. :N. V
‘J .I'fir'.

3.1

.};p
‘\

'V‘u
.. _

v

L " ill
1.11:“ ‘ .n" "I. ."-". -
'V. I II..(¢" .1? I. 1w' til!" "I
I I l' It'd" \‘..' _.
.I'V'I‘Vu'fl' 6‘:
I "-' "Cl‘n‘I IL, II 'JIF'...’ J,"
III." .I. | I'L' I

J

I . \

" ‘I ‘I'k‘fl:'\‘. ‘
1;. ,' '

.I,

u H“ ' "
U“ .‘m.

we MI. .,; .- I: .3
." ".3. .- -

 

JIM r JV».

 

i"; In

gum ‘ 3 1293 10809 4395

Inc

 

 

 

 

 

 

 

 

 

MIMI/III “mum IT I L

H
L-a— am- 'IIIfi i

e. O
a ‘ 2 '.T~‘.$’f.’_l’3' 6: 12mg)“

win.‘--‘§.'m~ -‘W‘Nw

. . .1
: t-u-e- wens}!- ’Jv;
'" uuu‘EU van-In“-

 

This is to certify that the

dissertation entitled

MECHANISMS OF AND BARRIERS TO SPIROPLASMA CITRI
INFECTION OF MACROSTELES FASCIFRONS (STAL)

presented by

Thomas Minster Mowry

has been accepted towards fulfillment
of the requirements for

Ph . D . degree in But omo logy

 

A

Major professor ’ .

Date November 21, 1985

 

MS U i: an Affirmative Action/Equal Opportunity Institution 0-12771

 

 

MSU

 

 

RETURNING MATERIALS:
Place in book drop to

 

 

 

 

LIBRARIES remove this checkout from
.—c_. your record. FINES will
be charged if book is
returned after the date
stamped below.
a . a.” -
:- .~I..J- ’
w’“ 3
0
no“ A

 

MECHANISMS OF AND BARRIERS TO SPIROPIASMA CITRI
INFECTION OF MACROSTELES FASCIFRONS (STAL)

by

Thomas Minster Mowry

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Entomology

1986

ABSTRACT

MECHANISMS OF AND BARRIERS TO SPIROPLASMA CITRI
INFECTION OF MACROSTELES FASCIFRONS (STAL)
by

Thomas Minster Mowry

Research was done to demonstrate that barriers to infection of

noncompetent Macrosteles fascifrons (Stal) by Spiroplasma citri exist

 

and to probe the infection mechanisms in competent leafhoppers. A
technique for sectioning fixed and frozen leathppers and preparation
of these sections for scanning electron microsc0py was developed. In-
ternal leathpper morphology was exceptionally well preserved and dis-
cernable. A colloidal gold-IgG conjugate that labelled the surface of
g. citri was used, along with ELISA, to assess the antigenic response
of the spirOplasma to glutaraldehyde fixation. The surface antigens of
§. citri were numerous and randomly distributed in the membrane.
Labelled antigens exhibited patching in unfixed cells, indicating that
they are mobile within the membrane. While ELISA showed a predictable
decline in overall antigenicity with increasing glutaraldehyde concen-
tration, cells labelled equally well at all fixative levels. Colloidal
gold-labelled cells were easily detected in the scanning electron mi-
crosc0pe in the backseatter mode. These data indicated that _S_. SEE}
could be detected within glutaraldehyde-fixed leathpper sections.
Twelve successive passages of the horseradish brittle root isolate

of §. citri, 24 through 35, were injected into adult leafhoppers. For

natural acquisition, adult leathppers were fed on §_. c_i_t_1_:i-infected
asters. After appropriate incubation periods, surviving leathppers
were caged individually on aster seedlings. Leafhoppers were removed
and frozen prior to spirOplasma isolation attempts from individual in-
sects. Phytotoxin activity of each passage was monitored. Membrane
preparations of passages 26 and 34 were analyzed with SDS-PAGE.

0f the injected leafhoppers, 3.6% transmitted g. citri while 50%
had acquired the pathogen. Of those naturally fed, 0.72 transmitted
and 5.4% acquired §, citri. 'These results indicate that both
mesenteronal and post-mesenteronal barriers to g. SEEEl infection and
transmission exist in g. fascifrons. The barriers involve different or
modified mechanisms for their penetration as seen by only one of the
naturally acquiring leathppers being able to transmit the pathogen.
None of the passages lost phytotoxin activity, indicating that in vitgg
passage probably reduces transmissibility rather than pathogenicity.
Membrane analysis revealed the loss of a protein from the membrane of
passage 34 relative to passage 26 which may be involved in binding of
the pathogen to host cell membranes prior to endocytosis and ultimate

completion of the transmission cycle.

DEDICATION

To my wife, Mary,
who has loved me with the love of Christ.
I pray He will bring her great reward,

because this result is unworthy of the sacrifice she has made.

ii

ACKNOWLEDGMENTS

I would.like to thank Dr. Mark Whalon, my Graduate.Advisor, for
persevering with and praying for me throughout the often tumultuous
days of this program. Neither of us yet realizes how important his
role has been, but his patience will be rewarded.

In addition, I thank Drs. Jim Bath, Ed Grafius, and Harold Miller
for their input into this research and encouragement to attain the
prize.

My good friends, Drs. Jim Miller and Karen Klomparens, deserve
special gratitude. Their support in both the good and desperate times
prevented some foolish decisions being made. If I can obtain a small

measure of their excellence and compassion, II will have been educated

indeed.
Finally, I would like to thank my daughters Teresa and Trista for
often doing without, yet fully supporting, me through six long years.

They are truly the glory of their father.

iii

TABLE OF CONTENTS

LIST OF TABLES . . . . . . . . .

LIST OF FIGURES. . . . . . . . .

GENERAL INTRODUCTION . . . . . .

Mycoplasmas as Plant Pathogens.

Characteristics of Spiroplasmas

Spiroplasma-Leafhopper Interactions

Literature Cited. . .

CHAPTER I:

Introduction. . . . . . . .
Materials and Methods . . .
Results . . . . . . . . . .
Discussion. . . . . . . . .
Literature Cited. . . . . .

CHAPTER II:

Introduction. . . . . . . .
Materials and Methods . . .
Spiroplasma Culture. .

Antiserum Production .

INVESTIGATION OF LEAFHOPPER INTERNAL
SCANNING ELECTRON MICROSCOPY

MORPHOLOGY VIA
OF FROZEN

EFFECTS OF GLUTARALDEHYDE FIXATION ON
OVERALL ANTIGENICITY OF

SECTIONS. .

SURFACE
SPIROPLASMA CITRI .

Preparation of Colloidal Gold-IgG Conjugate.

Fixation of Spiroplasma citri. . .

 

Colloidal 601d hbelling O O C O O I O O O O

Enzyme-linked Immunosorbent Assay (ELISA). .

iv

vi

vii

10
12
16
19

21

23
24
26
26
26
29
31
33

34

TABLE OF CONTENTS, continued

Results . . . . . . . . . . .
Discussion. . . . . . . . . .
Literature Cited. . . . . . .

CHAPTER III:

AND TRANSMISSION OF SPIROPLASMA CITRI. .

Introduction. . . . . . . . .

Materials and Methods . . . .

SpirOplasma and Leathpper Cultures. . . .

Passage and Injection of Spiroplasma citri

 

Acquisition of Spiroplasma citri from Plants

 

Isolation of SpirOplasmas.

Phytotoxin Assay . . . .
Membrane Preparation . .
Membrane ElectrOphoresis
Results . . . . . . . . . . .
Discussion. . . . . . . . . .
Literature Cited. . . . . . .

GENERAL CONCLUSION . . . . . . . .

EFFECTS OF IN VITRO PASSAGE ON THE PLASMA MEMBRANE

36
45

48

SO
51
54
54
54
55
57
57
58
59
61
67
7O
74

Table

Table

Table

Table

II-l.

III-1.

III-2.

III-3.

LIST OF TABLES

ELISA results for the horseradish brittle
root isolate of SpirOplasma citri fixed in
various concentrations of glutaraldehyde. .

Number of colony forming units (CFUs) of
the horseradish brittle root isolate of
Spiroplasma citri injected into Macrosteles
fascifrons. . . . . . . . . . . . . . . . .

 

Effects of in vitro passage on acquisition
and transmission of the horseradish brittle
root isolate of Spiroplasma citri by
injected Macrosteles fascifrons . . . . . .

 

Natural acquisition and transmission of the
horseradish brittle root isolate of
Spiroplasma citri by Macrosteles fascifrons

 

vi

43

56

62

63

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

II-l.

11-2.

11-3.

11-4.

11-5.

11-6 0

LIST OF FIGURES

Equipment necessary for sectioning fragile
specimens for scanning electron microscopy.

Processing of frozen sections for scanning
electron microscoPy . . . . . . . . . . . .

Frozen section of Scaphytopius acutus (Say)
processed through dehydration and critical
pOint dry1ng I O I O O O O O O O O O O O O O

 

Frozen section of Macrosteles fascifrons
(Stal) processed through lyophilization . .

 

pH isotherm for the gold colloid produced
by the sodium citrate method. . . . . . . .

IgG absorption isotherm for the gold
colloid produced by the sodium citrate
methOd O O O O O O O O O O O O O O O O I C O

Transmission.electron.micrographs of the
horseradish brittle root isolate of
Spiroplasma citri fixed in various
concentrations of glutaraldehyde and
labelled with a specific colloidal gold-IgG
conjugate . . . . . . . . . . . . . . . . .

 

Scanning electron micrographs of the
horseradish brittle root isolate of
§2i£22l2§22 sitzi fixed in 3-574

glutaraldehyde and labelled with a specific
colloidal gold-IgG conjugate. . . . . . . .

Transmission electron micrographs of the
horseradish brittle root isolate of
Spiroplasma citri unfixed and labelled with
a specific colloidal gold-IgG conjugate . .

Transmission electron micrographs of the
horseradish brittle root isolate of
§Ri£22l2§ee 9.12:1 fixed in 3-51
glutaraldehyde and pretreated with
nonspecific and specific IgG pmior to
labelling with a specific colloidal gold-

IgG conjugate . . . . . . . . . . . . . . .

vii

14

15

17

18

30

32

37

38

39

4O

LIST OF FIGURES, continued

Figure

Figure

Figure

Figure

Figure

11-7.

II-8.

II-9.

III-1.

III-2.

Transmission electron.micrographs of the
horseradish brittle root isolate of
§21£22l§§22 213.121 fixed in 3-574
glutaraldehyde prior to attempted labelling
with a goat anti-rabbit colloidal gold
conjugate (A) and unconjugated colloidal

gold(B). . . . . . . . . . . . . . . . . .

Scanning electron micrographs of sheep red
blood cells fixed in 2.0% glutaraldehyde
prior to attempted labelling with a
Spiroplasma citri-specific colloidal gold-
IgG conjugate . . . . . . . . . . . . . . .

 

Effects of glutaraldehyde fixation on the
enzyme-linked immunosorbent assay (ELISA)
for the horseradish brittle root isolate of
Spiroplasma citri . . . . . . . . . . . . .

 

Effects of in vitro passage on phytotoxin
production of the horseradish brittle root
isolate of Spiroplasma citri. . . . . . . .

 

 

SDS-PAGE analysis of purified membranes of
the horseradish brittle root isolate of
Spiroplasma citri from passages 26 and 34 .

 

viii

41

42

44

65

66

GENERAL INTRODUCTION

Mchplasmas as Plant Pathogens

The etiology of yellows-type plant diseases was for years con-
sidered to be viral (Whitcomb and Black 1982) until the historic report
of Doi et al. (1967) implicated mycoplasmas as possible causal agents.
Since then, two rather distinct groups of wall-free prokaryotic plant
pathogens have been defined: the pleomorphic mycoplasmalike organisms
(MLOs) and the helical Spiroplasmas. The MLOs remain an enigma, having
escaped in vitro isolation (Chen et al. 1982; Hayflick and Arai 1973)
and, therefore, the definitive proof that they are the causal agents of
the diseases with which they are associated. MLOs are, however, as-
sumed to cause such diseases as aster yellows, X-disease, clover phyl-
lody, and blueberry stunt (Nielson 1979).

Unlike the MLOs, nmny spirOplasmas affecting plants, arthrOpods,
and vertebrates have been isolated and maintained i_n m (Whitcomb
1980; 1981). Knowledge of spirOplasma existence is very recent, being
first recognized by Davis et al. (1972) in electron micrographs of corn
stunt-infected plant tissue. The first spiroplasma isolated in vitro
was a vertebrate pathogen, the suckling mouse cataract agent, but was
misidentified as a spirochete due to its morphology (Clark 1964). It
was the corn stunt spiroplasma (CSS) that was first isolated from plant
tissues, but this culture was neither maintained nor cloned (Chen and
Granados 1970). Subsequently, two separate groups isolated a spiro-
plasma from citrus stubborn-infected plant tissue that were maintained

in continuous culture (Fudl-Allah.et al. 1971; Saglio et‘al. 1971).

Markham et al. (1974) fulfilled Koch's postulates for this citrus spi-
roplasma, which was the first true mycoplasma proven to cause plant

disease. This organism has since been named Spiroplasma citri (Saglio

 

et al. 1973). It was again.two independent groups that.finally iso-
lated, amintained in continuous culture, and fulfilled Koch’s postu-
lates for the CSS (Chen and Liao 1975; Williamson and Whitcomb 1975).
To date, among the approximately 25 spirOplasmas available from the
American Type Culture Collection, only S. ElEEl and the CSS are phyto-

pathogenic.

Characteristics of SpirOplaslas

It is of considerable interest that the spiroplasmas represent a
unique group of microorganisms. They are prokaryotes exhibiting spiral
morphology and motility without the aid of a cell wall, as possessed by
spirochetes which demonstrate similar properties (Whitcomb 1981). Spi-
r0plasmas do possess cytoplasmic protein fibrils, ‘possibly related to
their morphology and motility, which are present in both helical and
nonhelical variants (Townsend and Archer 1983). In analyzing partially
purified membranes of these variants, Townsend et al. (1980) found that
a protein of molecular weight 39,000 was absent in the nonhelical
strain. These authors postulated that the missing membrane protein was
necessary for binding the cytoplasmic protein fibrils to the membrane
in order to complete the structure necessary for the spirOplasma’s he-
lical properties. Both variants of S. citri maintain phytOpathogenici-
ty (Townsend et al. 1977) and some have speculated that plant MLOs are
in fact noncultivable spiroplasmas (Davis 1974). However, it has been

shown that the spirOplasma-specific cytoplasmic protein fibrils are

3
absent in.MLOs infecting plants with several yellows diseases (Townsend
1983).

