of the staining solution on the

grid and appear with good contrast and resolution

(Figs. 29, 30 and 32). With the conventional PTA negative

staining method, virus particles were not concentrated at






-39-

the edge of the staining solution (Fig. 31), nor anywhere

else on the grid.



Thin Sectioning

Tissues examined by electron microscopy could be

directly compared to and correlated with those seen by

light microscopy when either the O/G or toluidine blue

methods described in this study were used. ORSV

inclusions were readily located by light microscopy after

the tissues were stained with the O/G combination and

subsequently could be processed for embedding and thin

sectioning. After ultrathin sections were made, the same

inclusions observed by light microscopy were also seen by

electron microscopy. However, the fine structure of the

cells and the inclusions at the ultrastructural level was

poorly resolved (Fig. 34). When Azure A was used, those

fine structures were even more poorly resolved. More

satisfactory results were obtained by using the toluidine

blue method. Semithin sections (0.5-1.0ium) were made

with an ultramicrotome and stained with 1% toluidine blue.

Although this dye stains cell structures in a nonselective

manner, the viral inclusions can still be recognized by

their characteristic morphology and can thus be

differentiated from cell organelles in these semithin






-40-


29 -


30-


Figure 29. Electron micrograph shows the accumulation
of VaMo-stained ORSV particles at droplet
edge (arrows) in Cattleya leaf extracts.
Bar = 200 nm.
Figure 30. Electron micrograph shows the accumulation
of VaMo-stained CyMV particles at droplet
edge (arrows) in Cymbidium leaf extracts.
Bar = 200 nm.






-41-


32 -


Figure 31.



Figure 32.


Electron micrograph shows the presence of
PTA-stained few CyMV particles at droplet
edge (arrows) from the same tissue as in
Fig. 30. Bar = 200 nm.
Electron micrograph shows the accumulation
of VaMo-stained CyMV and ORSV particles at
droplet edge (arrows) in high
concentration in an extract from a
Cymbidium leaf infected with both viruses.
Bar = 200 nm.






-42-


d-pp
*1- ^ .^ ^^


Figure 33.




Figure 34.


Light micrograph of ORSV-infected cells
showing a stacked-plate inclusion (SI,
side view) in semithin section (0.5 Amm
thick). Section is stained with toluidine
blue. Bar = 5 Am.
Electron micrograph of ORSV stacked-
plate inclusion (side view) in Cattleya
leaf cells stained with O/G
combination. Bar = 600 nm.






-43-


sections (Figs. 33, 35, and 38). Sections of these same

tissue areas can be obtained for electron microscopy by

taking an ultrathin section from the same block from which

the toluidine blue section is obtained. The same

inclusions can thus be located (Fig. 36 and 39) in these

ultrathin sections by their intracellular location in

relation to cell organelles noted by light microscopy.

Using this method, ORSV, CyMV and CMV inclusions composed

of virus particle aggregations (Figs. 34 and 36) were

located, as were BYMV inclusions, which were comprised of

cylindrical inclusions. Nuclear inclusions containing

rhabdovirus particles and granular electron dense materials

(Fig. 39) were also located by this method.

Immunofluorescence Microscopy

As described in the chapter 2, CyMV inclusions were

preserved by glutaraldehyde fixation. In this

investigation, it appeared that glutaraldehyde fixation

did not change the properties of CyMV inclusion

antigenicity since they still reacted with virus specific

antiserum (Figs. 40 and 41). In the newly emerged CMV-

infected leaves or flowers of Phalaenopsis, CMV crystal

inclusions were easily observed, although the size of

crystals varied (Fig. 42). In ORSV-infected Cattleya

tissues, the paracrystal and stacked-plate inclusions

fluoresced (Figs. 43, 44, 45 and 46). These virus-

infected tissues did not fluoresce when they were






-44-


Figure 35.




Figure 36.


Light micrograph of CMV-infected cell
showing the angular crystal (C) stained
with toluidine blue in semithin section
(0.5 Am thick). N:Nucleus, No: Nucleolus.
Bar = 5 Am.
Electron micrograph of the same CMV
crystal shown in Fig. 35. Bar = 545 nm.
Inset showing a portion of the same
crystal in higher magnification. Bar =
120 nm.





