DEVELOPMENT OF CYTOCHEMICAL METHODS
FOR THE STUDY OF
ASCOSPORE WALL BIOGENESIS AND MATURATION
%
By
DEMARIS E. LUSK
i
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF ^
DOCTOR OF PHILOSOPHY . i
UNIVERSITY OF FLORIDA ;
■ V ■
'1;^
ACKNOWLEDGEMENTS
In retrospect, I find that many people have helped me reach my goal to earn a
Ph.D. During my tenure as a student at the University of Florida I have studied with ail
of my committee members: Dr. Henry C. Aldrich, Dr. James Kimbrough, Dr. Walter S.
Judd, Dr. James F. Preston, and Dr. Dana Griffin. Each has had a hand in my training
and helped to sharpen my mind scientifically.
I have been fortunate to have had several outstanding non-committee mentors.
Foremost of this group is Dr. Greg Erdos (Department of Microbiology and Cell Science)
who put a considerable amount of time into my training in cytochemical techniques.
Dr. Steven Zam (Department of Microbiology Cell Science) and Robin Brigmon
(Department of Environmental Engineering Sciences) were responsible for my training
in hybridoma technology and methodology. Dr. Ross Brown (Department of Food
Science and Human Nutrition) gave me his time for an individualized course on
carbohydrate chemistry. Finally, Dr. William Stern guided me whenever I asked for it,
and helped to edit my work. -'
I appreciate the "hands on" help received from two of my peers: Robin Brigmon
who on occasion cared for my research hybridomas, and Dr. Mary Davis who helped
me to analyze ELISA data statistically.
I have been lucky to have friends with whom I could discuss science, and share
knowledge and from whom I could learn. Most notable of this group are Robin
Brigmon, Katy Gropp D.V.M. (Department of Physiology), Julia Wendt (Department of
Microbiology and Cell Science), Audrey Kalehua (Department of Neuroscience), Chi
Guang Wu (Department of Plant Pathology), and Dr. Wendy Zomlefer (Department of
Botany). Additionally, I wish to thank Gavin Goebel, my sister Vivian Cook, the late
Wendy Knowles, and Pamela Handley for their friendship, love, support and
encouragement.
I received two very important gifts for use in my research: a culture of
Ascodesmis sphaerospora from Dr. Kimbrough, and GS-II lectin-gold from Katy Gropp.
My research was supported by a grant from the Gas Research Institute, the University
of Florida Interdisciplinary Center for Biotechnology Research Electron Microscopy Core
Facility, and the Univerisity of Florida Department of Microbiology and Cell Science.
Ill
I^^'"
TABLE OF CONTENTS
ACKNOWLEDGMENTS I .• ii
LIST OF TABLES vi
LIST OF FIGURES vii
'\ ABSTRACT jx
CHAPTERS
1 INTRODUCTION 1
"i?; ; Ascospores 1
Ascosporogenesis 3
Chemistry of the Ascospore Wall 13
Chemistry of the Hyphal Wall 14
Study Proposal 29
Conclusions 31
2 STRATEGIES FOR TISSUE PREPARATION AND EMBEDDING 33
Literature Review 33
Tissue Preparation Strategies 40
3 DEVELOPMENT OF ANTIBODIES 44
Introduction 44
Materials and Methods 50
Results 54
Discussion 58
4 IMMUNOCYTOCHEMISTRY 61
Introduction 61
Materials and Methods 63
Results 68
Discussion 86
IV
'■^i-.
5 LECTIN CYTOCKPMISTRY 90
Introduction 90
Binding Specificities 91
Materials and Methods 96
Results 98
Discussion 116
6 CONCLUSIONS 119
Evaluation of Experimental Methods 119
Ascospore Wall Chemistry 121
Precursor Tracking 122
Maturation of the Ascospore Wall 122
APPENDICES 124
A FUNGAL CULTURE 124
B ISOLATION OF ASCOSPORE WALLS 125
C PROTOCOLS FOR HYBRIDOMA CONSTRUCTION AND CLONING 127
D FREEZING AND THAWING HYBRIDOMA OR SP2-0 CELLS 130
E LIGHT BREAK OF ASCOSPORES 132
F FIXATION PROTOCOL 134
G ANTIBODY LABELING 135
LITERATURE CITED 137
BIOGRAPHICAL SKETCH 153
■4
-A
i
LIST OF TABLES
Table 3.1. Mean optical densities for buffer wash, substrate, and secondary
antibody reatments.
56
Table 3.2. Least square means comparison of washing vs substrate and
antibody / substrate treatment.
57
Table 3.3. Least square means comparison of interaction of buffer type with
treatments.
57
Table 3.4. Mean optical densities for buffer control, immune mouse and test
mouse sera.
58
Table 4.1 . Tissue preparation and embedding 66
Table 5.1 . List of lectins and labeling protocol information 97
Table 5.2. Comparison of Con A and GS-II labeling on A. sphaerospora and
binding specificities.
117
Table 6.1. Comparison of wall labeling patterns 121
VI
♦•
■9:
-it'
LIST OF FIGURES
Figure 4.1. Serum labeling on A. sphaerospora 71
Figure 4.2. Serum labeling and buffer control on A. sphaerospora 72
Figure 4.3. 8F1 1 culture supernatant labeling on A. sphaerospora 72
Figure 4.4. Collage of 8F11 positive labeling 74
Figure 4.5. Determinant characterization for 8F1 1 '. 75
Figure 4.6. Developmental sequence with 8F1 1 labeling 76
Figure 4.7. Antibody 41 -1.1 labeling 79
■^; Figure 4.8. Pronase pretreatment with antibodies 1 2-2 and 41 -1 .1 80
■; . Figure 4.9. Antibody 12-2 labeling 83
Figure 4.1 0. Anti-A. sphaerospora ascospore wall antibody lableing on P. nigrella 85
Figure 5.1 . WGA labeling on A. sphaerospora 98
10 \. Figure 5.2. GS-II labeling on A. sphaerospora 100
/ :' , Figure 5.3. WGA labeling on P. nigrella 101
Figure 5.4. GS-II labeling on P. nigrella 102
Figure 5.5. WGA labeling with sugar control 103
X Figure5.6. LFA labeling on A. sphaerospora 104
- ■- Figure 5.7. LFA labeling on or around spent cells of A. sphaerospora 1 06
Figure 5.8. Con A labeling on A. sphaerospora 109
Figure 5.9. Con A labeling on P. nigrella Ill
, vli .
Figure 5.10. Con A labeling witha-mannosidase and/or pronase pretreatments 113
Figure 5.1 1 . Collage of lectin labelings on A. sphaerospora 1 1 5
VIII
Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
DEVELOPMENT OF CYTOCHEMICAL METHODS
FOR THE STUDY OF
ASCOSPORE WALL BIOGENESIS AND MATURATION
by
DEMARIS E. LUSK
August 1 991
Chairman: Dr. Henry 0. Aldrich
Major Department: Botany
Detailed morphological studies of the process of ascosporogenesis have been
well documented for several species of Ascomycetes. Biogenesis of ascospore wall
appears to be a de novo process which occurs between two unit membranes that
delimit the ascospore. Although morphological studies have provided a tremendous
amount of information about the process, neither biogenesis nor chemical maturation
events of these walls can be more than implied via morphology. The goals of this
project were to develop cytochemical methods with TEM detection, to improve our
understanding of ascospore wall biochemistry and developmental biology, and to
provide a foundation of information upon which further studies could be based.
One hybridoma-derived uncloned and two monoclonal antibody preparations
against mature ascospore walls of Ascodesmis sphaerospora were developed. The
ix
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•V. .• ,'
uncloned preparation, 8F1 1 , has demonstrated a late maturation event by highly specific
labeling of the primary wall layer in what appear to be very mature spores. Such an
event has never before been demonstrated for ascospores. Monoclonal antibody 41 -1 .1
clearly demonstrated the presence of a pronase sensitive antigen in the inner region of
the primary wall. Monoclonal antibody 12-2 identified an ascospore secondary wall
constituent and a cytoplasmic component.
The chemistry of ascospore walls was demonstrated as distinct from vegetative
cell walls by the labeling pattern of these antibodies and lectins tested. Con A labeled
primary and secondary spore walls and cytoplasmic components. WGA and GS-II
lectins labeled ascus walls and vegetative cell walls. GS-II also labeled a cytoplasmic,
electron-transparent component. LFA lectin was most specific for an external layer of
dead or interacting cells.
The results of this research provide an excellent springboard for further
developmental biology, biochemical/molecular structure, and fungal systematics
research. Although a probe capable of tracking wall material was not found, these
results are encouraging. Isolation and chemical analysis of antigens discovered here
would provide insight into the biochemistry of these walls. Morphometric analysis of
labeling could provide information about the wall molecular structure. Application of the
antibodies and lectins shown here to be cross-reactive with Pseudoplectania nigrella
could provide data for use in systematic research of the Pezizales.
.'■/i
CHAPTER 1
INTRODUCTION
Ascospores
Ascospores are the end product of the meiotic process in Ascomycete fungi.
The formation of ascospores has been described as epitomizing the sexual behavior
pattern diagnostic of the Ascomycotina (Beckett, 1981). Ascospores are important for
dissemination and survival in those species that produce them. As a propagule, they
are the sole mechanism for aerial dispersal for those species that do not produce
conidia. Airborne fungal spores, especially conidia, along with pollen and dust mites
are the most common and potent allergens. Additionally, characteristics of the ><,
ascospore have been used by taxonomists since the earliest ascomycete studies.
Despite the important position held in the life cycle, the importance of the biological , -, -■'■y
roles, the significance to human health, and the usefulness of their characteristics for
...» , .»
taxonomic work, ascospores have received considerably less research attention than
hyphae and conidia, especially in the areas of biochemistry and biogenesis.
Early light microscopy studies have provided hypotheses of ascospore origin
and development. Of these hypotheses, Harper's (1 897) "free-cell formation" has been
shown via ultrastructural studies to be correct. Studies of mature ascospore
morphology via light microscopy (e.g., LeGal, 1 947, 1 951 ) have been a starting point for 1
electron microscopy studies, as well as having provided invaluable data for taxonomic
application. In the area of cell biology, ascospore ontogeny (ascosporogenesis) has
i
interested researchers because it represents an unusual type of cytol<inesis and cell
differentiation. ,
Ultrastructural studies of ascosporogenesis came into vogue around the early I
1 960's, apparently culminated in Beckett's (1 981 ) synthesis on the subject, and continue
today. In that paper Beckett stated that the origin of wall materials, either as precursor
or assembled units, was unclear. Much more is known about chemistry and synthetic
machinery for hyphal wall constituents than for ascospore wall constituents. Synthesis
of hyphal wall components can occur either in situ or within the cell. In addition to
those potential synthesis sites, synthesis of ascospore wall components may also occur ^ — ?
in the ascus cytoplasm (epiplasm) that surrounds the developing spore. There is some
morphological evidence suggesting that that could in fact be the case for some wall
components. '■*
The very fact that ascospores are formed by the "free-cell" process within a ,^
walled cell makes them difficult to access for biochemical and biosynthesis study during
development. Isolation of ascospores at various stages of development for these types
of studies would be a more feasible pursuit if the ratio of spores per ascus, which is
•i
typically 8:1, were greater. Yet, the availability of ascospores for post-embedding
ultrastructure study has been demonstrated over and over again in the many
publications on the morphological aspects of ascosporogenesis. Following the trend
of post-embedding TEM study, and additionally testing various cytochemical reagents
and techniques, could provide the most expedient route to valuable chemical and
biogenesis information.
Just such a post-embedding cytochemical route has been pursued in this study.
The application of these techniques has provided data about the chemistry and biology .^
3
of ascospores that will improve our understanding of the biological processes involved.
This work has also provided a foundation of data upon which further work in the areas
of cell biology, developmental biology, and systematics can be based.
Ascosporoqenesis
Introduction
In general, for ascomycetes, the process of meiotic spore (ascospore)
production follows a pattern. First there is production of a dikaryotic ascus initial from
ascogenous hyphae, karyogamy, meiosis with concurrent production of spore delimiting
membranes (SDMs) often in the form of an ascus vesicle (Reeves, 1967; Carroll, 1967;
Rosing, 1982; Mims et al., 1990), then a post-meiotic mitotic nuclear division producing
8(1 n) nuclei, next an envelopment of the 1 n nuclei via constriction of the ascus vesicle
or otherwise, and finally ascospore wall formation between the SDMs concurrent with
maturation of the sporoplasm and vacuolation of the ascus cytoplasm (epiplasm).
Additionally, there may be further mitotic divisions of the 1 n nuclei after the intial 8 nuclei
have been delimited (Gibson & Kimbrough, 1988a, 1988b; Kimbrough et al., 1990). In
Beckett's (1981) review of ascospore formation literature, he stated that there are two
points of universal agreement amongst ascospore studies; those are "1) Nucleate
portions of the cytoplasm are delimited by an envelope of 2 unit membranes. 2)
Ascospore wall material is deposited between these 2 unit membranes which separate
as the spore matures."
Wall Nomenclature and Pattern of Wall Development
Once the spore delimiting membranes are in place around the nuclei, a wall
4
forms between them. The outer membrane is displaced from the spore plasma
membrane as the wall develops (Mims et al., 1990). No standardization of wall layer
terminology was proposed until Merkus (1973) and Beckett (1981) made efforts to that
effect. The wall material is laid down in two stages; in the first stage the primary wall
is laid down, and in the second stage a second layer is sometimes formed between the
primary wall and the outer delimiting membrane (e.g., Merkus, 1973; Gibson &
Kimbrough, 1988a, b; Kimbrough et al., 1990). Hohl and Streit (1975) did not find this
order of stepwise development in the wall of Neurospora lineolata. They found that after
the primary wall was laid down a secondary wall was formed to the inside, between the
primary wall and the spore plasma membrane (or inner delimiting membrane). This
seems to be the less common of the two methods. In both cases the second layer
deposited was called the secondary wall (Merkus, 1 973; Hohl & Streit, 1 975). The outer
delimiting membrane loosens, forming the so called "perisporal sac" prior to deposition
of secondary wall material.
Typically, periclinal (parallel to the wall inner surface) bands between the primary
and secondary wall layers are evident in micrographs of mature spores. Merkus (1973)
called these bands the epispore wall. Merkus' wall nomenclature seems complete,
clear, adequately descriptive, and efficient. Beckett (1981), in an effort to reduce the
confusion of wall nomenclature added to the confusion by defining the secondary wall
as "all subsequent wall material that is formed, either by modification of the primary wall
or by addition to it. . . ." It is clear that his secondary wall includes Merkus' epispore
wall. Merkus' wall nomenclature as just described is used through this work.
5 ,
Pre-wall Formation Events ' '^
Although some authors discuss crozier formation and development of the ascus
initial (e.g., Reeves, 1967; Zickler & Simonet, 1980; Rosing, 1982) and a few discuss §^
karyogamy as part of ascospore ontogeny (Leung & Williams, 1976), it is much easier ,:
to find information on the post-nuclear fusion processes of ascospore ontogeny (e.g.,
Carroll, 1967; Hohl & Streit, 1975; Merkus, 1976; Dyby & Kimbrough, 1987). At the
electron microscope level, crozier development as described for Chaetomium brasiliense ,
(Rosing, 1982), Myxotrichum deflexum (Rosing, 1985), and Pyronema domesticum :|
(Reeves, 1967) follows the stylized description usually taught as basic. Reeves (1967)
found young asci of P. domesticum with fusion nuclei to be rich in "long strands of
endoplasmic reticulum" and that basal vacuolation had been initiated. Leung & Williams
(1976) have provided a detailed description of the meiotic and post-meiotic mitotic ■ •'^
divisions in the asci of Pyricularia oryzae. There seem to be no striking abnormalities
in those divisions as described. Unfortunately, they gave no information on other ,;
cellular activities that occur simultaneously with those divisions. Zickler and Simonet
(1980) found in their experiments with sporulation deficient mutants of Podospora
anserina that any disturbance in the strict orientation of post-meiotic mitosis spindles
leads to irregular distribution of nuclei and afterward in the distribution of the ascospore.
They further stated that such disturbance is often associated with variation in the final
number of ascospores formed.
Over the years many suggestions for the origin of spore delimiting membranes
have been made. Beckett (1981) stated that all of the proposed methods of SDM
formation could be accommodated by the endo-membrane concept of Morre,
Mollenhauer, and Bracker (1971). In that same paper Beckett provided a table that
■■•-I
6
summarized the research to that date on the origin of these membranes. Structures
Implicated in the formation of SDIVIs include: the ascus plasma membrane, mesosomes,
myelin figures, cisternae of endoplasmic reticulum, endomembrane vesicles, and the >; :<
nuclear envelope.
Ascospore initiation appears to be dependent on the position of spore delimiting
membrane in relation to the haploid ascus nuclei. Two fundamentally different patterns
of spore initiation have been found. With a few exceptions, the Hemiascomycetes are
characterized by the direct envelopment of individual nuclei by membranes formed in
association with spindle pole bodies, while Euascomycetes form a discontinuous
membrane cylinder around the periphery of the ascus. This cylinder has two layers of
unit membrane and is the ascus vesicle. Zickler and Simonet (1980) observed ascus
vesicle formation in Podospora asernia even in the absence of live nuclei and concluded
that initiation and formation of ascus vesicles were independent of the nuclear divisions
and/or presence of spindle plaques. The ascus vesicle invaginates, giving rise to the
spore delimiting membrane. Typically, each nucleus present, and the adjacent
cytoplasm, are surrounded by the delimiting membrane and develop into ascospores. ,.
During these early phases of ascospore ontogeny changes in the epiplasm have
also been observed. In many Pezizalian fungi, Merkus (1975, 1976) noted the
development of "globular structures." Formation of these globular structures can begin
in the pre-meiotic ascus and continue through the spore delimiting stages. She
speculated that these are food reserves and stated that they do not seem to play a role
in the formation of walls (Merkus, 1975). A system of vacuoles at the ascus bases ''
exists, and an apical system of vacuoles begins forming around the period of the
second meiotic division (Reeves, 1 967). Niyo and coworkers (1 986) noted the presence -^
7
of vacuoles, ER, and lipid bodies in young asci. Microtubules are often noted in asci
before ascospore delimitation (Reeves, 1967; Beckett, 1981; Rosing, 1982, 1985; Dyby
& Kimbrough, 1987).
The Primary Wall
The primary wall formation is first evident in micrographs by a slight separation
between the inner and outer delimiting membranes. The delimiting membranes remain
appressed to the developing primary wall until it has apparently completed the
biosynthesis process (Merkus, 1973, 1974, 1975, 1976; Kimbrough & Gibson, 1990).
At the time of apparent completion, the outer delimiting membrane loosens signaling the
onset of secondary wall formation.
The appearance of the primary wall in electron micrographs is typically described
as electron-translucent or electron-transparent. In recent studies of mature spores (with
evident secondary wall and epispore layers), the primary walls have been shown to have
some electron-density (Kimbrough et al., 1990). Fibrillar orientation of the primary wall
in at least the case of Geopvxis carbonaria has been observed and described as
anticlinal (perpendicular to the wall inner surface) (Kimbrough & Gibson, 1990).
The exact origin of the primary wall material of ascospores remains to be
determined, and probably varies amongst the taxa. Reeves (1 967) and Rosing (1 982)
suggested ER that lies in close proximity to the spore plasma membrane as a
possibility. Rosing, in that same article, noted the appearance, increase in number, and
fusion of dark granules between the spore delimiting membranes in Chaetomium
brasiliense and suggested that those membranes actually synthesized the granules.
Merkus (1 976) suggested that spore plasma membrane plays a role in the synthesis of
8
primary wall material. These suggestions are supported by Beckett's (1 981 ) concluding
remark that "both spore plasm membrane and investing membrane play a role in
regulating wall formation in young ascospore initials." In Merkus' ascospore wall studies
(1973, 1974, 1975, 1976), she ruled out dictyosomes as a source and minimized the role
of lomasomes as a source of wall material. Reeves (1 967) found few lomasomes in
Pvronema domesticum asci. This seems to agree with Merkus' minimization. It has
additionally been suggested that small vacuoles originating in the sporoplasm may be
involved with primary wall synthesis (Kimbrough & Gibson, 1990; Wu & Kimbrough,
1991a, 1991b).
Merkus (1973, 1975) found that the dimensions of the primary wall layer varied
depending upon the fixative used. In further regard to ascospore shape she stated that
"the ascospores are rounded off before the primary wall is formed" in some species and
that "the ascospores round off during primary wall development" in other species. More
recently the primary wall of Gyromitra esculenta was said to confer the characteristic
shape seen in mature spores (Gibson & Kimbrough, 1 988b). It is unfortunate that these
types of data are not always noted because these data could be a useful taxonomic
characteristic. On the other hand, this type of deformation could be artifactural and
freeze fixation studies should preceed use of these data for taxonomy.
The Secondarv Wall
The loosening of the outer delimiting membrane forming the perisporal sac
signals the onset of secondary wall formation. In a few cases secondary wall formation
has been found to begin prior to completion of the primary wall (Merkus, 1976). With
few exceptions (e.g., Neurospora lineolata, Hohl & Streit, 1975), the secondary wall ,i|
,j(-v V..,;
•^
%
<
9
forms to the outside of the primary wall, between that wall layer and the outer delimiting
membrane. Undifferentiated material, sometimes described as fibrillar (Gibson &
Kimbrough, 1988b) or floccose (Kimbrough & Gibson, 1990), apparently accumulates
in the perisporal sac and then condenses onto the primary or epispore wall layer to form
the secondary wall (Merkus, 1 976). Beckett (1 981 ) concluded that "there is no common
pattern of development for the secondary wall formation." Certainly with the diversity
of ascospore ornamentation found in members of this class there can be little doubt to
the truth in that statement. Merkus (1 976) outlines seven different developmental groups
within the Pezizales alone. ' '
i ^ • t
Synthesis of secondary wall materials, either as precursor or final macromolecule,
could occur within the sporoplasm or at the spore plasma membrane, at the outer
delimiting membrane, or in the epiplasm. Gibson & Kimbrough (1988a) supposed
sporoplasm to be the primary or only source of material for the secondary wall. In
support of this supposition they argued that the epiplasm is isolated from the
developing wall whereas the sporoplasm remains in close contact. Despite this
argument, the sporoplasm as the sole or even primary source of secondary wall material
seems unlikely as these materials would have to traverse the existing primary wall.
Merkus (1976) felt it was unlikely that the sporoplasm has a function in secondary wall
'"■'4
formation and that the investing membrane might have an active role. Based on r"*)}
I
structural similarities of components within the epiplasm and the secondary wall, Merkus
(1976) found it highly probable that parts of the epiplasm are incorporated into the
secondary wall. Wu and Kimbrough (1 991 a, 1 991 b) provided morphological evidence
for diffusion, or movement otherwise, of materials from the epiplasm into the perisporal
sac. They proposed that these materials are involved in wall formation. Bellemer and
,*a
1
10
Melendez-Howell (1976) also suggested an active role on the part of the epiplasm.
Mechanisms controlling deposition and structure of the ascospore wall seem
unclear. Beckett (1981) presumed that the spore nucleus plays a major role in
controlling wall deposition and architecture. While the spore nucleus undoubtedly plays
such a role for the deposition of primary wall, and for secondary wall formed interior to
the primary wall (e.g., Neurospora lineolata), the situation is less clear for those fungi
in which the secondary wall forms to the outside of the primary wall. The question of
control of secondary wall formation is especially interesting as final form of
ornamentation is 1) due to this external wall layer and 2) is often diagnostic at the
species or genus level of fungi within the Pezizales.
The Epispore Wall
The epispore wall as seen in micrographs appears to be a periclinal band, or
more typically bands, of varing electron density. It is located between the primary and ;
secondary wall layers.
The timing of differentiation and source of material differentiated into this wall
layer are apparently variable. Differentiation of this layer could commence 1 ) after the
primary wall is formed but before secondary wall deposition begins, 2) during the
deposition of secondary wall material, or 3) after the secondary wall is complete.
Kimbrough & Gibson (1 988a) reported development of the epispore layer prior to the
deposition of secondary wall material for Helvella acetabulum. The bulk of available
data supports the differentiation of epispore layer(s) during deposition of secondary wall -, ,•
. .V-*c J
materials (in H. macropus and H. elastica, Gibson & Kimbrough, 1988a; in Gyromitra
esculenta, Gibson & Kimbrough, 1988b; Dyby & Kimbrough, 1987; in Geopyxis ,
ty-.
11
carbonaria. Kimbrough & Gibson, 1990; Kimbroughetal., 1990; in Ascobolus immersus.
and A. stictoldeus, Wu & Kimbrough, 1991a & b). No reports reviewed suggested that
differentiation of the epispore wall layer commenced after complete formation of the
secondary wall. It is obvious from the figures in the reviewed articles that differentiation
of the epispore layer(s) continues through the development of the secondary wall layer
and that epispore differentiation may not be complete until after the secondary wall is
apparently fully formed and mature.
Epispore wall constituents could be derived from the primary wall, secondary
wall, or laid down as new material prior to the deposition of secondary wall material.