Spiroplasmas are intracellular parasites, as are the MLOs, which
distinguishes them from the animal myc0p1asmas (Myc0plasmataceae). The
genome size of S. SlEEl is about 109 daltons, approximately double that
of the Myc0plasmataceae (Maniloff 1972; Saglio et al. 1973). This ad-
ditional genetic material may be necessary to code for the proteins
that may be involved in the mechanisms of intracellular parasitism as
well as those that maintain morphology and motility. In any event,
spiroplasmas demonstrate classical myc0plasma characteristics including
the lack of a cell wall with no reversion to walled forms, no murein
cell wall precursors, ability to pass through 0.22 pm filters, forma-
tion of umbonate colonies on solid media, and complete resistance to
penicillin (Bové and Saillard 1979).

In addition to motility, spiroplasmas exhibit an active chemotaxis
toward and away from such substances as amino acids and certain vita-
mins (Daniels et al. 1980). The prOperties of motility and chemotaxis
may play'an.important part in the adaptation of spiroplasmas to the
diverse habitats of plant phloem and leathpper tissues (Whitcomb
1981). I have observed that symptoms of S. giggi and the aster yellows
MLO in aster always appear first in the new crown growth, regardless of
the original inoculation point. This indicates that these organisms
may migrate to the root system from which they are redistributed

throughout the plant.

SpiroplaslarLeafhopper Interactions
PhytOpathogenic spiroplasmas are pathogenic to some of their leaf-

hopper vectors (Whitcomb and Williamson 1975; 1979), suggesting that

these insects may not be important in the natural maintenance of the
disease (Whitcomb 1981). Survival is reduced with S. w infection
of Circulifer tenellus (Baker) (Liu et al. 1983b) and CSS infection of
Dalbulus elimatus (Ball) (Madden and Nault 1983). In contrast, the
survival of Dalbulus maidis (DeLong and Wolcott), a principle vector of
the CSS is unaffected by the pathogen (Madden and Nault 1983). This
phenomenon allows speculation that the CSS is a mutualist in S. w,

as is apparently the case with the aster yellows MLO and Macrosteles

 

fascifrons (Stal) (Whitcomb and Williamson 1979). No such mutualism
has been demonstrated with S. 2123.1. and any of its identified leafhop~
per vectors.

S. 91331 is circulative and propagative in its leathpper vector,
S. tenellus (Liu et al. 1983a; 1983b). This means that S. glt_rimust
pass through, and possibly multiply in, at least three tissue types:
gut epithelium, hemolymph, and salivary acini (Whitcomb 1981). Al-
though not yet experimentally demonstrated, this is probably true for
the CSS and its leafhopper vectors. The molecular mechanisms that
allow for spir0plasma infection of these tissues have not been eluci-
dated. Liu (1982) speculated that S. gig; might enter the hemocoel
from the gut lumen by passing through the endOplasmic reticulum, which
is continuous from the outside to the inside of the epithelium. His
own electron microscopic evidence, however, does not support this idea.
Electron micrographs have shown S. gig; inside gut epithelial cells
and salivary acini (Liu et al. 1983b), indicating that the microorgan-
ism had to cross the plasma membrane. Because spirOplasma transmission
depends upon the circulative and propagative nature of the pathogen in

its vector, it is obvious that anything that blocks these processes

constitutes a barrier to transmission. Multiplication of S. girl in
the hemolymph of leafhoppers incapable of transmitting the pathogen
(Whitcomb et al. 1973) indicates that membranes may be the principle
barriers, physically and/or chemically.

Among the possible mechanisms for the entrance of pathogens into
host cells is receptor-mediated endocytosis (RME). In this mechanism,
the pathogen has a ligand on its surface that is recognized by a recep-
tor on the host cell membrane which, when bound by the ligand, initi-
ates a series of events that results in the endocytosis of the pathogen
(Lonberg-Holm 1981). A number of viruses engage in RME to infect their
respective hosts (Lenard and Miller 1983).

The attachment of Mycoplasma pneumoniae to host cell membranes

 

involves the ligand-receptor complex (Kahane et al. 1982), but endocy-
tosis does not occur as these pathogens are extracellular parasites
(Clyde 1979). In light of this, it is not unreasonable to hypothesize
that RME may be the mechanism of cell penetration used by spirOplasmas
in their leafhopper hosts. Research was initiated in an attempt to

test this general hypothesis.

LITERATURE CITED

Bové, J.M. and C. Saillard. 1979. Cell biology of spiroplasmas. Pp.
83-153 in: The Mycoplasmas, Vol. 3. R.F. Whitcomb and J.G.
Tully, eds. Academic Press, New York.

Chen, T.A. and R.R. Granados. 1970. Plant-pathogenic mycoplasmalike
organism: Maintenance in vitro and transmission to Zea mayS L.
Science 167: 1633-1636.

Chen, T.A. and C.H. Liao. 1975. Corn stunt spirOplasma: Isolation,
cultivation, and proof of pathogenicity. Science 188: 1015-1017.

Chen, T.A., J.M. Wells, and C.H. Liao. 1982. Cultivation in vitro:
Spiroplasmas, plant mchplasmas, and other fastidious, walled
prokaryotes. Phytopathol. Prokaryotes 2: 417-446.

Clark, H.F. 1964. Suckling mouse cataract agent. J. Infect. Dis.
114: 476-487.

Clyde, W.A., Jr. 1979. Myc0plasma pneumoniae infections of man. Pp.
275-306 in: The Mycoplasmas, Vol. 2. .JJS. Tully and R.F.
Whitcomb, eds. Academic Press, New York. 509 pp.

Daniels, M.J., J.M Longland, and J. Gilbart. 1980. Aspects of
motility and chemotaxis in spirOplasmas. J. Gen. Microbiol. 118:
429-436.

Davis, R.E. 1974. New approaches to the diagnosis and control of
plant yellows diseases. Pp. 289-302 in: Proceedings of the Third
International Symposium on Virus Diseases of Ornamental Plants.
R.H. Lawson and M.K. Corbett, eds.

Davis, R.E., J.F. Worley, R.F. Whitcomb, T. Ishijima, and R.L. Steere.
1972. Helical filaments produced by a mycoplasma-like organism
associated with corn stunt disease. Science 176: 521-523.

Doi, Y., M. Teranaka, K. Yora, and H. Asuyama. 1967. Mycoplasma- or
PLT group-like microorganisms found in the phloem elements of
plants infected with mulberry dwarf, potato witches' broom, aster
yellows, or Paulownia witches' broom. Ann. Phytopathol. Soc. Jpn.
33: 259-266.

Fudl-Allah, A.E.-S., E.C. Calavan, and E.C.K. Igwegbe. 1971. Culture
of a myc0plasma-like organism associated with stubborn disease of
citrus. Phytopathology 61: 1321. (abstr.).

Hayflick, L. and S. Arai. 1973. Failure to isolate mchplasmas from
aster yellows diseased plants and leathppers. Ann. N.Y. Acad.
Sci. 225: 494-502.

Kahane, I., M. Banai, S. Razin, and J. Feldner. 1982. Attachment of
myc0plasmas to host cell membranes. Rev. Infect. Dis. 4: $185-
$192.

Lenard, J. and D.K. Miller. 1983. Entry'of enveloped viruses into
cells. Pp. 119-138 in: Receptor Mediated Endocytosis. P.
Cuatrecasas and T. Roth, eds. Chapman and Hall, New York. 304 pp.

Liu, H.-Y. 1982. The transmission, multiplication and electron
microscopic examination of Spiroplasma citri in its vector,
Circulifer tenellus. Ph.D. Thesis, Univ. Calif., Riverside. 101

PP-

 

 

Liu, H.-Y., D.J. Gumpf, G.N. Oldfield, and E.C.Calavan. 1983a.
Transmission of Spiroplasma citri by Circulifer tenellus.
Phytopathology 73: 582-585.

 

 

Liu, H.-Y., D.J. Gumpf, G.N. Oldfield, and E.C. Calavan. 1983b. The
relationship of Spiroplasma citri and Circulifer tenellus.
Phytopathology 73: 585-590.

 

 

Lonberg-Holm, K. 1981. Attachment.of animal viruses to cells: An
introduction. Pp. 1-20 in: Virus Receptors, Part 2: Animal
Viruses. K. Lonberg-Holm and L. Philipson, eds. Chapman and
Hall, New York. 217 pp.

Madden, L.V. and L.R. Nault. 1983. Differential pathogenicity of corn
stunting mollicutes to leafhopper vectors in Dalbulus and Baldulus
species. Phytopathology 73: 1608-1614.

Maniloff, J. 1972. Cytology of the mycoplasmas. Pp. 67-91 in:
Pathogenic Mycoplasmas: A Ciba Foundation Symposium. Associated
Scientific Publishers, Amsterdam. 404 pp.

Markham, P.G., R. Townsend, M. Bar-Joseph, M.J. Daniels, A. Plaskitt,
and B.M. Meddins. 1974. Spiroplasmas are the causal agents of
citrus little-leaf disease. Ann. Appl. Biol. 78: 49-57.

Nielson, M.W. 1979. Taxonomic relationships of leafhopper vectors of
plant pathogens. Pp. 3-27 in: Leathpper Vectors and Plant
Disease Agents. K. Maramorosch and R.F. Harris, eds. Academic
Press, New York. 654 pp.

Saglio, P., D. Lafleche, C. Bonissol, and J.M. Bové. 1971. Culture 12
vitro des mycoplasmes associés au "Stubborn" des agrumes et leur

observation au microscope électronique. H.R. Hebd. Séances Acad.
Sci. Ser. D 272: 1387-1390.

Saglio, P., M. L’Hospital, D. Lafleche, G. Dupont, J.M. Bové, J.G.
Tully, and E.A. Freundt. 1973. sgroplasma gitri gen. and sp.
n.: A mycoplasma-like organism associated with "Stubborn" disease
of citrus. Int. J. Syst. Bacteriol. 23: 191-204.

 

Townsend, R. 1983. Myc0plasma-like organisms from plants with
"yellows" diseases lack a spiroplasma-specific antigen. .1. Gen.
Microbiol. 129: 1959-1964.

Townsend, R. and D.B. Archer. 1983. A fibril protein antigen Specific
to Spiroplasma. J. Gen. Microbiol. 129: 199-206.

 

Townsend, R., J. Burgess, and K.A. Plaskitt. 1980. Morphology and

ultrastructure of helical and nonhelical strains of Spiroplasma
citri. J. Bacteriol. 142: 973-981.

 

Townsend, R., P.G. Markham, K.A. Plaskitt, and.M.J. Daniels. 1977.
Isolation and characterization of a non-helical strain of
SpirOplasma citri. J. Gen. Microbiol. 100: 15-21.

 

Whitcomb, R.F. 1980. The genus Spiroplasma. Ann. Rev. Microbiol. 34:
677-709.

 

Whitcomb, R.F. 1981. The biology of spiroplasmas. Ann. Rev. Entomol.
26: 397-425.

Whitcomb, R.F. and L.M. Black. 1982. Plant and arthropod mycoplasmas:
A historical perspective. Pp. 40-81 in: Plant and Insect
Mycoplasma Techniques. M.J. Daniels and P.G. Markham, eds. John
Wiley and Sons, New York. 369 pp.

Whitcomb, R.F., J.G. Tully, J.M. Bové, and P. Saglio. 1973.
Spiroplasmas and acholeplasmas: Multiplication in insects.
Science 182: 1251-1253.

Whitcomb, R.F. and D.L. Williamson. 1975. Helical wall-free
prokaryotes in insects: Multiplication and pathogenicity. Ann.
N.Y. Acad. Sci. 266: 260-275.

Whitcomb. R.F. and D.L. Williamson. 1979. Pathogenicity of
mchplasmas for arthropods. Zbl. Bakt. Hyg., I. Abt. Orig. A 245:
200-221.

Williamson, D.L. and R.F. Whitcomb. 1975. Plant myc0plasmas: A

cultivable spiroplasma causes corn stunt disease. Science 188:
1018-1020.

CBAPIERI

INVESTIGATION OF LEAFHOPPER INTERNAL MORPHOLWY

VIA SCANNING ELECTRON NIQOSCOPY 0P FROZEN SBCTIWS

INTRODUCTION

The internal morphology of leathppers has been studied primarily
from the standpoint of individual organs and their function. Much of
this work was predicated upon the importance of these organs in the
transmission of plant pathogens or in secretion of substances toxic to
plant hosts (e.g., Gil-Fernandez and Black 1965; Nuorteva 1956). In
this regard, much emphasis has been placed upon the salivary glands and
associated secretory tissues (e.g., Raine and Forbes 1971; Raine et al.
1976; Sogawa 1965). Some researchers, however, have done more exten-
sive work on internal morphology. Dobroscky (1931) did a rather de-

tailed study of Macrosteles fascifrons (Stal) using histological stain-

 

ing and light microscopy. She concluded that the aster yellows agent
was not detectable in the leafhopper (Dobroscky 1929). More recently,
Gil-Fernandez and Black (1965) investigated the internal morphology of

Agallia constricta (Van Duzee), a vector of wound-tumor virus. Both of

 

these works, while done very well, were technologically limited and,
therefore, somewhat inconclusive. Dobroscky (1931) used light micro-
graphs with morphology left unlabelled, making it extremely difficult
to locate and identify internal structures. Gil-Fernandez and Black
(1965) published only schematic drawings resulting in problems when
trying to repeat their work with actual specimens under the microsc0pe.
Sogawa (1965) only examined salivary glands of various leathppers, but
published both light micrographs, schematic drawings, and tracings that
are very useful guides to the further study of these organs.

Definitive localization of plant pathogens within their leathpper

10

11

vectors requires that the insects be examined without organ dissection.
This is especially true if tissues acting as barriers to transmission
are to be identified. Liu et al. (1983) followed the course of

Spiroplasma citri infection of Circulifer tenellus (Baker) by dissect-

 

 

ing out various organs at progressive times and preparing them for ul-
trathin sectioning and transmission electron microsc0py (TEM). While
appropriate for studies of detailed ultrastructural relationships, this
approach leaves to speculation the actual progress of infection as all
tissues, e.g., hemolymph, cannot be examined in this manner. Sinha and
Chiykowski (1967) used a similar dissection method to detect the pres-
ence of the aster yellows MLO in various internal tissues of

Macrosteles fascifrons (Stal) by homogenizing dissected organs and in-

 

jecting the homogenate into noninfected leafhoppers and monitoring any
subsequent transmission. This approach, even carefully performed, can-
not completely avoid MLO contamination from other tissues and would,
therefore, be inappropriate for detection of barrier tissues. Using
frozen sections of whole insects and immunofluorescence, D01 (1970) and
D01 et al. (1967) localized Japanese encephalitis virus in its 9.2L?!
mosquito vectors. This method provides the kind of information neces-
sary for barrier tissue studies, but it may'not be feasible for use
with large numbers of specimens that would need to be examined to de-
tect noncompetent vectors.