-45-


Figure 37.


Figure 38.



Figure 39.


Light micrograph of nuclear inclusion (arrow)
induced by a rhabdovirus in Brassia leaf
cell stained with O/G. N:Nucleus. Bar = 5g m.
Light micrograph of nuclear inclusion (arrow)
induced by a rhabdovirus in semithin section
(0.5 aum thick) of Brassia leaf cell.
N:Nucleus. Bar = 5 um.
a) Electron micrograph of nuclear inclusion of
Brassia rhabdovirus. V:Virions. Rectangle
shows the location of nuclear inclusion.
Bar = 550 nm. b) Electron micrograph of
spokewheel aggregation of virus particles.
Bar = 225 nm.







-46-


40






















Figure 40.





Figure 41.


CyMV-infected Cymbidium leaf cells
photographed with epifluorescence
optics, showing the specific
fluorescence from virus aggregations
treated with TRITC-conjugated CyMV
antiserum. Bar = 5 Am.
The same field of Fig. 40 photographed
with visible light. Arrows show the
location of virus inclusion.
Bar = 5 Aim.







-47-


Figure 42. a) and b) CMV-infected Phalaenopsis
petal cells photographed with
epifluorescence optics, showing the
specific fluorescence from irregular
crystals treated with TRITC-conjugated
CMV antiserum. Bar = 5 ,um.







-48-


44 -.


Figure 43.





Figure 44.


ORSV-infected Cattleya leaf cells
photographed with epifluorescence
optics, showing the specific
fluorescence from paracrystal treated
with TRITC-conjugated ORSV antiserum.
Bar = 5 Aim.
The same field of Fig. 43 photographed
with visible light. P:Paracrystal.
Bar = 5 Am.






-49-


Figure 45.






Figure 46.


ORSV-infected Cattleya leaf cells
photographed with epifluorescence
optics, showing the specific
fluorescence from stacked-plate
inclusions treated with TRITC-
conjugated ORSV antiserum.
Bar = 5 Am.
The same field of Fig. 45 photographed
with visible light. SI:Stacked-plate
inclusion. Bar = 5 um.





-50-


incubated with TRITC-conjugated pre-immune serum or

heterologous antisera.

Protein A-Gold Labelling

Colloidal gold particles, 15 nm diameter, were

selectively labelled on CyMV, ORSV, CMV virions and BYMV

cylindrical inclusions in thin sections, which were

treated with homologous antisera and protein A-gold

complex, respectively (Figs. 47, 48, 49, 51, 53 and 55).

Very few gold particles were found on cell components not

containing virions or viral inclusions or on intercellular

spaces of infected tissue. Likewise, gold particles were

not found to be specifically reacted in tissue sections

treated with normal serum rather than homologous

antiserum, nor were they detected in sections exposed to

heterologous antisera (Figs. 50, 54 and 56). By this

method, virus-like or non-related virus particles are

easily distinguished, which is not possible by using

conventional thin sectioning techniques. In some orchid

cells, for example, some thread-like particles are easily

mistaken for CyMV virions, but these can be differentiated

because they do not show any specific gold labelling (Fig.

52).

Dilution of antisera affected the non-specific

labelling of gold particles. A high concentration of

antisera used for incubation resulted in more nonspecific

gold labellings (Fig. 47). For this reason, antisera

dilutions of 1/500 to 1/2000 were used in this study.





-51-


Figure 47.







Figure 48.


Electron micrograph of BYMV-infected pea leaf cell
showing cylindrical inclusions to which gold par-
ticles are attached. Arrows show the location of
nonspecific reaction where the cylindrical inclu-
sions are not present. Sections were incubated
with antiserum to BYMV cylindrical inclusions
(1/20 dilution) and labelled with protein A-gold.
Ch:Chloroplast, D:Dense body. Bar = 325 nm.
Same tissue and treatments as in Fig. 47, at an
antiserum dilution of 1/1000. Note that the gold
particles are more specifically associated with
the inclusion at this antiserum dilution than
shown in Fig. 47. D:Dense body.
N:Nucleus. Bar = 325 nm.