Merkus (1975, 1976) described the primary wall as differentiating into the epispore and
an endospore layer. While it is clear that she felt the parent material of the epispore to
be primary wall constituents as originally formed, more recent articles present evidence
for synthesis of new materials for this layer (in Helvella acetabulum, Gibson &
Kimbrough, 1 988a; in Ascobolus immersus, and A. stictoideus, Wu & Kimbrough 1 991 a
& b). In Helvella macropus the epispore was described as being evident soon after
secondary wall deposition began, and in H. elastica it was evident at the time
secondary wall deposition is evident (Gibson & Kimbrough, 1988a). Morphological
evidence is insufficient to determine the derivation of epispore wall materials. It is
possible that the epispore wall is an amalgamation of secondary and primary wall
material modified in situ by enzymes present in the wall or perisporal sac.
Ascopore Maturation
The appearance of ascospores during the initial phases of development is
distinctly different from that of a mature spore. This is the case for both sporoplasm and
M
12
the spore wall. In electron micrographs the sporoplasm is initially packed with
cytoplasmic components, such as mitochondria and ribosomes (e.g., Dyby &
Kimbrough, 1987; Kimbrough et a!., 1990), that are indicative of high levels of activity.
At the time they are delimited, ascospores are typically uninucleate. In the Helvellaceae
further mitotic nuclear divisions occur so that the mature spore is multinucleate (Gibson
& Kimbrough, 1988a, 1988b). Sometimes lipid droplets develop or coalesce during the
development and maturation processes (e.g., Gibson & Kimbrough, 1988a, 1988b;
Kimbrough et al., 1990). At about the time the epispore wall layers are forming, the
sporoplasm appears to condense (Kimbrough et al., 1990). Most strikingly, the
membranes of the sporoplasm take on a negative appearance as compared to earlier
stages. In what appears to be the most mature spore state while still in the ascus, the
sporoplasm is typically missing from the section. This most probably indicates poor
infiltration and/or polymerization of the resin. This problem is likely to be the result of
changes in the spore wall that seal it from the external environment.
The primary and secondary wall layers are apparently constructed sequentially.
The primary wall as it appears at the time the perisporal sac forms has been called
mature (Gibson & Kimbrough, 1988b). This wall layer would not actually be mature if
in fact the primary wall undergoes further change before the spore is expelled. Merkus'
(1976) hypothesis regarding the differentiation of epispore wall from the primary wall
Implied such immaturity of the primary wall. Slight staining differences observed ■,
(Kimbrough et al., 1990) between the just formed primary wall and the primary wall of
mature spores are also suggestive of post-formation changes within this wall layer.
The appearence of the epispore wall changes from a single layer to several
layers in the most mature spores observed (e.g., Gibson & Kimbrough, 1988a;
Kimbrough et al., 1990).
!!
13
The appearance of the secondary wall changes as deposition progresses,
especially when ornaments are formed. Sometimes, as the secondary wall develops
differential staining of fibrillar material occurs (Kimbrough et al., 1990) and/or electron-
translucent lacunae (Kimbrough et al., 1 990; Kimbrough & Gibson, 1 990) are evident in |
developing ornaments and wall thickenings. J
In conclusion, possible maturation processes within the wall could produce 1)
a change in the staining properties of the primary wall, 2) layering of the epispore, 3)
"^
iihy
differential staining of the secondary wall, and 4) a change in the permeability of the M
.-Cm
wall. -'n
Chemistrv of Ascospore Wall
Very little information on the chemical composition, or nature, of ascospore wall . j /^
layers is available. The only information found on the chemistry of ascospores of non-
yeast species is based on cytochemical experimentation primarily with silver periodate
stain. Silver proteinate stain demonstrates the presence of periodate sensitive
carbohydrates. Periodate sensitive carbohydrates are those that possess residues with
vicinal diols. Glucans, mannans, and galactans with (1-3) linkages are insensitive to
periodate and will not stain with with the silver proteinate staining procedure. Likewise
chitin, because of the N-acetyl substitution on carbon 2, is insensitive to silver periodate.
Sensitive pyranosyls would have (1 -4) or (1 -6) linkages and be unsubstituted. Based
on the negative results of silver proteinate staining experiments, Dyby & Kimbrough
(1987) concluded that the primary wall of those fungi studied (Peziza spp.) is primarily
composed of (1-3) glucan rather than chitin or other polysaccharides. Similar staining
and conclusions were drawn for Geopyxis carbonaria (Kimbrough & Gibson, 1990). "" ;
Gibson and Kimbrough found the prinnary walls of Gyromitra esculenta (1988b) and
Helvella spp. (1 988a) to have some affinity for silver proteinate and they suggested the
presence of chitin. These conclusions are incorrect in being both more specific than,
and at variance with known carbohydrate sensitivities for periodate. "^
The outer edge of the secondary wall of Peziza spp., and the inner band of the
epispore wall stained positively with silver proteinate (Dyby and Kimbrough, 1 987). They
speculated that the secondary wall ornaments consisted of lipids, protein, glycoprotein, |
and possibly chitin. In Geopyxis carbonaria (Kimbrough & Gibson. 1990) and Gyromitra -j
esculenta (Gibson & Kimbrough, 1988b), there was no evident staining by silver
proteinate in the secondary wall layers. Merkus (1973) felt that the secondary wall was - ^ , ^.^
" *'r 'J
formed via deposition of membranous fragments in a homogeneous matrix. - ' -^ >
No work specifically on biochemistry of non-yeast ascospores appeared in a
recent text on the subject of fungal wall biochemistry (Kuhn et al., 1990). A great deal "|
more is known about hyphal walls than ascospore walls. Research on the structure,
biochemistry, synthesis, and even genetics of hyphal walls is available. './^
Chemistry of the Hvphal Wall
Functions of the Fungal Wall
The cell walls of fungi function in every aspect of fungal life. Fungal morphology can
vary to meet functional needs by a change in wall construction (Bartnicki-Garcia, 1 968).
These cell walls provide a structural barrier that is resistant to lysis by competing
microflora or host defenses, prevents disruption of the protoplast by free water, and
maintains cellular form. A variety of enzymes have been found in hyphal walls. The
^
'■/J5
15
walls are the site of recognition systems (e.g., self-self and self-host) and mediate
adherence. They undoubtedly help prevent desiccation, but may additionally act as a
filter and ion exchanger (Reiss, 1 986). The many functional aspects and dynamic nature
of cell walls have prompted some researchers to recognize cell walls as organelles
(Mauseth, 1988).
Wall Chemistry
Hyphal walls have been reported to be 80%-90% polysaccharide (Farkas, 1 979;
Zonneveld, 1971; Bartnicki-Garcia, 1968). This characteristic is in common with gram-
positive bacterial and plant cell walls (Peberdy, 1990). Various glucans (Wessels, 1986;
Zonneveld, 1971), chitin (Wessels, 1986; Bartnicki-Garcia, 1968), chitosan (Mol &
Wessels, 1987), other homo- and heterpolysaccharides, glycoproteins (Gorin, 1985;
Johnston, 1965), and peptido-polysaccharides (Gander, 1974) make up the
carbohydrate fraction of hyphal walls. These polysaccharides are composed of amino
sugars, hexoses, hexuronic acids, methyl pentoses, and pentoses (Farkas, 1979).
Bartnicki-Garcia (1 968) stated that "at least 1 1 monosaccharides" are reported to occur
in hyphal walls; but only D-glucose, N-acetyl-glucosamine, D-mannose, D-galactose and
D-galactosamine are consistently found in the Ascomycetes, with the latter two sugars
being more-or-less characteristic of this class of fungi. On the basis of their presumed
function and physical form, cell wall components can be divided into two major
categories: skeletal and matrix. Additionally, a gel-like (or glycocalyx) layer surrounding
hyphae has been described (Wessels, 1986).
Skeletal elements are crystalline or microfibrillar in form, and consist primarily of
chitin, and/or crystalline beta-glucans (6(1-3) linked homopolymer; Farkas, 1979). It is
"t'l
16
important to note that some researchers report protein (s) as always being associated
with chitin, and further, that this association is in a regular or crystalline fashion (Neville,
1975; Blackwell, 1982). However, Rudall (1969), on whose information Neville and
Blackwell base this stated association, reported a protein-chitin association for the
crystalline chitin of crustacean, insect, and spider cuticles, but that glucan(s) of 6(1-3)
and 6(1-6) linkages are the principal protein-associated substance(s) in fungi. Glucans
of these same linkages, although with a higher degree of 6(1-6) branching, probably
make up the gel-like layer that surrounds the hyphae (Wessels, 1986; Peberdy, 1990). (<*
The matrix is then the remainder of wall components; amorphous homo- and 4
Ml
heteropolysaccharides, glyco-conjugates, proteins, and lipids or lipo-conjugates.
i
Survey of Methods i
Current knowledge about the architecture and chemistry of hyphal walls is 1
founded in three basic research methods. These are 1) degradation (extraction, "H
digestion) followed by chemical analysis and/or shadow casting TEM of the surface, 2)
localization via cytochemistry and transmission electron microscopy techniques, and 3)
immunological studies. Additionally, morphological studies, especially those examining ^
changes associated with altered nutritional or environmental conditions, have
contributed to current understanding of these walls.
Degradation of walls appears to be accomplished most often by chemical (alkali,
acid, etc.) extraction, but some investigators report the use of enzyme digestions. In ^
- i
general, after wall isolation and any desired preparatory steps (e.g., treatment with ^
boiling diethyl ether then diethyl ether-ethanol-HCI for removal of lipids, Zonneveld,
1971 ; or treatment with hot phenol and water, 9:1 v/v, for removal of RNA and protein . -^
17
impurities, Johnston, 1 965), chemical extractions begin with hot water or/then mild alkali
(e.g., 5% KOH), followed by acid hydrolysis of the soluble fractlon(s). More severe alkali
and acid treatments are then applied to the initially insoluble residue to further
fractionate the wall components. Between each step there is commonly a separation
of supernatant from residue and wash(es) of the residue.
A major portion of fungal cell walls are soluble in hot water, phenol, and/or alkali.
At least two fungal polysaccharides, lichenin (a 6(1 -4) and 6(1 -3) linked glucopyranose
polymer) and nigeran (a glucopyranose polymer with alternating a (1-3) and a (1-4)
linkages) are soluble in hot water. The latter is partially characterized by its solubility
in water according to Aronson (1981). Wessels (1986) describes glucans with 6(1-3)
and 6(1-6) as being "more or less" soluble in water. Hearn and coworkers (1989)
studied only the water soluble fraction of Aspergillus fumigatus mycelia (including
cytoplasm) and found predominantly galactomannans and glucans.
Cell wall outer layers are "as a rule" soluble in dilute alkali according to Wessels
(1986). Often extraction procedures begin with alkali, or with hot water, as pointed out
previously. Some glucans are soluble in dilute alkali but not in hot water. The
differences in glucan structure associated with hot water solubility /insolubility appear to
be slight. For example, pseudonigeran (a glucopyranose polymer of consecutive a (1 -3)
with interspersed a (1-4) linkages) is not soluble in hot water but is soluble in alkali,
whereas nigeran (a glucopyranose polymer of alternating a (1 -3) and a (1 -4) linkages) is
characterized by its water solubility (Gorin, & Spencer, 1968). Additionally, Wessels
(1986) pointed out that water soluble 6-(1 -3)-6-(1 -6) glucans have longer (1-6)-6-linked
branches than those that are water-insoluble/alkali-soluble, although some of these
glucans remain insoluble under either of these conditions. In their comparison of
I
i .<
' '^^
A
18
polysaccharides obtained from water extraction and those of alkali extraction, Hearn and
coworkers (1989) reported "marked differences in the contents of non-reducing end-
units of or -D-Man(p) and 6-D-Gal(f)." These differences are primarily number of units per
side chain.
Mol and Wessels (1 987) described "most" yeast wall fractionations as beginning
with a "rigorous" alkali step to remove mannans and proteins. Zonneveld (1971) found
a considerable portion of the wall (22% dry weight of complete wall) in this fraction.
Galactomannans (e.g., Gorin, 1985) and other heteropolysacchrides (e.g., Johnston,
1965) and glycoprotein conjugates (e.g., Mahadevan & latum, 1967) are commonly
found in both alkali and water (Hearn et al., 1989) fractions. Acid hydrolysis is the final
step before quantitative analysis of either of these fractions. Zonneveld (1 971 ) used 2%
hydrochloric acid at 100°C for an hour to hydrolyze these fractions. Mahadevan and
Tatum (1965) initially used 3N hydrochloric acid to hydrolyze the carbohydrates, then
did a second treatment with 6N hydrochloric acid to hydrolyze proteins.
Treatment of the alkaline-insoluble fraction with hydrochloric or sulfuric acid
(e.g., 40% HgSO, (v/v) at 4°C for 18 hr, then diluted and boiled 3 hr) is thought to
hydrolyze all the remaining glycosidic bonds, except chitin, leaving chitin as a final
residue (Zonneveld, 1 971 ). Nitrous acid is also commonly used. It is said to specifically
attack non-acetylated glucosamine residues and depolymerize glucosamine-containing
polymers (Stagg & Feather, 1973; Mol & Wessels, 1987; Davis & Bartnicki-Garcia, 1984).
Enzymes have been useful in carbohydrate degradation/dissection of whole
walls, and/or wall fractions for component analysis, elucidation of glycosidic bond type,
and localization. Mahadevan and Tatum (1965) used crude enzyme complexes from
snail gut (known to contain chitinase, carbohydrases, and proteases) and Aspergillus
1
•*S
^j
M
19
niger (known to contain cellulase) for degradation of cell walls and various wall fractions
produced by chemical treatment. These results were then compared with the chemical
hydrolysis data for their conclusions regarding the importance of various wall
constituents in maintaining the wild-type colonial morphology in Neurospora crassa.
Novaes-Ledieu and Mendoza (1981) used (3(1-3)-glucanase, isolated from Rhizopus
arrhizus. to confirm the presence of predominantly B(1-3) linkages in a glucan of the
alkali-insoluble fraction. Mol and Wessels (1987) used chitinase from Serratia
marcescens to establish a glucan-glucosamine link and thus the presence of
glucosaminoglycan in the walls of Saccharomyces cerevisiae. Examples of enzyme
localization uses are discussed later.
Various analytical methods are used to ascertain molecular structure, hydrolysate
composition, linkage information, and other relevant data. Various chromatographic/
electrophoretic (e.g., thin-layer, Zonneveld, 1 971 ; thin-layer and HPLC, Briza et al., 1 986;
gas-chromatography, Stagg & Feather, 1973; SDS-PAGE, Hearn et al., 1989),
colorimetric/spectrophotometric(e.g., Mol & Wessels, 1987; Novaes-Ledieu & f^endoza,
1981; Zonneveld, 1971, 1972; Mahadevan & latum, 1965, 1967), optical-rotation
analysis (e.g., Zonneveld, 1971; Johnston, 1965), infrared spectrometric (e.g., Briza et
al., 1988; Novaes-Ledieu & Mendoza, 1981), and various NMR (e.g., GLC-MS, Hearn et
al., 1 989; NMR, Briza et al., 1 986, 1 988; C-n.m.r., Gorin & lacomini, 1 984) methods have
been used to determine hydrolysate composition and linkage information. Paper
chromatography (immobility of polymer/mobility of primed residue) has even been used
to monitor chitosan synthesis (Davis & Bartnicki-Garcia, 1984). X-ray crystallography,
or diffraction (e.g., Rudall, 1969; Blackwell, 1982) has been used for determining the
1
1
.i
%
20
structure of relatively insoluble residues. This technique has been used to verify the
presence of such structures as crystalline chitin.
Localization of wall components via light and electron microscopy techniques
provides visual information on which to base models of wall structures. Fluorescence
(light) microscopy using autofluorescence (e.g., Briza et al., 1986), fluorescent stains
(e.g., Briza et al., 1 988), and fluorescent-labelled conjugates (e.g., Briza et al., 1 988) has
been used to determine presence and in some cases (such as yeast bud scar) location
of inner and outer wall layers. Sequential enzyme digestions followed by shadow
casting TEM at each step has provided extensive insight regarding wall architecture
(Hunsley & Burnett, 1970; Burnett, 1979). TEM of specimens prepared only for
morphology (e.g., Dute et al., 1989) provides general information on which initial
hypotheses and further studies can be based. TEM of sections labelled with gold- "i
conjugated lectins and enzymes has, in some cases, resulted in evidence of various wall
components residing within specific wall layers (Benhamou, 1988, 1989). Some lectins
and their binding specificities are given in chapter 5.
Two immunological strategies have been employed for analysis and identification
of wall components. The so called "blind" approach uses whole fungi, isolated wall '
fragments or fractions (e.g., Young and Larsh, 1982) and the direct approach, which
employs pure antigen as immunogen (e.g.. Green et al., 1980). The blind approach has
the advantages of requiring less effort in preparation of immunogen, and the produced
monoclonals can then be used to isolate the antigenic molecules in relatively pure form
for further analysis. Through these methods mural mannan, galactomannan, and protein
antigens have been isolated (Reiss, 1986). These methods will be discussed in greater
detail in chapter 3.
\ ^A
t'/y.'S
21
fungal wall morphology (Burnett, 1979; Zonneveld, 1971). In combination with other
preparatory and analytical methods, this approach can be put to use in wall studies.
An example of such a study is Zonneveld's (1973) substitution of the glucose analog,
2-deoxy-glucose, for glucose to determine the role(s) of a (1-3) glucan in vegetative
growth and sexual morphogenesis.
The Carbohydrates
As previously stated, hyphal walls are mainly composed of various
carbohydrates, including chitin. The presence of chitin in ascomycete hyphal walls was
established over 20 years ago (Aronson, 1965; Bartnicki-Garcia, 1968). More recently
chitin was said to be "the most characteristic component of fungal walls" (Wessels,
1 986). It accounts for a significant portion of the wall in some fungi (e.g., about 1 0% in
Neurospora crassa, Burnett, 1979; and 9-13% in Aspergillus niqer, Johnson, 1965). The
presence of chitin in conidia and ascospores is highly probable, but neither so well, nor
ubiquitously, established. However, the occurrence of chitin in crustaceans, insects,
and spiders, as well as fungal hyphae, prompted Rees (1977) to suggest that this
polymer may be "more abundant in nature than cellulose."
Chitin is a B-(1-4) linked polymer of N-acetylglucosamine. Although chitin is
usually considered to be a homopolymer, non-acetylated residues may occur (Rudall,
1969; Wessels, 1986). Crystallization occurs when single chitin polymers pack, or pile,
side-by-side and form numerous, regular, inter-polymer CO— NH hydrogen bonds
(Rudall, 1969; Rees, 1977). Three crystalline forms of chitin (a-, 8-, and 5f-) are known
(Rudall, 1969). The 8- form is made up of chains piled in parallel orientation to one
another while the a- form is of antiparallel orientation, and the li- form (Fig. 8) has both .^
■I
22
parallel and antiparallel polymer components. The a- form Is the most stable (Rudall,
1969), and the form present in fungal chltin (Rudall, 1969; Wessels, 1986).
Rudall (1 969) describes fungal chltin as "spirally wound fibrils." An alternate term
for Rudall's fibril is microfibril, and this latter term appears to be more widely used. More
recent researchers find the relationship between crystallinity and microfibrillar structure
not so clear-cut (Wessels, 1986). In fact, according to Wessels (1986), associated 6-
glucan may prevent "formation of perfect crystallites of chitin." In fungal hyphae these
microfibrils are interwoven forming a rigid web which is capable of retaining its shape
even after removal of matrix materials (Burnett, 1979). This led Burnett (1979) to
conclude that chitin performs "a genuine skeletal function." It is important to recognize
that chitin may not be the major contributor of mechanical strength and stability for all
fungi that are considered to be Ascomycetes. It has been suggested that in
Saccharomyces cerevisiae a portion of the chitin present is not found in crystalline form,
and that crystalline chitin may not be the primary element of mechanical strength in this
fungus (Mol & Wessels, 1987).
Complete deacetylation of chitin polymers produces homopolymers of
glucosamine, or chitosan. There may be a range of deacetylated polymers from chitin
to chitosan present in fungal walls (Rudall, 1969; Mol & Wessels, 1987). Studies have
shown biological deacetylation of chitin to be the mode of chitosan formation (e.g.,
Davis & Bartnicki-Garcia, 1984). Incomplete deacetylation may cause imperfections in
the crystalline structure and allow water penetration of the resultant pseudo-chitin
(Rudall, 1969).
Chitosan has been found in the walls of Zygomycetes (Bartnicki-Garcia, 1968),
non-reproductive and non-lamellae fruit-body cells of the Basidiomycete species
23
Aqaricus bisporus (brunnescens) and A. campestris (Novaes-Ledieu & Mendoza, 1981),
Sacchromyces cerevisiae cells in early stationary growth phase (Mo! & Weasels, 1987),
and the ascospore walls of yeast strain AP3 (Briza et al., 1988). In terms of taxonomic
groups in which chitosan can be found, this polymer is probably more widespread than
early reviews indicate (e.g., Bartnicki-Garcia, 1968), but it may be restricted in the type
of cell in which it occurs.
The glucans (D-glucopyranosyl polymers) are also important in terms of their
abundance and function in hyphal walls. Up to 25% (w/w) of Neurospora crassa walls
are composed of glucan (Burnett, 1979). The glucans known from ascomycete walls
include 6(1 -3), B(1 -6), S(1 -3)-B(1 -6), a (1 -3), a (1 -3)-a (1 -4) linked, and possibly a (1 -4)
linked D-glucopyranosyl.
Although the presence of cellulose (6(1-4)-D-glucopyranosyl) in the cell walls is
typical for some fungi such as the Oomycetes (Bartnicki-Garcia, 1968), it is
characteristically absent in Ascomycetes. Within the Ascomycetes the presence of
cellulose has only been documented in species of the non-Pezizalian ascomycetes
Europhium and Ophiostoma (Aronson, 1981). Chitin and B(1-3) linked glucans provide
the mechanical support for fungal cells that cellulose does for higher plant walls. , ,. V!
Pure B(1-3) glucan, or those polymers with infrequent B(1-6) linkages, can
crystallize into microfibrils (Burnett, 1979). The extent to which B(1-3)-B(1-6) glucans can ' '<
crystallize seems to be dependent on the frequency of B(1-6) branches (Burnett, 1979).
Glucans of this type with a high frequency of B(1 -6) linkages are presumably more
amorphous than those with a low frequency. Amorphous molecules are generally
considered to be matrix components. This type of mixed linkage glucan has been
found in notable quantities in alkali-insoluble wall fractions of Sacchromyces cerevisiae
24
(Mol & Wessels, 1987), associated with chitin (Wessels, 1986; Rudall, 1969) and In
Aspergillus niqer (Stagg & Feather, 1973).
Zonneveld (1971) has shown the presence of a(1-3)-glucopyranosyl in
Aspergillus nidulans, and demonstrated its importance in the fructification elsewhere
(Zonneveld, 1973). These a -(1-3) glucans are generally considered to be linear
(Aronson, 1981). In a earlier study of A. niqer Johnston (1965) reported a wall fraction
of predominantly a -(1-3) linked glucose residues. This glucan was found in the alkali-
soluble fraction (S-glucan; Zonneveld, 1971), implying that this too is a matrix
component. Wessels (1986) indicated that a-(1-3)-D-glucan occurred in the alkaline
soluble fraction of both Ascomycete and Basidiomycete walls. This glucan has been
shown to have a characteristic rodlet-form in the outer wall region of the Basidiomycete
Schizophvllum commune, but at least for Neurospora crassa. no evidence of this form
has been found (Burnett, 1979). Rodlet structures have also been demonstrated by
freeze fracture techniques in the condial walls of Scopulariopsis brevicaulis (Cole &
Aldrich, 1971) and teliospore walls of Neovossia horrida (Nawaz & Hess, 1987).
Although no chemical data were given for those teliospores (Nawaz & Hess, 1 987),
rodlets in conidial walls are described as proteinaceous (Hashimoto et al., 1976).
The term "mycodextran" was coined by Dox and Neidig (1914) for the glucan of
alternating a (1 -3) and a (1 -4) linkages they isolated from Penicillium expansum. This
J
glucan now goes by the name nigeran. Johnston (1965) found this glucan in the hyphal ."j
walls of A. niger. Based on Johnston's data, Gorin (1968) reported this component to
represent 26-42% of the total wall, Aronson (1 981 ) reported 4-6%, and by this author's
rough calculation from that data, 10%. For A. nidulans Zonneveld (1971) reported that
few, if any q: (1 -4) glycosidic linkages exist. This large discrepancy between species may
iM
m
mannan as a glycoprotein with 2 distinct carbohydrate moieties; one with a-D-(1-6)
backbone and a-D-(1-3) linked branches, the other with only a -(1-2) linkages.
Trichosporon aculeatum has a branched mannan in which all the linkages of yeast
mannan exist, but more than 5 consecutive a -D-(1 -2) linkages were never found (Gorin,
'^
..1
25
be actual, or it may be due to culture conditions, or methodology. Gorin (1968) found
that A. niger grown with starch rather than glucose as the primary carbon source had
predominantly (87%) a (1-3) linkages (pseudonigeran). Gorin (1968) also stated,
apparently contrary to Johnston's (1 965) written opinion, that pseudonigeran was the
glucan present in A. niger rather than both nigeran and pseudonigeran because neither ^,J
were soluble in hot water. Pseudonigeran is thought to be more widespread
taxonomically than nigeran (Aronson, 1981). Zonneveld (1972, 1973) found a (1-4) i^,..!
linked glucose residues in the alkali-insoluble fraction along with 6(1-3), 6(1-6),
mannose-galactose polymers, and chitin. Horikoshi and lida (1964) reported a glucan ]
of a (1 -3) and a (1 -4) linked residues, but gave no indication of the proportions of these
linkages within the polymer. Aronson (1981) stated that a (1-4) linkages between - .^
glucansto heteropolysaccharides (e.g., 6-glucan-galactomannorhamnan in Fusicoccum
amygdali) "are unquesionably significant" as they knit various polysaccharides into larger
wall complexes. No reports of a consecutively ct (1 -4) linked glucan were found in this
literature search and review.