Preparation of fragile biological specimens for studying internal
morphology in the scanning electron microscope (SEM) can be difficult
when trying to preserve spatial and ultrastructural integrity. Sec-
tioning methods often produce unusable products. For example, freeze

fracturing follows planes of natural weakness (Postek et al. 1980)

12

making it almost impossible to obtain reproducible sections from com-
plex organisms. Moreover, because of the extreme brittleness of tissue
frozen in liquid nitrogen, serial sections or sections through specific
areas cannot be cut from the same specimen. Use of the cryostat par-
tially alleviates these problems, but obtaining the desired sectioning
plane can still be very difficult due to problems in precise orienta-
tion of the embedded specimen. Due to infiltration problems, embedding
specimens in various polymers used for ultrathin sectioning is usually
not feasible with many relatively large or nonpermeable samples which
may be used for morphological studies. Resin digestion to restore to-
pography is harsh and often destroys fine detail. In addition, embed-
ding media and/or digestion may chemically modify the specimen so that
immunological labels cannot be used (Hemming et a1. 1983).

Herein I describe a rapid, simple, and inexpensive method of pre-
paring leafhoppers for examination of internal morphology using the
scanning electron microscope. The results overcome most of the diffi-
culties encountered with the light microscopic techniques mentioned

above.

‘NAIERIALS ANDMNETHODS

The leathppers used in this work were ScaphytoPius acutus (Say)

 

and Macrosteles fascifrons (Stal) reared separately in the greenhouse

 

on red clover (Trifolium pratense L.) and barley (Hordeum vulgare L.),

 

respectively. Adults were fixed in 2% (v/v) glutaraldehyde in phos-
phate buffered saline (PBS; 0.01M phosphate + 0.15M NaCl, pH 7.4) for 2
hrs at room temperature and washed three times for 30 min each in PBS

prior to embedding.

13

The materials required for sectioningiand'handling the insects
are; a sectioning stage, an injector razor blade clamped in a hemo-
stat, 2 cm long pieces of teflon tubing with an ID equal to the OD of
the SEM stub being used, and small pieces of microscope lens tissue
(Fig. 1a). The liquid nitrogen-cooled sectioning stage was constructed
from a block of styrofoam by cutting a hole in the center large enough
to accommodate a 5 cm deep X 7 cm diameter crystalizing dish that
serves as a reservoir for the liquid nitrogen. Around this hole a
groove was cut to allow a 9 cm diameter petri dish to rest upside down
on top of the crystalizing dish and flush with the styrofoam surface.
Ten min after filling the crystalizing dish with liquid nitrogen, the
petri dish.stage:had reached its coldest temperature. Subsequently
applied drape of Tissue-TekR thoroughly hardened within 15 sec.

Leathppers were embedded in Tissue-TekR by placing a drOp on the

cold stage and quickly inserting the insect before the medium hardened.

In these experiments, the dorsal surface was left exposed to facilitate

sectioning orientation (Figp 1b). Any orientation may be selected but

the specimen must be inserted into the Tissue-TekR before it hardens
for successful sectioning. Longitudinal sections approximately 0.25 mm
thick were cut using a teflon-coated injector razor blade clamped in a
hemostat (Fig. 1a) and transferred to PBS-wetted lens tissue positioned
on top of 2 cm long pieces of 3/8 inch ID teflon tubing (Fig. 2a).
Another piece of PBS-wetted lens tissue was sandwiched over the first
and both were fastened to the tubing with wire (Fig. 2b). This sand-
wich formed the means for handling the fragile sections during a buffer
wash to remove the Tissue-TekR, dehydration through a graded ethanol

series, and subsequent critical point drying.

 

Figure I—1.——Equipment necessary for sectioning fragile specimens for
scanning electron microscopy. (A) Tools for sectioning
include sectioning stage, injector razor blade clamped in
a hemostat, pieces of teflon tubing, lens tissue for
forming sandwich (see text), and rack to hold tubing
throughout the procedure. (B) Frozen sectioning stage
with leafhopper embedded in Tissue-Tek’ (arrow) ready for
sectioning.

 

Figure I—2.-—Processing of frozen sections for scanning electron micro—
scopy. (A) Steps necessary in forming section sandwich
include transfer of frozen sections to wet lens tissue
already draped over teflon tubing, covering sections with
second piece of lens tissue, securing lens tissue with
wire, and trimming both wire and lens tissue. (B) Com-
pleted section sandwich ready for dehydration and critical
point drying. (C) Critical point driednsections being
mounted by inserting stub with Tubecoat“ up through the
teflon tubing. (D) Mounted sections (arrow) with upper
layer of lens tissue removed.

16

Following critical point drying, the wire was removed and a 3/8
inch OD aluminum stub with TubecoatR on the upper surface was gently
pushed up the teflon tubing until it contacted the lens paper sandwich
(Fig. 2c). Care was taken to put just enough TubecoatR on the stub to
seep through the paper and contact the sections without flooding them
and thereby ruining morphology. When the TubecoatR had dried, the
upper layer of lens paper was carefully removed leaving the sections
attached to the stub with the lower layer of lens paper, the excess of
which was trimmed off with a razor blade. (Fig. 2d).

In addition to dehydration and critical point drying, sections
were also lyophilized. Following the buffer wash, the sandwich was
rinsed in distilled water, removed from the tubing, and spread on fil-
ter paper in a 9 cm diameter petri dish. The tap layer of tissue paper
was removed and the sections were frozen at -25 C prior to placing them
in a Virtis Preservator 120 lyophilizer for 24 hrs. LyOphilized sec-
tions were carefully attached directly to the stubs with TubecoatR.

All specimens were sputter coated with approximately 30 nm of gold

and examined in a JEOL JSM 35C scanning electron microsc0pe operated at

15 kV.

RESUETS
Sectioned, dehydrated, and critical point dried S. Sggggg prepared
using this method demonstrated well preserved ultrastructure and easily
identified tissues and organs (Fig. 3). Sectioned and lyophilized M.

fascifrons also demonstrated organ and tissue integrity and spatial

 

relationships which were well maintained (Fig. 4). Thoracic flight

musculature was readily discernable as were abdominal digestive

l7

 

Figure I—3.-—Frozen section of Scaphytopius acutus (Say) processed
through dehydration and critical point drying. (A) Whole
body section showing intact head, thorax, and abdomen with
associated internal morphology. (B) Salivary gland.

(C) Intracellular ultrastructure of salivary gland. AL,
anterior lobe; CC, coagulated cytoplasm; DM, direct flight
muscles; FG, foregut; HG, hindgut; LC, longitudinal con—
nective; LM, longitudinal flight muscles; PL, posterior
lobe; SG, salivary gland; SGr, secretory granule; SubG,
subesophageal ganglion; Squ, supraesophageal ganglion;
TC, Tubecoat‘.

 

Figure I-4.--Frozen section of Macrosteles fascifrons (Stal) processed

through lyophilization. (A) Whole body section showing
intact head, thorax, and abdomen with associated internal
morphology. (B) Salivary gland. (C) Intracellular
ultrastructure of salivary gland. LL, lateral lobe; HT,
Malpighian tubules; SD, salivary duct; T, testes. See
Figure 3 for common abbreviations.

19

tissues. More delicate organs, such as neural ganglia and salivary
glands, were structurally intact and very easy to locate. In addition,
intracellular ultrastructure was well preserved as evidenced by the

acinous salivary gland cells (Figs. 3c and 4c).

DISCUSSION
These results are especially significant for S. acutus which
underwent the relatively rigorous treatment of critical point drying.
Although organ and intracellular ultrastructure were not affected, de-
hydration and critical point drying did result in clearing out the
specimen. This was due to the removal of most, if not all, of the
hemolymph from body cavities by the ethanol dehydration.

The sections of M. fascifrons showed the effects of ice crystal

 

formation during refreezing and lyophilization as evidenced by the
voids in the salivary glands (Fig. 4b). In addition, the entire inter-
nal morphology was less discernable due to the presence of hemolymph in
the body cavities. While lyophilized sections are adequate for gross
morphology, zit appears that dehydrated and critical point dried sec-
tions are preferable for more detailed analyses, making this handling
technique particularly useful for such fragile specimens.

The method of sectioning and handling the subsequently fragile
sections described here is an effective technique that overcomes sever-
al problems encountered when processing sectioned specimens for the
scanning electron microscope. It offers the ability to section in the
desired plane, as well as to obtain multiple and reproducible sections
from the same specimen. The cost of materials is insignificant and no

specialized equipment is necessary prior to the drying stage.

20

Moreover, it is possible to process the sandwiched sections through
other procedures before drying, e.g., substrate- or antibody-specific
labelling with substances detectable in X-ray and/or backscatter
analyses.

Several things should be kept in mind when attempting to section
specimens with this method. Do not totally cover the specimen with
Tissue-TekR as this will prevent selecting the desired sectioning
plane. Keep the sections: 1) frozen on the sectioning stage while
sectioning a specimen, or 2) moist on the tissue paper once sectioning
is completed. Even partial air drying can badly distort delicate tis-
sues. The 1eafhoppers used in this work ranged from 3 to 6 mm long but
there is no 3 priori reason why this technique would not work equally

well on larger or smaller specimens of any tissue type.

LITERATURE CITED

Dobroscky; I.D. 1929. 'Is the aster-yellows virus detectable in its
insect vector? Phytopathology 19: 1009-1015.

Dobroscky, I.D. 1931. Morphological and cytological studies on the
salivary glands and alimentary tract of Cicadula sexnotata
(Fallen), the carrier of aster yellows virus. Contrib. Boyce
Thompson Inst. 3: 39-58.

 

Doi, R. 1970. Studies on the mode of deveIOpment of Japanese
encephalitis virus in some groups of mosquitoes by the fluorescent
antibody technique. Jpn. J. Exp. Med. 40: 101-115.

Doi, R., A. Shirasaka, and M. Sasa. 1967. The mode of development of
Japanese encephalitis virus 111 the mosquito Sglgg
tritaeniorhynchus summorosus as observed by the fluorescent
antibody technique. Jpn. J. Exp. Med. 37: 227-238.

 

Gil-Fernandez, C. and L.M. Black. 1965. Some aspects of the internal

anatomy of the leathpper Agallia constricta (Homeptera:
Cicadellidae). Ann. Entomol. Soc. Amer. 58: 275-284.

 

Hemming, F.J., P. Mesguich, G. Morel, and P.M. Dubois. 1983.
Cryoultramicrotomy versus plastic embedding: Comparative

immunocytochemistry of rat anterior pituitary cells. .J. Micros.
131: 25-34.

Liu, H.-Y., D.J. Gumpf, G.N. Oldfield, E.C. Calavan. 1983. The
relationship of SpirOplasma citri and Circulifer tenellus.
PhytOpathology 73: 585-590.

 

 

Nuorteva, P. 1956. Notes on the anatomy of the salivary glands and on
the occurrence of proteases in these organs in some leafhoppers
(Hom., Auchenorrhyncha). Suomen Hyon. Aikakausk. 22: 103-108.

Postek, M.T., K.S. Howard, A.H. Johnson, and R.L. McMichael. 1980.
Scanning Electron Microsc0py. A Student's Handbook. Ladd Research
Industries. Burlington, VT. 305 pp.

Raine, .1. and A.R. Fbrbes. 1971. The salivary syringe of the
leafhopper Macrosteles fascifrons (Homoptera: Cicadellidae) and
the occurrence of myc0p1asma-like organisms in its ducts. Can.
Entomol. 103: 110-116.

 

Raine, J., A.R. Forbes, and F.E. Skelton. 1976. Myc0plasma-like
bodies, rickettsia-like bodies, and salivary bodies in the
salivary glands and saliva of the leafhopper Macrosteles
fascifrons (HomOptera: Cicadellidae). Can. Entomol. 108: 1009-
1019.

 

 

21

22

Sinha, R.C. and L.N. Chiykowski. 1967. Multiplication of aster
yellows virus in a nonvector leathpper. Virology 31: 461-466.

Sogawa, R5 1965.' Studies of the salivary glands of rice leafhoppers
I. Morphology'and histology. Jpn. J. Appl. Entomol. 2001. 9:
275-302.

GAPTERII

EFFECTS OF GLUTARALDEHYDE FIXATION ON SURFACE AND OVERALL

ANTIGENICITY OP SPIROPLASMA CITRI

23

INTRODUCTION
Spiroplasmas have been subjected to several immunological investi-

gations. The enzyme-linked immunosorbent assay (ELISA) has been

applied to the detection of both Spiroplasma citri and the corn stunt

 

spiroplasma (CSS) in plant and insect hosts (Bové et al. 1979; Clark et
a1. 1978; Raju and Nyland 1981; Saillard et al. 1978). The metabolism
inhibition (MI) test, which measures the acidic metabolites released
into the culture medium, omiginally developed to measure antibody to
mycoplasmas (Taylor-Robinson et al. 1966) has been applied to spiro-
plasmas for the same purpose (Williamson et al. 1979). A new tech-
nique, the spiroplasma deformation (DF) test, which assesses the mor-
phological effects of antibody binding, was deve10ped exclusively for
the study of spiroplasmas (Williamson et al. 1978). Both the MI and DF
tests have been used in combination to assess relationships among the
many strains of spiroplasmas (Davis et al. 1979; Williamson et al.
1979). From this has arisen the concept of the serogroup for separat-
ing the spiroplasmas serologically and at least six serogroups contain-
ing 24 strains are currently recognized (Whitcomb 1980).

Relatively little is known about the antigenic prOperties of the
cell membrane of spirOplasmas. Most, if not all, of the antigens com-
mon to the various serogroups are thought to be cytoplasmic (Whitcomb
1980). For many animal mchplasmas, membrane proteins are the primary
cell antigens, although membrane lipids, in the form of glycolipids,
also elicit immune responses (Razin et al. 1972). Many important anti-
gens of plasma membranes are glyc0proteins. The membrane of S. 53523;

contains approximately 2% carbohydrate (Razin et al. 1973) and most of

24

25

this is probably bound to proteins (Bové and Saillard 1979). This,
coupled with the fact that the membrane is also about 48% protein, sug-
gests that the surface of S. 51351 is highly antigenic.