-52-


L^t1r
fek4 L<


Figure 49.





Figure 50.


Electron micrograph of pinwheels showing
gold particles attached to the arms in
higher magnification. Micrograph
represents the same tissue and treatments
as described in Fig. 48. D:Dense body.
Bar = 120 nm.
Same tissue and treatment as in Fig.
49, except that 1/500 normal serum
was used. Arrows showing the locations
of non-specific reaction. Bar = 240 nm.


^ ilk








-53-


4
1%. -. ., ,. TJ'


Figure 51.





Figure 52.


Electron micrograph of virus aggregations
in CyMV-infected Cymbidium leaf cell
showing the presence of gold particles on
their surface. Section is incubated with
CyMV antiserum (1/2000 dilution) and
labelled with protein A-gold. Bar = 260 nm.
Electron micrograph of virus-like
structures in the same antiserum-treated
section as in Fig. 51, but showing no
specific reaction. Same treatment as in
Fig. 51. Bar = 320 nm.









-54-


''- .- -

a o*
..*. .. "

I.^- I..
, .. .-. ;. a


@e roS
.* -* *:* *
A-.:" '.; ...


m
*..*.-.. : **; V.


^ L* A **S
k.14*^. .-..

9**
q73


Figure 53.







Figure 54.


Electron micrograph of virus aggregations
of ORSV-infected Cattleya leaf cell
showing the presence of gold particles on
their surface. Section is incubated with
ORSV antiserum (1/1000 dilution) and
labelled with protein A-gold. Bar = 300 nm.
Same area as in Fig. 53, except
incubation is with CyMV antiserum
(1/1000 dilution). Bar = 300 nm.


J 4
k_, ,






-55-


Figure 55.





Figure 56.


Electron micrograph of virus particles
distributed within the cytoplasm of CMV-
infected Phalaenopsis leaf cells. Section
was incubated with CMV antiserum (1/1000
dilution) and labelled with protein A-gold
Pd:Plasmodesma. CW:Cell wall. Bar = 350 nm.
Same tissue and treatments as in Fig. 55,
except that incubation was with normal serum
(1/500 dilution). Note the background of
nonspecific reaction (arrows). CW:Cell wall.
Bar = 350 nm.






-56-


Discussion

Negative staining using PTA has been used extensively

to detect plant viruses (Hitchborn and Hills, 1965; Doi et

al., 1969). However, large grid areas need to be scanned

(time-consuming) in checking for the presence of virus

particles. VaMo negative staining provides a much more

efficient way to detect the infections of CyMV and ORSV

and other viruses (Boothroyd and Israel, 1980). Using

this method, as little as 6 minutes from grid preparation

to examination may be all that is necessary for a

diagnosis. The resolution and contrast of the VaMo stain

compare favorably with those noted for PTA.

Processing tissues for electron microscopy is

extremely labor-intensive. Since virus-induced inclusions

are sometimes widely scattered in host tissues, success in

locating them by this conventional technique is often

limited. The procedure of thin sectioning described here

permits electron microscopy of materials selected by light

microscopy. For example, Russo et al. (1982) failed to

find crystalline inclusions in broad bean true mosaic

infected tissues by using electron microscopic methods

exclusively. However, and contrary to their conclusions,

such inclusions were later found when light microscopic

methods were employed (Ko, unpublished). One of the chief






-57-

advantages of light microscopy is the short time required

for tissue preparation and the extensive areas that can be

scanned. Also, many viruses induce characteristic

inclusions which can readily be recognized by this method

on the basis of morphology, staining reaction, and

location. However, the low resolution of light microscopy

is a major limiting factor in the interpretations of the

inclusions seen by this method. The O/G and toluidine

blue methods described in this study provide the solution

to this problem, since the same (O/G) or adjacent

(toluidine blue) sections can be examined by both light

and electron microscopy. The O/G was less satisfactory

for this purpose, since the fine structures of cells and

inclusions were adversely affected by its staining process.