Mannose is commonly found in fungal wall digestions (Bartnicki-Garcia, 1968).
Apparently homopolymers occur, but mannose is more often described as a constituent
of heteropolysaccharides and glycoprotein conjugates. Yeast mannan has been
r^ Jlfrntdl
o
described as having an a-D-(1-6) backbone, and Q:-D-(1-3) and a-D-(1-2) branches ,, J
(Gorin, 1968) of two to five residues (Reiss, 1986). Farkas (1979) described yeast
^
26
1 968). In Candida albicans cell wall mannans with a -D-(1 -2) and Ck: -D-(1 -6) linkages are
major antigens (Gorin, 1968; Reiss, 1986). Unlike the previous mannan, these antigenic
mannans have furanosyl, as well as pyranosyl residues (Gorin, 1968).
Galactose, like mannose, is commonly found in fungal walls, but in this case is
neither present in all fungi, nor even all Ascomycetes. Both furanosyl and pyranosyl
residues occur in galactan homopolymers (Gorin, 1968), and heteropolysaccharides
(Gorin, 1985). Apparently galactose is more abundant in heteropolysaccharides.
Galactocarolose is an example of galactan from Penicillium charlesii. Gorin (1968)
described this molecule as a linear oligomer (9-1 0 residues) of a -D-(1 -5)-galactofuranose
(Gorin, 1968). Galactocarolose has also been described as a degradation product of
peptidophospho-galactomannans (Salt & Gander, 1985; Preston & Gander, 1968).
Phosphorylated residues of both galactose and mannose have been found in
fungal walls (Gorin, 1968). These residues, and 2-amino-2-deoxy-D-galactose, have
been described as occurring as components of "exocellular" polymers (Gorin, 1 968).
The exact position and role in (or outside) the wall is unclear.
Wall Proteins and Glycoproteins , , , ' '
Proteins are an obvious component within the wall since amino acids are
commonly found in wall fractionations (Gorin, 1985; Novaes-Ledieu, & Mendoza, 1981 ;
Zonneveld, 1971; Johnston, 1965; Mahadevan, & Tatum, 1965). Wall proteins occur
both glyco-conjugated (Hearn, et al., 1989; Salt, & Gander, 1985; Aronson, 1981), and
apparently unconjugated (Farkas, 1979). Rosenberger (1976) found fungal walls to be
1 0-1 5% protein after extensive washings and considered this protein to be a structural
component. Glycoprotein in the wall may be a component in a supra-molecule capable
» .*.
■•Vi«t
1
■4
'■>S£S
■ h;i
%
•?l
•J
^
27
of sealing in unbound wall materials (Farkas, 1 979). Mural glycoproteins may participate
in cell-cell recognition, cell dfferentiation, and mating (Tanner, 1990).
Farkas (1979), Reiss (1986) and Kuhn & Trinci (1990) described the wall as the
location of a number of enzymes. In hyphae, some of the mural enzymes undoubtedly -,;
play a role in the provision of nutrients (Kuhn & Trinci, 1990). Mural enzymes fall into -*^^u
two major categories; the proteinases and B(1 -3)glucanases (Reiss, 1986). Other
enzymes known to occur murally are invertase, acid phosphatase (Farkas, 1979; Reiss, '■■'''•
'I
1990) and 13(1 -4)xylanase (Notario et al., 1979).
A mannan-protein complex has been described as the matrix component in ]
yeasts (Peberdy, 1990). Aronson (1981) provided another example in that some 10%
of Pyricularia orvzae wall was said to be "proteohetero-glycan." When purified, this
molecule was determined to be 91 % carbohydrate and 9% protein. The polysaccharide
portion had an a -(1-6) mannopyranosyl main chain with (1-2) linked glucomannan or
galactomannan side chains (Aronson, 1981).
■jm
M
f "
Wall Lipids • '' '
Bartnicki-Garcia (1 968) presented evidence supporting lipid(s) as a bona fide wall
component. Cell walls of Aspergillus niger have been reported to be 2-7% lipid .:
(Johnston, 1 965). No further information was found on its relationship(s) with other wall ..«
components, conjugate partner(s), or roles within the wall. i
Locations of Synthesis Enzymes
Various enzymes have been isolated which are involved in the synthesis of wall
components. Publications on these enzymes began appearing in 1957 with Glaser and
28
Brown's (1 957) description of chitin syntiiesis in fungi. The bulk of chitin syntlietase has
since been found to be attached to the plasma membrane (Duran et a!., 1975; Kang et
al., 1985). Furthermore, isolated intact membranes have been shown to synthesize
chitin on the external face of those membranes in vivo (Cabib et al., 1983).
Chitosan synthesis has been characterized as a chitin deacetylation process
(Davis & Bartnicki-Garcia, 1984). Interestingly, only 37% of the chitin deacetylase was
associated with the extracellular fraction. The remainer was associated with the
particulate (14%) and soluble 20000g supernatant (49%) fractions (Araki & Ito, 1975).
The other structural carbohydrate known to occur in fungi is 6(1 -3)glucans. 8(1 -
3)glucan synthase has, like chitin synthetase, been found to be a membrane bound
enzyme. Further, it has been described as an integral, trans-plasma membrane enzyme
(in Neurospora crassa, Hrmova et al., 1 989; in Mucor rouxii, Fevre et al., 1 990; Peberdy,
1990). Activity of B(1-3)glucan synthase has also been found in association with both
endoplasmic reticulum and plasma membrane fractions (in, Saprolegnia monoica, Fevre,
1984).
Glycosyl transferases would also be involved in construction of wall
carbohydrates. These enzymes might be expected to occur in the cytosol and indeed
the mannosyltransferases have been found in the cytosolic particulate fraction of
Crvptococcus laurentii (Schutzbach & Ankel, 1972).
The occurrence of glycoproteins in the wall has been previously mentioned.
Tanner (1990) described glycoproteins as occurring only in special cellular
compartments including the cell wall, and organelles involved in glycoprotein systhesis,
i.e., endoplasmic reticulum, Golgi complex, and secretory vesicles. Mannoprotein
formation in yeasts has long been thought to be a process involving much or all of the
endomembrane system (Farkas, 1979).
>?
"'-'Ski
Ml
W
^^■
M
29
Study Proposal
Hypotheses
Very little is known about the chemistry, biosynthesis, or maturation process of
ascospore walls. The bulk of fungal research in these areas has focused on hyphae.
This may be due in part to the fact that the developing ascospore is difficult to access *h
in comparison to hyphae. Yet, this type of research would add greatly to our
understanding of the biology of this group of organisms. Jl
The most fundamental questions are that of chemistry and biosynthesis of J
ascospore walls and their constituent layers. Toward answering such questions some -^
researchers have published a limited amount of cytochemical data. Those experiments
have provided infomation of a general, non-specific nature. Nevertheless, it is important
to make assumptions and/or hypotheses about the specific chemical nature so that
appropriate experimental designs may be created. Thus, it is necessary in this case to
apply the information available on hyphae to develop hypotheses. The original study
proposal used available information to just such an end.
Skeletal elements in hyphae consist primarily of chitin and/or B(1-3)glucan
(Farkas, 1979). It not inconceivable that some mannans could play a strucural role.
Thus, it was hypothesized that structural elements of ascospores are most likely to be
chitin and/or 6(1-3)glucan and less likely to be mannan. This hypothesis is supported
by the fact that the synthesis enzymes for chitin and B(1-3) glucan have been found to
be located in plasma membranes (Duran et al., 1975; Kang et al., 1985; Hrmova et al.,
1989; Fevre et al., 1990). Further, there is strong evidence indicating that the spore
delimiting membranes are derived from the ascus plasma membrane (Mims, 1990).
Due to the potential diversity of matrix constituents no hypothesis regarding
^' ■'^.
:"'>■
4
^
•riij
■>:
^
30
specific conponents was put forward in the original proposal. Although, a hypothesis
of general similarity (i.e., Ho: these wall systems will have some shared components)
was forwarded. In terms of classes of molecules, it is likely that proteins and
glycoproteins are generated at or in the endomembrane system in either the epiplasm ^
or sporoplasm of ascospores. Further, it is also probable that some matrix components
arrive at the delimiting membranes in vesicles of the endomembrane system.
As no structure similar to ascospore secondary walls has been described for
hyphae, no hypotheses for common constituents could be proposed. Morphological
evidence seems to indicate that at least the major components of this wall layer are
synthesized at the outer delimiting membrane or in the epiplasm. Based on
morphological evidence for the vesicular epiplasm origin of secondary wall components
and highly probable endomembrane orgin of some matrix components it was
hypothesized that given the appropriate probe, it would be possible to track wall j
materials not synthesized in situ. This hypothesis is not directly testable and therefore
was only a secondary goal of this research.
Maturation events in the ascospore walls undoubtly occur. Minimally, such an
event is required to fulfil the sealing function necessary for survival of the spore. It was
hypothesized that maturation events would be documentable using the proposed
cytochemical techniques. Again, this hypothesis is not directly testable and therefore
was considered to be a secondary goal of this research.
Using the chemical and biosynthesis information available on hyphae it was
possible to select commercially available probes to test the chemical similarity
hypotheses. Monoclonal antibodies developed against either hyphal or ascospore walls
could be used for the same purpose. Anti-ascospore wall antibodies were of particular
interest as they could also provide evidence for unique chemistry of the spore wall.
-<i
31
Materials and Methods J
Due to the repetitious nature of developing protocols, and the immunological
requirement for large quantities of antigen relatively free of contaminating wall materials,
the research organism must 1) produce ascocarps readily in culture, 2) sporulate ^
prolifically, and 3) not produce conidia. Ascodesmis sphaerospora meets these criteria,
was available, and was thus proposed for use and used as the research organism.
Development of cytochemical techniques and protocols specific for elucidation
of biochemical and biosynthesis (biogenesis) information for ascospores was the
primary goal of this research. Basic technology for such work (cytochemical stains;
.i
carbohydrate, lectin, enzyme, and immuno-, cytochemistry; and use of secondary
probes) was outlined by Aldrich and Todd (1966). More specific information on these
techniques was also readily available and is reviewed in chapters 2, 4, and 5. i
Carbohydrates are an obvious target of this research and thus lectin and immuno-
cytochemical techinques were proposed as the initial and primary focus of the
techniques research. Use of enzyme-digestion and enzyme-probe techniques was also .,^ .
■_f'.' " -.
proposed as a third line of techniques research. ^
Production of anti-spore wall antibodies was seen as necessary for successful -i
'. - ' -'"J?.
completion of this project. Technology for preparation of monoclonal antibodies from
mice spleen (and other sources) is well described in the literature and is reviewed in ' --^,
J
J]
chapter 3. '
Conclusions
As proposed, this study was seen to have the potential to produce data that
could increase our knowledge of the chemistry and our understanding of the biology
32
Of ascosporogenesis. The procedures developed would be applicable to other fungi
and had the potential for addressing other biological questions. Thus, as proposed, it : ' S
was felt that this work had great potential for provision of a foundation of data for future
research on the biology of Ascomycete fungi. *
4
CHAPTER 2
STRATEGIES FOR
TISSUE PREPARATION AND EMBEDDING
Literature Review
Introduction
Perhaps one of the most difficult steps of any long-term experimental project is
the preparation of material for experiments that are temporally far removed. This can
be a critical problem for cytochemical experiments where tissue may only be available
on rare occasions, or in limited amounts. The fixation and resin embedding of tissue
immortalizes it, but also changes it irreversibly. Pre-embedding experiments are
sometimes the most appropriate route. The possibility of pre-embedding experiments
which exists for some tissues are out of the question here because of the impermeable
nature of cell walls.
The success of cytochemical experimention such as proposed for this study, is
relatively dependent on the condition of target molecules. If the changes incurred : ; ji'^
■ •■■'• ■ - y?:
during tissue processing significantly alter potential target molecules, then cytochemical ''
experiments to detect such molecules can, and probably will, be rendered ineffective
(e.g., Craig & Goodchild, 1982; Eldred et al., 1983; Erdos & Whitaker, 1983; Hardham, J
1
1985). Bendayan (1989b) reported that the tissue components should retain their 3-
dimensional configuration in order to be recognized by enzyme probes. The ascospore ]
constituents that are the potential target molecules include carbohydrates, proteins and
glycoproteins. It is important therefore to understand how tissue processing might affect
33
34
these molecules specifically prior to the actual tissue processing and cytochemical
experimentation.
Tissue processing involves fixation of the material, sometimes a secondary "ii
fixation, dehydration, infiltration of a resin, and polymerization of that resin. Significant ,.^
changes at the molecular level can occur during any of these processing steps. Each r'^^
of these steps, including typically used reagents and potential resultant molecular
changes are reviewed below. The extent to which tissue processing alters the biological ,,^
configuration of macromolecules varies (Bendayan, 1989a). Therefore, one must (
j
develop fixation and dehydration protocols and chose an embedding medium optimal i^
' '.'.I
for the cytochemical probe, and more specifically for its target molecule.
Fixation
The goal of fixation is to kill and stabilize cell structures. This should be done '1
rapidly so that a minimum of autolytic (postmortem) damage occurs. Fixation of
biological material is often done in two steps. The primary fixation is most typically done
with glutaraldehyde, and/or formaldehyde and/or acrolein. The secondary fixation is ^'A
done with osmium tetroxide after the primary fixation and buffer washes. ' .;.
Glutaraldehyde, or a mixture of glutaraldehyde and formaldehyde, is probably the ' . ^ . '
most commonly used primary fixative for electron microscopy. Glutaraldehyde is a -
dialdehyde and very effectively stabilizes proteins via irreversible cross-linking. Hayat ^
(1 981 ) stated that no other fixative has surpassed the ability of glutaraldehyde to cross-
link proteins and preserve tissue proteins for electron microscopy. This fixative
introduces both intra- and intermolecular cross-links in proteins but is unable to cross-
link low concentrations of proteins (Hayat, 1 981 , 1 986). Glutaraldehyde reacts with the ^
i
35
c -amino group of lysine, N-terminal amino groups, a -amino groups of free amino acids,
protein associated DNA, and tlie 1° amino groups of etiianolamine containing
phospholipids (Hayat, 1 986; Sternberger, 1 986). Most lipids (other than phospholipids), ''*
myelin, and glycoproteins are not fixed by glutaraldehyde (Hayat, 1 986). Glycoproteins . ^
are said to be "immobilized" by glutaraldehyde. Glutaraldehyde is not thought to interact
with carbohydrates (McLean and Nakane, 1974). For good morphological preservation
of biological material primary fixation with 2%-3% glutaraldehyde (v/v) in buffer for 1-2
hr at 4°C or room temperature is usually adequate. Low concentrations of J
glutaraldehyde are recommended for immunocytochemistry (especially with monoclonai
antibodies, Beesley, 1 989) and enzyme cytochemistry (Bendayan, 1 989a) since retention
of biological configuration can be altered by this fixative. Loss of antigenicity or receptor
■'ii
integrity during dehydration and infiltration may be reduced by glutaraldehyde (Craig & -1
Goodchild, 1982). DeWaele and coworkers (1983) reported that some glutaraldehyde i
in the fixative solution enhances the permeablility of the cell surface membranes. This
would be particularly beneficial for pre-embedding experiments. The concentration of
glutaraldehyde in the fixative solution could be less relevant when the receptor site is
carbohydrate in nature.
Formaldehyde can also be used as the sole primary fixative but this is not
recommended for good ultrastructural preservation (Hayat, 1981, 1986). Unlike ,.
glutaraldehyde, it is a mono-aldehyde and its reactions with proteins and other cellular ^ i,
1
components are at least partly reversible. It penetrates tissue rapidly and in that respect 1
•I
is superior to glutaraldehyde. Cross-linking of protein is slow with formaldehyde (Hayat,
1986). It reacts with free amino groups, hydroxyl, caroxyl, sulfhydryl, and peptide
bonds. Formaldehyde is a poor fixative for lipids and actually degrades some types ,^4
36
of lipids (Hayat, 1986). If only this fixative is used, lipids may be extracted during
dehydration.
Acrolein is a monoaldehyde which can be used as a fixative. It is an extremely
reactive, flammable, volatile, and toxic (respiratory, ocular mucosa, and skin irritant)
reagent (Hayat, 1981). It reacts rapidly with free amino groups and is superior to
formaldehyde for cross-linking protein (Hayat, 1986). This aldehyde is bifunctional by
virtue of its double bond. It also reacts with carboxyl, imidazol, and substrates that bear
sulfhydryl or thiol groups and is thought to react with fatty acids (Hayat, 1981 & 1986).
Mixtures of aldehydes are recommended (Hayat, 1986) because they often
produce superior ultrastructure preservation.
Secondary fixation with osmium tetroxide is commonly used for routine
morphological work. Osmium tetroxide has two major advantages for morphological
work; 1) it is a heavy metal salt and imparts contrast to those molecules and structures
it stabilizes, and 2) it stabilizes unsaturated fatty acids by oxidizing the available double
bonds (Hayat, 1 986). Thus, osmium tetroxide is the fixative of choice for stabilizing and
visualizing membranes. In addition to its action on lipids, it also cross-links proteins to
a small degree (Hayat, 1 986). Osmium tetroxide is said to denature the a -helix regions
of membrane proteins (Lenard & Singer, 1968).
'■, /■ •.•■It ^ t
Additives to Primary Fix • *. ' -a' ■ '\J ■■
Additives to the primary fixative solution such as picric acid (Stefanini et al., 1 967;
Dae et al., 1982) periodate-lysine (Hixson et al., 1981; McLean and Nakane, 1974:
Pollard et al., 1 987), and tannic acid (Stirling, 1 989) have been recommended to improve
morphology without loss of antigenic or binding site receptivity (Stirling, 1990).
■:-l
'^
• 1
*.
li.
37
Dehydration
Dehydration is a requirement for proper infiltration and polymerization of plastic
resins. While the epoxy resins are hydrophobic and will not tolerate any water, the
acrylic resins are water tolerant (Newman, 1 987). The dehydrant should be compatible
with the resin, inert to biological material, and should not denature molecular
components (Stirling, 1990). Ethanol has been reported to fulfill these requirements
(Carlemalm et al., 1982), yet lipid extraction (Weibull et al., 1983) and dimensional
changes (Boyde et al., 1977) have also been reported to occur when ethanol
concentrations exceed 70%. Specimens for ultrastructure study typically employ epoxy
resins and are dehydrated in ethanol series through 1 00% followed by acetone washes.
Acetone may also be used with the acrylic Lowicryl resins. Kellenberger and coworkers
(1 987) reported freeze-substitution experiments where 3% glutaraldehyde in acetone and
infiltration with acetone diluted Lowicryl were used. At low temperatures extraction does
not appear to be a problem. Conversely, acetone should be avoided when the acrylic
resin LR White is employed as this solvent may interfere with the polymerization process
(Stirling, 1990). When LR White resin is to be used dehydration through only 70%
ethanol has been recommended to avoid the detrimental effect of higher concentrations
(Newman & Jasani, 1984; Newman, 1987; Newman & Hobot, 1987).
Resins and Polymerization
As early as 1 962 it was suggested that the media could exert a "differential effect
by differences in the way in which they combine with reactive groups of proteins and
nucleic acids, and possibly by differences in the penetrability of the insoluble polymers
by the enzymes" (Leduc & Bernhard, 1962). The two problems related to resins are 1)
. ."M
".^^i
:ji
' > '4%
38
preservation of binding site receptivity (antigenicity) within the tissue and 2) steric ^\
hindrance of the probe (Causton, 1 984). These problems demand close attention to the ^
chemical reactivity of cured resin, the curing process itself and to the degree of cross-
linking achieved during the curing process. The success of EM detection also requires ]
the resin be stable in an electron beam. Causton (1984) recommended epoxy cross- '?^'.-??l
linked systems or cross-linked hydrophilic acrylics for best results and greatest flexibility
of technique. iji
Another potential problem discussed by Newman and Hobot (1 987) is that of
extraction of tissues by the resins. Polymerization by chemical acceleration of the resin
was the solution they suggested and demonstrated (Newman & Hobot, 1 987). The rate
of diffusion of the accelerator into the tissue is an obvious limiting factor.
Araldite, Epon, and Spurr are the epoxy resins used for electron microscopy.
They all have the advantage of being stable in the beam and the disadvantages of a
high degree of cross-linking not only with resin components but also with peptide ''
groups, and of being hydrophobic. An additional disadvantage of Araldite is that the ;,
component, diglycidyl ether of bisphenol A, is a large molecule and has a slow rate of ^ -^
diffusion into tissue (Causton, 1984). Epon and Spurr resins are less viscous than ' ^
Araldite and provide improved diffusion properties. Spurr resin has the highest rate of
diffusion of all these epoxy resins (Causton, 1984). \ / \^.- < •,- - ' ' :" ^ ..'^
Similar to glutaraldehyde, cross-linking of resin to peptide groups may disrupt
specific receptor requirements of the molecular probes. Such cross-linking may also
alter the way in which a section is cleaved from the block and thus alter the amount of
surface area available for cytochemical interaction with the tissue. Kellenberger and
coworkers (1987) have shown the relief of Epon sections to be smoother than that of
'^
iuS
1^
v^
39
Lowicryl sections. They further suggested that the cleavage where co-polymerization
does not exist will follow the interfaces between resin and proteins whereas the
cleavage will preferentially not follow such interfaces where co-polymerization does
exist. Essentially, a cleavage which follows the resin/protein interface is preferable
because binding sites are laid open (Kellenberger et al., 1987).
The characteristic hydrophobicity is imparted to these resins by alkane (RCHg)
side chains (Causton, 1984). Newman and Jasani (1984) described the epoxies as
impermeable to aqueous solutions at neutral pH and thus antibodies are isolated from
the antigens by a hydrophobic barrier. Treatment with oxidizing agents such as ;l
hydrogen peroxide, periodic acid (periodate) or potassium permanganate produces
hydrophilic groups, thus distroying the hydrophobic barrier. These treatments may also
oxidize target molecules and therefore are best avoided (Causton, 1 984; Newman &
-*
"4
Jasani, 1984). ,"-^
The acrylic resins are the Lowicryls (K4M, HM20) and LR White. The great
advantage these resins have over the epoxy resins is that they are hydrophilic (Newman,
1987). Thus hydration sensitive receptor sites are more lil<ely to be retained, the need
for the potentially detrimental oxidation treatment is supposedly eliminated and the
mildest curing conditions can be chosen (Causton, 1984).
Newman and Hobot (1987) reported that these hydrophilic resins swell in
aqueous solution and that this swelling is dependent on the degree of cross-linl<ing.
They further postulate, as Kellenberger and coworkers (1 987) did for Lowicryl section
"relief", that this swelling may improve receptor site accessibility.
Lowicryls can be cured with UV-light as well as with chemical accelerators. They
are very mobile at low temperatures and thus infiltration and polymerization can be <
1
40
done at low temperatures. Although ultrastructural preservation is improved by low
temperature methods, Newman (1 987) pointed out that this does not automatically imply
improved preservation of antigenicity. Causton (1984) stated that Lowicryl "has no
special features that make it especially suited to electron microscopy."
LR White resin can be cured with UV-light, heat, and chemical accelerators
(Newman, 1987). Newman and Jasani (1984) reported that best results for post-
embedding cytochemistry were obtained with this resin when it had a slow (50° C) heat
cure. Later, Newman and Hobot (1987) described catalytic polymerization at room
temperature and at 0° C to be a further improvement. This work was done with human
pituitary and rat kidney tissue, not a tissue with cell walls where the rate of penetration
of the accelerator would be a more critical factor. Newman and Hobot (1 987) reported
gelling of chemically accelerated resin within approximately 7 minutes. It is doubtful that
the accelerator could completely infiltrate both ascus and ascospore walls that rapidly.
Tissue Preparation Strategies
Introduction
The cytochemical study proposed in chapter 1 requires use of post-embedding
methods. The principal probes proposed for use in this study were antibodies, lectins
and possibly enzymes; the potential target molecules were protein, glycoprotein, and
carbohydrate in nature. It is clear from the preceding literature review that tissue
processing and embedding inevitably causes a reduction in the receptivity of some '
binding sites due to loss of or damage to tissue elements. The trade off between
morphology and labelability has long been recognized. In fact, the issue was resolved
by Leduc and Bernard (1962) via acceptance of artifacts and poor morphology for the _-^^
-;^
41
contribution to our knowledge of ultrastructural cinemistry tiieir experiments could
provide. Similar acceptance of poor morphology was proposed as a starting place for
a study.
Fixation
The choice of fixative can be critical, especially for use of protein binding probes.