Immunological methods offer the most efficient and sensitive means
of investigating spiroplasmas within their hosts. In fact, some sort
of specific label.is necessary to examine spiroplasmas within.their
leathpper vectors as helical morphology is not always expressed in
these hosts (Whitcomb and Williamson 1975). While undoubtedly correct
in their conclusions, Liu et al. (1983) offered only circumstantial

evidence that the microorganism identified within Circulifer tenellus

 

(Baker) was S. EEEEE' When sectioned leafhoppers are subjected to
scanning electron microscopy (as described in Chapter 1) in order to
detect spiroplasma infection, a specific label is indispensable as non-
helical cells would be indistinguishable from numerous other particles
within the specimen. Antibodies conjugated to various substances are
generally the labels of choice.

Detection and localization of spiroplasmas within their leafhopper
vectors requires chemical fixation for at least two reasons. First,
the morphological integrity of the host must be preserved in order to
locate and identify infected tissues. Second, the spatial distribution
of the spiroplasma must remain unchanged during the preparation of the
specimen for examination. Because glutaraldehyde fixation fulfills the
first requirement, it is reasonable to assume that the second will be
met as well. Caution must be exercized when using immunological labels
as glutaraldehyde fixation often reduces or destroys antigenicity (Van
Ewijk et a1. 1980). Virtually no information exists regarding the an-

tigenic response of spiroplasmas to glutaraldehyde fixation. It was

26

necessary to investigate this before attempting to label S. citri

within its vector, Macrosteles fascifrons (Stal).

 

MATERIALS AND METHODS

SpirOplaama Culture

The horseradish brittle root isolate of Spiroplasma citri was ob-

 

tained from Dr. C.E. Eastman, Illinois Natural History Survey, univer-
sity of Illinois, Champaign, Illinois. It was the sixth clone in the
20th passage and designated BR6-P20. The spiroplaana was cultured aer-
obically in LD8 medium (Lee and Davis 1984) at 31-32 C. Phenol red was
added to the medium to monitor growth and only log phase cultures were

used, assumed to be when the color of the medium just changed to

yellow.

Antiserum Production

Antigen was prepared by distributing 2000 ml of log phase BR6-P23
into 250 ml bottles and centrifuging at 16,000g for 60 min. The pel-
lets from eight bottles were each resuspended in 10 ml phosphate buff-
ered saline (PBS; 0.01M sodium phosphate + 0.15M NaCl, pH 7.4) and two
suspensions were pooled in each of four 40 ml centrifuge tubes. The
tubes were kept overnight at 4 C. An additional 10 m1 of PBS was added
to each tube and all were centrifuged at 15,000g for 30 min. The pel-
lets were resuspended in 5 ml PBS each, pooled, and sonicated for 30
sec using a Blackstone Ultrasonic generator fully tuned at 25% power.
Total protein was assayed by the method of Lowry et al. (1951) and the
antigen was stored in 1 ml aliquots at -25 C.

A New Zealand White rabbit was ear-bled for 10 ml blood as a

27

nonspecific serum source. After bleeding, the rabbit was ether anes-
thetized and injected with BR6-P23 antigen. Two ml antigen consisting
of approximately 14 mg protein was emulsified with 2 ml Freund’s com-
plete adjuvant and injected at four intramuscular hip and two subcutan-
eous back locations. Three weeks later, the rabbit was boosted in the
same manner except the back injections were omitted. One week follow-
ing the first boost, the rabbit was ear bled for 50 ml of blood. At 6
and 7 weeks following this first bleeding, the rabbit was again given
the same booster injections. At 10 and 19 days after the final boost,
the rabbit was ear bled for 50 ml of blood.

The blood from each bleeding was kept at room temperature for 60
min to allow clot formation and then held overnight at 4 C to shrink
the clot. The serum was decanted and centrifuged at 5000g for 30 min
to remove cellular debris. The clarified serum was passed through a
0.22 pm filter, aliquoted, and stored frozen at -25 C.

All antisera were titered using the MI test (Williamson et al.
1979) performed in microtiter plates. A log phase culture of BR6-P23
was used as antigen and LD8 medium was used to make all antiserum and
antigen dilutions. A 1:81 dilution of antiserum and the following di-
lutions of antigen were prepared: 1:2, 1:4, 10-1, 10'2, 10'3, 10‘4,
and 10'5. One hundred pl of LD8 medium was added to all wells in
columns 1 through 9, 11 and 12. One hundred fifty pl was added to the
wells in column 10 and these were used as medium control wells. To all
wells in column 1, rows A through H, 50 pl of the 1:81 antiserum dilu-
tion was added, producing a 1:243 dilution in the first well of each
row. Threefold dilutions of antiserum were made by serially transfer-

ring 50 pl across each row through column 9, but not columns 10, 11,

28

and 12. The 50 pl to be transferred out of the wells in column 9 was
discarded, leaving 100 pl in all wells of the plate except those in
column 10. A 50 pl amount of each antigen dilution, one dilution for
each row, was added to wells 1 through 9, 11, and 12. Columns 11 and
12 were, therefore, antiserum-free and served as antigen control wells.
At this point, a fresh vial of guinea pig complement (GIBCO, Inc.) was
rehydrated with PBS and an 8% solution prepared by mixing 0.8 ml com-
plement and 9.2 ml LD8 medium. This solution was passed through a 0.45
pm filter and 50 pl was added to all wells in the plate. The plate was
incubated at 31-32 C and read for a color change from red to yellow for
up to 8 days. The metabolism inhibition titer was expressed as the
highest antiserum dilution to prevent a color change at the highest
antigen dilution that produced a color change in the control wells.

The IgG fraction was purified from the highest titer antiserum by
ammonium sulfate precipitation and DEAE-SephacelR chromatography. Half
strength PBS was used to dilute 3.4 ml of antiserum to 10 ml and 10 ml
saturated ammonium sulfate was added. The mixture was gently stirred
for 60 min at room temperature and the precipitate was collected by
centrifugation at 5000g for 15 min. The precipitate was dissolved in
10 ml 1/2X PBS and the precipitation repeated twice more. The final
precipitate was dissolved in 2 m1 l/2X PBS and passed through a Sepha-
dexR G-25 column equilibrated in 1/2X PBS to remove ammonium sulfate.
After the void volume, 2.8 ml was collected and this was passed through
a DEAE-SephadexR column. The first 24, 1 ml fractions were pooled and
sufficient 1/2X PBS was added to bring the absorbance at 280 nm to 1.4
(91 mg/ml). The purified IgG was aliquoted into 1 ml portions and

stored at -25 C.

29

Preparation of Colloidal Gold-IgG Conjugate

The gold colloid was prepared by a modification of the method of
Frens (1973). To 500 ml of boiling 0.01% (w/v) HAuCl4 was added 12.5
ml of 11 N33C6H807 and the mixture was refluxed for 30 min. The color
of the colloid changed from an initial pale yellow to deep blue-black
to red-orange, indicating the reaction endpoint, within 5 min. The
additional reflux time insured the reaction reached 100% so as to pre-
vent unnecessary flocculation. The colloid was allowed to cool and
stored at 4 C. This procedure produced a colloid with an average par-
ticle size of 18 nm.

Conjugation of the gold colloid to purified rabbit IgG was per-
formed at pH 7.6 (Geoghegan and Ackerman 1977). A pH curve was con-
structed by placing 6.5 ml of the gold colloid in a 10 ml beaker and
measuring the pH after successive additions of 5 pl 0.2M K2C03 (Figure
1). From this, it was determined that 5.4 pl of 0.2M K2003 per ml of
colloid was necessary to bring the pH to 7.6.

The Optimum amount of IgG necessary to stabilize the gold colloid
during conjugation was determined by constructing an IgG absorption
isotherm using a variation of the method of Geoghegan and Ackerman
(1977). An aliquot of purified IgG was passed through a SephadexR
column equilibrated in distilled-deionized-distilled water (3D-H20) to
remove salts that would cause flocculation of the gold. To 11 , 2 ml
portions of the gold colloid, at pH 7.6, was added the following,pg
amounts of desalted IgG: 0, 4, 8, 12, 16, 20, 24, 28, 32, 36, and 40.
After 1 minute, 0.2 ml of 10% (w/v) NaCl was added which causes unsta-
bilized gold colloids to flocculate, indicated by a blue color. The

absorbance at 580 nm was plotted against the amount of IgG added

30

 

1 0.0 jr 1 ' V I ‘i 1 ‘ T ‘

 

 

 

5.0 l l l 1 l l l l l

o 10 20 3b 40 so
pl 0.2M choa/as ml Colloidal Gold

 

Figure II-1.--pH isotherm for the gold colloid produced by the sodium
citrate method (see text). From this curve, it was
determined that 5.4 pl of 0.2M K2c03 per ml colloidal
gold was necessary to bring the pH to 7.6.

31

(Figure 2) and the amount of IgG necessary to stabilize the colloid was
determined from where the curve becomes asymptotic to the abscissa. To
assure complete stabilization, 10 pg IgG/ml gold colloid was used for
conjugation.

Approximately 225 ml of gold colloid was centrifuged at 500g for
15 min to remove aggregates and 210 ml of the supernatant was trans-
ferred to a beaker for conjugation to IgG. Using desalted IgG, 2.1 mg
in 3.1 ml was added with gentle stirring. After 2 min, 2 ml of 1%
(w/v) polyethelene glycol (PEG; MW=20,000) was added to further stabi-
lize the conjugate. After a further 1 min of mixing, the conjugate was
centrifuged at 13,500g for 35 min. The very loose pellet was saved in
about 1 ml of the supernatant and resuspended in PPP buffer (0.01M so-
dium phosphate + 4% [w/v] polyvinylpyrrolidone + 0.2 mg/ml PEG). The
conjugate was passed through a 0.22 pm filter, brought to 40 ml with
PPP buffer, and stored at 4 C. Prior to every use, the conjugate was

centrifuged at 500g for 15 min to remove aggregates.

Fixation of Sjiroplas-a citri

Glutaraldehyde concentrations of 7.0, 6.0, 5.0, 4.0, 3.0, 2.0,
1.0, 0.5, 0.2, and 0.0% (v/v) were prepared in 7% (w/v) sorbitol (to
maintain prOper osmolality). To 15 ml portions of log phase BR6-P24
was added an equal amount of fixative producing final concentrations of
3.5, 3.0, 2.5, 2.0, 1.5, 1.0, 0.5, 0.25, 0.1, and 0.0%. These were
incubated at room temperature for 60 min followed by centrifugation at
15,000g for 20 min. The pellets were resuspended in 10 ml 0.1M NH4CI
in PBS and incubated at room temperature for 30 min to block any free

glutaraldehyde reactive groups. Ten ml of PBS was added and the tubes

32

 

 

 

 

 

0.6 -
0
0
5 L
E
2 0.2 »
b
0 a 16 24 52 1 40

pg IgG/2 ml Colloidal Gold

Figure II-2.--IgG absorption isotherm for the gold colloid produced by
the sodium citrate method (see text). From this curve,
it was determined that 10 pg of IgG per ml colloidal
gold would safely stabilize the colloid.

33
recentrifuged as above. The pellets were resuspended in PPP buffer for

colloidal gold labelling or PBS for ELISA.

Colloidal Gold Labelling

Fixed spiroplasmas in PPP buffer were centrifuged at 15,000g for
20 min in a swinging bucket rotor. The pellets were resuspended in 0.5
m1 PPP buffer, 0.5 m1 of colloidal gold-IgG conjugate previously dilu-
ted 1:4 added, and the tubes incubated at room temperature for 30 min.
A drop of this reaction mixture was placed directly on parlodion-coated
copper grids and allowed to stand for 30 min at 31 C and 100% relative
humidity. Excess fluid was drawn off, the grids allowed to air dry,
and negatively stained with 2% (w/v) ammonium molybdate adjusted to pH
7.5. All grids were examined in a Philips 201 transmission electron
microscope at 60 kV. In some cases, a drop of the reaction mixture was
placed on highly polished carbon planchets, incubated, and dried as
above. After attaching,the planchets to aluminum stubs with Tube-
coatR, they were examined in a JEOL JSM 35C scanning electron micro-
scope at 20 kV in both secondary and backscatter electron modes. The
secondary electron mode produces the topographical images normally as-
sociated with scanning electron microscopy. The backscatter electron
mode produces images based upon the molecular weight differences of the
elements that comprise the specimen.

Unfixed spiroplasmas were also labelled with colloidal gold. A
portion of the colloidal gold-IgG conjugate was diluted 1:4 and brought
to 7% (w/v) with solid sorbitol. Equal volumes of this reagent and log
phase BR6-P24 were mixed, incubated at room temperature for 60 min, and
applied to grids as above. Negative staining was accomplished using 2%

(w/v) ammonium molybdate in 3.5% (w/v) sorbitol adjusted to pH 7.5.

34

The sorbitol was included to maintain osmolality and prevent gross mor-
phological changes and/or cell lysis.

To assess the specificity of the colloidal gold label, fixed spi-
roplasmas were treated with free IgG preparations prior to labelling.
After blocking free aldehyde groups with 0.1M NH401, spiroplasmas fixed
in 3.5% glutaraldehyde concentrations were made to 25, 20, 15, 10, 5,
1, 0.5, 0.1, 0.05, 0.01, 0.005, and 0.001 pg/ml with nonspecific, puri-
fied from preimmune serum, and specific IgG in 1/2X PBS. After incuba-
tion at 37 C for 4 hrs, 10 ml of 1/2X PBS was added and the mixtures
were centrifuged at 15,000g in a swinging bucket rotor. Labelling was
then performed as above. Pretreatment with 1/2X PBS alone was used as
a control. As additional controls, spirOplasmas fixed in 3.5% (v/v)
glutaraldehyde were subjected to attempted labelling with unconjugated
gold particles and also with a goat anti-rabbit IgG-colloidal gold con-
jugate. Finally, an attempt was made to label sheep red blood cells
fixed in 3.5% (w/v) glutaraldehyde with the S. ESEES-specific colloidal

gold-IgG conjugate produced here.

Enzyme-linked Inmunosorbent Assay (ELISA)

The double antibody sandwich ELISA described by Clark and Adams
(1977) was used to measure the overall antigenic response of S. SEEEA
to glutaraldehyde fixation. Fixed spiroplasmas from 10 ml of culture
were suspended in 2 ml of PBS and sonicated for 30 seconds as above.
Total protein was assayed by the method of Lowry et al. (1951) and, for
ELISA testing, dilutions of 1:10, 1:50, 1:100, 1:500, and 1:1000 were
made. Spiroplasmas subjected to the entire fixation procedure, but

without glutaraldehydeg and those prepared directly from culture were

35

used as controls.