Therefore, the toluidine blue method was used in this

study to reveal the fine structure of inclusions. In this

study, orchid infections caused by ORSV, CyMV, CMV, BYMV

and the Brassia rhabdovirus were readily studied by this

approach. Toluidine blue is not a selective dye,

however, it facilitates locating the inclusion-containing

areas for ultrathin sectioning.

Fluorescent antibodies have been used extensively in

studying animal virus infections. However, their use in

the study of plant viruses has been very limited. This is

partly because the removal of non-specific staining of

plant tissues by fluorescent conjugates is very difficult,






-58-

and standard absorption procedures are inadequate. In

some cases, the bright autofluorescence of lignified cell

walls of certain tissues interferes with observation and

photography of specific fluorescence in tissues. Yet

another problem in this procedure is that it is difficult

to obtain satisfactorily thin (15,um or below) sections of

plant tissues. In this study, non-specific staining was

eliminated by incubating TRITC-conjugated antisera with

healthy tissue extracts before staining infected tissues.

With healthy tissue extracts and conjugated antibody

prepared in advance, the procedure from diseased plant to

microscopic examination can be completed in less than 2

hours. This is much shorter than the time required for

the immunofluorescence procedures described by others

(Mumford and Thornley, 1977; Nishiguchi et al., 1980; Rao

et al., 1978).

Ferritin and colloidal gold labelled antibodies are

the most common probes used for localization at the

electron microscopic level (Gildow, 1982; Martelli and

Russo, 1984; Baker et al., 1985). Labelling of ultrathin

sections with colloidal-gold conjugated antibodies is

generally accomplished with less nonspecific binding than

when ferritin is used (De Mey, 1984; Baker et al., 1985).

In animal virology, several studies have been performed

using the protein A-gold approach in order to trace the

place of synthesis, the post-translational glycosylation,






-59-

and the pathway taken by specific viral proteins in

infected cells.(Green et al., 1981; Griffiths et al.,

1982, 1983). It has also been used to reveal the

antigenic sites in different cellular compartments of

infected cells (Garzon et al., 1982; Roth, 1983, 1984).

These studies have demonstrated that the protein A-gold

technique represents a valuable approach for medical

diagnosis (Garzon et al., 1982). It has also been

introduced as an alternative to other techniques for the

ultrastructural localization of antigenic sites (Bendayan,

1984). In plant virology, immunogold labelling was first

applied to barley stripe mosaic virus by using gold-

immunoglobulin (IgG) complexes for Lowicryl-embedded

tissues (Lin and Langenberg, 1983). It was also used to

identify viral antigens in suspensions (Giunchedi and

Langenberg, 1982; Lin, 1984; Louro and Langenberg, 1984).

However, some complexity is involved in using Lowicryl and

the gold-IgG complex (Causton, 1984; Newman and Jasani,

1984, Louro and Lesemann, 1984). The problems associated

with the use of Lowicryl can be circumvented by using LR

White resin instead. The polymerization of LR White does

not require the very low temperatures and ultraviolet

light necessary for the polymerization of Lowicryl

.(Causton, 1984). This study demonstrates that the higher

temperature for polymerization of LR White does not

interfere with the specificity of immunogold labelling






-60-

for orchid viruses. The time period for the

polymerization of LR White is also much shorter than for

Lowicryl. In addition, this embedding medium possesses

another advantage of being very easy to use.

The protein A-gold technique is simple, sensitive,

reliable, and gives specific labelling of high reso-

lution. When compared with other immunocytochemical

techniques, it has repeatedly provided superior results

(Bendayan, 1984; Beesley et al., 1982). It also dis-

plays several advantages: antibodies raised in different

mammalian species can be used; the preparation of

colloidal gold and the formation and purification of the

protein A-gold complex are quite simple procedures; and

the use of gold particles as the electron-dense marker

allows for easy identification of the labelled structures,

and makes the technique suitable for double labellings

(Roth, 1983). The reliability of the technique and its

wide range of applications in electron microscopy have

been clearly demonstrated (Bendayan, 1984). It is usually

very difficult to identify the spherical virus particles

scattered within the cells in ultrathin sections. This

study has shown that with the immunogold technique,

spherical virus particles can be differentiated. In BYMV-

infected pea tissues, there are some thin plate inclusions

which may be misidentified as BYMV cylindrical inclusions.