Antibody and enzyme probes are used for detection of protein and glycoprotein
molecules. To some extent it may be possible to increase an antibody's ability to -5
1
recognize a glutaraldehyde fixed molecule by light fixation of the immunogen prior to
its use. Light fixation in this case would be fixation with 0.5% glutaraldehyde for 30
minutes on ice. It is then possible to use tissues fixed with at least 0.5% glutaraldehyde,
and possibly up to 2% glutaraldehyde (Erdos, personal communication) with those
antibodies. Unfortunately, for use enzyme probes a lowered concentration or no .'
glutaraldehyde in the fixative solution is typically required. Generally, a combination of
glutaraldehyde and formaldehyde is recommended for post-embedding cytochemistry
(e.g., 0.1%-1% glutaraldehyde with 2%-4% formaldehyde, Stirling, 1990; Roth, 1983;
DeWaele et al., 1983). Acrolein, or mixes with acrolein were not recommended in any
of the literature reviewed here. Use of osmium tetroxide post-fixation is not
recommended in general where post-embedding cytochemistry is to be used because
of its adverse effect on antigenicity and receptor site reactivity (Bendayan, 1989b;
Stirling, 1990). When osmium tetroxide is used it is recommended to pre-treat sections ^
with a saturated solution of periodate (Bendayan & Zollinger, 1 983; Bendayan 1 984a,
1984b, 1989a, 1989b). This treatment, in turn, could damage some carbohydrates and
oxidize alkanes. Stirling (1990) recommended preparing tissue with a number of • <>«Bi
%
42
different fixations. Following the recommendations of Erdos (personal communication)
and Stirling (1990), the following preparations were proposed as an adequate start-up
system;
1) Fixation of the immunogen as previously described, and embedding an
alioquot of this preparation for TEM use.
2) Fixation of lightly broken spores that have not been fractionated and retain
some cytoplasm with 0.5% glutaraldehyde and 4% formaldehyde.
3) Fixation of ascocarps with several combinations of glutaraldehyde and
formaldehyde including: 0% glutaraldehyde with 4% formaldehyde, 0.5% with 4%,
1% with 2%-4%, and 2% with 2%.
4) Fixation of ascocarps with 2% glutaraldehyde and 2% formaldehyde then
post-fixed with osmium tetroxide (for comparative morphology). "■%
Use of additive(s) to the fixatives was omitted from this plan. It was felt that the various
fixative mixtures proposed would provide enough variation for initial screening of probes
and testing of protocols.
■4
Resins and Dehydration *^"
Lowicryl K4M, LR White, Spurr, and Epon resins were available for this study.
Lowicryl K4M with low temperature infiltration and polymerization provides the greatest
advantages for post-embedding cytochemical experimentation. Unfortunately, the .i|
experimental organism has brown spores which are impenetrable to UV radiation for
polymerization. Chemical acceleration in a low temperature environment is possible with
both K4M and LR White, but local temperatures may be variable and could potentially
exceed an acceptable limit. Additionally, the cytoplasm of the spore may not obtain an
43
adequate amount of accerator to polymerize properly. LR White polymerized in a 50-
60°C oven thus appeared to be the best choice for this work. The potential for heat
damage to potential binding sites was recognized, and accepted as part of the
cytochemial reagent screening procedures.
LR White will tolerate up to 1 2% water in the tissues and still polymerize (Stirling,
1989). Dehydration through 95% alcohol is therefore not necessary. So as not to push
the limits of the resin to a critical point, and keeping in mind that the stock alcohols
used for dehydration may contain slightly less alcohol than the label suggests due to
evaporation, dehydration through 95% alcohol (ethanol) was proposed.
Newman and Hobot (1 987) recommended a rather short infiltration period with
several changes of fresh 100% resin to avoid or reduce extraction. This recom-
mendation was not followed because of the diffusion limits potentially imposed by the
cell walls. A series of dilutions in 95% ethanol followed by several changes of 100%
resin were proposed for the infiltration process.
Spurr resin rather than Epon was proposed for use with samples prepared for
morphological study. Spurr resin is less viscous than Epon and therefore can infiltrate
tissues with greater ease than Epon. Although Epon provides better morphology, for
fungi and other other organisms with heavy cell walls, infiltration is the more critical
factor. Spurr resin will not polymerize properly if water is present in the tissue, and
therefore tissues used for this purpose needed to be dehydrated through ethanol and
acetone before the infiltration process began.
CHAPTER 3
DEVELOPMENT OF ANTIBODIES
Introduction
Antibodies
Antibodies, or immunoglobulins, are glycoproteins that make up the fraction of ^
blood plasma called gamma globulin. Immunoglobulins are produced when a chemical,
or chemicals, recognized as foreign is present in the body. It is part of the immune
response. The specific chemical an antibody is made against and will bind to is called
an antigen. That part of the antigen molecule which is actually bound by the antibody
is the antigenic determinant or epitope and is typically 5-7 residues of a polymer
(Goding, 1986). A single foreign molecule can have several antigenic sites. For
example, lysozyme has 8 predominant antigenic sites (Sercarz et al., 1 974). The binding
of an antibody to its target epitope on the antigen is highly specific.
In their classic paper, Kohler and Milstein (1975) introduced a way to construct
hybrid B-lymphocyte/myeloma cells (hybridomas) which can make antibodies. All of the
antibodies produced by a single hybridoma clone have the same amino acid sequence
and hence have the same binding properties (Edwards, 1981). These are called
monoclonal antibodies. They can be selected for a predefined specificity and thus have ^
become a valuable laboratory tool, although they have not diminished the need for
polyclonal antibodies.
Antiserum developed against an antigen typically contains antibodies to a
44
45
number of antigenic determinants on that target antigen. These antibodies are not
derived from a single genotype of B-lymphocyte and are therefore called polyclonal
antibodies.
As laboratory tools, there are pros and cons to both polyclonal and monoclonal
antibodies. When an antigen is purifiable, polyclonal antibodies are often preferred.
They will provide a precise identification of their target antigen whereas monoclonal
antibodies are unable to distinguish between a group of different molecules which all
bear the appropriate antigenic determinant (Edwards, 1981). Additionally, development
of polyclonal antibodies requires much less work than development of monoclonals.
A great deal of time spent "cell farming" and preforming hundreds or even thousands
of tests is typically required to develop a usable monoclonal hybridoma cell line and
antibody preparation (Goding, 1986), Conversely, if an antigen is not purifiable, or is
unknown at the onset of experiments, monoclonal antibodies make the identification,
assay, marking and purification of that antigen possible (Edwards, 1981). For
immunocytochemical experimentation the best polyclonal antiserum tends to be inferior
to monoclonal antibodies in terms of unwanted background (Mason et al., 1983).
Immunogens
When whole cells or isolated cell walls are used as immunogen, there are many
different potentially antigenic molecules present. Typically, in a molecularly diverse
immunogen some of the molecules present will be more antigenic than other molecules
present. The term "immunodominant" is sometimes used to describe this phenomenon
(Mason et al., 1983). This greater antigenicity, or immunodominance, results in a
stronger response to these molecules. Therefore, one cannot assume that antibodies
46
will be produced against a particular molecule of interest if several other molecules are
presented at the same time. On the other hand, if very little is known about a chemically
complex system, like the ascospore walls in the present study, any information gained
by this so called "blind approach" (Mason et al., 1983) can increase our knowledge of
the chemistry and biology of the system. In fact, the blind approach has been promoted
as a valuable tool for cytochemical research in cases where little is known about the
chemistry of a system (Sternberger, 1986). For the study of fungal antigens, Reiss
(1986) promoted the use of whole cells and/or wall fragments as immunogen.
Fungi and Fungal Walls as Antigens
In a recent review of fungal infections, fungi are described as poor antigens
(Khardori, 1989). Host non-specific and innate defense mechanisms such as intact
skin, mucus membranes, indigenous microbial flora, and the fungicidal activity of certain
cell types are apparently of greater importance than antibodies in protection against
opportunistic fungal infections (Khardori, 1989). The status (health) of the host rather
than the pathogenic properties of the fungus influence contraction and severity of fungal
diseases (Khadori, 1 989). Reiss (1 986) further specified chronic fungal infections as the
result of defects in immunoregulation controlled by thymic functions.
Despite this low antigenicity, there are a number of reports, particularly in the
medical literature of monoclonal antibody development against fungal antigens (e.g., for
Telletiasp., Banowetz et al., 1984; for Ophiostoma ulma, Benhamou & Ouellette, 1986;
for Phytophthora cinnamomi, Hardham et al., 1 985, 1 986; for Candida albicans. Brawner
& Cutler, 1986a, 1986b; Hopwood et al., 1986; Hospenthal et al., 1988; for Candida
tropicalis. Reiss et al., 1986b; for Aspergillus fumigatus. Ste-Marie et al., 1990). The
'■•fl
47
specific antigens and/or antigenic determinants reported for fungi include: peptido-L-
fucomannan (Miyazaki et al., 1980), a high molecular weight glycoprotein of
Phytophthora cinnamomi (Guber & Hardham, 1988). Candida tropicalis mannan (Reiss
et al,, 1986b), oligogalactoside side chains and mannopyranosyl side chains of a
Asperqillus fumigatus galactomannan (Ste-Marie et al., 1990), and M-protein of
histoplasmin from Histoplasma capsulatum (Reiss et al., 1986a). The major surface
antigens of fungi are thought to be mannans because; 1) Con A lectin agglutination of
C. albicans, which is inhibited by methyl-a-mannoside, 2) localization on surface of C.
albicans by the silver proteinate method, 3) ultrastructural localization with Con A on
surface of Sporothrix schenckii, 4) chemical analysis after digestion of Histoplasma
capsulatum walls with various glucanases, and 5) mannans were detected in fractions
of C. albicans walls that had been extracted with cold dilute alkali (Reiss, 1 986).
Production
Information and discussion on the production of monoclonal antibodies is
abundant and easily found in immunology text books (e.g., McMichael & Fabre, 1982;
Coding, 1986), review articles (e.g., Edwards, 1981; Mason et al., 1983), and articles
pertaining to specific antigens (e.g., Reiss et al., 1986a, 1986b; Ste-Marie et al., 1990).
At this point in time too much information is available to adequately review the subject
as a whole, and thus only a few references shall be discussed.
Two immunization processes, in vitro and in vivo, are currently used to develop
antibody producing cell lines. The in vitro techniques for development of antibody were
first deomonstrated by Mishell and coworkers (1 967). Basically, these techniques differ
from the in vivo techniques by immunization of non-immune B-lymphocytes (B-cells) in
4
*
■•1
48
culture rather than immunization of a mouse (or other mammal). The in vitro method
is preferable when antigen is limited (Pardue et al., 1983), or a weak antigen is of
interest (Pardue et a!., 1983; Borrebeck & Moller, 1986; Brams et al., 1987). For this
study these advantages were not sufficient to warrant the extra time required to learn
the techniques or additional time spent developing a specific protocol and "cell farming."
The in vivo method requires that mice be immunized. When whole cells or cell
fragments are of interest, 1-5 x 10^ cells or parts per immunization is recommended
(Bastin et al., 1 982; Mason et al., 1 983). There is diversity in the literature as to the most
appropriate immunization schedule (Prabhakar et al, 1984). A final intraperitoneal or
intravenous immunization 3 days prior to removal of the spleen is the most universally
accepted procedure (Prabhakar et al., 1984).
After mice have been immunized for a sufficient period of time their serum can
be tested for the presence of antibodies against the antigen of interest. If such
antibodies are present, a fusion of spleen derived B-lymphocytes and a appropriate
myeloma cell type can be made to produce hybridoma cells. Fusion of the cells can
be accomplished using Sendai virus (Kohler & Milstein, 1 975), or with polyethyleneglycol
(PEG; Mason et al., 1983; Prabhakar et al., 1984). During the fusion process it is
possible for B-cell/B-cell and myeloma/myeloma fusions to occur. Some cells may
remain unfused. Selective media are used to prevent these unwanted cell types from
growing and perhaps overgrowing the hybridomas (Mason et al., 1983). Although
Mason and co-workers (1 983) described the process of cell fusion as being inefficient
in that only a small minority of cells undergo fusion, fusions can yield hundreds to
thousands of individual hybrid cell lines (Edwards, 1981). Of the cell lines produced by
a fusion only about 10% produce antibody to the antigen used to immunize the mouse
(Edwards, 1981).
49
Once colonies of hybridomas begin to expand rapidly or fill about 1/2 of any size
well, the culture supernatant can be tested for the presence of antibodies. Several
assays have been described for detection of antibodies in hybridoma culture
supernatant including radioimmunoassay (RIA), enzyme-linked-immunosorbent-assay
(ELISA) and immunofluorescence (FA). The choice of assay has been described as
"extremely crucial" because the assay can greatly affect the selection of antibodies with
different specificities (Prabhakar et al., 1984).
ELISA
ELISA is a commonly used assay for detection of antibodies in hybridoma culture
supernatant. It has the advantage over RIA's in not requiring any radioactive reagents.
It usually requires less preparative effort than cytochemical methods such as the FA.
The ELISA is based on the premise that an immunoreagent (e.g., antibody, 1
antigen) can be immobilized on a carrier surface while retaining its activity or capicity
for binding (Voller & Bidwell, 1980). Typically the process requires adsorption of the
relevant antigen to wells of plastic microtiter plates, incubation with the test samples,
incubation with an enzyme-labeled antibody which is directed against the antibody in
the test sample (e.g., goat-anti-mouse antibody), incubation with the appropriate enzyme
substrate, then stopping and photometric determination of the reaction (Voller & Bidwell,
1980).
Antibody Containing Products
The antibody-containing products are serum, ascites fluid, and hybridoma culture
supernatant. Immune-mouse blood is collected at the time a sacrifice is made. This
I
50 ,'^"'
serum provides a positive control while serum from a non-immune, or normal, mouse ' '^
provides negative controls for both ELISA and EM screenings of hybridoma culture
supernatants.
:1
Hybridoma culture supernatant contains a sufficient quantity of antibody (5- :
25^,g/ml; Edwards, 1981) to not only allow ELISA screening, but also be a useful source
for cytochemistry.
Ascites fluid is produced by planting hybridoma cells in the intra-peritoneal cavity . y;;|
of a live mouse. There they typically grow, divide and promote production of fluid ]
(ascites) which has a high titre of antibody (0.1-1mg/ml; Edwards, 1981). In addition to
the antibody of Interest, ascites fluid will contain a number of non-specific serum
immunoglobulins which can produce background staining in immunocytochemical
experiments (Mason et al., 1 983). Based on the Mason's arguement, ascites will not be '"'^
used in this project. , |
Materials and Methods
Fungal Cultures •.:
Cultures of Ascodesmis sphaerospora (culture #260) were kindly donated by Dr. ►>
J.W. Kimbrough. Cultures were grown on corn meal/malt extract/yeast extract (CMMY)
medium. No unusual treatment or growth conditions were required, although petri
plates were sealed with parafilm so that moisture would condense on the lids and trap
expelled ascospores. The exact formula for CMMY and growth conditions are given in
Appendix A. Cultures were allowed to sporulate for 1 to 3 weeks before the ascospores
were collected and processed.
A
•A-
"*!
^<
51 '
Isolation of Immunoaen
Ascospores that collected in the condensation on the lids were swept together
with a rubber policeman and transferred to flask with a Pasteur pipet. The spores were j
allowed to settle to the bottom and the excess fluid removed. The spores were then ^
broken in a Braun homogenizer and separated from the cytoplasmic components by
centrifugation over a steep sucrose gradient. The spore fragments were washed several
times, fixed with 0.5% glutaraldehyde and 1 % formaldehyde, washed several times more '^
with buffer (cacodylate once, then PBS) before they were used as immunogen. Details 'm
of the entire process are given in Appendix B. ^ ;
The concentration of wall fragments per ml was determined by making three 10-
fold serial dilutions, then counting fragments on a hemacytometer. Antigen was
prepared twice. The first preparation contained approximately 4.5 x 1 0^ parts per ml and
the second contained about 7x10^ parts per ml. Mice were given approximately 2-3.5
X lO'' parts per immunization.
Immunization
Two sets of mice were immunized. From the first set of 4, a single mouse was
sacrificed after two months of regular immunization (once every other week) and a final
boost three days prior as described in Appendix C. A second mouse was sacrificed
after that initial two months, a 3 month break, and another 2 months of regular
immunization and pre-sacrifice boost. The other mice in this set died due to unforseen
circumstances. When these were lost, immunization of another set was begun
immediately so that mice would be available whenever needed.
i.-f-
52 ; .
Only one mouse was used from the second set of 3. Its immunization schedule
■'■■■ '"'---i
was similar to that of the second mouse from the first set.
Hybridoma Production and Cloning . ^ ^
'I
The complete protocol used for hybridoma production and cloning, including ''"^-''_
materials, is given in Appendix C. A total of 3 fusions were performed.
Feeder Cells
Use of feeder cells (which can be peritoneal exudate cells, thymocytes or B-cells
of a normal mouse) is recommended especially with when transferring hybridomas to
a larger volume or cloning (Prabhakar et al., 1984; Coding, 1986). Thymocyte and/or
B-cells were used for these purposes here. The thymus and/or spleen was removed
from young mice and pushed through a metal mesh screen. They were then diluted to
approximately 1 0^ cells per ml in appropriate media and either plated out at 1 0O^x I per
well of a 96-well plate or placed in a sterile growth flask and cultured as for hybridomas.
^>'^
«
I
I
ELISA
The indirect-method as outlined by Voller and Bidwell (1980) and was used with
alterations as needed for this antigen/antibody system. Those alterations were: 1) the -.:,
antigen was dried down onto the plate in a 37°C oven overnight, 2) test samples
contained mouse antibody rather than human, 3) both PBS and tris (0.02M)/high salt
(0.5M)/tween (0.1% v/v; THST) antibody buffering systems were tried, 4) the secondary
>
enzyme labeled antibody was alkaline-phosphatase-goat-anti-mouse IgC/IgM specific
X^ „. :. (Jackson Laboratory), and 5) the enzyme substrate was p-nitrophenylphosphate. Plates
' : H ' . ' . ■
r iJk ' ' * '■ . ..; ■■'
53
were read on a SKT Labinstruments EasyReader SF Plus microtiter plate reader using
a 405 nm filter.
It became apparent after several screenings that the background on the assays
done was unacceptably high. Two experiments were designed and run to demonstrate
this statistically. The first experiment run was without a primary antibody and tested
both substrate alone and secondary antibody with subsequent substrate step. These
variables were nested within a buffer wash trial comparing PBS with THST. The second
experiment compared buffer negative controls, immune-mouse serum and normal-mouse
serum. SAS statistical analysis of this data was done using PC SAS version 6.03 (SAS
institure Inc.) program, on an IBM PS 2 computer.
EM Screening
EM screening methods are given in Appendix F, and are discussed in chapter
4.
Collection and Storage of Products
Immune-mouse blood was collected at the time spleen was removed. Heparin
was added to prevent coaggulation. The blood was heated to 60°C and held at that
temperature for 30 minutes to inactivate serum proteases and other enzymes. The cells
were removed by centrifugation (Dynac centrifuge, speed setting 90-100, for 30
minutes). Serum was pipeted off and further diluted with (1 M) PBS with 1 % w/v sodium
azide for a final dilution of 1/10. Serum was then frozen and stored at -80°C. Normal
mouse serum was handled in exactly the same manner.
When small volumes of culture supernatant were harvested but not used
M
54
immediately, the supernatant was stored in microfuge tubes at either 4°C, -20°C, or -
80°C depending on the projected time of use. Several of these went through several
freeze/thaw cycles and were damaged.
Hybridoma cells were removed from large volumes by centrifugation (Dynac
centrifuge, speed setting 5, for 8-1 0 minutes). Sodium azide was added (0.1 % w/v) and
then the supernatant was aliquoted, frozen and stored at -80°C.
The protocol for preparation and freezing of hybridomas in given in Appendix D. ^
Results ^:'
Immunoqen j.i
It should be noted that the concentrations (fragment particles/ml) determined
using the hemacytometer for wall fragments per ml are not highly reliable. The reasons
'I
are: 1) The shape of fragments was very inconsistent and some of them may have
been smaller than can be resolved on the light microscope when using a hemacyto-
meter. 2) They were additionally diluted by sticking to plasticware, glassware, and each
other, hence reducing the apparent count. -
Immunization
The second and third fusions used mice that had an extended immunization
regime. These mice had greatly enlarged spleens. Si
Hybridoma Production and Cloning
Three fusions were preformed. The first fusion required 4 - 96-well plates and
the second and third required 9 plates each. The increase in plates required per fusion j
-4
'^^'-w.
■ : ; ■ ^/ . . 55
was most likely due to an increase in the number of cells per spleen. Additionally, the
B-cells were removed from the spleen by the syringe method (Appendix C, method 2)
in the last two fusions. This is the more gentle way to release the B-cells and survival
of B-cells was probably improved.
All 3 fusions had a high hybridoma recovery rate. Between 70%-90% of the wells
had viable hyridomas after 2 weeks of feeding with selective media. A majority of those
tested positive with ELISA testing.
Over 100 mixed cell cultures from fusions were screened using EM techniques.
Of these cultures six were targeted for expansion and coloning.
Products
Three highly useful antibody-containing culture supernatants were identified and
proven in EM screening. It was possible to clone only 2 of the three. The third was lost
in the first expansion into a 24-well plate.
Hybridoma cells from over 100 mixed cell cultures and several vials all of the
cloned cell lines were frozen for future use.
ELISA
SAS data is given in tables 3.1-3.4. Optical densities of the 3 treatments 1 ) buffer
washes only, 2) substrate treatment, and 3) both secondary and substrate treatments
are compared using log-transformation of raw data and the least-square mean test in
table 3.2. These data demonstrate that the optical density of the substrate only
treatment was significantly different from that of the buffer washes. Further, this analysis
shows that the optical density when secondary antibody treatment was included was
• ^*
',■■ •>;
56
significantly different from the substrate only treatment. It is evident from the means
based on raw data (table 3.1) that these significant differences are due to increasing
optical density. This indicates that a statistically significant increase in background
optical density occurs at each of these later steps in the ELISA assay protocol.
"XT,
Table 3.1 Mean optical densities for buffer wash, substrate, and secondary antibody
treatments.
Tab. 3.2 Tab. 3.3
Std. Dev. LS # LS #
Trtm.
NObs.
Mean
PBS
w/ 2° AB
8
0.5035
w/ sub.
PBS
w/o 2° AB
8
0.0489
w/o sub.
PBS
w/o 2° AB
8
0.0764
w/ sub.
THST
w/ 2° AB
8
0.3030
w/ sub.
THST
w/o 2° AB
8
0.0558
w/o sub.
THST
w/o 2° AB
8
0.0896
w/ sub.
0.0782
0.0034
0.0018
0.0718
0.0079
0.0093
1
1
5
6
.y
sub., substrate
2° AB, secondary antibody
Tab., table
LS #, least square mean number
w/, with
w/o, without
57
Table 3.2. Least square means comparison of washing vs substrate and antibody/
substrate treatment. (Data log transformed; x < 0.05 indicates a
significant difference.)
Pr > |T| HO: LSMEAN (i)=LSMEAN (j)
i/i 1
2
3
1
0.0001
0.0001
2 0.0001
0.0001
3 0.0001
0.0001
Table 3.3. Least square means comparison of interaction of buffer type with
treatments. (Data log transformed; x < 0.05 indicates a significant
difference.)
Pr > |T| HO: LSMEAN (i) = LSMEAN 0)
c ■.- </
i/1
1
2
3
4
5
6
1
0.0001
0.0001
0.0001
0.0001
0.0001
2
0.0001
0.0001
0.0001
0.0636
0.0001
3
0.0001
0.0001
0.0001
0.0001
0.0228
4
0.0001
0.0001
0.0001
,
0.0001
0.0001
5
0.0001
0.0636
0.0001
0.0001
0.0001
6
0.0001
0.0001
0.0228
0.0001
0.0001
ii
• *.A;
4
•^
The type of buffer used for washing also affects the amount of background
optical density. As would be expected, there was no statistically significant difference . ;^
..."
in the optical densities of the two buffer controls (table 3.3, LS# 2 vs 5). On the other
hand, when antibody was tested, there was a statistically significant difference in optical
density readings between the PBS and THST buffer washes (table 3.3, LS# 1 vs 4). i^
From the raw data means (table 3.1 : 0.5035 vs 0.3030) it is apparent that THST buffer ' "^
washes reduced the background in the system.
The mean optical densities of the sera are given in table 3.4. The high standard
deviations of the serum means are the result of pooling data from 1/500 and 1/1000
serum dilutions. The mean optical densities for normal-mouse serum in both buffer
58
wash systems were high enough to be considered positive for anti-wall antibodies.
Each of those means are over 1 .5 standard deviation units greater than the buffer
control (which did have the secondary antibody treatment).
Table 3.4: Mean optical densities for buffer control, immune mouse
and test mouse sera.
■ U^
Trtm.
N Obs.
PBS
PBS
4
PBS
IM
4
PBS
NM
4
THST
THST
4
THST
IM
4
THST
NM
4
IM = immune mouse
NM = normal mouse
Mean
0.3692
2.4285
1 .0942
0.3518
2.0472
0.7628
Std. Dev.
0.0391
0.1749
0.3609
0.0350
0.3359
0.2314
4
Discussion
Hvbridoma Production
The great success of these fusions in terms of hybridoma recovery and apparent
production of anti-fungal antibodies (via ELISA testing), especially the second fusion,
was overwhelming and many lines were lost to poor management and inexperience.
'A
•r.
59
All 3 of the antibody preparations used came from the second fusion. The third fusion
was done primarily in an effort to reproduce antibody 8F1 1 . The effort was apparently
unsuccessful but it also provided an opportunity to try a different management system.