An alkaline phosphatase-IgG conjugate was prepared by mixing 0.8
ml (= 4.4 mg) of the enzyme (Sigma Type VII-S) with 1.8 ml (8 1.8 mg)
of S. SEES-specific IgG. This mixture was dialyzed three times
against 1000 ml of PBS, twice for 60 min and once overnight, at 4 C.
The dialysate (1.85 ml) was made to 0.06% with glutaraldehyde and
allowed to stand at room temperature for 4 hrs. The conjugate was di-
alyzed as above to remove excess glutaraldehyde, solid bovine serum
albumin added to 5 mg/ml, and stored at 4 C. For use, the conjugate
was diluted 1:1000 with conjugate buffer (PBS + 2% [w/v] polyvinylpyr-
rolidone + 0.2% [w/v] ovalbumin, pH 7.4). Coating IgG was prepared by

diluting S. citri-specific IgG to 1 pg/ml with coating buffer (15mM

Na2C03 + 35mM NaHCO3 + 0.02% [w/v] NaN3: pH 9-6)-

ELISA was performed by first adding 200 p1 of coating IgG to the
wells of microtiter plates and incubating for 4 hrs at 37 C. The
plates were washed three times for 3 min each with PBS-Tween (PBS +
0.05% [v/v] Tween-20, pH 7.4). After shaking the plates dry, 200 pl of
the various fixed, sonicated, and diluted spiroplasma preparations were
added to appropriate wells and incubated overnight at 4 C. The plates
were washed, as above, and 200 pl of conjugate was added to all wells
followed by incubation for 4 hrs at 37 C. After a final washing, 200
pl of p-nitrophenyl phosphate at 1 mg/ml in substrate buffer (9.7%
[v/v] diethanolamine + 0.02% [w/v] NaN3: PH 9-8) was added and the
plates incubated for 30 min at room temperature. The reaction was

stOpped by adding 50 pl of 3M NaOH and the absorbance read at 405 nm.

RESUETS

The effects of glutaraldehyde fixation on morphology and surface
labelling of S. glggi are presented in Figure 3. At all fixative con-
centrations, helical morphology was preserved and cell lysis or "bleb"
formation was prevented. No quantitative difference in gold labelling
was visually detected at any glutaraldehyde concentration. The gold
particles were distributed randomly over the entire surface of the
cells, with only limited patchiness.

Colloidal gold-labelled S. 51351 was observed in the scanning
electron microsc0pe (Figure 4). .Although visible in the micrographs,
spiroplasmas were invisible on the CRT screen of the microscope when in
secondary electron mode. SpirOplasmas were found by searching the car-
bon planchets in backscatter electron mode.

unfixed spiroplasmas lost helical morphology even under the mild
conditions used for gold labelling and negative staining (Figure 5).
Cells often remained filamentous, but blebs were always present. The
gold particles were always clustered in the bleb area of the cell with
none of the filaments showing any label.

S. SEES-specific IgG applied before the colloidal gold blocked
labelling, but nonspecific IgG did.not (Figure 6). Below 5 pg/ml,
little blocking was observed. Goat anti-rabbit IgG-colloidal gold con-
jugate and unconjugated colloidal gold both failed to label S. gig;
(Figure 7), nor did the S. ESEES-specific colloidal gold-IgG conjugate
label the fixed sheep red blood cells (Figure 8).

ELISA results showed that increasing glutaraldehyde concentration
decreased overall antigenicity of S. 2135; (Table 1 and Figure 9).

Above a fixed spirOplasma protein concentration of approximately 2.5

36

37

35

20

O5

 

Figure II—3.-—Transmission electron micrographs of the horseradish
brittle root isolate of Spiroplasma citri fixed in
various concentrations of glutaraldehyde and labelled
with a specific colloidal gold—IgG conjugate. Cells
were negatively stained with 2% ammonium molybdate.
The number indicate the percent glutaraldehyde used
for fixation. Bars are 1 pm.

 

 

Figure II-4.-—Scanning electron micrographs of the horseradish brittle
root isolate of Spiroplasma citri fixed in 3.5% glutar—
aldehyde and labelled with a specific colloidal gold—IgG
conjugate. The specimens are uncoated with B and D
being the backscatter images of the secondary images A
and C, respectively. Bars are l um.

39

 

Figure II-5.-—Transmission electron micrographs of the horseradish
brittle root isolate of Spiroplasma citri unfixed and
labelled with a specific colloidal gold-IgG conjugate.

Cells were negatively stained with 2% ammonium molybdate.
Bars are 1 pm.

 

4O

 

 

 

.O5NS .OSS‘
. T' ,5
A 3%
_ #4;
-L
.O1NS .013
.OO5NS .0053

   

001A} .0018

Figure II-6.--Transmission electron micrographs of the horseradish
brittle root isolate of Spiroplasma citri fixed in 3.5%
glutaraldehyde and pretreated with nonspecific and spec-
ific IgG prior to labelling with a specific colloidal
gold-IgG conjugate. Cells were negatively stained with
2% ammonium molybdate. Blocking of the colloidal gold
label was attempted using the IgG concentrations shown,
25NS = 25 ug/ml nonspecific IgG, 258 = 25 ug/ml specific
IgG, and so forth. Bars are 1 pm.

41

 

 

Figure II—7.-—Transmission electron micrographs of the horseradish
brittle root isolate of Spiroplasma citri fixed in 3.5%
glutaraldehyde prior to attempted labelling with a goat
anti-rabbit colloidal gold conjugate (A) and unconjugated

colloidal gold (B). Cells were negatively stained with
2% ammonium molybdate. Bars are 1 um.

42

 

Figure II-8.--Scanning electron micrographs of sheep red blood cells
fixed in 2.0% glutaraldehyde prior to attempted labelling
with a Spiroplasma citri-specific colloidal gold-IgG

 

conjugate. Cells were carbon coated. Bars are 1 um.

43

Table II-1.--ELISA results for the horseradish brittle root isolate of
Spiroplasma citri fixed in various concentrations of

 

glutaraldehyde.

 

 

Absorbancea at 405 nm

 

 

Percent for_given protein concentration (pg/ml)
Glutaraldehyde 25 5.0 2.5 0.5 0.25
3.5 1.47(.05) 0.54(.04) 0.33(.02) 0.12(.01) 0.09(.01)
3.0 1.53(.07) 0.58(.05) 0.36(.02) 0.12(.01) 0.09(.01)
2.5 1.58(.10) 0.63(.03) 0.37(.03) 0.12(.01) 0.09(.01)
2.0 1.68(.04) 0.73(.03) 0.46(.02) 0.14(.01) 0.10(.00)
1.5 1.73(.04) 0.76(.05) 0.50(.02) 0.15(.02) 0.11(.01)
1.0 1.77(.02) 0.93(.09) 0.60(.02) 0.18(.01) 0.12(.00)
0.5 1.81(.02) 1.39(.08) 0.94(.05) 0.27(.01) 0.18(.02)
0.25 1.80(.02) 1.71(.04) 1.32(.04) 0.40(.05) 0.24(.01)
0.10 1.79(.02) 1.81(.03) 1.75(.04) 0.68(.03) 0.39(.04)
0.0 1.79(.03) 1.80(.03) 1.81(.02) 1.79(.01) 1.73(.02)

 

aMean (and standard deviation) of 4 wells from 4 different plates.

44

 

 

 

E
g 1.2 P 4
O
3..
O P -l
0
C
G
‘9
o 0.8 ' ‘
C)
.D
< 1
F 4
0.4 - -

 

 

0 0.5 1.0 115 2.0 2.5 an 3.5
96 Glutaraldehyde

Figure II-9.--Effects of glutaraldehyde fixation on the enzyme-linked
immunosorbent assay (ELISA) for the horseradish brittle
root isolate of SpirOplasma citri. Shown are the
results for 25 pg/ml (O——O) and 0.5 pg/ml (O-—-O)
spiroplasma protein.

45
pg/ml, S. g_i_t_1;i antigens produced strong ELISA reactions at all fixa-
tive concentrations. At 0.25 pg/ml, the lowest protein concentration
to produce a maximum ELISA response with the unfixed preparation, fixed

S. citri antigens were undetectable above 1% glutaraldehyde.

DISCUSSION

These results demonstrate that the colloidal gold labelling of S.
SEEEA was specific and not due to residual reactivity of glutaraldehyde
in fixed preparations. This, and the fact that the colloidal gold did
not label sheep red blood cell membranes, indicates that spirOplasmas
may be detected within glutaraldehyde-fixed complex host tissues with-
out extensive background labelling due to the fixative. Mbre impor-
tant, however, is the maintenance of surface antigenicity at all glut-
araldehyde concentrations as this fixative often destroys cell surface
immunoreactivity at concentrations higher than 0.1% (Van Ewijk et al.
1980). Accurate localization of S. gigs; within its leafhopper vectors
depends upon good fixation of the host tissues and, apparently, the
spiroplasma can withstand this relatively harsh treatment.

Glutaraldehyde fixation and colloidal gold labelling have revealed
certain properties of the S. 21551 membrane. The fixed cells (Figure
3)showed surface antigens to be numerous and distributed over the en-
tire plasma membrane. The unfixed cells (Figure 5) demonstrated that
the surface antigens are mobile in the membrane as antibody binding
caused a capping phenomenon characteristic of many mammalian cells
(Eisen 1980). Indeed, this may be the basis of the spiroplasma defor-
mation test described by Williamson et al. (1978). In any event, these

membrane properties have great importance if ligand-receptor mechanisms

46

are the means of spiroplasma invasion of host cells, as ubiquity and
mobility of the ligand would facilitate binding of the host cell recep-
tor. This antigen mobility undoubtedly plays a role in the change in
morphology of S. 2123; within different hosts.

ELISA revealed that surface antigenicity is only a small part of
the overall antigenic makeup of S. 31331. This is supported by the
fact that even when undetectable with ELISA, surface labelling was un-
diminished. Apparently, glutaraldehyde can pass across the plasma mem-
brane to attack cytoplasmic antigens, which are much more sensitive to
the fixative as they are not protected by being embedded in the mem-
brane. This antigenic disparity is important in the preparation of
antisera to spiroplasmas. If intended for use in simply detecting spi-
roplasmas, e.g., with ELISA, then antigen should be prepared for injec-
tion so as to insure presentation of cyt0plasmic antigens, even though
this may reduce the immunogenicity of membrane antigens. Samples to be
tested should be prepared to expose cytoplasmic spiroplasma antigens to
the test antiserum. I earlier found that sonication of test samples
more than.doubled the resulting ELISA.reactions, apparently because
cytoplasmic antigens were released (Mowry 1982).

In conclusion, these experiments demonstrate that glutaraldehyde-
fixed S. ElELi. remains surface immunoreactive despite a predictable
decline in overall antigenicity. This fortuitous result makes it pos-
sible to detect S. Ei££1.13 fixed host tissues using immunological
methods, specifically colloidal gold labelling. My preliminary experi-
mentation has indicated that the S. ESEES-specific colloidal gold label
does not bind nonspecifically'to leafhopper tissues prepared by the

method described in Chapter I. The task at hand is to combine these

47

two techniques for the detection of spiroplasma plant pathogens within
their leafhopper vectors to identify sites of and barriers to infection

(see General Conclusion).

LITERATURE CITED

Bové, J.M., G. Moutous, C. Saillard, A. Fos, J. Bonfils, J.-C.
Vignault, A. Nhami, M. Abassi, K. Kabbage, B. Hafidi, C. Mouches,
and G. Viennot-Bourgin. 1979. Mise en évidence de SpirOplasma
citri, l'agent causal de la maladie du (stubborn) des agrumes dans
7 cicadelles du Maroc. C.R. Acad. Sci. Ser. D 288: 335-338.

 

Bové, J.M. and C. Saillard. 1979. Cell biology of spiroplasmas. Pp.
83-153 in: The Myc0plasmas, Vol. III. R.F. Whitcomb and J.G.
Tully, eds. Academic Press, New York. 351 pp.

Clark, H.F. and A.N. Adams. 1977. Characteristics of the microplate
method of enzyme-linked immunosorbent assay for the detection of
plant viruses. J. Gen. Virol. 34: 475-483.

Clark, H.F., C.L. Flegg, M. Bar-Joseph, and S. Rottem. 1978. The
detection of Spiroplasma citri by enzyme-linked immunosorbent
assay (ELISA). Phytopathol. Z. 92: 332-337.

Davis, R.E., I.-M. Lee, and L.R. Basciano. 1979. Spiroplasmas:

Serological grouping of strains associated with plants and
insects. Can. J. Microbiol. 25: 861-866.

Eisen, H.N. 1980. Immunology. Harper and Row, Philadelphia. 547 pp.

Frens, G. 1973. Controlled nucleation for the regulation of the

particle size in monodisperse gold suspensions. Nature Phys. Sci.
241: 20-22.

Geoghegan, W.D. and G.A. Ackerman. 1977. Adsorption of horseradish
peroxidase, ovomucoid and anti-immunoglobulin to colloidal gold
for the indirect detection of concanavalin A, wheat germ
agglutinin and goat anti-human immunoglobulin G on cell surfaces
at the electron microscope level: A new method, theory and
application. J. Histochem. Cytochem. 25: 1187-1200.

Lee, I.-M. and R.E. Davis. 1984. New media for rapid growth of

Spiroplasma citri and corn stunt spiroplasma. PhytOpathology 74:
84-89.

Liu, H.-Y., D.J. Gumpf, G.N. Oldfield, and E.C. Calavan. 1983. The
relationship of Spiroplasma citri and Circulifer tenellus.
Phytopathology 73: 585-590.

 

3Lowry, O.H., N.J. Rosebrough, A.L. Farr, and.R.J. Randall. 1951.

Protein measurement with the Folin phenol reagent. J. Biol. Chem.
193: 265-275.

48

49

Mowry, T.M. 1982. Leafhopper sampling in Michigan peach orchards and
serological detection of a spiroplasma associated with X-disease
in plant and insect tissue. Master's Thesis, Mich. St. Univ.,
East Lansing. 134 pp.

Raju, E.C. and G. Nyland. 1981. Enzyme-linked immunosorbent assay for
the detection of corn stunt spiroplasma in plant and insect
tissue. Curr. Microbiol. 5: 101-104.