The composition of thin plates cannot be recognized by







-61-

conventional electron microscopy of thin sections.

However, they can be differentiated from cylindrical

inclusions by gold labelling without any difficulty.

Thin sectioning, VaMo negative staining,

immunofluorescence microscopy, and protein A-gold labelling

described in this chapter confirmed that light microscopy

was an accurate and useful method for detecting orchid

viruses.











CHAPTER 4
CONCLUSIONS


Light microscopy using the Azure A and O/G stain was

shown to be a useful diagnostic tool for some orchid

viruses. For orchid viruses which cannot be detected by

conventional methods bioassayy, leaf dip electron

microscopy and/or serology), this technique is the only

practical alternative for this purpose. It also has the

advantage of being fast and inexpensive. It is an

important technique for developing countries which cannot

afford expensive facilities. In addition to diagnosis,

this technique is very useful in the selection of tissues

for ultrathin sectioning, for monitoring viral inclusion

purification, and the selection of antisera for

identifying the specific level of viruses (Christie and

Edwardson, 1977; Hiebert et al., 1984).

Orchids posed several problems which had to be

overcome to make the light microscope techniques

successful. For thick-leaved orchids, paradermal sections

could be conveniently made, whereas, for thin-leaved

orchids, sandpaper abrasion was preferable to expose cells

for staining. For instances where cell walls were

exceptionally thick, heating at 60 C was necessary to


-62-






-63-

facilitate stain penetration in ORSV-infected tissues.

ORSV, unlike other tobamoviruses, required a longer period

of heating to facilitate Azure A stain penetration of

virus aggregates. Rhabdoviruses infecting Brassia and

Cymbidium required heating at 60 C and treatment in Triton

X-100 to facilitate staining inclusions contained within

the nucleus. Presently, this is the only practical

technique to detect the rhabdovirus infection. Since

rhabdoviruses have been considered as important as ORSV

and CyMV (Lesemann and Marwitz, 1983), light microscopy

could be an important tool for their detection.

Interpretation of the light microscopy results was

facilitated by the development of the electron microscopic

techniques improvised for this research. The same

inclusions seen in tissues stained with O/G or toluidine

blue for light microscopic examination were also observed

by electron microscopy. The viral nature of inclusions

seen by light microscopy was also confirmed by in situ

examination of tissues directly labelled with TRITC-

conjugated antisera or indirectly using protein A-gold.

Nonspecific antibody binding was eliminated in

fluorescence microscopy by pre-incubating TRITC-conjugated

antisera with healthy tissue extracts.

Immunogold labelling has been used to identify the

viruses in ultrathin sections (Lin and Langenberg, 1983)

and leaf dips (Louro and Lesemann, 1984; Lin, 1984). In






-64-

addition to providing confirmatory data regarding the

viral nature of inclusions, this procedure is potentially

valuable for the in situ examination of viruses occurring

in low titers, and distinguishing viral from non-viral

intracellular structures. As shown in this study, the

procedure proved useful for capsid and nonstructural

proteins.

LR White resin was suitable for embedding tissues

infected with plant viruses. The use of this resin

facilitated immunogold labelling studies considerably

because it is much less labor intensive than the

previously used resins, such as epoxy resin or even

Lowicryl resin.

Another labor-saving improvisation developed in

this study was a multi-grid staining apparatus involving

simple equipment available in all reasonably equipped

scientific laboratories. The device could house at

least 10 grids simultaneously for staining and

processing for electron microscopy without individual

handling of the grids.

VaMo was an effective negative stain for orchid

viruses in leaf dip diagnosis. This stain promoted the

congregation of virions to the edge of the droplet on the

grid. Therefore, only the droplet edge needed to be

scanned in the electron microscope. The stain contrast

and resolution of VaMo compared favorably with that of







-65-

PTA, and its use is therefore to increase the accuracy of

orchid virus diagnosis, while reducing the time needed for

scanning at the same time.

A labor-saving procedure useful for ORSV and CyMV

bioassay to indicator plants was also developed. The

technique obviates the need for using a mortar and pestle

for trituration and thereby is much more convenient to the

specific indexing needs for orchid growers, who must index

numerous samples.