In the first 2 fusions ELISA testing began as soon as hybridoma colonies filled 1/4 to 1/2
a well in the 96 well plates and those wells which tested positive were immediately
expanded and cloned. In the third fusion, after an intial growth period, cells from 8 wells
were transferred into a single well of a 24 well plate and allowed to expand before
testing. After 2-5 days growth in the 24 well plate, 100|il of culture supernatant was
harvested for testing and the cells were frozen. This method required about 3-4 weeks
of growth with only 10-20 hr of labor a week to take cells from fusion to freezer, vs 2-3
months of 40-60 hr per week labor of the previous method required. The supernatant
could then be stored and tested at a convienent time. After testing, cells could be
brought out of the freezer in small numbers, cultured, retested, expanded, and cloned
at a convenient time. Athough time has not permitted further work with the cells from
the third fusion, they are available.
ELISA
Several problems were encountered with the ELISA system for this antigen. The
wall fragments are heavy and sticky which made preparing the plates difficult and time
consuming. These factors also made the particle count per well unreliable. Background
from the secondary antibody-enzyme conjugate was sufficient to make some negative
results appear positive or hide low concentrations of antibody that might be expected
from a colony which is just establishing itself. Normal-mouse serum also appeared to
contain reactive antibodies with this system, whereas in the EM screening, no significant
v-^^
^'4
■,'-.. ti<^
•a
60
labeling occurred. For these reasons the ELISA was found to be not only a great deal
of trouble but an ineffective assay system for this antigen.
Mason and coworkers (1 983) and Sternberger (1 986) expressed a preference for
■3
EM screening of hybridoma culture supernatants when the final use is to be :,;;:.|
immunocytochemistry. Mason and coworkers (1983) based their preference on the *^
arguements that 1 ) monoclonal antibodies which react strongly in one assay procedure
do not always give satisfactory results in another unrelated assay system, and that 2) -'^4
■i
the results from immunocytochemical techniques are inherently more informative
(providing not only +/- results, but specific background and localization data). Based
on these opinions, arguements and experience with the ELISA, any future screening for
anti-ascospore antibodies will be done using cytochemical techniques. It is further
suggested that cytochemical techniques be used when screening for antibodies against
any fungal wall system if the antigen is wall fragments and the intended final use is
immunocytochemical. Assessment of the value of the labeling information then
becomes a part of the screening process.
• <;
.^
CHAPTER 4
IMMUNOCYTOCHEMISTRY
Introduction
Brief History of Immunocvtochemistry
The practice of cellular localization began in the 1830's with Raspail's
"microchemistry," or chemical analysis In combination with microscopic examination
(Raspail, 1830). The immunological approach in histochemistry (light level cell
chemistry) was introduced by Coons and coworkers (1941). They used fluorescent
conjugated antibodies to identify sites of antigen-antibody reaction at the light
microscopic level. Development of an electron-dense marker was necessary for
immuno-labeling to be applied to electron microscopy (immuno-cytochemistry). Singer
(1959) introduced the use of ferritin as an electron-dense marker. Nakane and Pierce
(1 966) described the application of horse-radish-peroxidase (HRP) and diaminobenzidine
(DAB) reaction to histochemistry. Immunogold techniques, and use of gold as as
electron-dense marker for electron microscopy were introduced by Faulk and Taylor
(1971) and later by Romano and coworkers (1974). Lectin-gold techiques for
microscopy were described in articles such as Roth (1983). The avidin-biotin-gold
system (Tolson et al., 1981) and enzyme-gold techniques (e.g., Bendayan, 1981, 1982)
were also described in the early 1980's. In the past 20 years histochemistry has
progressed from the use of stains which are capable of identifying classes of molecules
such as deoxyribonucleic acids to the use of probes and techniques which are highly
61
)
^ W^
r
4
62
specific for particular substrates and that can demonstrate subcellular location. Causton
(1984) described immunocytochemistry as potentially being the most demading of all
the staining techniques.
Immunolabelino of Fungi
Several publications have used immunocytochemistry to examine fungi.
Localization of ligninperoxidase in Phanerochaete chrysosporium is reported by Daniel
and coworkers (1989). Ste-Marie and colleagues (1990) report development of 2 anti-
Asperaillus fumiqatus monoclonal antibodies. The first, MAbI , labeled the inner cell wall
of hyphae and conidia, and intracellular membranes. The second, MAb40, bound
hyphal and conidial walls more diffusely and intracellular membranes less intensely.
This second antibody was also found to recognize the cell walls of Candida albicans :J. ■:!*
serotype A. Brawner and Cutler (1986b) demonstrated variable expression of cell
surface antigens in Candida albicans during spore germination using 2 monoclonal
antibodies (H9 and C6). Phvtophthora cinnamoni zoospore encystment was found to
be induced by specific lectin and antibody binding to the cell surafce (Gubler &
Hardham, 1988). Undoubtedly, more publications exist, particularly in the medical -M
literature. Reiss (1986) described localization studies as an important step
subsequent to the development of antibodies and characterization of their determinants.
His work has primarily been in the field of medical mycology.
Charaterization of Antigenic Determinants
Most naturally occurring antigens are proteins and carbohydrates including
glycoproteins and glycolipids (Coding, 1986). In any case, some idea of the nature of
H^
63
the determinant is desirable. A number of simple tests have been used with ELISA,
Western blots, and thin layer chromatography. These include heat treatment, proteinase
treatment (notably pronase and trypsin), and periodate treatment (Goding, 1986).
Proteins are typically sensitive to proteinase and heat but not periodate, while the
converse typically is true for carbohydrates. Yet these tests are not absolutely
diagnostic since some proteins resist digestion by proteinases and some carbohydrates
are insensitive to periodate. Additionally, the amino acids tyrosine, tryptophan and
methionine may react with periodate (Geoghegan et al., 1982; Yamasaki et al., 1982).
Periodate has successfully been used as a pretreatment for antibody labeling on
sections of osmocated tissue (Bendayan & Zollinger, 1983). These authors
demonstrated improved labeling with this pretreatment, but it should be noted that the
antigens of interest were proteinaceous.
Materials and Methods
Experimental Organism
The choice of Ascodesmis sphaerospora as the experimental organism was
explained in chapter 1 . Conditions under which it was grown and spores harvested
were given in chapter 3. For electron microscopy both ascospores and apothecia were
harvested and prepared for study. Spores were collected in the same manner as for
preparation of immunogen but were handled differently thereafter as explained below.
Apothecia were monitered for development and harvested just after spores were noticed
in the water droplets on the petri dish lid; usually 10-14 days after inoculation. At this
point it was thought that most of the ascospore developmental stages would be
represented, yet the culture was relatively young and active.
.,1
'^
1 .
-^
. -f .
64
Tissue Preparation
Ascospores were prepared for electron microscopy by an inital fixation step and -■
then "gently" breaking them with vortex and glass beads followed by another fixation
step, dehydration, and infiltration. The detailed protocol is given in Appendix E. "f
i
A general protocol for fixation, dehydration, infiltration, embedding and infiltration
of apothecia is given in Appendix F. Blocks of agar (approx., 0.5cm x 1cm-2cm x
0.25cm-0.5cm) were cut from cultures of sporulating A. sphaerospora and prepared. .. %^*i
Additionally, pelleted apothecia were prepared by flooding plates with the various ]
reagents up to 75% or 95% ethanol step of dehydration. At this point apothecia were ~ <^
scraped off of the agar and treated as a suspension and pelleted between every step
thereafter.
Table 4.1 demonstrates the specific variations in fixation, dehydration, and resin ' .'\\
used to prepare material for electron microscopy. ^.
Pseudoplectania nigrella, used for comparitive work was collected in the Oregon *~-
coastal range in March, 1 990. Sections of apothecial tissue were fixed for one hour on
ice in the field immediately upon collection. One set of tissue samples was fixed with ;
2% glutaraldehyde and 2% formaldehyde (block 225-A'). This set was later split, and
half was post-fixed with osmium tetroxide (block 225-A). Another set of tissue samples
was fixed with 1% glutaraldehyde and 2% paraformaldehyde (block 225-B). Tissue
samples not post-fixed were embedded in LR White (blocks 225- A & A'). Dehydration J
and further processing was as in the protocol given in Appendix F. Post-fixed tissue
was embedded in Spurr resin. Sections for EM labeling experiments were cut from
block 225-B.
65
Sectioning
Pale gold to silver (70-90 nm) sections were cut on a RMC MT6000-XL
microtome or a LKB 8800 ultramicrotome III. For cytochemical experiments sections
were placed on formvar (0.25%-0.3% powder w/v in ethylene dichloride) coated 75 or
100 mesh nickel grids. Formvar coated 75 mesh copper grids were used for
morphological study.
- ■■-'/"-
The Antibodies
The development and cloning of the three antibodies tested were the subject of
chapter 3. The antibodies used for immunolabeling were in the hybridoma culture
supernatant. Culture supernatants were diluted 3/4, 1/2 and 1/4 in either PBS or THST
buffer for labeling experiments. The three antibodies primarily used will be hereafter
referred to as 8F1 1 , 1 2-2, and 41 -1 .1 . The latter two are monoclonal.
Normal-mouse and immune-mouse serums, and PBS and THST buffers were
used as controls.
Experimental Immunocvtochemistry
Several general types of experiments were performed: screening of hybridoma
culture supernatants, testing of positive and negative serum controls, monoclonal jj
labeling with special attention given to finding developmental sequences, determinant
characterization, and determinant unmasking.
A general protocol, with special notes for determinant and unmasking steps, is
given in Appendix E. This general protocol was established as effective by
experimenting with positive and negative serum control on sections cut from various
£♦3
66
Table 4.1: Tissue preparation and embedding.
BLOCK NUIVIBER
1
SP
1
2
3
4
5
Immunogen
X
Spores (LB)
X
AC undisturbed
X
X
X
X
X
AC pelleted
30min. 1°fix
X
X
X
45 min. 1°fix
X
60 min. 1°fix
X
X
X
0% G / 4% F
X
X
0.5% G / 0% F
X
0.5% G / 2% F
X
0.5% G / 4% F
1 % G / 2% F
X
X
1%G/3-4%F
2% G / 2% F
X
Os04
X
95% EtOH
X
X
X
X
X
X
Acetone
X
LR White
X
X
X
X
X
X
Spurr's
X
LB, ligiitly broken
AC, ascocarp
G, glutaraldehyde
F, formaldehyde
67
Table 4.1 continued.
BLOCK NUMBER
11
12
13
14
15
16
26
Immunogen
Spores (LB)
AC undisturbed
AC pelleted
X
X
X
X
X
X
X
30 min. 1°fix
X
X
45 min. r fix
60 min. 1°fix
X
X
X
X
X
0% G / 4% F
0.5% G / 0% F
0.5% G / 2% F
0.5% G / 4% F
X
X
1%G/2%F
1%G/3-4%F
X
X
2% G / 2% F
X
X
X
Os04
X
X
95% EtOH
X
X
X
X
X
X
Acetone
X
j LR White
X
X
X
X
X
X
! Spurr's
X
v-'l.
.■ ■> • -JM
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blocks with 1/100, 1/1000 and 1/10,000 serum dilutions. All of the experiments more or
less follow that protocol with major differences being the particular blocks (tissue
preparation) used.
Sections used for screening of hybridoma culture supernatants were cut from the
SP block. This block was chosen for this purpose because the material was prepared
■,^fi
« »": ■ ■•■ • ■<■■{■:
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68
most like the immunogen (I block), yet the wall sturucture is less disrupted and it retains •" ■
cytoplasm so that cross-reactivity could be monitored. Further, because this boick is
a pellet of spores less trimming and facing was required than was necessary for the
apothecial blocks, thus reducing time and effort required to section.
Initial experiments with monoclonal antibodies were done using sections from
blocks 4, 1 4 and 1 5.
Determinant and unmasking experiments were also done using sections from a
variety of blocks, but most typically from the SP block to save time in preparing
sections. These experiments required a pretreatment of sections with saturated
periodate for 30 to 60 minutes at room temperature. Alternatively, tissues in section
were digested with 1 % (w/v) pronase in PBS for 60 minutes.
Specific deviations from the general protocol and origin of sections are given in
the individual figure captions and noted in the results section where appropriate.
Evaluation of Labeling
All evaluations of labeling are qualitative rather than quantitative. Qualitative -'^j$
evaluation is sufficient for gross determination of specificity, background, and labeling
density.
Results
Serum Labeling
Immune-mouse serum labeled all parts of the ascospore wall, from the spore
plasma membrane edge to the furtherest tip of secondary wall ornament (figs. 4.1 A &
4.2B). Ascus and vegetative cell walls were labeled to a much lesser extent (fig.4.2A).
■ • ' ■- ■/ '.■■■'..
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69
Additionally, sporoplasm, epiplasm (ascus cytoplasm), and vegetative cell cytoplasm
components were specifically labeled. Conversely, normal-mouse serum did not ' -^
rl
specifically label any part of the fungus, and the background labeling was minimal (fig. -'i
4.1 C). The buffer negative control also had minimal background labeling (fig. 4.2B).
Antibody Screening
Between 200 and 300 culture supernatant screenings were done during the
processes of identification and cloning of anti-wall antibody-producing hybridomas. Not
only were 8F11, 12-2 and 41-1.1 identified and used, but 5 other monoclonal lines from
culture 12 and 2 others from culture 41-1 were identified. Enough culture supernatants '';
from these later lines exist for further testing with them when desired.
None of these screenings were done with periodate- or pronase-pretreated
sections. It is now obvious from the pretreatment results with antibodies 1 2 and 41
(figs. 4.7, 4.8 & 4.9), both cloned and uncloned, that some of the supernatants that had
only scanty but apparently specific labeling may have actually been quite good if the
sections had been pretreated.
Antibodv 8F1 1
Although not a monoclonal, antibody 8F1 1 performed as specifically on sections
as the monoclonals similarly tested. It labeled the primary wall and sporoplasmic
vesicles (fig. 4.3A). Cell wall labelling was typically restricted to the outer 2/3 to 3/4 of -
this wall layer (figs. 4.3A, 4.4A & B). From the experiments performed it is impossible
to determine if the wall and vesicle labeling are the result of the same antibody.
Demonstration of identical antibody labeling on the cell wall and vesicles could be
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A) labeling of immune-mouse serum (from second fusion, diluted 1/1000);
B) buffer negative control. «
Figure 4.3. 8F1 1 culture supernatant labeling on A. sphaerospora.
A) Labeling on the ascospore wall and sporoplasmic vesicles (pointers);
B) Labeling on the vegetative wall, including septum.
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A & C) positive control, without pretreatment;
B) pretreated with periodate;
D) pretreated with pronase.
accomplished by competing off the anti-wall antibodies with clean wall preparation (such
as that used for immunogen) prior to incubation of the section(s). This experiment was
not preformed due to the limited quantity of this antibody preparation.
The antigenic determinant was both periodate and pronase sensitive (fig. 4.5),
suggesting a glycoprotein antigen or conformational determinant, or release of the
antigen from the sections. A conformational determinant, in this case, could occur
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when a protein and carbohydrate were closely associated, but not covalently bound
together. The antigenic determinant must be exposed in section, rather than buried in
the wall as no pretreatment of the sections was required for labeling.
This antibody preparation labeled ascospores only in the late stages of the
developmental sequence (fig. 4.6).
•1
Anitbody 41-1.1
This monolonal antibody labeled an inner (sporoplasmic) layer of the primary
ascospore wall (fig. 4.7). Labeling was evident in every developmental stage examined.
Figure 4.8. Pronase pretreatment with antibodies 1 2-2 and 41 -1 .1 ,
A) antibody 41-1.1;
B) antibody 1 2-2;
C) buffer negative control with pronase pretreatment.
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sections through the primary wall. This suggests that the determinant was somehow
buried in the section.
This antibody has been found to work well with tissue that has been fixed with
2% glutaraldehyde and post-fixed with osmium tetroxide.
Antibody 12-2
Monoclonal antibody 12-2 specifically labeled the secondary wall and a
sporoplasmic component (fig. 4.9). A complete developmental sequence was not
present in sections thus far tested for labeling with this antibody.
The antigenic determinant was neither periodate nor pronase sensitive, and in
fact both pretreatments improve labeling (fig 4.8B).
Interspecies Cross-Reaction
Antibodies 1 2-2 and 41 -1 .1 were tested for labeling on Pseudoplectania niqrella.
These antibodies did cross-react with this species although they did not label the walls.
Antibody 12-2 does not apparently label any part of the ascopore wall, but quite
specifically labeled the sporoplasm as it does in Ascodesmis sphaerospora (fig. 4.1 OB).
Antibody 41-1.1 specifically labeled a component within the perisporal sac, although this
material does not seem to condense on the wall as there was no wall labeling (fig.
4.1 OA). P. niqrella is the only other species these antibodies have been tested on to
date.
r^'i
81
The antigenic determinant is pronase sensitive (fig. 4.8A) and periodate (fig. 4.7A) ' '
insensitive. This suggests a protein, proteinecous hapten, or glycoprotein antigen. -- , ^
Notably, labeling of sections not treated with periodate was only seen on tangential > :4
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Discussion
Serum Labeling
Serum labeling is important because it demonstrates potential of the mouse B-
cells and can demonstrate cross-reactivity with other wall systems. Labeling is evident I
on all areas of the ascospore wall. This indicates that either one immunodominant - -
antigen occurs throughout the entire wall or that antibodies are being made to various
antigens in every layer of the ascospore wall. The latter would appear to be the case,
since antibody preparations 8F11, 12-2, and 41-1.1 all label different areas of the wall,
and all
have different sensitivities to periodate and pronase.
Ascus and vegetative walls were labeled with immune serum. This would
indicate cross-reactivity and common antigens (or at least determinants) in these wall
systems if no vegetative wall contaminated the immunogen. It is quite possible that a
very small amount of hyphal wall material from germinated spores was present in the
immunogen. A conclusive statement of cross-reactivity cannot be made at this time for
that reason.
Antibody 8F1 1
Antibody 8F11 labeling definitively demonstrates a late maturation event in the
ascospore primary wall layer. Interestingly, labeling did not built up through the
developmental sequence, but appears quite suddenly. The vesicle labeling was
concurrent with and as sudden as the wall labeling. Relative to the developmental
process, the time that 8F11 labeling appears coincides with the first appearance of
fixation and infiltration artifacts in the sporoplasm. It seems quite possible that the ' ^
appearance of this determinant at this time is either a result of a "sealing" process, or
. -A
i
I
87
part of that process. While it is possible that the antigen is inaccessible during early ,
development, such inaccessibility is in direct contrast with the sealing/protective function
and seems unlikely.
There was no strong evidence suggesting how the 8F1 1 determinant got into its
position (outer 2/3 to 3/4) within the primary wall layer. The appearance of 8F11
labeling at this late maturation stage could represent an addition to the wall or an in situ
modification of the wall. If this is an addition to the wall, where are the synthesis
enzymes and precursors situated? The labeling in the outer area of the primary wall
layer with no apparent migration across the inner zone of the primary wall or across the \
'i
•)
secondary wall would suggest that the synthesis enzymes are in the wall. If the vesicle ■
labeling was due to the same determinant, precursor packaging could be suggested,
but the synthesis enzyme would also have to be present within the vesicle, since the i
labeling pattern was just as sudden for the vesicles as it was for the wall. If a wall <
\ -
constituent is enzymatically modified then enzymes would have to be in situ for both "?
walls and vesicles. On the other hand, there could be a physical or physicochemical !
factor such as hydration/dehydration and/or incorporation of divalent cations
■I
responsible for modification. An in situ modification seems the more probable of these '
i
two suggested processes because of the suddenness of labeling and greater diversity .^^
of ways for a modification to occur. ''
Antibodv 41-1.1
One of the most interesting aspects of labeling with antibody 41-1.1 is that a
pretreatment of sections with periodate is required for labeling in all but tangential
sections. What the periodate did to the plastic or to the ascospore wall that resulted
:H
in improved labeling is the obvious question. At least on Epon resin, periodate does
not appear to remove, or etch away the resin (Bendayan & Zollinger, 1 988). This tissue
was not post-stained with osmium tetroxide, so this is not a question of unmasking
determinants by removal of this fixative. Conversely, periodate is known to react with
sensitive carbohydrates by opening up the pyranosyl units. Between these points, and
the labeling of tangential sections through the primary wall, it seems most probable that
the antigenic determinant was buried in wall carbohydrates. It would appear that the '^
free path between wall molecules is a major influence on the outcome of this antibody's
diffusion into the section. This is similar to Causton's (1 984) description for epoxy resin 1
crosslinking density.
Antibodv8F11 and 41-1.1
It was obvious from the labeling patterns of these two antibodies, when taken
together (figs. 4.4, 4.6A & 4.7A), that there are distinct layers within the primary walls
and that these layers have different constituents.
s
■:l
Antibody 12-2
It is very interesting that labeling was improved by both periodate and pronase
pre-treatment of the sections. This may be the result of an alteration of the antigenci ,
• -" '-'1
molecule before the mouse immunoglobulin responce ensued. The immune system's
first response to a fungal invasion is a killer cell response and use of lytic enzymes
(Reiss, 1 986). It is possible that the mouse killer response slightly altered this molecule -M
before the immune response proceeded to production of immunoglobulins and that
periodate and pronase pretreatments of sections in some way mimicked that alteration.
ia«*^^i-'
89
Work with this antibody did not get far into the morphological aspects. In one
labeling test it appeared that the determinant might be sensitive to glutaraldehyde.
Sections from block 15 (2% glutaraldehyde in the fixative) did not label as well as
sections from block 4 (1% glutaraldehyde in the fixative), but conclusions can not be
drawn as yet because the sections from block 15 were unsatisfactory. Until the
morphology is improved it will not be possible to determine if this antibody Is an
appropriate probe for tracking precursors. It does label both epiplasm and sporoplasm
components and thus shows potential for being such a probe.
Highly specific labeling of sporoplasmic components, but not wall components,
was also demonstrated in Pseudoplectania niqrella. Further work with this antibody and
analysis of cellular labeling patterns could provide both phylogenetic and biological
information for large number of Pezizales. Generalization to the order is substantiated
by the fact that Ascodesmis and Pseudoplectania are distantly related. They are
members of different suborders within the Pezizales. Antibody 41-1 .1 shows the same
type of research potential, but perhaps slightly more limited as cytoplasmic components
were not strongly labeled.
^
^•1
CHAPTER 5
LECTIN CYTOCHEMISTRY
Introduction
Lectins are carbohydrate-binding proteins (or glycoproteins) of non-immune
origin which agglutinate cells and/or precipitate glycoconjugates (Goldstein et al., 1 980).
As of 1 986 no purified lectin had been shown to exhibit enzymatic activity (Goldstein &
Poretz, 1986).
Lectins can be classified into carbohydrate binding groups (Goldstein & Poretz,
1986; Benhamou, 1989b). These groups are: mannose/glucose binding, N-acetyl-
glucosamine binding, N-acetyl-galactosamine/galactose binding, sialic acid binding, and
L-fucose binding (Goldstein & Poretz, 1986). Some workers have made a distinction
between N-acetyl-galactosamine and galactose binding groups (e.g., Benhamou,
1989b). With the exceptions of sialic acid and L-fucose, all of these carbohydrate
groups are known to occur in fungi and were discussed in chapter 2.
The lectins within each of these categories differ markedly with respect to their
anomeric specificity (Goldstein & Poretz, 1986). Further, this specificity has been
attributed to the sterochemical fit between complementary molecules (Sharon & Lis,
1989). Carbohydrates are bound noncovalently by lectins (Sharon & Lis, 1989). Each
lectin differs with respect to its cross-reactivity with other sugars. Lectins further differ
in number of glycosyl units their binding sites can accommodate. Some lectins appear
S^
m
90
91
to only bind one glycosyl unit winile others, such as WGA, have an extended binding site
capable of accommodating 2-5 residues (Goldstein & Poretz, 1986).
For this study a lectin kit was purchased that had a representative lectin for each
binding group. Additionally, GS-II lectin was obtained as a gift from Dr. Katie Gropp.
These lectins, their binding specificities, and previous uses in fungal research are
reviewed below.
Binding Specificities
WGA / GS II
WGA (wheat germ agglutinin; Triticum vulgare) and GS-II (Griffonia simplicifolia)
lectins label N-acetylglucosamine, although in totally different ways. Both of these
lectins should detect chitin, but GS-II is not commonly used for this purpose. WGA has
been used as a probe for chitin in several pulbications (e.g., Benhamou & Ouellette,
1986; Benhamou, 1988; Simmons, 1989).
WGA is a dimeric carbohydrate-free protein (Goldstein & Poretz, 1986). It
apparently has a binding site which consists of 4 adjacent subsites (A-B-C-D; Allen et
al., 1973). Allen and coworkers (1973) envisioned sites A, B, and C as accommodating
N-acetylglucosamine while the D site could accommodate other glycosides. The B site
was further described as being able to handle a residue with a C-3 substitution, as in
N-acetylmuramic acid. This lectin has been shown to have an affinity for a number of
various oligomers, but far and away its greatest affinity is for pentamers, tetramers, and
trimers of N-acetylglucosamine (Goldstein & Poretz, 1986). N-acetylglucosamine is the
only simple sugar tested that binds to WGA (Allen et al., 1973). It has been suggested
that the monomer binds to subsite C (Allen et al., 1 973). Neither glucosamine nor 6(1 -4)
^
J' i
92
polymers of glucosamine (chitosan) bind with WGA (Goldstein & Poretz, 1986). WGA
has been shown to have an affinity for sialic acids (Goldstein et al., 1975; Mandal &
Mandal, 1990) which appears to be due to the similarity in configuration of the sugars
(Monsigny et al., 1980).