Razin, S., M. Hasin, Z. Ne’eman, and S. Rottem. 1973. Isolation,
chemical composition, and ultrastructural features of the cell
membrane of the mycoplasma-like organism Spir0plasma citri. J.
Bacteriol. 116: 1421-1435.

 

Razin, 8., IL. Kahane, and J. Kovartovsky. 1972. Immunochemistry of
mchplasna membranes. Pp. 93-122 in: Pathogenic Myc0plasmas. A.
Ciba Foundation Symposium. Associated Scientific Publishers,
Amsterdam. 404 pp.

Saillard, C., J. Dunez, O. Garcia-Jurado, A. Nhami, and J. Bové. 1978.
Détection de Spiroplasma citri dans les agrumes et les pervenches
par la technique immuno-enzymatique (ELISA). C.R. Acad. Sci. Ser.
D 286: 1245-1248.

 

Taylor-Robinson, D., R.H. Purcell, D.C. Wong, and R.M. Chanock. 1966.
A colour test for the measurement of antibody to certain
mycoplasma species based upon the inhibition of acid production.
J. Hyg. 64: 91-104.

Van Ewijk, W., R.C. Coffman, and I.L. Weissman. 1980. Immunoelectron
microscopy of cell surface antigens: A quantitative analysis of
antibody binding after different fixation protocols. Histochem.
J. 12: 349-361.

Whitcomb, R.F. 1980. The genus Spiroplasma. Ann. Rev. Microbiol. 34:
677-709.

 

Whitcomb, R.F. and D.L. Williamson. 1975. Helical wall-free
prokaryotes in insects: Multiplication and pathogenicity. Ann.

Williamson, D.L., J.G. Tully, and R.F. Whitcomb. 1979. Serological
relationships of spiroplasmas as shown by combined deformation and
metabolism inhibition tests. Int. J. Syst. Bacteriol. 29: 345-
351.

Williamson, D.L., R.F. Whitcomb, and J.G. Tully. 1978. The
spiroplasma deformation test, a new serological method. Curr.
Microbiol. 1: 203-207.

cam III

EFFECTS OF IN VITRO PASSAGE ON THE PLASMA MEMBRANE

AND TRANSMISSION OF SPIROPLASMA CIEI

50

INTRODUCTION
The discovery that mchplasmalike organisms were the possible
etiological agents of several yellows-type plant diseases (Doi et al.
1967) initiated an intensive search for presumptive mchplasmas causing
diseases for which no pathogen had yet been found (Whitcomb and Black
1982). This resulted in the isolation and fulfilling of Koch’s postu-
lates for two plant pathogenic mchplasmas, the corn stunt spiroplasma

(Chen and Liao 1975; Williamson and Whitcomb 1975) and Spiroplasma

 

giggi (Markham et al. 1974). At that time, leafhoppers were the only
known vectors of the yellows disease pathogens. Another search, which
still continues, began for the leafhoppers responsible for transmitting
plant spiroplasmas.

Only four leafhoppers are currently known to transmit S. REESE

from plant to plant. These are Scaphytopius nitridus (DeLong)

 

(Kaloostian et al. 1976) , S. acutus delongl Young (Kaloostian et al.

 

1979), Circulifer tenellus (Baker) (Kaloostian et al. 1976; 1979;

 

Oldfield et al. 1976; Liu et al. 1983a), and Macrosteles fascifrons

 

(Stal) (O’Hayer et al. 1983). Two other leafhoppers, Euscelidius

 

variegatus (Kirsch) and Euscelis plebejus (Fallén), have been shown to
be experimental vectors, i.e., the pathogen is acquired by injection
(Markham et al. 1974; Markham and Townsend 1979). S. SiEEl was detect-
ed by the enzyme-linked immunosorbent assay (ELISA) in seven other
field-collected leafhoppers (Bové et al. 1979). In membrane feeding
experiments, four additional leafhoppers and one membracid were able to
acquire, but not transmit, S. SE32 (Rana et a1. 1975). It appears

that while a number of leafhoppers may be capable of acquiring and

51

52

maintaining the pathogen, a relatively few are able to transmit it to
plants.

S. 91531 is circulative and propagative in its leafhopper vector,
S. tenellus (Liu et al. 1983a; 1983b). This means that the pathogen
must pass through at least three tissue types before transmission can
occur: gut epithelium, hemolymph, and salivary acini (Whitcomb 1981).
This is certainly true for the other natural vectors of spiroplasmas.
The mechanisms involved in this passage, especially across membranes,
are largely unknown. Gildow (1985) demonstrated that barley yellow
dwarf virus (BYDV) adsorbed specifically to the plasma membrane of
hindgut epithelium in its aphid vector. This initiated coated pit de-
velopment and eventual endocytosis of the virion within a coated vesi-
cle. Gildow postulated that specific adsorption of BYDV to gut epithe-
lium may account for the specificity of virus transmission among
aphids. Liu (1983b) observed S. ElEEl partially embedded in the wall
of gut epithelial cells and within "vesicles" between the epithelium
and the basement.membrane. This spiroplasma.has also been observed
within vesicles inside cells of monolayer cell culture (Markham 1982).
These reports all argue for receptor-mediated endocytosis as a mechan-
isnlof leafhopper infection, particularly'in light of the fact that
coated pits, which are clusters of receptors, form in response to bind-
ing of receptors by ligands on the invading particle (Willingham and
Pastan 1983). It has been suggested that spiroplasmas pass between the
cells of gut epithelium and salivary acini via the cell junctions as
they have been observed in these locations (Markham 1982). This does
not, however, account for the multiplication of spiroplasmas in the

cells of these organs and may reflect an inadvertant deposition of the

53

pathogen. In any event, phytOpathogenic spiroplasmas do enter the
cells of their insect vectors and it would appear that the mechanisms
involved are both molecular and specific.

The phytopathology of S. 2133; is not well understood, but is gen-
erally felt to arise from a toxin produced by the pathogen. The major
acidic metabolite of S. 5155; is lactic acid (Hawthorne and Vandemark
1977), but it is probably not involved pathologically at the concentra-
tions normally encountered (Daniels 1979a). Two metabolites have ex-
hibited phytotoxicity, one being neutral (Daniels and Meddins 1974) and
the other acidic (Daniels 1979a). Both are low molecular weight sub-
stances that are very unstable and, therefore, have not been isolated.
How continuous l2 ygggg passage of S. ElEEl affects production of these
toxins is unknown.

The plasma membrane of S. SHEA has been fairly well character-
ized. Gross analysis revealed that the membrane is about 30% lipid and
48% protein (Razin et al. 1973), by weight, with carbohydrates consti-
tuting about 2.2% (Whitcomb 1980). ElectrOphoretic analysis has de-
tected 16 membrane proteins (bands) of which spiralin is the dominant
member, comprising approximately 22% of total membrane protein (Whit-
comb 1980; Wroblewski et al. 1977). The metabolic and structural roles
of these membrane proteins have not been determined, although spiralin
may be involved in maintenance of spiral morphology as antispiralin
antibodies deform the organism (Wroblewski 1978).

The response of S. glggi to S3 XEEEB culture and passage may
supply information about 13 1312 mechanisms of infection and survival.
The adaptation of microorganisms to the culture environment often leads

to the inability to complete their natural biological cycle or, in the

54

case of pathogens, the loss of pathogenicity (attenuation). By compar-
ing modified and unmodified cultures, it is sometimes possible to
detect macromolecular changes that may be involved in the lost function
due to continuous culture. The purpose of this research was to assess
the effects of $2 yiggg passage of S. 91531 related to its pathogenic-
ity and/or transmissibility and to discover any concurrent macromolecu-
lar changes in the plasma membrane. It was also an objective to pro-

vide evidence for transmission barriers in the leafhopper vector.

INATERIAleANDINETEODS

Spiroplasma and Leafhopper Cultures

The horseradish brittle root isolate of S. EEEEA was obtained from
Dr. C.E. Eastman, Illinois Natural History Survey, University of Illi-
nois, Champaign, Illinois. It was the sixth clone in the 20th passage
and designated BR6-P20. The spiroplasma was cultured aerobically in
LD8 medium (Lee and Davis 1984) at 30-31 C.

The aster leafhopper, Macrosteles fascifrons (Stal), was used as

 

the vector of S. citri. It was maintained on barley (Hordeum vulgare
L.) in the greenhouse under a 16 hr day supplemented with fluorescent

lights. Only adults of both sexes were used in these experiments.

Passage and Injection of Spiroplasma citri

Passage 24 was obtained by inoculating 1 ml of passage 23, thawed
from frozen storage, into 99 ml of fresh LD8 medium. This was the
starting point because it had been previously determined through leaf-
luopper injection that passage 24 was as pathogenic as any earlier pas-

sage. Each passage was initiated by inoculating 10 ml of the previous

55
passage in log phase growth into 90 ml fresh LD8 medium and culturing
for 48 hrs. At the end of this time, a 10 ml aliquot was removed for
leathpper injection. For each passage, approximately 100 leafhoppers
were anesthetized with carbon dioxide and immobilized on an upside down
glass Petri dish with Parafilm'MR stretched to the breaking point.
Each insect was injected through the Parafilm with glass needles using
an ISCOR microapplicator. The inoculum consisted of 0.1-0.2 pl of cul-
ture containing the number of colony forming units (CFUs) listed in
Table 1 and was injected between the ventral abdominal sclerites.

Fresh LD8 medium was injected into approximately 100 leafhoppers as a

control. After injection, leafhoppers were held for 14 days on asters
(Callistephus chinensis Nees) cv. American Branching in a growth cham-
ber under 16 hrs of fluorescent light per 24 hr period at 27-28 C.
Following this incubation period, all surviving leafhoppers were trans-
ferred individually to 4-5 wk old aster test plants for an inoculation
access period of 7 days under the same light/temperature regime. Upon
removal of the insects, the test plants were held in the greenhouse up

to 10 wks for symptom development.

Acquisition of Spiroplasma citri from Plants

 

Mollicute-free adult leafhoppers were caged in groups of 50 on S.
giEES-infected asters for 14 days in a growth chamber under the above
conditions. Survivors were transferred to healthy asters for 7 days
and then individually to 4-5 wk old aster test plants for an additional
7 days. After removing all insects, the test plants were held in the

greenhouse up to 10 wks for symptom develOpment.

56

Table III-1.--Number of colony forming units (CFUs) of the horseradish
brittle isolate of SpirOplasma citri injected into

Macrosteles fascifrons.

 

 

0.1-0.2 pl of undiluted culture.

Leafhoppers were injected with
Values were obtained by

plating 25 pl of appropriately diluted culture on solid
LD8 medium and counting subsequent colonies.

 

 

 

Passage No. CFUs/mla CFUs Injectedb

24 NBC NBC

25 4.9 x 109 4.9 x 105 - 9.9 105
26 6.9 x 109 6.9 x 105 - 1.4 106
27 9.1 x 109 9.1 x 105 - 1.8 106
28 1.5 x 109 1.5 x 105 - 2.9 105
29 1.9 x 1010 1.9 x 106 - 3.8 106
30 2.3 x 109 2.3 x 105 - 4.6 105
31 2.3 x 1011 2.3 x 107 - 4.6 107
32 2.3 x 1010 2.3 x 106 - 4.6 106
33 5.8 x 109 5.8 x 105 - 1.2 106
34 2.1 x 1011 2.1 x 107 - 4.2 107
35 2.6 x 109 2.6 x 105 - 5.2 105

 

8Mean of 3 plates. Values are for

bIn 0.1-0.2 61.

cNot done.

undiluted cultures.

57
Isolation of Spiroplasmas

All surviving leafhoppers from both injection and plant acquisi-
tion experiments were frozen at -25 C. Injected insects were held fro-
zen up to 5 wks and the plant-fed leathppers held overnight. Isola-
tion of spiroplasmas from each individual leafhopper was performed
using a modification of the method of Markham et al. (1983) . The in-
sects were surface sterilized by immersion in 70% (v/v) ethanol for 2
min followed by two rinses in sterile distilled water. Each was then
thoroughly ground in 2 ml LD8 medium. The homogenate was passed
through a 0.45 pm filter into a sterile tube and incubated at 30-31 C
for 4 wks.

Spiroplasma isolation attempts were made from all test plants
showing S. 21331 symptoms and some selected nonsymptomatic plants using
a modification of the method of Bové et al. (1983). A 2 4cm piece of
stem tissue was surface sterilized by immersion in 70% (v/v) ethanol
for 2 min followed by 5 min in 1% (w/v) sodium hypochlorite. After
three rinses in sterile distilled.water, the pieces were cut up and
crushed in 3 ml LD8 medium and allowed to stand for 15 min at room
temperature. The homogenate was filtered through a 0.45 pm filter into
sterile tubes containing the necessary amount of LD8 medium to make
both 1:4 and 1:20 dilutions of the original filtrate. The tubes were

incubated at 30-31 C for up to 4 wks.

Phytotoxin Assay
For each passage of S. citri, phytotoxins were assayed using a
modification of the method of Daniels (1979a). Newly Opened leaves of

broad bean (Vicia faba L.) cv. Long Pod Fava were surface sterilized by

 

immersion in 70% (v/v) ethanol followed by two rinses in sterile

58

distilled water. The leaves were placed upside down on sterile, wet
gauze pads and.the lower epidermis removed with fine point forceps.
Using a sterilized cork borer, 3 mm diameter pieces were cut from the
leaves and floated, stripped side down, on the surface of sterile 2%
(w/v) sorbitol until used (no more than 1 hr).

From each passage, 12 ml of culture was centrifuged at 15,000g for
20 min, the supernatant transferred to a small beaker, and the pH mea-
sured. The clarified culture supernatant was passed through a 0.22
pm filter and dilutions made aseptically with sterile 2% (w/v) sorbitol
to produce supernatant concentrations of 100, 90, 80, 70, 60, 50, 40,
30, 20, and 10%. LD8 medium was made to 100, 50, and 10% for use as
controls. Two ml of each dilution was placed in 30 X 10 mm culture
dishes and two disks of prepared broad bean leaves were floated,
stripped side down, on the surface of the fluid. The dishes were
covered and incubated overnight at 37 C. The next morning, the diluted
supernatants were aspirated off and the leaf disks washed twice in dis-
tilled-deionized-distilled water (3D'H20). After removing as much
water as possible, the leaf disks were extracted for 1 hr with 100%
methanol followed by two washes in 3D-H20. The disks were appropriate-
ly arranged on moist filter paper and photographed using Panatomic-XR

negative film.