APPENDIX I
A SIMPLIFIED BIOASSAY TECHNIQUE FOR
CYMBIDIUM MOSAIC AND ODONTOGLOSSUM RINGSPOT VIRUSES



Although there are a number of advanced techniques

available to detect orchid viruses, bioassay still remains a

popular, inexpensive, and reasonably reliable technique for

detecting Odontoglossum ringspot (ORSV) and Cymbidium mosaic

(CyMV) infections (Lawson and Ali, 1975; Lawson and Brannigan,

in press; Wisler and Zettler, in press; Zettler et al., 1984).

Gomphrena globosa.is the laboratory indicator plant for ORSV

while Cassia occidentalis is used for CyMV.

In the normal bioassay method, clean mortar and pestles

are used to extract juice from the orchid leaf sample, often in

the presence of a buffer solution (Lawson and Ali, 1975; Lawson

and Brannigan, in press). This juice is then rubbed onto the

leaves of indicator plants, which have previously been dusted

lightly with a fine abrasive (usually 300-600 mesh

Carborundum). The abrasive produces microscopic wounds which

facilitate entry of the virus particles into the inoculated

leaves. To avoid carry-over of inoculum, the mortar and pestle

must be thoroughly cleaned (preferably sterilized) between

samples.


-66-







-67-


Recent studies have shown that the time-consuming step of

using a mortar and pestle can readily be circumvented by using

a razor blade incision to obtain inoculum. The new technique

is relatively easy. A razor blade is used to slice the leaf

sample just below and parallel to the leaf surface, thereby
2
exposing wounds about 3.0 cm Immediately after cutting, the

wounded orchid leaf surfaces can be gently rubbed by hand onto

the leaves of indicator plants. If the wounded surfaces are

dry, they can be misted slightly with water before inoculation.

As in juice inoculations, the leaves of indicator plants should

be rubbed firmly but gently to prevent damage, which could

interfere with test results.

Most assayists rub the two cotyledonary leaves (or the

first set of true leaves) of young Cassia seedlings, whereas

for Gomphrena, the fifth or sixth pairs of true leaves are

inoculated. A fresh cut of the orchid leaf sample should be

made for each Cassia cotyledon or Gomphrena leaf rubbed to

assure an abundant supply of inoculum. Before proceeding to

the next orchid sample, a fresh razor blade or an old one that

has been sterilized should be used. Sterilization can be done

conveniently by dipping the razors into 70% ethanol and

flaming them. After inoculations, excess orchid leaf juice and

Carborundum can be gently rinsed from the inoculated leaves

with water. As might be expected, symptoms observed on

indicator plants inoculated by the razor blade method are






-68-

identical to those resulting from the mortar and pestle method,

described previously (Lawson and Ali, 1975; Lawson and

Brannigan, in press). Discrete dark brown localized lesions are

typical of CyMV infections on Cassia cotyledons, whereas the

lesions induced by ORSV on Gomphrena leaves are light tan in

color. The lesion number in different tests will vary

according to the virus concentration in the orchid sample, the

condition of the indicator plants, and the skill of the

inoculator. Results can be recorded about one week after

inoculation if greenhouse temperatures are between 80-90 F. At

lower temperatures, slightly longer times may be required.

Both the razor blade and the mortar and pestle techniques

are reasonably reliable methods of virus transmission. In a

comparative test involving 16 Cassia and 6 Gomphrena seedlings

inoculated by each method, an average of 12 lesions/Cassia

cotyledon and 74 lesions/Gomphrena leaf were recorded when

razor blades were used in comparison to 5 lesions/Cassia

cotyledon and 43 lesions/Gomphrena leaf for the other method.