GS-II lectin is a tetramer of apparently identical subunits with one binding site
each (Ebisu et al., 1986). It is a glycoprotein with aproximately 4% carbohydrate
(Goldstein & Poretz, 1986). This lectin binds best to N.N'-diacetylglucosamine and
N,N',N"-triacetylglucosamine, although it has also been shown to precipitate with rabbit-
liver glycogen and, to a lesser extent, with Saccharomyces cervisiae mannan (Ebisu et
al., 1978). GS-II will bind both a and 6 anomers (Ebisu et al., 1978). GS-II lectin differs
from WGA in that it does not bind to internal 8(1-4) linked N-acetylglucosamine and
does not appear to possess an extened binding site for contiguous 8(1-4) linked
residues, Goldstein and Poretz (1986) described this lectin as being of particular
interest because it is the only lectin that interacts with terminal nonreducing Ck:- or B-N-
acetylglucosamine.
Benhamou and Ouellette (1986) localized N-acetylglucosamine in the walls of
Ascocalyx abietina with WGA lectin. Cell wall labeling with this lectin was also found in
both Ophiostoma ulmi and Verticillium albo-atrum (Benhamou, 1 988). Bonfante-Fasolo
and coworkers (1 990) used WGA lectin to localize chitin in vegetative cell walls and both
WGA and chitinase-gold for localization in the spore walls of Glomus versiforme
(Bonfante-Fasolo et al., 1986). Chitin in the bud scars of several yeasts was shown
using WGA (Simmons, 1989).
^>
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LFA ■'■'i
LFA lectin is derived from the slug Umax flavus. It is apparently a proteinaceous
dimer (Goldstein & Poretz, 1 986). It is specific for sialic acids (Miller et al., 1 982). Sialic
acid binding lectins are ubiquitous among invertebrates (IVIandal & Mandal, 1990). The
sialic acids are a family of about 30 derivatives of N-acetyl or N-glycolyl neuraminic
acids (Mandal & Mandal, 1990).
Using LFA lectin Benhamou and Ouellette (1 986) found labeling of lipid bodies
and a fibrillar network surrounding the fungal cells. In V. albo-atrum this lectin was also
found to label lipid bodies, but in O. ulmi there was intense labeling of the cytoplasm
and weak labeling on organelles, plasma membrane, walls and septa (Benhamou, 1 988).
v. , Con A
Concanavalin A (Con A) is a carbohydrate-free metalloprotein with 4 subunits.
Each subunit contains one Ca^^ and one Mn^^ ion (Goldstein & Poretz, 1986).
Con A was first reported to precipitate with glycogen and yeast mannan by
^ Sumner & Howell (1 936). Goldstein and coworkers (1 965) found that a polysaccharide
.^ J <; , must have a minimum of approximately 1 0%-1 5% non-(1 -6) linkages for Con A interact
f. .■-,■;
with it and form a precipitate. Even at high concentrations of Con A, linear
■ ^ polysaccharides were not found to precipitate (Goldstein et al., 1965). Yeast mannan
had almost 5 times the turbidity of glycogen and Goldstein and coworkers (1965)
J suggested that this may be due to the extensive branching of the molecule (34%).
<; Manners and Wright (1962) reported an approximately linear relationship between the -^A
extent of branching of a glycogen-like polysaccharide and the resultant turbidity. In
general, Con A can be described as binding to internal mannose (especially at
■l
t ;
94
branching points) and external glucose residues (Debray et al., 1981; Goldstein &
Poretz, 1986).
It is also noteworthy that the binding of Con A to sugar monomers, dimers or
polysaccharides is dependent on the concentration of Con A in solution (Goldstein et
al., 1965; Smith et al., 1968).
Con A was found to bind zoospores of Phytophthora cinnamomi and it was
concluded via the localation of its binding that these zoospores have a glycocalyx is
(Hardham, 1989), and further that binding of this lectin can induce encystment of the
zoospores (Hardham & Suzaki, 1986). This lectin has been shown to bind to the ceil
walls, electron-dense inclusions, and septal associated Woronin bodies of 0. ulmi, and
cytoplasm of V. albo-atrum (Benhamou, 1988).
PNA
Peanut lectin (PNA) is a tetrameric, carbohydrate-free protein. This lectin
possesses an extended binding site which is specific for D-galactopyranosyl end-
groups. In hemagglutination inhibition experiments this lectin was most effectively
inhibited by Gal6(1-3)NAcGal (Pereira et al., 1976; Goldstein & Poretz, 1986). This
disaccharide was found to be the most complementary to the binding site (Goldstein
& Poretz, 1986). PNA was also inhibited by Gal6(1-6)Glc, GalB(1-4)Glc, galactosamine,
and methyl a -galactoside (Pereira et al., 1976).
To localize a -D-galactose in 0. ulmi and V. albo-atrum Benhamou (1988) used
the castor bean (Ricinus communis) lectin, RCA-I. RCA-I is capable of binding
monomers and homodimers of D-galactose and lactose (Goldstein & Poretz, 1 986). Cell
95
walls of y. albo-atrum were specifically labeled while the walls of O. ulmi were unlabeled
(Benhamou, 1988).
DBA
This lectin is derived from Dolichos biflorus (horse gram). It is agglutinates blood
group A specifically (Goldstein & Poretz, 1986). It is a tetrameric glycoprotein (Carter
& Etzler, 1975) which has 2 binding sites per molecule (Etzler et al., 1981), and is
dependent on divalent metal ions for carbohydrate-binding activity (Kocourek et al.,
1 977; Borrebaeck et al., 1 981 ). In hemagglutination inhibition experiments NAcGala (1 -
3)NAcGala (1 -3)Gal6(1 -4)Gal6(1 -4)Glc and NAcGala (1 -3)NAcGal were found to be the
most effective inhibitors (Baker et al., 1 983). This lectin also binds the Q; anomers of N-
acetylgalactosamine and galactose (Goldstein & Poretz, 1986).
Benhamou and Ouellette (1986) used lectin from Helix pomatia (Roman snail),
HPA, to localize a-Nracetylgalactosamine on Ascocalyx abietina. HPA has been
described as binding to NAcGala (1-3)NAcGal with greater affinity than a-NAcGal, but
also as a valuable probe for detection of terminal, nonreducing a-NAcGal (Goldstein &
Poretz, 1986). The walls of this fungus were labeled, especially the external layer of old
cells. Benhamou (1988) using that same lectin found walls of V. albo-atrum to be
labeled but that O. ulmi walls were not labeled. P. cinnamomi cysts labeled with HPA,
but zoospores were unlabeled (Hardham, 1989).
!•■.,. . UEA-I
Ulex europaeus lectin, UEA-I, agglutinates O type blood (Goldstein & Poretz,
1986). It appears to be a carbohydrate-free metalloprotein dimer (Horejsi & Kocoure,
96
1 974). UEA-I binds a -L-fucose monomer and terminal non-reducing a -L-fucose (1 -3) or
(1-6) linked to N-acetylglucosamine (Goldstein & Poretz, 1986; Benhamou & Ouellette,
1986). Apparently this lectin can discriminate between trisaccharides that differ slightly
in the nature of the penultimate residue due to an extended binding site (Goldstein &
Poretz, 1986).
In Ascocalyx abietina. Benhamou and Ouellette (1986) found lipid bodies to be
strongly labeled while ail other organelles and cytoplasm labeled very slightly with UEA-I
lectin.
Materials and Methods
Fungal Cultures and Other Fungi
Details of fungal sources, culture and collection were given in Appendix A and
chapter 4.
Tissue Preparation for EM
Tissue preparation and sectioning were explained in chapter 4 and appendices
E & F. Both lightly broken spores (block SP) and apothecial tissue (blocks 5 & 1 5) were
used. Additionally, Pseudoplectania niqrella fixed with 2% glutaraldehyde and 2%
formaldehyde and embedded in LR White was tested for labeling with some of the
lectins.
Blood of types 0 and A were fixed in 2% glutaraldehyde and 2% formaldehyde,
for 30 minutes on ice, dehydrated through ethanol series to 95%, and embedded in LR
White resin.
1
1-'
Table 5.1 : List of lectins and labeling protocol information.
» •
Lectin
WGA
Method
indirect
Buffer
PBS
Glycopro.-Gold
ovomucoid-gold
GS-II
direct
PBS-CaCl2-2H20
-
LFA
indirect
tris-saline
fetuin-gold
Con A
indirect
tris-saline
horse radish peroxidase-gold
PNA
direct
PBS
-
DBA
direct
PBS
-
UEA
direct
PBS
-
. ..-I
97
Reagents and Cvtochemical Labeling
A lectin gold staining kit was obtained from EY Labortories (catalog #LGS-01)
which contained all the necessary components including lectins listed above,
glycoprotein-gold, buffers and sugar inhibitors. GS-II lectin (EY Lab., catalog #GP-2402)
was also obtained. PBS (0.1 5M) with calcium chloride (0.9mM) buffer and N-
acetyglucosamine (EY Lab., #LGS-01) sugar inhibitor were used with the GS-II lectin.
An instruction manual with protocols and recommended dilutions was included in the
kit. Those instructions were followed, although the dilutions of both lectin and
glycoprotein-gold (where used) had to be adjusted for optimal labeling. The protocols
provided by EY are similar to those given by Roth (1989).
In one of the GS-II experiments a pre-treatment with periodate was done to
demonstrate sensitive sugars. The methodology is identical to that used with antibodies
as described in appendix G.
'.:-;'*!
98
Results
PNA, DBA and UEA-I
PNA, DBA, and UEA-I did not label the fungi and bacteria tested. UEA-I and DBA
did not label sections of the appropriate agglutinating blood type.
WGA / GS-II
Both WGA and GS-II lectins labeled the ascus wall but not the ascospore walls
of Ascodesmis sphaerospora mgs fs 1 &5.2). This same labeling pattern was apparent
on Pseudoplectania niqrella for both lectins (figs 5.3 & 5.4). Both lectins labeled
vegetative cell walls to some extent. Interestingly, GS-II did not label P, niqrella
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Figure 5.1. WGA labeling on A. sphaerospora.
A) WGA test;
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paraphysis walls but did label vegetative cells in the excipular layers below the
hypothecium (fig. 5.4A & C). Paraphyses were not labeled with WGA either.
In both species the WGA labeling was increased with what should have been the
sugar negative control (fig. 5-5). EY Labs provided N-acetylglucosamine for this
purpose. No other negative control was immediately available and therefore none was
tried.
In addition to specific labeling of the ascus walls, GS-II lectin also specifically
labeled electron transparent areas within both the ascus and vegetative cells (fig. 5.2).
Tt
102
Figure 5.4. GS-II labeling on P. niqrella.
A) ascus, ascospore and paraphysis labeling;
B) cells of the excipular layer;
C) sugar negative control.
103
Figure 5.5. WGA labeling with sugar control.
A) sugar control on A. sphaerospora:
B) sugar control on A. sphaerospora:
C) WGA without sugar on A. sphaerospora:
D) sugar control on P. nigrella:
E) WGA without sugar on P. nigrella.
LFA
Labeling of LFA was evident over the cytoplasm of ascospores of A.
sphaerospora (figs. 5.6). This labeling did not appear to be specific. Specific labeling
occurred in restricted areas around spent asci and on the outside of older cells (figs.
5.7).
'^.
104
<::.
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i
Figure 5.6. LFA labeling on A. sphaerospora.
A) on an ascospore (1/80 lectin dilution);
B) on an ascospore (1/40 lectin dilution);
C) sugar negative control (1/80 lectin dilution) on an ascospore.
Con A
Con A labeling on A. sphaerospora was different than for the previous lectins.
In this case the lectin labeled the ascospore walls strongly (fig. 5.8 A-C) and an inner
layer of vegetative cell walls or the plasma membrane of these cells (fig. 5.8D & 5.1 OB)
and a similar area on the ascus walls (fig. 5.1 1 A), but nothing more on these later wall
systems (figs. 5.8A & D, and 5.11 A & B). Electron transparent areas within vegetative
cells and asci were labeled when higher concentrations of the lectin were used (fig.
5.8D).
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In contrast, Con A labeling on the ascospore walls of Pseudoplectania niqrella
occured only in the secondary wall region (fig. 5.9A). Additionally, the outer layer of the
ascus wall, or glycocalyx-like region, was specifically labeled (fig. 5.9A).
Ascospore secondary walls of Ascodesmis sphaerospora seem to label more
and more strongly as the spore matures (fig. 5.8C & 5.1 1 A vs 5.8A & B). Interestingly,
the ascospore secondary wall is strongly labeled on older spores which are still within
the ascus, but not on those which have been expelled (such as on in the sections of
lightly broken spores, see fig. 5.10).
The background seemed to be a slight problem using the dilutions of lectin and
glycoconjugate recommended by the manufacture. In fact much better results, in terms
of specific labeling without apparent background, were obtained using twice the
recommended dilution (figs. 5.8D at 1/40 & 5.1 1 B at 1/80). At these higher dilutions the
electron-transparent areas were not well labeled (fig. 5.1 IB). This change in labeling
pattern might be expected because a similar concentration dependent pattern was
found in precipitation studies (Manners & Wright, 1962; Goldstein, 1965).
In a digestion experiment with a-mannosidase (Sigma #M-1266) Con A was
shown to be labeling a-mannan in the ascospore walls of Ascodesmis sphaerospora.
Figure 5.1 OA demonstrates the Con A positive control that received no pretreatment
other than buffer washes. In comparing figure 5.1 OA with 5.1 OB, which demonstrates
pretreatment with a-mannosidase, there appears to be a slight decrease in labeling of
the primary wall, and both figures appear to have background labeling on the plastic.
To test for Con A recognition of any a-mannosidase that may have not washed off in
the rinses between these two treatments a pronase digestion to remove the a-
mannosidase was also preformed. Figure 5.1 OC demonstrates the pronase control (with
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only the pronase pretreatment) and figures 5.1 OE and F demonstrate the results of the
double digestion. The obvious, great reduction in labeling of the primary wall in the
double digestion (figs. 5.10E & F) in comparison with the Con A control (fig. 5.10A) and
the single digestions (figs. 5.1 OB & C), indicates that Con A labeling primarily due to the
presence of mannan in the wall.
Discussion
Wall Chemistry
The presence of chitin in the cell walls is one of the characteristics of
Ascomycetes (Barknicki-Garcia, 1968). WGA has been used successfully in the past to
localize chitin. In both Ascodesmis sphaerospora and Pseudoplectania niqrella ascus
but not ascospore walls were labeled (figs. 5.1 & 5.3). This pattern was also observed
with GS-II lectin in both these species (figs. 5.2 & 5.4). It is most probable that no part
of the spore walls of these species contains chitin.
If chitin is absent from these ascospores walls then they must have some other
structural polymer(s). Strong labeling of the primary wall with Con A (figs. 5.8A-C &
5.1 1 A) is suggestive of a -mannan filling the structural role. The mannosidase digestion
provides more definitive evidence for presence of Ck; -mannan within the wall (fig. 5.9).
Some labeling is observed on the wall after digestion (fig. 5.9B) but this was expected
as this enzyme is an exoenzyme incapable of degrading past branching points and Con
A preferentially binds at branch points (Manner & Wright, 1962; Goldstein, 1965). While
an argument could be made for Con A labeling a glucose polymer in these spore walls,
this now appears far less likely than mannan labeling. Additionally, if a glucose
117
if ir ' '
Table 5.2: Comparison of Con A and GS-II labeling on Ascodesmis
sphaerospora and binding specificities.
Spore Glycogen Gly-
Lectin Wall Ascus Areas Man, cogen N-AcGIc
Con A + ± + ++ + ±
GS-II - + + ' ± + + +
-, not labeled or bound
±, slightly labeled or bound
+, specifically labeled, binding standard
+ + , binding greater than standard
polymer was present one might expect to see GS-II labeling as well because of similar
glycogen binding of these lectins (table 5.2).
Glycogen Labeling
Both Con A and GS-II lectins have been reported to bind glycogen (Con A,
Sumner & Howell, 1936; GS-II, Ebisu et al., 1978). Both of these lectins label electron-
transparent areas within asci and vegetative cells (Con A, fig. 5.8D, & GS-II, figs. 5.2A
& C, 5.1 1 A & B) that was initially thought to be an artifact of poor fixation. To further
substantiate the chemistry of these electron-transparent areas, sections to be labeled
with GS-II were pretreated with periodate (fig. 5.1 1 E vs F). While cell walls continued
to label as would be expected for N-acetylglucosamine, the electron-transparent areas
no longer labeled as would be expected for glycogen. Thus it is concluded that these
electron-transparent areas are not artifacts but glycogen.
Methods Notes
It has been clearly shown that a lectin can label more than one carbohydrate
type within the fungal cell (GS-II lectin; N-acetylglucoseamine in the walls, and glycogen
118
within the cell; fig. 5.1 OD & E vs F). The sugar negative control for this lectin was not
labeled on either site. An unlabeled sugar negative control alone therefore can not be
used to determine the particular carbohydrate being labeled when a lectin has multiple
specificities. r^
It was pointed out in the results that the WGA labeling was increased by use of
the supposed sugar negative control which was N-acetylglucosamine. No other sugar
control was readily available. Nevertheless it would appear that label continued to be
specific as the overall pattern continued to be similar to that of GS-II and WGA without 1
1
sugar. The fact that labeling increased while continuing to be specific suggests that the
WGA was primed by the monomer. In inhibition studies (Goldstein et al., 1 975) N,N',N"-
triacetylchitotriose was found to be almost 30 times more effective than N,N'-
diacetylchitobiose. If the monomer does bind to subsite C as Allen and coworkers
(1973) suggested it would seem that the lectin would be able to identify and bind
available N,N'-diacetylchitobiose units in the A and B site up to 30 times more readily.
Bonfonte-Fasolo and coworkers (1 986) described an orientation of chitin fibrils within
fungal cell walls. That varying orientation would lend itself to exposure of differing
lengths of chitin fibril in section. If WGA is primed by the monomer an increase in
labeling on exposed chitobiose, probably proportional to the amount exposed, would
be expected.
CHAPTER 6
CONCLUSIONS
Evaluation of Experimental Methods
Tissue Preparation
In general fixatives with less than 1% glutaraldehyde were not required, even with
the antibodies. Fixative with 2% glutaraldehyde was most commonly used with the ^
lectins, and worked quite well. Pelleting the apothecial was quite helpful in reducing the •,;:
time required for block trimming and facing. Although scrapping the apothecia off the
agar disrupted the older hyphal cells and mature asci, it was far more convenient than
using unpelleted apothecia.
Immunological
Evaluation of immunological techniques were given at the end of Chapter 3. In
review, two main points were made. First, a hybridoma management plan was
presented which would allow testing and work to proceed at a pace limited by the EM
work rather than the needs of billions of hybridoma cells in log growth phase. Second,
the ELISA test was found to be inadequate for this ascospore wall antigen because of
background problems.
Cvtochemical
The first priority was to try to examine and compare the chemistry of the
119
■if ,'■
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s ^\
■J
' '120 J ■
ascospore walls with that of hyphal walls. Thus, the majority of experiments have been
with lightly fixed tissues for optimal labeling conditions, even though good ultrastructural
preservation was sacrificed. This was particularly necessary for working with unknown
determinants of antibodies. Now that labeling has been proven with various probes, i
further work on both morphology and biological definition of the binding sites will be - ■
possible.
The antibodies have proven themselves finicky as to pretreatment of sections to
obtain good labeling results and buffering systems required to reduce background ^
labeling. i
The lectins have proven themselves problematic in regard to background
labeling, not labeling even on positive controls, apparent lectin priming resulting in
increased labeling with the sugar negative control, and labeling more than one type of ;
sugar on the sections. To an extent, problems with background and lack of labeling
have been resolved by altering the dilutions of the primary probe and/or glycoprotein-
gold (indirect method). It appears that increasing the dilution of Con A lectin reduces v a
specific labeling on the electron transparent areas, which are presumably glycogen, i
without greatly reducing ascospore wall labeling. This is not an unexpected result as
similar results have been found in precipitation and inhibition studies.
Evaluation of labeling was qualitative rather than quantitative. This type of
evaluation has been sufficient for this study. From qualitative analysis there appears to
be some linearity to the labeling patterns of antibody 8F1 1 (fig. 4.4 & 4.6A). Quantitative
analysis using morphometric techniques must be done to make a well-founded
statement about such labeling patterns. Although detailed analysis of labeling patterns
was not a goal of this study, it can be done at some point in the future with micrographs
taken for this study.
.';i
^
121
Ascospore Wall Chemistry
Any hypothesis regarding a generalized chemical similarity has been disproven
with every specific probe used in this study (table 6.1). Immune-sera showed some
cross-reactivity but the possibility of immunogen contamination renders that data
unacceptable as proof of chemical similarity. From the information gained in this study
it appears that the various wall systems in fungi are chemically distinct. The correlation
of structure and function seems to once again be reinforced by experimental data.
Table 6.1 : Comparision of wall labeling patterns.
Trtm.
A. SDhaerospoi
Ascospore
V 2° Ascus
[a
Vea.
P. niqrella
Ascospore
r 2° Ascus
Veq.
Serum
-1-
-1- -1-
+
0
0 0
0
8F11
-1-
-
-
0
0 0
0
41.1-3
-1-
-
-
-
-
-
12-2
-
-1-
-
-
-
-
WGA
-
+
+
-
+
+
GS-II
-
+
+
-
-»-
+
Con A
-1-
-t-
±
-
+ +
-
LFA
-
±
±
0
0 0
0
-, no labeling
-I-, specific label
±, labeling variable
0, not tested
or questionable
"• { -
In the proposal it was hypothesized that skeletal elements of the ascospore wall
would be chitin or 6(1-3) glucan, or possibly mannan. WGA and GS-II lectin labeling
(figs. 5.1 , 5.3, 5.2 & 5.4) demonstrate chitin in the ascus wall and a lack of detectable
quantities of chitin in the ascospore wall of both Ascodesmis sphaerospora and
Pseudoplectanianiqrella. Con A labeling, especially in combination with :- -mannosidase
digestion (figs. 5.11), on A. sphaerospora suggests a mannan structural polymer, but
does not exclude 6-glucan playing such a role solely or in combination with mannan.
Conversely, P. nigreM ascospore walls were only labeled in the secondary wall layer, .d
and thus mannan would not appear to be the structural polymer in the primary wall of -i
this species. By elimination, it would appear in this case that a glucan or glucans are '
the skeletal elements in this case, but a strong statement to that effect can not be made
until there is positive evidence. Such an apparent chemical difference between these two
species is not unexpected considering the distant relationship of these fungi.
Precursor Tracking • 1*
In the proposal it was hypothesized that given the appropriate probe it should 1
be possible to track wall materials not synthesized in situ. Antibody 12-2 has shown i
promise of being such a probe, but poor morphology has interfered with proving it. '
Further work towards this goal would be worthwhile.
Maturation of the Ascospore Wall • i
Antibody 8F1 1 has definitively shown a maturation event in the primary wall. '
Labeling with this antibody does not occur until the ascospore is apparently complete
and relatively mature (fig. 4.4 & 4.6).
123
A further point can be made concerning the literature on ascosporogenesis. The
use of the word "mature" has been used in at least two different contexts in regard to
the primary wall. Gibson and Kimbrough (1988) describe the time of primary wall
maturity as when the outer delimiting membrane moves away from the primary wall.
Later in that same paper they discuss a change in silver proteinate staining over time
and stated that the staining change indicated a change in the primary wall chemistry.
It would appear that they have inadvertently contradicted themselves if "mature" is
considered a final product rather than termination of a obvious deposition process.
Labeling by antibody 8F11 strongly supports the hypothesis of continued chemical
changes in the primary wall after the termination of obvious deposition and apparent
morphological maturity. Thus, the primary wall has been shown not to be in final, or
fully mature form, until very late in the process of ascosporogenesis.
\H ■
*»"■ - APPENDIX A
J
FUNGAL CULTURE
Fungus
Ascodesmis sphaerospora stock culture (# 260) was obtained as a gift from Dr.
Kimbrough.
Corn Meal Malt Yeast Medium (CMMY)
8.5g corn meal agar (BBL, 11132)
4.0g bacto-agar (Difco, 0140-01)
0.5g yeast extract (BBL, 1 1 929)
0.5g malt extract (Difco, 01 86-02-4)
500 ml deionized H2O
Protocol
1. Ingredients are placed in a 1000ml flask.
2. Cover flask with aluminum foil and sterilize,
(autoclave for 20 minutes)
3. Pour sterile medium about 3mm-5mm deep in 100mm x 15 mm sterile plastic petri
dishes (Fisher, 8-757-13).
4. Let sit overnight.
5. Inoculate plates with about a 0.5cm-1cm square from a previous culture.
6. Seal plates with parafilm.
7. Grow at room temperature with a source of natural light.
124
APPENDIX B
ISOlvMION OF ASCOSPORE WALLS
For Immunization and EM
1 . Inoculate plates of CMMY agar with Ascodesmis sphaerospora.
2. Grow up for 2-3 weeks in natural light.
3. Collect spores off lids with rubber policeman and dHp and place in flask or other
suitable clean storage container.
4. Break spores in Braun homogenizer.
a. Fill jar about 1/3 full of beads.