Membrane Preparation

Plasma membranes were isolated from log phase cultures of BR6-P26
and BR6-P34 using a modification of the method of Razin et al. (1973).
One 1 of log phase culture was centrifuged at 15,000g for 40 min in 250

ml bottles. The pellets were resuspended in 10 ml 0.25M NaCl,

59

distributed into two 36 ml tubes and centrifuged at 15,000g for 20 min
in a swinging bucket rotor, as were all subsequent centrifugations.
The pellets were again resuspended in 10 ml 0.25M NaCl, held on ice,
and sonicated three times for 30 sec each, with 30 sec intervals on
ice, using 8 BlackstoneR Ultrasonic generator fully tuned at 25% power.
The suspensions were transferred to new tubes, filled with 0.25M NaCl,
and centrifuged at 34,000g for 30 min. The pellets were resuspended in
15 ml of 0.02M MgClz containing 10 pg/ml deoxyribonuclease (Sigma Type
DN-25) warmed to 37 C and incubated for 15 min at 37 C. Fifteen m1 of
0.02M ug012 was added and the tubes centrifuged at 43,100g for 30 min.
The membranes were alternately washed three times each with cold low
strength phosphate buffered saline (PBS; 0.01M sodium phosphate + 0.05M
N301: PH 7-5) and C01d 3D'H20. The final pellets were resuspended in
about 5 ml PBS (0.01M sodium phosphate + 0.15M NaCl + 0.005M EDTA, pH
7.4). Total protein was measured with the method of Lowry et al.
(1951) and the volume adjusted to a protein concentration of 1 mg/ml.

The membranes were stored in 1 ml aliquots at -25 C until used.

Membrane Electrophoresis

Membranes were prepared for electrophoresis by centrifuging 2 ml
(= 2 mg)lof purified material at 43,100g for 30 min and resuspending
the pellet in 1 ml of sample buffer (0.0625M Tris-HCl, pH 6.8 + 10%
[v/v] glycerol + 2% [w/v] SDS + 5% [v/v] 2-mercaptoethanol + 0.002%
[w/v] bromophenol blue). This suspension was heated at 100 C for 3 min
to completely denature all proteins. After cooling to room tempera-
ture, 25 pl aliquots (= 50 pg total protein) were layered on the gel
for analysis. Molecular weight standards (Sigma MW-SDS-200) were pre-

pared and used in the same manner, except that 10 p1 was applied to the

60

gel. The standards were: carbonic anhydrase (MW 29,000), ovalbumin
(MW 45,000), bovine serum albumin (MW 66,000), phosphorylase B (MW
97,400), B-galactosidase (MW 116,000), and myosin (MW 205,000).

Sodium dodecyl sulfate polyacrylamide gel electrOphoresis (SDS-
PAGE) was carried out in the discontinuous buffer system of Laemmli
(1970). The resolving gel consisted of a 5-20% (w/v) acrylamide gradi-
ent and a 1 cm stacking gel of 3.75% (w/v) acrylamide was used. Run-
ning conditions were a constant current of 15 mA until the bromOphenol
blue dye front reached the resolving gel (approximately 2 hrs) at which
time the current was increased to 30 mA until the dye front was about 1
cm from the bottom of the gel (5-6 hrs). The analysis was done using a
BidRad ProteanR vertical electrophoresis device.

Separated proteins were visualized by the double staining tech-
nique of Dzandu et al. (1984). Each gel was fixed in 500 ml 40% (v/v)
methanol/10% (v/v) acetic acid for 1 hr at room temperature followed by
two 30 min.washes in 500 ml 10% (v/v) ethanol/5% (v/v) acetic.acid.
The gel was equilibrated in 200 ml of oxidizer (3.4 mM potassium di-
chromate + 3.2 mM nitric acid) for 10 min followed by three 5 min
washes in 500 ml distilled water. The gel was bathed in 200 m1 of
silver reagent (20 mM silver nitrate) followed by three 1 min washes in
distilled water. Band development was accomplished by immersing the
gel in three changes of 200 ml of develOper (0.28M N32C03,+ 0.008%
[w/v] paraformaldehyde) at 37 C for 20 min each. After a 1 min wash in
distilled water, the gel was immersed in 200 ml 10% (v/v) acetic acid.
The gels were photographed on a fluorescent light box using Kodacolor
negative and Kodachrome slide films with a orange FL-B filter.

After the first photography, each gel was immersed in 200 ml of

61

counterstain (0.1% [w/v] Coomassie brilliant blue R-250 + 25% [v/v]
methanol + 7.5% [v/v] acetic acid) for 1 hr. Gels were destained over-

night in 25% (v/v) methanol/7.5% (v/v) acetic acid and photographed as

above.

RESULTS

The M. fascifrons used in these experiments were able to acquire

 

‘S. glggi via injection and subsequently transmit the pathogen (Table
2). Transmission was poor, never exceeding 11.6% (passage 26). Acqui-
sition, however, was much higher, reaching 89% in passage 27. No
transmission occurred after passage 30, even though acquisition re-
mained at high levels for all but passage 35. Regression analysis of
percent transmission on passage number revealed a negative relationship
(y = 33.003 - 0.998x) with a slope significantly different from zero at
P<.005 (F = 18.89 with 11 and 1 degrees of freedom). Regression analy-
sis of percent acquisition on passage number did not produce a lepe
significantly different from zero (F = 1.62 with 11 and 1 degrees of
freedom). None of the control leafhoppers injected with LD8 medium
either acquired or transmittequ. £3551. Only one of the leafhoppers
given an acquisition access period on diseased plants was able to
transmit S. giggl, and a small number were able to acquire the pathogen
in each of the four experiments (Table 3).

A total of 17 plants developed S. SALE; symptoms, which were iden-
tical to those described by O’Hayer et a1. (1983). Symptoms first ap-
peared in young leaves, which showed chlorosis and stunting and often
become asymmetric. Three symptomatic plants suddenly wilted and died

before spiroplasma isolation could be attempted. Of the remaining 14

62

Table III-2.--Effects of 12 vitro passage on acquisition and transmis-

 

sion of the horseradish brittle root isolate of
SpirOplasma citri by injected Macrosteles fascifrons.
After injection, leafhoppers were given a 14 day incuba-
tion period on healthy asters and then caged individually
for 7 days on 4-5 week old aster test plants. Acquisi-
tion was confirmed by isolation of spiroplasmas from
individual insects.

 

 

 

 

Fraction of Fraction of

 

 

 

Leafhoppers Transmittinga Leathppers Acquiringb

Passage No. Treatedc:d Controle Treatedc:d Control8
24 1/9 0/4 4/6 0/4
25 3/43 0/56 15/38 0/50
26 5/43 0/30 22/36 0/25
27 2/44 0/72 33/37 0/63
28 0/18 0/17 7/14 0/15
29 1/38 0/31 12/32 0/28
30 4/66 0/36 3/49 0/29
31 0/46 0/32 13/35 0/30
32 0/65 0/40 43/57 0/38
33 0/41 0/40 24/29 0/33
34 0/16 0/30 4/16 0/27
35 0/11 0/55 0/11 0/51

 

aNo. transmitting/no. surviving incubation period.

bNo. producing spiroplasma culture/no. surviving inoculation period.

cInjected with 3 x 105-5 x 107 CFUs in 0.1-0.2 .41.

dFor transmitting leafhoppers: y = 33.003 - 0.998x, r2 = 0.654, slope
significantly different from zero at P<.005.
For acquiring leafhoppers: y = 135.353 - 2.977x, r2 = 0.140, lepe
not significantly different from zero.

eInjected with 0.1-0.2 pl LD8 medium.

63

Table III-3.--Natural acquisition and transmission of the horseradish

brittle root isolate of Spiroplasma citri by Macrosteles
fascifrons. Treated leafhoppers were given a 14 day

 

 

 

acquisition access period on diseased asters followed by
a 7 day incubation period on healthy asters. Controls
fed only on healthy asters. Leafhoppers were caged
individually on 4-5 week old asters for a 7 day
inoculation access period. Acquisition was confirmed by
isolation of spirOplasmas from individual insects.

 

 

 

 

 

Fraction of Fraction of
Leafhoppers Transmittinga LeafhoPpers Acquiringb
Experiment No. Treated Control Treated Control
1 0/32 0/5 1/29 0/3
2 0/37 0/5 2/35 0/4
3 1/33 0/5 3/32 0/4
4 0/34 0/5 1/33 0/4

 

aNo. transmitting/no. surviving incubation period.

bNo. producing spiroplasma culture/no. surviving inoculation period.

64

plants, 13 produced spiroplasma cultures and isolation was not per-
formed on the symptomatic plant from natural transmission.

All 12 S. 91331 passages involved in these experiments exhibited
phytotoxin activity in the culture supernates (Figure 1). Some varia-
bility was observed in the assay, which is probably attributable to the
age and physiological condition of the leaves used. Even though young,
newly opened leaves were used throughout, not all were exactly the same
age. The pH of the culture supernates varied from 6.92 to 7.36, but
this was apparently unrelated to toxin activity.

SDS-PAGE analysis of S. EEEEE membrane proteins revealed many more
proteins and/or subunits than previously reported for this spiroplasma
(Wroblewski et al. 1977) (Figure 2). The double staining technique
demonstrated the presence of sialoglyc0proteins (yellow bands) in the
membrane of S. EEEEA’ which has not been reported before. Of interest
is the largest and most densely stained band just below the MW 29,000
marker protein. It is most likely spiralin, which has a molecular
weight of 26,000 (Wroblewski 1978). This band was stained yellow with
the silver stain, indicating that it is a sialoglyc0protein which con-
tradicts Wroblewski (1978) who stated that spiralin contains no carbo-
hydrate. Further investigation is necessary to resolve this
discrepancy.

Passage 26 membrane analysis showed a conventional Coomassie bril-
liant blue-sensitive protein (approximate MW 19,000) that was apparent-
ly lost in passage 34 (Figure 2, arrows). In addition, there appears
to be a reduction in some sialoglyc0proteins in passage 34 relative to
passage 26, as indicated by the less intense yellow color at the band

Llocation just above the deleted protein.

65

“-7“ ‘03..) alt-P” fi'7.“

III as an I. a. as to so to 10 100 so no 20 to an to so no 10
*IllllD IIDIIDllllIllIIIlII - (1' cl loo .. CD I. IID1II
.I O

“on: fl.‘.,1 lfl-m ‘07.”
an I) In n1 co x1 to :n a: 10 “N ”1 ID 1' 1' ii ‘0 II II In

Que-00000300»-.. ° "' 0'0

 

O. 0.

III - '2‘ pl 0 7.1) out - '11 pl . 6.96 _

A“ Q Q N U ” U D I II 100 90 fl ,0 00 30 60 n 10 10

‘0' 1D CI'III1II IIDFI‘Inu 0 II 0- 1| 'CIIOI II 4 « .

C

.- . m .- - 2.. "-1";- ‘ f” ' .-;:_’:,

HI II II II C. II to II 10 ul Ill I! II Ill (I II CD II ll ll

0 ‘OilIII1IIIIII C. Owl 9‘ Ch Ill I'll-I I. II-«n: on ..
1r; . _ mil

an. - 12.4. N: - 7.23 7-.- '0 8.- _ -

100 90 80 70 00 50 m an 2" 1‘ . I I 3 C I O I I 3

Cl 1.. IIDGIDl-D Cllllllifi '- 0» OD lIDIII«I."IlIIlIIIIL ,

a O .
m-m plot“ Ins-m pith”
100 ’0 N N 69 w 60 ” 20 10 m ’0 I) 70 t. a 40 n 20 10
a 00>¢H|10010> Oil!!! 0 Illlll1ll IllIIIlII Cll.l'
o .1 .24

Figure III—1.--Effects of $3 vitro passage on phytotoxin production of
the horseradish brittle root isolate of Spiroplasma
citri. Two 3 cm broad bean leaf disks were incubated
overnight at 37 C in the percent culture supernate
indicated. Controls, second row of disks, were incubated
in diluted LD8 medium. Blackened disks indicate toxin

activity.

 

Figure III-2.—-SDS—PAGE analysis of purified membranes of the horseradish
brittle root isolate of §PEI92¥§§EE.Q§EEZ from passages 26
and 34. Resolving gels were 5—20% acrylamide gradients
with a 1 cm 3.75% stacking gel. Run conditions were 15 mA
for ca. 2 hrs followed by 30 mA for ca. 6 hrs. Figures A
and C show results of silver staining for sialoglyc0proteins
while B and D show the same gels, respectively, counter—
stained with Coomassie Brilliant blue. Arrow indicates
band missing in passage 34.

DISCUSSION
These results agree with those of O’Hayer et al. (1983) as to the

relative inefficiency of M. fascifrons in transmitting S. citri. They

 

diverge, however, in that this leafhopper colony is even less efficient
at naturally transmitting the pathogen. This is probably related to
the difference in biotype and/or length of time the leafhopper has been

in culture, over seven years. In any event, M. fascifrons remains a

 

poor vector of S. £12£1°

The pathogenicity of S. 9155; appears to be dependent upon phyto-
toxin activity (Daniels 1979a; 1979b; 1983). If true, then the primary
effect of continuous £2 23359 passage is on transmissibility, rather
than on pathogenicity, as toxin activity was not lost in any of the
passages investigated here. It is likely that the toxins involved are
excreted end products of necessary metabolic reactions, hence, they are
not lost with passage. Several circulative viruses, including wound-
tumor virus, potato yellow-dwarf virus, and pea enation mosaic virus,
have lost transmissibility after prolonged propagation within host
plants without passage through their insect vectors (Liu et al. 1973;
Wolcyrz and Black 1957; Bath and Chapman 1967; Tsai and Bath 1974).
The transmissibility of the corn stunt spirOplasma also declines with
repeated Sgrxiggg passage (Whitcomb and Williamson 1979). It is appar-
ent that transmissibility'is a less stable characteristic of insect
vectored plant pathogens than is pathogenicity.

Microorganisms adapt very rapidly to their environment. Cell-free
culture may present a situation where S. SEEEA must give up some meta-
bolic activity in order to adapt and maximize growth. The cell-free

environment certainly obviates the need to maintain mechanisms of cell

67

68

penetration. The regression analyses supports the idea that continuous
12 ylggg passage results in reduced transmissibility but does not
affect the ability of the spiroplasma to survive within the leafhopper.
It seems reasonable, then, to speculate from these data that continuous
i2_yS££2 passage causes S. SiEEl to lose its ability to infect the
necessary leafhopper tissues that would effect transmission. Because
nontransmissible passages of S. ElEEl were able to survive and probably
prOpagate within the leafhopper, it is likely that the mechanisms pre-
sumably lost involved membrane translocation, i.e., the spiroplasma
could not enter the necessary cells that ultimately lead to transmis-
sion, viz., the salivary acini or other barrier tissues en route.