The razor blade technique is recommended as a convenient

assay method for ORSV and CyMV, especially when large numbers

of orchid samples must be indexed. However, such

recommendations cannot be made for some of the other viruses

known to infect orchids. This is particularly true of those

viruses which may occur in low concentrations in their orchid

hosts or cannot be transmitted by rub-inoculation (e.g., the

Cypripedium filamentous and Masdevallia isometric viruses)






-69-

(Lawson and Brannigan, in press). Orchidists inexperienced at

rub-inoculation should include a few noninoculated Cassia and

Gomphrena seedlings in their controls. Sometimes other

factors, such as pollution damage, can induce lesions on Cassia

or Gomphrena plants which can be confused with those caused by

virus (Lawson and Brannigan, in press). Inoculating some indi-

cator plants with an orchid sample known to be free of virus is

recommended so that possible inoculation damage can be

distinguished from viral lesions.












APPENDIX II
AN EFFICIENT PROCEDURE FOR STAINING LARGE NUMBERS
OF ELECTRON MICROSCOPE GRIDS



This paper describes a simple tubular device which can be

used to stain large numbers of electron microscope grids

simultaneously. Staining and rinsing of ultrathin sections

mounted on individual grids are time-consuming and tedious if

grids are handled individually according to the usual

techniques recommended for electron microscopy (Hayat, 1970).

In addition, the repeated use of forceps may damage the support

films and grids. Therefore, several attempts have been made

for simultaneous staining of many grids (Chen, 1973; Fisher,

1972; Godkin, 1977; Gorycki, 1978; Hiraoka, 1972; Robertson and

Roberts, 1971). However, most of these devices are either

difficult to construct or use.

These studies revealed that any polyethylene tubing with an

exterior diameter of 3 mm can be used. Tubes such as those

found in ordinary commercial ball-point pens (with an interior

diameter of 1.5 mm) were most satisfactory for this purpose.

All it requires is that a few holes must be cut into the tubing

so that grids can be inserted and anchored while being stained

and rinsed (Fig. 57). The diamond-shaped and circular holes

are made using a needle-point scalpel. The tubes should be


-70-






-71-

thoroughly cleaned before inserting the grids. The

polyethylene tube is then placed into a glass tube (3.5 mm

interior diameter).

To retain their proper orientation, the grids are

carefully inserted, one at a time, into the polyethylene tube

as it is being placed into the glass tube. When in place, and

prior to staining, grids are moistened by rinsing them 5-10

seconds in deionized water. Rinsing can be accomplished by

using an intravenous set (e.g., IV Set, McGaw Laboratories,

Inc., Sabana Grande, Puerto Rico 00747) to supply a jet of

water sufficiently fine to enter the glass and polyethylene

tubes (Fig. 58). After rinsing, excess water can be eliminated

from the tubes by gently tapping the tubes on a solid surface

covered with filter paper or shaking them by hand. For

staining, the tubes are attached to a 2 ml pipette pump (pi.

pump 2000, Glasfirn Giessen, D 6300 Giessen, West Germany)

(Fig. 59). As fluids are drawn up into the glass tube, care

should be taken to avoid air bubbles possibly present on the

grids.

These staining and rinsing devices have been used

successfully for over one year. The tubes can be reused, but

must be cleaned thoroughly between operations. Labs using this

technique are able to handle many grids simultaneously. In

our experience, the grid holder provides a simple, efficient

way for staining thin sections and has reduced mechanical

damage and contamination of thin sections.





-72-


I J 4 ,,* *, ]
I I
10mm

57












^-~~.H9-^kl


Figure 57.




Figure 58.

Figure 59.


Diagram of polyethylene tube showing the side
for insertion of grids through diamond-
shaped openings (left). The grids are
anchored in the small round holes directly
beneath the diamond-shaped holes (right -
reverse side of tube).
Intravenous set used to rinse grids con-
tained within polyethylene tube.
Two ml pipette pump used to apply stains
to grids.