Beads: Potters Co. #P-008 (about 0.17mm)
b. Add spore suspension to about 1/2 full.
c. Add 3 drops of tributyl phosphate (antifoam agent).
d. Stop the jar, and put a little glycerin on jar body.
e. Place in homogenizer, adjust CO^, and shake for 2-3 minutes.
f. Filter off homogenate and wash the beads with dHgO.
5. Centrifuge at 10,000 rpm (15,890 g) for 10 minutes over a 10% over 25% over 50%
sucrose step gradient. Sucrose is dissolved in dKO. (Centrifuge: Beckman model
J2-21)
6. Remove and discard supernatant.
7. Resuspend pellet in dHjO.
8. Centrifuge at 5,000 rpm (3,970 g), for 5 minutes.
9. Repeat steps 6, 7 & 8 at least once more, the last time in 0.1 M cacodylate buffer.
10. Suspend the pellet in 0.5% glutaraldehyde and 1% paraformaldehyde in 0.1 M
cacodylate buffer. Fix for 30 minutes on ice.
125
126
11. Centrifuge at 2,000 rpm, for 5 minutes. (Centrifuge: Damon Clinical model CL)
12. Remove and discard supernatant. For preparation of immunogen proceed to step
13 of this protocol. For embedding proceed to Appendix C, step 13.
13. Resuspend pellet in 0.1 M cacodylate buffer, and let stand for 15 minutes.
14a. For immunogen, repeat steps 11, 12, & 13 at least twice more using sterile PBS
buffer (0.1 5M, pH 7.0-7.4).
14b. For ELISA and EM, repeat steps 11 through 13 once or twice more using dHjO.
15a. Suspend in PBS buffer.
15b. For ELISA, suspend in dHjO.
15c. For EM, precede to Appendix D step 15.
1 6. Store at 4° C.
1 7. Estimate concentration of wall fragment suspension by the hemacytometer counting
method. (America Optical, bright line hemacytometer)
if
APPENDIX C
PROTOCOLS FOR HYBRIDOMA CONSTRUCTION
AND CLONING
Mouse immunization
1. Young, healthy, female, Balb-c mice were selected from the stock collection.
2. Inoculations were made every 2 weeks for a minimum of 2 months.
a. The first 2 immunizations were sub-cutaneous in the neck region with 0.1 ml
of 50% Freund's (SIGMA, St. Louis, MO.) complete adjuvant and 50%
resuspended broken spore-wall stock suspension (4.5-7 x 10^ parts per ml).
b. All further immunizations were intra-peritoneal with 1 ml of 50% Freund's
incomplete adjuvant and 50% resuspended broken spore-wall stock.
3. A final pre-fusion boost (as in 2B above) 2-3 days prior to sacrifice.
Cell fusion
1 . Grow SP2-0 myeloma cells to log phase in a 75ml culture flask. This usually takes
about 2-3 days in 75ml culture flask.
2. Sacrifice mouse, and save blood for serum.
3. Dissect out spleen.
4. Break up spleen cells by one of the following methods:
Method 1 :
a. Mash spleen through a sterile screen into DME media.
b. Pull cells into syringe through an 18 gauge needle and expel through a 21
gauge needle.
c. Pull cells into syringe through an 1 8 gauge needle and expel through a 23
gauge needle.
127
128
d. Pull cells into syringe through an 1 8 gauge needle and expel through a 25
or 26 gauge needle into centrifuge tube.
Method 2:
a. Set spleen on sterile screen above a sterile beaker.
b. Clip a small piece of tissue off the larger end with sterile scissors.
c. Place 18 gauge needle on a syringe and fill it with DME media.
d. While holding the spleen with forceps, insert the needle in the uncut end and
force the media through the spleen.
e. Repeat steps c and d until the spleen in more or less empty of cells. , i
f. Transfer cells from beaker to a centrifuge tube.
5. Transfer SP2-0 cells to centrifuge tubes.
6. Balance all centrifuge tubes (those with spleen and SP2-0 cells) and centrifuge them
at about lO.OOOrpm for 10 minutes. Discard spleen cell supernatant and save the
SP2-0 cell supernatant (= conditioned media).
7. Resuspend spleen and SP2-0 each in 5 ml of DME media.
8. Count live cells of each suspension, determine concentration of live cells.
9. In a sterile centrifuge tube mix cells at a rate of 1 0/1 live spleen/live SP2-0 cells.
10. Add 1 ml PEG dropwise over 1 minute period.
1 1 . One minute later add 1 ml DME slowly while gently swirling.
12. One minute later add 2ml DME slowly while gently swirling.
13. Two minutes later add 4 ml DME slowly while gently swirling.
14. Four minutes later add 10 ml DME slowly while gently swirling.
15. Centrifuge at 10,000 rpm for 10 minutes. Discard supernatant.
16. Resuspend cells in DME with 10%-25% horse serum (or non-serum substitute such
as CPSR).
17. Count cells and dilute further with that medium as needed to obtain 100-200 cells
per well (see step 18).
'\
129
18. Place 50 or 100 111 cell suspension per well into 96 well culture plates.
19. Place in a controlled environment (37°C, and 5% humidity) incubator.
Cell selection
1 . Twelve to eighteen hours after plating the fusion cells feed them an equal amount (50
or loop, I) of 2X HAT media.
2. Feed approximately every 5 days thereafter. The first 2 of these feedings with 1 X HAT
in 55% DME / 20% horse serum (or CSPR) / 25% SP2-0 conditioned media, and
thereafter with same media minus the HAT.
Cloning
Method 1 :
1 . Grow cells up into large volume.
2. Count cells and dilute to approximately 1x10^ cells per ml media.
3. Prepare 2 - 96 well plates by placing 100|,i.l media in each well.
4. Place 100|J,I cell suspension in each well of the first column on the first plate.
5. Place 100|J,I from wells of the first column into the wells of the second column, etc.,
until serial dilutions have been made across all the wells of both plates.
6. Monitor for growth of single colonies per well.
Method 2:
1 . Prepare plates as in step 3 above.
2. Transfer 100^,1 of cell suspension from a target well (of 96 or 24 well plate) to a well
in the first column of the first cloning plate.
3. Proceed to step 5 above.
APPENDIX D
FREEZING AND THAWING HYBRIDOMA OR SP2-0 CELLS
Freezing Media
1 0% v/v horse serum or CSPR
1 0% DMSO
balance feed medium (eg. DME with 5%-20% horse serum or CSPR and 0%-25%
conditioned medium)
Freezing Protocol
1 . Prepare cryovials.
2. Spin down cells at low speed (Dynac centrifuge, speed setting 5, for 8-10 minutes).
3. Remove medium and save if needed.
4. Add 1ml-2ml of freeze media for each 5ml-15ml of media removed. (This is not
terribly critical.)
5. Place immediately in a freezer (-80°C) or liquid nitrogen. (If immediately placed in a
-20°C freezer transfer to -80°C or liquid nitrogen as soon as possible and not more
than 30-60 minutes.)
Thawing Protocol
1. Thaw cells as quickly as possible (at about 37°C).
2. Add to a centrifuge tube with 10ml-15ml of DME.
3. Centrifuge as in step 2 above.
4. Remove DME from cell pellet and discard.
5a. Repeat steps 2-4 of desired.
130
131
5b. Suspend pellet in 10ml-15ml of feed medium.
6. Grow up in a small cell culture flask (e.g., 25 ml) under typical conditions (Appendix
C).
APPENDIX E
LIGHT BREAK OF ASCOSPORES
Fixative
0.5% glutaraldehyde
2% paraformaldehyde
in cacodylate buffer (approx. 0.1 M)
Protocol
1. Collect spores, suspend in dHjO and place in a microfuge tube.
2. Centrifuge at 1 ,000 g for 3 minutes. (Fisher Centrifuge, model 59)
3. Remove and discard supernatant.
4. Resuspend pellet in fix, and place on ice for 15 minutes.
5. Centrifuge at 1 ,000 g for 3 minutes.
6. Remove and discard excess fix.
7. Add glass beads to approximately 1 :5, v:v, spores:beads, then gently mix.
8. Add additional fix until all beads are just moist, and vortex for 100-120 seconds
(setting 9).
9. Add additional fix so that liquid level is well above the beads and gently suspend the
spores.
10. Once the beads have settled, use Pasteur pipet to remove the spore suspension,
being careful not to pick up any beads.
1 1 . Place spore suspension on ice for 1 5 minutes. (Total fix time is about 45 minutes.)
12. Centrifuge at 1,000 g for 1 minute.
13. Resuspend in 0.1 M cacodylate buffer, and let stand at room temperature for 15
minutes.
14. Repeat steps 12 and 13 twice more, the last time resuspending in dHp.
132
<' , 133
1 5. Dehydrate in ethanol series to 95% EtOH.
16. Infiltrate witli, and embed in LR White resin.
Ref.: Olson, L.W. 1978. Preparation of "difficult" spores, and sporangia for electron
microscopy. In: M.S. Fuller-(ed.), Lower Fungi In The Laboratory, Dept. of Botany, Univ.
of Georgia, publisher.
^li;
APPENDIX F
FIXATION PROTOCOL
Fixative
The various combination of glutaraldehyde/formaldehyde concentrations are given in
table 4.1 . In all cases the final concentration of cacodylate buffer used was 0.1 M.
Protocol
1 . Cut out pieces of agar with greatest density of apothecia, trimming off excess depth,
so that final dimensions are 0.5cm x 1cm-2cm x 0.25cm-0.5cm.
2a. Place pieces of agar in fixative solution.
2b. Alternatively, skip step 1 and flood plates with fixative.
3. Fix for 30-60 minutes on ice.
4. Wash for 30 minutes using 3 changes of 0.1 M cacodylate buffer.
5. Wash for 10 minutes in dHgO (optional).
6a. Dehydrate through ethanol series to 95% for LR White Resin.
6b. Dehydrate through ethanol series through 100% and acetone for Spurr's resin.
7. Infiltrate with, and embed in the appropriate resin increasing the resin concentration
in 25% v/v increments.
8. Polymerize resin at 50-60°C for 12-24 hours.
134
APPENDIX G
ANTIBODY LABELLING
Protocol
0. Pretreatment (see below).
1 . Block, 5% milk, with or without 1 % sodium azide, in PBS. (Carnation Co.)
2. Wash, briefly in PBS, and/or blot.
3. Primary antibody, 1 hour at room temperature, diluted in PBS or Tris/high salt/tween
20 (THST).
4. Washes, about 5 times for total of 50 minutes in buffer (PBS, or first wash in THST,
then PBS. Blot.
5. Gold-conjugated secondary antibody, 1 hour at room temperature. (EY Labs, goat-
anti-mouse IgG/IgM, 15 nm gold, diluted to 1.25 |ig/ml.)
6. Washes, about 5 times for total of 50 minutes in PBS. Blot.
7. Washes, 2 or 3 times for total of 30 minutes in dHjO. Blot, and view, post-stain, or
store.
Pre-treatment experiments
1. Sodium meta-periodate etching:
a. Float grids on saturated NalO, in dH^O, at room temperature for 1 hour.
(Fisher Scientific Co., lot 716131.)
b. Washes, 3 times for total of 30 minutes in dHgO. Blot.
c. Begin above labelling protocol, or proceed with Pronase digest.
2. Pronase digest:
a. Float grids on 1% w/v solution of Pronase in PBS buffer, for 1 hour at 37°C.
(Calbiochem, B grade, 45,000 PUK units/gm.)
135
136
b. Washes, 3 times for total of 30 minutes in PBS. Blot.
c. Begin labelling protocol.
M
t..^
Vl'
I
Literature Cited
Aldrich, H.C., and W.J. Todd. 1986. Ultrastructure Techniques for
Microorganisms. Plenum Press. New York.
Allen, A.K., A. Neuberger and N. Sharon. 1973. The purification, composition
and pecificity of wheat-germ agglutinin. Biochem. J. 131:155-162.
Araki, Y. and E. Ito. 1975. A pathway of chitosan formation in Mucor rouxii.
enzymatic deacetylation of chitin. Eur. J. Biochem. 55:71-78.
Aronson, J.M. 1965. The cell wall, jn: G.C. Ainsworth and A.S. Sussman (eds.).
The Fungi: An Advanced Treatise, volume I. Academic Press, New York,
pp. 49-76.
Aronson, J.M. 1981. Cell wall chemistry, ultrastructure, and metabolism, in:
G.T. Cole and B. Kendrick (eds.). Biology of Conidial Fungi. Academic
Press, New York 2:459-507.
Baker, C.A., S. Sugii, E.A. Kabat, R.M. Ratcliffe, P. Hermentin, and R.U. Lemieux.
1983. Immunochemical studies on the combining sites of Forssman
reactive hemagglutanins from Dolichos biflorus. Helix pomatia and
Wistaria floribanda. Biochemistry 22:2741-2750.
Banowetz, G.M., E.J. Tirone, and B.B. Krygier. 1984. Immunological
comparisons of teliospores of two wheat bunt fungi, Tilletia species,
using monoclonal antibodies and antisera. Mycologia, 76:51-62.
Bartnicki-Garcia, S. 1968. Cell wall chemistry, morphogenesis, and taxonomy
of fungi. Annu. Rev. Microbiol. 22:87-108.
Bastin, J.M., J. Kirkley, and A.J. McMicheal. 1982. Production of monoclonal
antibodies. In: A.J. McMichael and J.W. Fabre (eds.). Monoclonal
antibodies in clinical medicine. Academic Press Inc., New York, pp. SOS-
SI 7.
Beckett, A. 1981. Ascospore formation, jn: G. Turina & H.R. Hoh (eds.). The
Fungat Spore: Morphogenetic Controls. Academic Press, New York pp.
107-129.
137
138
Beesley, J.E. 1989. Immunocytochemistry of microbiological organisms: A
survey of techniques and applications. Tech. Immunocytochem. 4:67-
93.
Bellemere, A. and L.-M. Melendez-Howell. 1976. Etude ultrastructure comparee
des I'ornamentation. externe de la parol des ascospores de deux
Pezizales: Peziza fortinii n. sp., recoltee qu Mexique, et Aleuria aurantia
(Oed. ex Fr.) Fuckel. Revue Mycol. 40:3-19.
Bendayan, M. 1981. Ultrastructural localization of nucleic acids by means of
nuclease-gold complexes. J. Histochem. Cytochem. 29:531-541.
1982. Double immunocytochemical labeling applying the protein
A-gold technique. J. Histochem. Cytochem. 30:81-85
1984a. Protein A-gold electron microscopic
immunocytochemistry: Methods, applications and limitations. J. Electron
Microsc. Tech. 1:243-258.
• 1984b. Enzyme-gold electron microscopic cytochemistry: A new
affinity approach for the ultrastructural localization of macromolecules.
J. Electron Microsc. Tech. 1:349-372.
• 1989a. Protein A-gold and protein G-gold postembedding
immunoelectron microscopy, jn: H.A. Hayat (ed.) Colloidal gold:
Principles, methods, and applications, volume 1. Academic Press, New
York, pp. 33-94.
1989b. The enzyme-gold cytochemical approach: A review. In:
(M.A.Hayat, ed.) Colloidal gold: Principles, methods, and applications,
Vol. 2. Academic Press, New York, pp. 117-147.
Bendayan, M. and M. Zollinger. 1983. Ultrastructural localization of antigenic
sites on osmium-fixed tissues applying the protein A-gold technique. J.
Histochem. Cytochem. 31(1):101-109.
Benhamou, N. 1988. Ultrastructural localization of carbohydrates in the cell
walls of two pathogenic fungi: A comparative study. Mycologia
80(3): 324-337,
• 1989a. Ultrastructural study of galacturonic acid distribution in
some pathogenic fungi using gold-complexed Aplvsia depilans gonad
lectin. Can. J. Microbiol. 35:349-358.
• 1989b. Preparation and application of lectin-gold complexes. In:
M.A.Hayat (ed.). Colloidal gold: Principles, methods, and applications,
Vol. 1. Academic Press, New York, pp. 95-143.
Benhamou, N., and G.B. Ouellette. 1986. Ultrastructural localization of
glycoconjugates in the fungus Ascocalyx abietina, the scleroderris canker
agent of conifers, using lectin-gold complexes. J. Histochem. Cytochem.
43{7): 855-867.
Blackwell, J. 1982. The macromolecular organization of cellulose and chitin.
in: R.M. Brown, Jr. (ed.). Cellulose and Other Natural Polymer Systems:
Biogenesis, Structure, and Degradation. Plenum Press, New York, pp.
403-428.
Bonfante-Fasolo, P., A. Faccio, S. Perotto, and A. Schubert. 1990. Correlation
between chitin distribution and cell wall morphology in the mycorrhizal
fungus Glomus versiforme. Mycol. Res. 94(2): 157-1 65.
Bonfante-Fasolo, P., B. Vian, and B. Testa. 1986. Ultrastructural localization of
chitin in the cell wall of a fungal spore. Biol. Cell 57:265-270.
Borrebaeck, A.C.K., B. Lonnerdal, and M.E. Etzler. 1981. Metal ion content of
Dolichos biflorus lectin and effect of divalent cations on lectin activity.
Biochemistry 20:41 1 9-41 22.
Borrebaeck, A.C.K., and S.A. Moller. 1986. In vitro immunization. Effect of
growth and differentiation factors on antigen-specific B cell activation and
production of monoclonal antibodies to autologous antigens and weak
immunogens. J.Immunol. 1 36(1 0):371 0-371 5.
Boyde, A., E. Bailey, S.J. Jones, and A. Tamarin. 1977. Dimensional changes
during specimen preparation for SEM. SEM/IITRI 1:507-508.
Brams, P., D.E. Pettijohn, M. Brown, and L. Olsson. 1987. In vitro B-lymphocyte
antigen priming against both non-immunogenic and immumogenic
molecules requiring low amounts of antigen and applicable in hybridoma
technology. J. Immunol. Methods 98:11-22.
Brawner, D.L, and J.E. Cutler. 1986a. Ultrastructural and biochemical studies
of two dynamically expressed cell surface determinants on Candida
albicans. Infect. Immun. 51(1):327-336.
. . 1986b. Variability in expression of cell
surface antigens of Candida albicans during morphogenesis. Infect.
Immun. 51(1):337-343.
Briza, P., G. Winkler, H. Kalchhauser, and M. Breitenbach. 1986. Dityrosine is
a prominent component of yeast ascospore wall. J. Biol. Chem.
261 (9): 4288-4294.
Briza. P., G. Winkler, and M. Breitenbach. 1988. Chemical composition of the
yeast ascospore wall. J. Biol. Chem. 263(23): 11 569-1 1574.
139
140
Burnett, J.H. 1979. Aspects of the structure and growth of hyphal walls. In:
J.H. Burnett, and A.P.J. TrincI (eds.). Fungal Walls and Hyphal Growth.
Cambridge University Press, Cambridge, pp. 1-26.
Cabib, E., B. Bowers, and R.L. Roberts. 1983. Vectorial synthesis of a
polysaccharide by isolated plasma membranes. Proc. Natl. Acac. Sci.
USA 80:3318-3321.
Cabib, E., S.J. Silverman, A. Sburlati, and M.L Slater. 1990. Chitin synthesis In
Yeast (Saccharomvces cervisiae) |n: P.J. Kuhn, A.P.J. Trinci, M.J. Jung,
M.W. Goosey, and LG. Copping (eds.). Biochemistry of cell walls and
membranes in fungi. Springer-Verlag, New York, pp. 31-42.
Carlemalm, E., R.M. Garavito, and W. Villiger. 1982. Resin development for
electron microscopy and an analysis of embedding at low temperature.
J. Microsc. 126:123-143.
Carroll, G.C. 1967. The Ultrastructure of ascospore delimitation in Saccobolus
karverni. Jour. Cell Bio. 33:218-224.
Carter, W.G., and M.E. Etzler. 1975a. Isolation and characterization of subunits
from the predominant form of Dolichos biflorus lectin. Biochemistry
14:2685-2689.
. 1975b. Isolation, characterization, and subunit
structures of multiple forms of Dolichos biflorus lectin. J. Biol. Chem.
250(7): 2756-2762.
Causton, B.E. 1984. The choice of resins for electron Immunocytochemistry.
in: J.M, Polak and I.M. Varndell (eds.) Immunolabelling for electron
microscopy. Elsevier Science, New York, pp. 29-36.
Cole, G.T., and H.C. Aldrich. 1971. Ultrastructure of conidogenesis in
scopulariopsis brevicaulis. Can. J. Bot. 49:745-755.
Coons, A.H., H.J. Creech, and R.N. Jones. 1941. Immunological properties of
an antibody containing a fluorescent group. Proc. Soc. Exp. Biol.
47:200-202.
Craig, S. and D.J. Goodchild. 1982. Post-embedding immunolabelling. Some
effects of tissue preparation on the antigenicity of plant proteins. Eur. J.
Cell Biol., 28:251-256.
Dae, M.W., M.A. Heymann, and A.L Jones. 1982. A new technique for
perfusion fixation and contrast enhancement of foetal lamb myocardium
for electron microscopy. J. Microsc. 127:301-305.
141
Daniel, G., T. Nilsson, and B. Pettersson. 1989. Intra- and extracellular
localization of lignin peroxidase during the degradation of solid wood and
wood fragments by Phianerochaete chrysosporium by using transmission
electron microscopy and immuno-gold labeling. Appl. Envir. Microbiol.
55(4):871-881.
Davis, LL. and S. Bartnicki-Garcia. 1984. Chitosan synthesis by the tandem
action of chitin synthetase and chitin deacetylase from Mucor rouxii.
Biochem. 23:1065-1073.
Debray, H., D. Decout, G. Strecker, G. Spik, and J. Montreuil. 1981. Specificity
of twelve lectins towars oligosaccharides and glycoproteins related to N-
glycosylproteins. Eur. J. Biochem. 117:41-55.
DeWaele, M., J. DeMey, M. Moeremans, M. DeBrabander, and B. VanCamp.
1983. Immunogold staining method for the detection of cell surface
antigens with monoclonal antibodies. Immunochemistry 2:1-23.
Dox, A.W., and R.E. Neidig. 1914. The soluble polysaccharides of lower fungi.
I. Mycodextran, a new polysaccharide in Penicillium expansum. J. Biol.
Chem. 18:167-175.
Duran A., B. Bowers, and E. Cabib. 1975. Chitin synthetase zymogen is
attached to the yeast plasma membranes. Proc Natl. Acad. Sci., USA
72:3952-3955.
Dute, R.R., J.D. Weete, and A.E. Rushing. 1989. Ultrastructure of dormant and
germinating conidia of Aspergillus ochraceus. Mycologia 81(5}: 772-782.
Dyby, S.D. and J.W. Kimbrough. 1987. A comparative ultrastructural study of
ascospore ontogeny in selected species of Peziza (Pezizales;
Ascomycetes). Bot. Gaz. 148(3):283-296.
Ebisu, S., P.N.S. Iyer, and I.J. Goldstein. 1978. Equalibrium dialysis and
carbohydrate binding studies on the 2-acetamido-2-deoxy-D-
glycopyranosyl-binding lectin from Bandeiraea simplicifolia seeds. Carb.
Res. 61:129-138.
Edwards, P.A.W. 1981. Some properties and applications of monoclonal
antibodies. Biochem. J. 200:1-10.
Eldred, W.D., C. Zucker, H.J. Karten, and S. Yazulla. 1983. Comparison of
fixation and penetration enhancement techniques for use in ultrastructural
Immunocytochemistry. J. Histochem. Cytochem., 31:285-292.
Erdos, G.W. and D. Whitaker. 1983. Failure to detect immunocytochemically
reactive endogenous lectin on the cell surface of Dictyostelium
discoideum. J. Cell Biol. 97:993-1000.
!«k
Etzler, M.E., S. Gupta, and C.A.K. Borrebaeck. 1981. Carbohydrate binding
properties of the Dolichos biflorus lectin and its subunits. J. Biol. Chem.
256(5): 2367-2370.
Faulk, W.P. and G.M. Taylor. 1971. An immunocolloid method for the electron
microscope. Immunochemistry 8:1081-1083.
Farkas, V. 1979. Biosynthesis of cell walls of fungi. Microbio. Rev. 43(2): 117-
1 44. , -
Fevre, M. 1984. Action of nucleotides on membrane bound and solubilized 8-
glucan syntheases from Saproleania monoica. In: C. Nombela (ed.).
Microbial cell wall synthesis and autolysis. Elsevier Scientific,
Amsterdam, p. 131.
Fevre, M., V. Girard, and P. Nodet. 1990. Cellulose and 6-glucan synthesis in
, ,',': ■; , Saproleania. in: P.J. Kuhn, A.P.J. Trinci, M.J. Jung, M.W. Goosey, and
L.G. Copping (eds.). Biochemistry of cell walls and membranes in fungi.
Springer-Verlag, New York, pp. 97-108.
^v: Gander, J.E. 1 974. Fungal cell wall glycoproteins and peptido-polysaccharides.
~, Annu. Rev. Microbiol. 28:103-119.
4 - Geoghegan, K.F., J.L Dallas, and R.E. Feeney. 1980. Periodate inactivation of
ovotransferrin and human serum transferrin. J. Biol. Chem. 255:11429-
11434.
Gibson, J.L. and J.W. Kimbrough. 1988a. Ultrastructural observations on
Helvellaceae (Pezizales). Ascosporogenesis of selected species of
Helvella. Can. J. Bot. 66:771-783.