No first hand evidence was obtained in this research that would
elucidate the leafhopper infection mechanisms used by S. EEEEl’ How-
ever, when considering the possible events involved in cell penetra-
tion, it is obvious that membrane-membrane interactions occur first and
some mechanism of endocytosis must take place. Whether this mechanism
is receptor-mediated, enzymatic, or physical, the host cell must not be
destroyed, especially in the case of salivary acini as living cells are
necessary for eventual secretion of the pathogen. Because S. 21221 has
been visualized in membranous "vesicles" within host tissues (Liu et
al. 1983b), receptor-mediated endocytosis may be involved in transcel-
lular transport. This necessitates a host membrane receptor as well as
a spiroplasma membrane ligand. The loss and reduction of certain mem-
brane proteins in passage 34 of S. 21331 may indicate the loss or re-
duction of a ligand necessary for host membrane binding. The altera-
tion of a soluble protein in a nontransmissible strain of pea enation

mosaic virus relative to a transmissible strain may have represented a

69

ligand necessary for membrane binding (Clarke and Bath 1977). More-
over, the nontransmissible strain apparently could not enter the sali-

vary gland of the pea aphid vector, Acrythosiphon pisum (Harris), after

 

injection into the hemocoel (Harris et a1. 1975). These data are very
similar to those presented here and both support a ligand-receptor
mechanism of membrane translocation. This work certainly has not elim-
inated this possibility and the testing of the receptor-mediated
endocytosis hypothesis for infection of specific leafhopper tissues
merits further investigation.

The differential acquisition of S. citri by M. fascifrons by in-

 

jection.and natural feeding indicates that more than one barrier to
transmission exists. Of all the injected leafhoppers from which spiro-
plasma isolations were attempted, 50% produced cultures. Only 5.4% of
the leafhoppers fed on diseased plants had acquired the spirOplasma.
This speaks strongly for a mesenteronal barrier to S. EEEEE infection
in noncompetent (nontransmitting) leafhoppers. The fact that many in-
jected leathppers were able to maintain viable spiroplasmas indicates
that a post-mesenteronal barrier exists as well, possibly the salivary
acini membranes. In addition, because only one of the naturally
acquiring leafhoppers transmitted the pathogen, the two barriers are
probably mutually exclusive, i.e., the mechanism for penetration of the
one is different from that for the other. This stands to reason as one
would expect the membrane macromolecular profiles of salivary acini and
gut epithelium to be substantially different, necessitating different

or modified mechanisms of penetration.

LITERATURE CITED

Bath, J.E. and R.R. Chapman. 1967. Differential transmission of two
pea enation mosaic virus isolates by the pea aphid, Acyrthosiphon
pisum (Harris). Virology 33: 503-506.

 

Bové, J.M., G. Moutous, C. Saillard, A. Fos, J. Bonfils, J.-C.
Vignault, A. Nhami, M. Abassi, K. Kabbage, B. Hafidi, C. Mouches,
and G. Viennot-Bourgin. 1979. Mise en évidence de SpirOplasma
citri, l'agent causal de la maladie du (stubborn) des agrumes dans
7 cicadelles du Maroc. C.R. Acad. Sci. Ser. D 288: 335-338.

 

 

Bové, J.M., R.F. Whitcomb, and R.E. MCCoy. 1983. Culture techniques

for spiroplasmas from arthropods. Methods in Mycoplasmology 2:
217-223.

Chen, T.A. and C.H. Liao. 1975. Corn stunt spiroplasma: Isolation,
cultivation, and proof of pathogenicity. Science 188: 1015-1017.

Clarke, R.C. and J.E. Bath. 1977. Serological properties of aphid-
transmissible and aphid-nontransmissible pea enation mosaic virus
isolates. Phytopathology 67: 1035-1040.

Daniels, 1M.J. 1979a. .A simple technique for assaying certain
microbial phytotoxins and its application to the study of toxins
produced by Spiroplasma citri. J. Gen. Microbiol. 114: 323-328.

 

Daniels, M.J. 1979b. The pathogenicity of mycoplasmas for plants.
Zbl. Bakt. Hyg., I Abt. Orig. A 245: 184-199.

Daniels, M.J. 1983. Mechanisms of spiroplasma pathogenicity. Ann.
Rev. Phytopathol. 21: 29-43.

Daniels, M.J. and B.M. Meddins. 1974. The pathogenicity of
SpirOplasma citri. Les Myc0plasmes, Colloq. INSERM 33: 195-200.

Doi, Y., M. Teranaka, K. Yora, and H. Asuyama. 1967. Mycoplasma- or
PLT group-like microorganisms found in the phloem elements of
plants infected with mulberry dwarf, potato witches’ broom, aster
yellows, or Paulownia witches' broom. Ann. PhytOpath. Soc. Jpn.
33: 259-266.

Dzandu, J.K., M.E. Deh, D.L. Barratt, and C.E. Wise. 1984. Detection
of erythrocyte membrane proteins, sialoglyc0proteins, and lipids
in the same polyacrylamide gel using a double-staining technique.
Cell Biol. 81: 1733-1737.

Gildow, F.E. 1985. Transcellular transport of barley yellow dwarf
virus into the hemocoel of the aphid vector, Rhopalosiphum padi.
Phytopathology 75: 292-297.

 

70

71

Harris, R.F., J.E. Bath, G. Thottappilly, and C.R. Hooper. 1975. Fate
of pea enation mosaic virus in PEMV-infected pea aphids. Virology
65: 148-162.

Hawthorne, J.D. and P.J. Vandemark. 1977. Metabolic studies of
Spiroplasma citri. Annual Meeting of the American Society for
Microbiology, Abstract 133.

 

Kaloostian, C.H., G.N. Oldfield, E.C. Calavan, and R.L. Blue. 1976.
Leafhoppers transmit citrus stubborn disease to weed host. Calif.
Agric. 30: 4-5.

Kaloostian, C.H., G.N. Oldfield, H.D. Pierce, and E.C. Calavan. 1979.
Spiroplasma citri and its transmission to citrus and other plants
by leafhoppers. Pp. 447-450 in: Leafhopper Vectors and Plant
Disease Agents. K. Maramorosch and R.F. Harris, eds. Academic
Press, New York. 654 pp.

 

Laemmli, U.K. 1970. Cleavage of structural proteins during the
assembly of the head of bacteriophage T4. Nature 227: 680-685.

Lee, I.-M. and R.E. Davis. 1984. New media for rapid growth of
SpirOplasma citri and corn stunt spiroplasma. Phytopathology 74:
84-89 0

 

Liu, H.-Y., D.J. Gumpf, G.N. Oldfield, and E.C. Calavan. 1983a.
Transmission of Spiroplasma citri by Circulifer tenellus.
PhytOpathology 73: 582-585.

 

 

Liu, H.-Y., D.J. Gumpf, G.N. Oldfield, and E.C. Calavan. 1983b. The
relationship of Spiroplasma citri and Circulifer tenellus.
Phytopathology 73: 585-590.

 

 

Liu, H.-Y., I. Kimura, and L.M. Black. 1973. Specific infectivity of
different wound-tumor virus isolates. Virology 51: 320-326.

Lowry, O.H., N.J. Rosebrough, A.L. Farr, and R.J. Randall. 1951.
Protein measurement with the Folin phenol reagent. J. Biol. Chem.
193: 265-275.

Markham, P.G. 1982. The behaviour of spiroplasmas in leafhoppers: A
review. Congress of the International Organization of
Mycoplasmologists, Tokyo.

Markham, P.G., T.B. Clark, and R.F. Whitcomb. 1983. Culture
techniques for spiroplasmas from arthropods. Methods in
Myc0plasmology 2: 217-223.

Markham, P.G. and R. Townsend. 1979. Experimental vectors of
spirOplasmas. Pp. 413-445 in: Leafhopper Vectors and Plant
Disease Agents. K. Maramorosch and K.F. Harris, eds. Academic
Press, New YOrk. 654 pp.

72

Markham, P.G., R. Townsend, M. Bar-Joseph, M.J. Daniels, A. Plaskitt,
and B.M. Meddins. 1974. SpirOplasmas are the causal agents of
citrus little-leaf disease. Ann. Appl. Biol. 78: 49-57.

O'Hayer, K.W., G.A. Schultz, C.E. Eastman, J. Fletcher, and R.M.
Goodman. 1983. Transmission of SpirOplasma citri by the aster
leafhopper Macrosteles fascifrons (HomOptera: Cicadellidae).
Ann. Appl. Biol. 102: 311-318.

 

 

Oldfield, G.N., C.H. Kaloostian, H.D. Pierce, E.C. Calavan, A.L.
Granett, and R.L. Blue. 1976. Beet leafhopper transmits citrus
stubborn disease. Calif. Agric. 30: 15.

Rana, C.L., C.H. Kaloostian, G.N. Oldfield, A.L. Granett, E.C. Calavan,
H.D. Pierce, I.M. Lee, and D.J. Gumpf. 1975. Acquisition of
Spiroplasma citri through membranes by homopterous insects.
Phytopathlogy 65: 1143-1145.

Razin, S., M. Hasin, Z. Ne'eman, and S. Rottem. 1973. Isolation,
chemical composition, and ultrastructural features of the cell
membrane of the myc0plasma-like organism SjiroPlasma citri. J.
Bacteriol. 116: 1421-1435.

 

Tsai, J.H. and J.E. Bath. 1974. The loss of transmissibility of two
pea enation mosaic virus isolates by the pea aphid, Acyrthosiphon
pisum (Harris). Proc. Amer. Phytopathol. Soc. 1: 115-116.

 

Whitcomb, R.F. 1980. The genus Spiroplasma. Ann. Rev. Microbiol. 34:
677-709.

 

Whitcomb, R.F. 1981. The biology of spiroplasmas. Ann. Rev. Entomol.
26: 397-425.

Whitcomb, R.F. and L.M. Black. 1982. Plant and arthrOpod mycoplasmas:
A historical perspective. Pp. 40-81 in: Plant and Insect
Mycoplasma Techniques. M.J. Daniels and P.G. Markham, eds. John
Wiley and Sons, New York. 369 pp.

Whitcomb, R.F. and D.L. Williamson. 1979. Pathogenicity of
mchplasmas for arthrOpods. Zbl. Bakt. Hyg., I. Abt. Orig. A 245:
200-221.

Williamson, D.L. and R.F. Whitcomb. 1975. Plant mycoplasmas: A

cultivable spiroplasma causes corn stunt disease. Science 188:
1018-1020.

Willingham, M.C. and I.H. Pastan. 1983. Receptor-mediated
endocytosis: General considerations and morphological approaches.
Pp. 1-17 in: Receptor-Mediated Endocytosis. P. Cuatrecasas and
T. Roth, eds. Chapman and Hall, New York. 304 pp.

Wolcyrz, S. and L.M. Black. 1957. Origin.of‘vectorless strains of
potato yellow-dwarf virus. Phytopathology 47: 38. (Abstr.).

73

Wroblewski, H. 1978. Spiralin: Its topomolecular anatomy and its
possible function in the SpirOplasma citri cell membrane. Zbl.
Bakt. Hyg., I Abt. Orig. A 241: 179-180. (abstr.).

Wroblewski, H., K.-E. Johansson, and S. Hjertén. 1977. Purification
and characterization of spiralin, the main protein of the
SpirOplasma citri membrane. Biochim. BiOphys. Acta 465: 275-289.

GENERAL CONCLUSION
This research has supplied information as well as posed questions
for future investigations into the nature of insect-plant pathogen in-
teractions. The combination of scanning electron microscopy of leaf-

hopper internal morphology and immunological detection of Spir0plasma

 

gi££l_could.not be accomplished in this work. Finding S. SiEEl in
leafhopper sections necessitates coating specimens with something other
than gold, as gold coating would obliterate the gold label on the spi-
roplasma rendering it undetectable with backscatter analysis. Carbon
and aluminum coatings were used in lieu of gold, but these were not
able to provide enough conductivity to the specimens and much charging
occurred, resulting in poor images in the normal secondary electron
mode. Pursuit of this approach to plant pathogen localization in vec-
tor tissues is a logical and necessary complement to Chapters I and II.
It offers a relatively fast and accurate means of identifying vector
tissues that are barriers to plant pathogen infection in noncompetent
insects. Analysis of these tissues and subsequent comparison to con-
current tissues in competent vectors may provide important information
as to the mechanisms of vector infection and, hopefully, means of
manipulating those mechanisms for disease control.

This project revealed some important information about the mem-
brane of S. ElEEi: including the ubiquity and mobility of membrane an-
tigens and the previously unknown complexity of membrane proteins.
These data indicate that membrane interactions with host cells may be
more sophisticated than is currently thought. Certainly the reduction

and loss of membrane proteins with continuous S2 vitro passage that is

74

75

coincidental with the loss of transmissability is cause to further
probe these phenomena with a view toward elucidating the mechanisms
involved. This will mean working out systems to investigate, e.g., if
ligand-receptor binding occurs or if enzymes exist that attack host
cell membranes in a highly specific manner. I have succeeded in par-
tially purifying a membrane fraction from the mesenteronal tissue of

ScaphytOJius acutus (Say), which was chosen because of its large size.

 

Much work is yet to be done to fully characterize this fraction, but,
once completed, this approach should be applicable to the smaller leaf-
hoppers that transmit S. glgi. 12 m reaction of purified leafhop-
per membranes with those of S. $32.21 may reveal specific protein bind-
ing between the preparations. This is one suggested means of testing a
receptor-ligand hypothesis of pathogen attachment to host membranes.
For success, however, it will be necessary to select a much more effi-

cient vector of S. citri than Macrosteles fascifrons (Stal), viz.,

 

Circulifer tenellus (Baker).

 

Finally, this work has demonstrated that barriers to the transmis-

sion of S. citri do exist in M. fascifrons. Moreover, these barriers

 

are not identical and in combination tend to insure nontransmission.
This may account for the leathppers known to harbor S. Egg natural-
ly, but which lack the ability to transmit the pathogen. If these bar-
riers can be defined and ultimately introduced into competent vector
populations, a new avenue to control of insect-vectored plant pathogens

will be opened .

GQN STQTE UNIV. L

nICHI
ll
3

l I ll l Wlllllllflzl’ll

12931080

llllllr

l.