APPENDIX III
COMPARATIVE DIAGNOSES OF ORCHID VIRUSES
BY LIGHT MICROSCOPY, ELECTRON MICROSCOPY, AND SEROLOGY


Diagnostic Methods*

Orchid Foliar
Samples Symptoms Light Electron Serology
(Genus) Microscopy Microscopy


Phaius
Cattleya
Cattleya
Cattleya
Odontocidium
Odontoglossum
Odontoglossum
Odontoglossumn
Odontoglossum
Wilsonara
Phalaenopsis
Phalaenopsis

Phalaenopsis

Phalaenopsis
Odontocidium

Odontocidium
Oncidium

Oncidium
Odontoglossum
Cattleya
Phalaenopsis
Vanilla
Cattleya
Vanilla
Cattleya
Phalaenopsis
Cattleya
Encyclia
Phalaenopsis
Vanda


Mosaic
Necrosis
NE**
NE
Mottle
Mottle
Mottle
Mottle
Mottle
Mottle
Chlorosis
Chlorosis
Necrosis
Chlorosis
Necrosis
NE
Mottle &
Necrosis
Mottle
Mottle &
Necrosis
Mottle
Mottle
NE
NE
Mottle
NE
NE
NE
NE
Necrosis
Mottle
NE
NE


CyMV, ORSV
CyMV


Rhabdovirus
Rhabdovirus
Rhabdovirus
Rhabdovirus


CyMV
Rhabdovirus

CyMV
-






CyMV
Rhabdovirus

ORSV
CyMV,
Potyvirus

Rhabdovirus
CyMV
CyMV, ORSV
Potyvirus
ORSV
ORSV
CyMV
CyMV
CyMV, ORSV
Potyvirus
Potyvirus


CyMV, ORSV
CyMV


CyMV


CyMV

CyMV


ORSV
CyMV



CyMV
CyMV, ORSV


ORSV
CyMV
CyMV
CyMV, ORSV


CyMV, ORSV
CyMV, ORSV


CyMV


CyMV

CyMV


ORSV
CyMV



CyMV
CyMV

ORSV
ORSV
CyMV
CyMV, ORSV
CyMV, ORSV
-


-73-






-74-

*For light microscopy, Azure A and O/G stains were used (see
chapter 2). For electron microscopy, 2% potassium
phosphotungstate (pH 6.9, containing 0.025% bacitracin)
was used for negative stain. For serology, the SDS immuno-
diffusion test as described by Wisler et al. (1982) was
used. The CMV, ORSV, and CyMV antisera were as described
in chapter 3. The BYMV capsid and nuclear inclusion
antisera used were obtained from C.-A. Chang.

**NE: Non-evident symptom.












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BIOGRAPHICAL SKETCH

Nan-Jing Ko was born in Changhwa, Taiwan, on November

5, 1948. He received his primary and secondary education

at the Sanchuan Elementary and Changhwa Junior and Senior

High Schools, Changhwa. In 1971 he obtained a Bachelor of

Science degree in plant pathology from the Plant Pathology

Department, National Chung Hsing University at Taichung.

After two years' military service as a Second Lieutenant,

he returned to the Plant Pathology Department of the

National Chung Hsing University where he earned his Master

of Science degree in 1977. He received a one-year

fellowship from the National Science Council of the

Republic of China from August 1979 to July 1980, of which

he spent seven months at the Ohio State University and

five months at the University of Florida. During that

time, he worked with Dr. 0. E. Bradfute and Dr. J. R.

Edwardson, respectively. In 1982 he received a two-year

fellowship from the National Science Council of the Republic

of China. In January 1983 he entered the University of Florida

to continue work towards a Doctor of Philosophy degree

in plant pathology. He is an associate professor in the

Plant Pathology Department of the National Chung Hsing

University. After graduation he will continue in his current

position at the National Chung Hsing University, Taiwan.


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I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in sco nd quality, as a
dissertation for the degree of Doctor of sophy.



Francis W. Zettler, Chairman
Professor of Plant Pathology

I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.



Jr n R. Edwardson, Cochairman
Professor of Agronomy

I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.



Ere'st Hieber
Professor of Plant Pathology

I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.



Dan E. Purcifull
Professor of P1 at Pathology

I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.



Henr C. Aldrich
Professor of Microbiology
and Cell Science







I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.



Thomas J,-Sheehan
Professor of Ornamental
Horticulture

This dissertation was submitted to the Graduate Faculty of the
College of Agriculture and to the Graduate School and was
accepted as partial fulfillment of the requirements for the
degree of Doctor of Philosophy.


May 1985


Dean, Q1lege of Ag* Ilture




Dean for Graduate Studies and
Research






































UNIVERSITY OF FLORIDA
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