■ , ■ . • 1988b. Ultrastructural observations on
r , Helvellaceae (Pezizales). II. Ascosporogenesis of Gvromitra exculenta.
Can. J. Bot. 66:1 743-1 749.
Glaser, L. and D.H. Brown. 1957. The synthesis of chitin in cell free extracts of
Neurospora crassa. J. Biol. Chem. 228:729-742.
Coding. 1986. Monoclonal antibodies: Principles and practice (2nd edition).
Academic Press, N.Y.
Goldstein, I.J., S. Hammarstrom, and G. Sundblad. 1975. Precipitation and
carbohydrate binding specificity studies on wheat germ agglutinin.
' ■ ■' . Biochim. Biophys. Acta 405:53-61.
Goldstein, I.J., C.E. Hollerman, and J.M. Merrick. 1965. Protein-carbohydrate
interaction I. The interaction of polysaccharides with concanavalin A.
Biochim. Biophys. Acta 97:68-76.
143
Goldstein, I.J., R.C. Hughes, M. Monsigny, T. Osawa, and N. Sharon. 1980.
What should be called a lectin? Nature (London), 285:66.
Goldstein I.J., and R.D. Poretz. 1986. Isolation, Physicochemical
characterization, and carbohydrate-binding specificity of lectins. Jn: (I.E.
Liener, N. Sharon, .and I.J. Goldstein, eds.) The lectins: Properties,
functions, and applications in biology and medicine. Academic Press,
Inc., New York, pp. 33-247.
Gorin, P.A. 1985. Structural diversity of D-galacto-D-mannan components
isolated from lichens having ascomycetous mycosymbionts.
Carbohydrate Research 142:253-267.
Gorin, P.A.J, and M. lacomini. 1984. Polysaccharides of the lichens Cetraria
islandica and Ramarlina usnea. Carbohydrate Res. 128:1 1 9-1 32.
Gorin, P.A.J. , and J.F.T. Spencer. 1968. Structural chemistry of fungal
polysaccharides. Adv. Carbohydr. Chem. 23:367-417.
Green, J.H., W.K. Harrell, J.E. Johnson, and R. Benson. 1980. Isolation of an
antigen from Blastomvces detmatitidis that is specific for the diagnosis
of blastomycosis. Curr. Microbiol. 4:293-296.
Guber, F. and A.R. Hardham. 1988. Secretion of adhesive material during
encystment of Phytophthora cinnamomi zoospores, characterized by
immunogold labelling with monoclonal antibodies to components of
peripheral vesicles. J. Cell Sci. 90:225-235.
Hardham, A.R. 1985. Studies on the cell surface of zoospores and cysts of the
fungus Phytophthora cinnamomi: The influence of fixation on patterns of
lectin binding. J. Histochem. Cytochem., 33(2}: 11 0-1 18.
1989. Lectin and antibody labelling of surface components of
spores of Phytophthora cinnamomi. Aust. J. Plant Physiol. 16:19-32.
Hardham, A.R. and E. Suzaki. 1986. Encystment of zoospores of the fungus,
Phytophthora cinnamomi, is induced by specific lectin and monoclonal
antibody binding to the cell surface. Protoplasma 133:165-173.
Hardham, A.R., E. Suzaki, and J.L. Perkin. 1985. The detection of monoclonal
antibodies specific for surface components on zoospores and cysts of
Phytophthora cinnamomi. Exp. Myclo. 9:264-268.
. 1986. Monoclonal antibodies
to isolate-, species-, and genus-specific components on the surface of
zoospores and cysts of the fungus Phytophthora cinnamomi. Can. J.
Sot. 64:311-321.
••1
144
Harper, R.A. 1897. Kerntheilung und zellbildung im ascus. Jahrbuch fur
wissenschaftliche Botanik 30:249-284.
Hashimoto, T., CD. Wu-Yuan, and H.J. Blumenthal. 1976. Isolation and
characterization of the rodlet layer of Trichophyton mentaqrophytes
microconidial wall. J. Bacteriol. 127:1543-1549.
Hayat, M.A. 1 981 . Fixation for electron microscopy. Academic Press, New
York.
1986. Basic techniques for transmission electron microscopy.
Academic Press, New York, pp. 41 1
Hearn, V.M., B.L Griffins, and P.A. Gorin. 1989. Structural analysis of water-
soluble fractions obtained from Aspergillus fumiqatus mycelium.
Glycoconjugate J. 6:85-100.
Hixson, D.C., J.M. Yep, J.R. Glenney, T. Hayes, and E.F. Walborg. 1981.
Evaluation of periodate/lysine/paraformaldehyde fixation as a method for
cross-linking plasma membrane glycoproteins. J. Histochem. Cytochem.
29:561-566.
Hohl, H.R., & W. Streit. 1975. Ultrastructure of ascus, ascospore, and ascocarp
in Neurospora lineolata. Mycologia 67:367-381 .
Hopwood, v., D. Poulain, B. Fortier, G. Evans, and A. Vernes. 1986. A
monoclonal antibody to a cell wall component of Candida albicans.
Infect. Immun. 54(1):222-227.
Horejsi, V. & J. Kocourek. 1974. Studies on phytohemagglutinins. XVII. Some
properties of the anti-H specific phytohemagglutinin of the Furze seeds •. .i
(Ulex europaeus L). Biochem. Biophys. Acta 336:329-337. i'^
Horikoshi, K., and S. lida. 1964. Studies of the spore coats of fungi I. Isolation • " ■']
and composition of the spore coats of Aspergillus orvzae. Biochim. 1
Biophys. Acta 83:197-203.
Hospenthal D.R., A.L Rogers, and G.L Mills. 1988. Development of
amphotericin B lipsomes bearing antibody specific to Candida albicans.
Mycopathologia 101:37-45.
Hrmova, M., C.S. Taft, and C.P. Selitrennikoff. 1989. 1 ,3-6-D-Glucan synthase
of Neurospora crassa: Partial purification and characterization of
solubilized enzyme activity. Exp. Mycol. 13:129-139.
Hunsley, D., and J.H. Burnett. 1 970. The ultrastructural architecture of the walls
of some hyphal fungi. J. Gen. Microbiol. 62:203-218.
145
j\>, Johnston, I.R. 1965. The composition of the cell wall of Aspergillus niqer.
Biochem. J. 96:651-658.
Kang, M.S., J. Au-Young, and E. Cabib. 1985. Modification of yeast plasma
membrane density by Concanavalin A attachment. Application to study
of chitin synthetase, distribution. J. Biol. Chem. 260:12680-12664.
Kellenberger, E., M Durrenberger, W. Viliiger, E. Carlemalm, and M. Wurtz. 1987.
The efficiency of immunolabel on Lowicryl sections compared to
theoretical predictions. J. Histochem. Cytochem. 35(9): 959-969.
Khardori, N. 1989. Host-parasite interaction in fungal infections. Eur. J. Clin.
Microbiol. Infect. Dis. 8(4):331-351.
Kimbrough, J.W., and J. L Gibson. 1990. Geopyxis carbonaria. Ultrastructural
and cytological observations of apothecial tissues of Geopyxis carbonaria
(Pezizales, Ascomycetes). Can. J. Bot. 68:243-257.
Kimbrough, J.W., C.G. Wu, and J.L Gibson. 1990. Ultrastructural observation
on Helvellaceae (Pezizales, Ascomycetes). IV. Ascospore ontogeny in
selected species of Gvromitra subgenus Piscina. Can. J. Bot. 68:317-
328.
Kocourek. J., G.A. Jamieson, T. Votruba, and V. Horejsi. 1977. Studies on
phytohemagglutinins. I. Some properties of the lectins of horse gram
seeds (Dolichos biflorus L.). Biochem. Biophys. Acta 500:344-360.
Kohler, G. and C. Milstein. 1975. Continuous cultures of fused cells secreting
antibody of predefined specificity. Nature 256:495-497.
Kuhn, P.J. and A.P.J. Trinci. 1990. Cell walls and membranes in fungi - an
introduction, jn: Kuhn, P.J., A.P.J. Trinci, M.J. Jung, M.W. Goosey, and
LG. Copping (eds). 1990. Biochemistry of cell walls and membranes in
fungi. Springer-Verlag, New York, pp. 1-4.
Kuhn, P.J., A.P.J. Trinci, M.J. Jung, M.W. Goosey, and LG. Copping (eds).
1990. Biochemistry of cell walls and membranes in fungi. Springer-
Verlag, New York.
Leduc, E.H. and W. Bernhard. 1962. Water-soluble embedding media for
ultrastructural cytochemistry, jn: R.J.C. Harris (ed). The interpretation of
ultrastructure. Academic Press, New York, pp. 21-45.
LeGal, M. 1947. Recherches sur les ornamentations sporales des
Discomycetes opercules. Ann. Sci. Nat. 1 1 ser. Bot. 7:73-297.
• 1951. Les Discomycetes de Madagascar. Paris.
146
Lenard, J. and S.J. Singer. 1968. Alterations of the conformation of proteins in
red blood cell membranes and in solution by fixatives used in electron
microscopy. J. Cell Biol. 37:117-121.
Leung, H. and P.H. Williams. 1987. Nuclear division and chromosome behavior
during meiosis and ascosporogenesis in Pyricularia oryzae. Can. J. Bot.
65:112-123.
Mahadevan, P.P., and E.L. latum. 1965. Relationship of the major constituents
of the Neurospora crassa cell wall to wild-type and colonial morphology.
J. Bact. 90(4): 1073-1 081.
1 967. Localization of structural polymers
in the cell wall of Neurospora crassa. J. Cell Biol. 35:295-302.
Mandal, C. and C. Mandal. 1990. Sialic acid binding lectins. Experimentia
46:433-441.
Manners, D.J., and A. Wright. 1962. a-1,4-glucosans. XIV. The interaction of
Concanavalin-A with glycogens. J. Chem. Soc. 1962(4): 4592-4600.
Mason, D.Y., J.L Cordell and K.A.F. Pulford. 1983. Production of monoclonal
antibodies for immunocytochemical use. Immunocytochemistry 2:175-
216.
Mauseth, J.D. 1988. Plant anatomy. Benjamin Cummings Pub. Co., MenIo
Park, CA.
McLean, I.W. and P.K. Nakane. 1974. Periodate-lysine-paraformaldehyde
fixative: A new fixative for immunoelectron microscopy. J. Histochem.
Cytochem. 22:1077-1083.
McMichael, A.J. and J.W. Fabre (eds.). 1982. Monoclonal antibodies in clinical
medicine. Academic Press, New York.
Merkus, E. 1973. Ultrastructure of the ascospore wall in Pezizales
(Ascomycetes) - I. Ascodesmis microscopica (Crouan) Seaver and A.
Nigricans van Tiegh. Persoonia 7(3):351 -366.
• 1974. Ultrastructure of the ascospore wall in Pezizales
(Ascomycetes - II. Pyronemataceae sensu Eckblad. Persoonia 8(1): 1 -22.
• 1975. Ultrastructure of the ascospore wall in Pezizales
(Ascomycetes) - III. Otideaceae and Pezizaceae. Persoonia 8(3):227-
247.
■ 1976. Ultrastructure of the ascospore wall in Pezizales
(Ascomycetes) - IV. Morchella Helveilaceae. Rhizinzceae. Thelebolaceae.
and Sarcoscvphaceae. General discussion. Persoonia 9(1):1-38.
Miller, R.L, J.F. Collawn, and W.W. Fish. 1982. Purification and macromolecular
properties of a sialic acid-specific lectin from the slug Umax flavus. J.
Biol. Chem. 257:7574-7580.
Mims, C.W., E.A. Richardson, and J.W. Kimbrough. 1990. Ultrastructure of
ascospore delimitation in freeze substituted samples of Ascodesmis
nigricans (Pezizales). Protoplasma 156:94-102.
Mishell, R.L, and R.W. Dutton. 1966. Immunization of normal mouse spleen cell
suspensions in vitro. Science, 153:1004-1006.
Miyazaki, T., T. Yadomae, H. Yamada, O. Hayachi, I Suzuki, and Y. Ohshima.
1980. Immunochemical examination of the polysaccharides of
mucorales. In: P.A. Sandford, and K. Matsuda (eds.). Fungal
Polysaccharides. American Chemical Society, Washington D.C., pp. 81-
94.
Mol, P.C. and J.G.H. Wessels. 1987. Linkages between glucosamino-glycan
and glucan determine alkali-insolubility of the glycan in walls of
Saccharomvces cervisiae. FEMS Microbiology Letters 41(1):95-99.
Monsigny, M., A.-C. Poche, C. Sene. R. Maget-Dana, and F. Delmotte. 1980.
Sugar-lectin interactions: How does wheat-germ agglutinin bind
sialoglycoconjugates? Eur. J. Biochem. 104:147-153.
Morre, D.J., H.H. Mollenhauer, and C.E. Bracker. 1971. Origin and continuity
of golgi apparatus. In: J. Reinert, and H. Ursprung (eds.). Results and
problems in cell differentiation: Origin and cell continuity of cell
organelles. Springer-Verlag, New York, 2:82-126.
Nakane, P.K. and G.B. Pierce. 1966. Enzyme-labeled antibodies: Preparation
and application for the localization of antigens. J. Histochem. Cytochem.
14:929-931.
Nawaz, M.S. and W.M. Hess. 1987. Ultrastructure of Neovossia horrida
teliospores. Mycologia 79(2): 173-1 79.
Neville, A.C. 1 975. Biology of the arthropod cuticle. Springer-Verlag, New York.
Newman, G.R. 1987. Use and abuse of LR White. Histochem. J. 19:118-120.
Newman, G.R. and J.A. Hobot. 1987. Modern acrylics for post-embedding
immunostaining techniques. J. Histochem. Cytochem. 35(9): 971 -981.
w
148
Newman, G.R. and B. Jasani. 1984. Post-embedding immunoenzyme
techniques. In: J.M. Polak and I.IVI. Varndell (eds.). Immunolabelling for
electron microscopy. Elsevier Science, New York, pp. 53-70.
Niyo, K.A., H.S. McNabb Jr., and L.H. Tiffany. 1986. Ultrastructure of athe
ascocarps, asci, and ascospores of Mvcosphaerella populorum.
Mycologia 78(2): 202-2 12.
Notario, V., T.G. Villa, and J.R. Villaneuva. 1979. Cell wall associated 1,4-beta-
D-xylanase in Cryptococcus albidus var. aerius: in situ characterization
of the activity. J. Gen. Microbiol. 114:415-422.
Novaes-Ledieu, M. and C.G. Mendoza. 1981. The cell walls of Aqaricus
bisporus and Aqaricus campestris fruiting body hyphae. Can. J.
Microbiol. 27(8}: 779-787.
Pardue, R.L, R.C. Brady, G.W. Perry, and J.R. Dedman. 1983. Production of
monoclonal antibodies against calmodulin by in vitro immunization of
spleen cells. J. Cell Biol. 96:1149-1154.
Peberdy, J.F. 1990. Fungal cell walls - A review. In: P.J. Kuhn, A.P.J. Trinci,
M.J. Jung, M.W. Goosey, and L.G. Copping (eds.). Biochemistry of cell
walls and membranes in fungi. Springer-Verlag, New York, pp. 5-30.
Pereira, M.E.A., E.A. Kabat, R. Lotan, and N Sharon. 1976. Immunochemical
studies on the specificity of the peanus (Arachis hypogaea) agglutinin.
Carbohydr. Res. 51:107-118.
Pollard, K., D. Lunny, C.S. Holgate, P. Jackson, and C.C. Gird. 1987. Fixation,
processing, and immunochemical reagent effects on preservation of T-
lymphocyte surface membrane antigens in paraffin-embedded tissue. J.
Histochem. Cytochem 35(11): 1329-1 338.
Prabhakar, B.S., M.V. Haspel, and A.L Notkins. 1984. Monoclonal antibody
techniques applied to viruses, in: K. Maramorosah and H. Koprowski
(eds.). Methods in virology, volume VII. Academic Press Inc., New York,
pp. 1-18.
Preston, J.F. and J.E. Gander. 1968. Isolation and partial characterization of the
extracellular polysaccharides of Penicillium charlesii. I. Occurrence of
galactofuranose in high molecular weight polymers. Arch. Biochem.
Biophys. 124:504-512.
Raspail, F.V. 1830. Essai de chime microcopique appliquee a la physiologie.
Paris.
Rees, D.A. 1977. Polysaccharide shapes. Chapman and Hall, London.
149
Reeves, F., Jr. 1967. Ascospore formation in Pyronema domesticum.
Mycologia 59:1018-1033.
Reiss, E. 1986. iVIolecular immunology of mycotic and actinomycotic infections.
Elsevier Scientific, New York.
Reiss, E., J.B. Knowles, S.L Bragg, and L Kaufman. 1986a. Monoclonal
Antibodies against the M-protein and carbohydrate antiges of
histoplasmin characterized by the enzyme-linked immunoelectrotransfer
blot method. Infect. Immun. 53(3):540-546.
Reiss, E., L DeRepentigny, R.J. Kuykendall, A.W Carter, R. Galindo, P.Auger,
S.L. Bragg, and L Kaufman. 1986b. Monoclonal antibodies against
Candida tropicalis Mannan: Antigen detection by enzyme immunoassay
and immunofluorescence. J. Clin. Microbiol. 24(5): 796-802.
Romano, E.L., C. Stolinski, and N.C. Hughes-Jones. 1974. An antiglobulin
reagent labelled with colloidal gold for use in electron microscopy.
Immunochemistry, 1_7:521 .
Rosenburger, R.F. 1976. The cell wall, in: J.E. Smith and D.R. Berry (eds.),
The filamentous fungi, volume 2. Wiley, New York, pp. 328-344.
Rosing, W.C. 1982. Ultrastructure of Ascus and ascospore development in
Chaetomium brasiliense. Mycologia 74(6):960-974.
1985. Fine structure of cleistothecia, asci, and ascospores of
Mvxotrichum deflexum. Mycologia 77(6): 920-926.
Roth, J. 1983. Application of lectin-gold complexes for electron microscopic
localization of glyconjugates on thin sections. J. Histochem. Cytochem.
31:987-999.
. 1989. Postembedding labeling on lowicryl K4M tissue sections:
Detection and modification of cellular components. Methods Cell Biol.
31:513-551.
Rudall, K.M. 1969. Chitin and its association with other molecules. J. Polymer
Sci. C28:83-102.
Salt, S.D. and J.D. Gander. 1985. Variations in phosphoryl substituents in
extracellular peptidophosphogalacto-mannans from Penicillium charlesii
G.Smith. Exp. Mycol. 9:9-19.
Schutzbach, J.S. and H. Ankel. 1 972. Mannosyltransferases from Cryptococcus
laurentii. Methods in Enzymology 28:553-560.
150
Sercarz, E., B. Bonavida, A. Miller, R.J. Sclblendki, and J.A. Stratton. 1974.
Immune response to Lysozyme: Limited heterogeneity caused by
restricted T cells. In: E.F. Osserman, R.E. Canfield, and S. Beychok
(eds.). Lysozyme. Academic Press, New York, pp. 143-152.
Sharon, N. and H. Lis. 1989. Lectins as cell recognition molecules. Science
246:227-234.
Simmons, R.B. 1989. Comparison of chitin localization in Saccharomvces
cerevisiae and Crvptococcus neoformans. and Malassezia spp. Mycol.
Res. 93(4): 551 -553.
Singer, S.J. 1959. Preparation of an electron dense antibody conjugate.
Nature (London) 183:1523-1524.
Smith, E.E., Z.H.G. Smith and I.J. Goldstein. 1968. A turbidimetric study of the
interaction of concanavalin A with amylopectin and glycogen and some
of their enzymic and chemical degradation products. Biochem. J.
107:715-724.
Stagg, CM. and M.S. Feather. 1 973. The characterization of a chitin-associated
D-glucan from the cell walls of Aspergillus niqer. Biochim. Biophys. Acta
320:64-72.
Stefanini, M., C. DeMartino, and L. Zamboni. 1967. Fization of ejaculated
spermatozoa for electron microscopy. Nature 216:173-174.
Ste-Marie, L, S. Senechal, M. Boushira, S. Garzon, H. Strykowski, L Pedneault,
and L. DeRepentigny. 1990. Production and characterization of
monoclonal antibodies to cell wall antigen of Aspergillus fumiqatus.
Infection and Immunity, 58(7):21 05-21 14.
Sternberger, LA. 1986. Immunocytochemistry. Churchill Livingstone. New
York.
Stirling, J.W. 1989. Ultrastructural localization of lysozyme In human colon
eosinophils using the protein A-gold technique: Effects of tissue
processing on probe distribution. J. Histochem. Cytochem. 37(5): 709-
714.
. 1990. Immune- and affinity probes for electron microscopy: A
review of labeling and preparation techniques. J. Histochem. Cytochem.,
38(2): 145-1 57.
Sumner J.B. and S.F. Howell. 1936. The identification of the hemagglutinin of
the jack bean with concanavalin A. J. Bacteriol. 32:227-237
151
Tanner, W. 1990. Synthesis and function of glycosylated proteins in
Saccharomvces cerevisiae. in: P.J. Kuhn, A.P.J. Trinci, M.J. Jung, M.W.
Goosey, and LG. Copping (eds.), Biochemistry of cell walls and
membranes in fungi. Springer-Verlag, New York, pp. 109-118.
Tolson, N.D., B. Boothroyd,^nd C.R. Hopkins. 1981. Cell surface labelling with
colloidal gold particulates: The use of avidin and staphylococcal protein
A-coated gold in conjunction with biotin and Fc-bearing ligands. J.
Micorsc. 123:21 5-220.
Voller, A., and D.E. Bidwell. The enzyme linked immunosorbent assay, volume
2: A review of recent developments with abstracts of microplate
applications. Microsystems, Guernsey, U.K.
Weibull, C, A. Christiansson, and E. Carlemalm. 1983. Extraction of membrane
lipids during fixation, dehydration and embedding of Acholeplasma
laidlawii - cells for electron microscopy. J. Microsc. 129:201-207.
Wessels, J.G.H. 1979. Wall structure and growth in SchizophvUum commune,
in: J.H. Wessels & A.P.J. Trinci (eds.). Fungal walls and hyphal growth
pp. 27-48.
. 1986. Cell wall synthesis in apical hyphal growth. International
Review of Cytology 104:37-79
Wu, C.G. and J.W. Kimbrough. 1991a. Ultrastructural studies on
ascosporogenesis in Ascobolus stictoideus (Pezizales, Ascomycetes).
Can. J. Bot. (in press).
1991b. Ultrastructural studies on
ascosporogenesis in Ascobolus immersus (Pezizales, Ascomycetes).
Mycologia (in press).
Yamasaki, F.B., D.T. and R.E. Feeney. 1982. Periodate oxidation of methionine
in proteins. Anal. Biochem. 125:183-189.
Young, K.D. and H.W. Larsh. 1982. Production and characterization of a
hybridoma-derived antibody to Blastomvces dermatitidis. J. Clin.
Microbiol. 15:204-207.
Zickler, D. and J.-M. Simonet. 1980. Identification of gene-controlled steps of
ascospore development in Podospora anserina. Exp. Mycol. 4:191-206.
Zonneveld, B.J.M. 1971. Biochemical analysis of the cell wall of Asperaillus
nidulans. Biochim. Biophysi. Acta 249:506-514.
4' ■'
ti- ; -■
152
. 1972. Morphogenesis in Aspergillus nidulans: the
significance of a -1 ,3 glucanase for cleistothecium development. Biochim.
Biophys. Acta 273:174-187.
. 1 973. Inhibitory effect of 2-deoxyglucose on cell wall a -1 ,3
glucan synthesis and cleistothecium development in Aspergillus nidulans.
Developmental Biology 34:1-8.
BIOGRAPHICAL SKETCH
My given name Is Demaris Ellen Lusk, but most people know me as "Dee." I was
born in Eugene, Oregon on 19 June 1957. I grew up on a small family farm near
Eugene. There I learned to garden, can, cook, and raise both dairy and beef stock. I
spent many hours walking in the neighboring woods and valleys. During these walks
I gained a appreciation for the land and its many plant and animal residents.
I graduated from Winston Churchill High School in 1 975. A year after graduation
I enrolled at Lane Community College. While attending this college I decided on a
program in plant sciences. After two years of basic course work at Lane Community
College I transferred to Oregon State University in Corvallis, Oregon. There I earned a
Bachelor of Science degree and a Master of Science degree, both in Botany. My
masters thesis work was in the area of fungal systematics. I came to the University of
Florida to continue my studies of fungi and their biology. Additionally, I had hoped to
learn techniques of electron microscopy. I have achieved these goals and more here
at the University of Florida and I am grateful for the outstanding training I have received.
153
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Henry AldriiHr Chairman
Professor of Microbiology and Cell
Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor oW^hilosophy. y
^mes Preston
rofessor of Microbiology and Cell
Science
i^rc
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
James Kimbrough y.
Professor of Plant Pathology^
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Walter Judd
Professor of Botany''
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor ofpliilosophJ.
Dbna Griffin III
Professor of Bote
This dissertation was submitted to the Graduate Faculty of the Department of
English in the College of Liberal Arts and Sciences and to the Graduate School and was
accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy.
August 1 991 '-y)^acUhr^ C^^^^^<^^^'^-^-^
Dean Loctefnart
Graduate School
r4r
4
I
UNIVERSITY OF FLORIDA
3 1262 08554 0374
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