DEVELOPMENT OF CYTOCHEMICAL METHODS
FOR THE STUDY OF
ASCOSPORE WALL BIOGENESIS AND MATURATION
DEMARIS E. LUSK
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
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 all
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
I received two very important gifts for use in my research: a culture of
Ascodesmis sphaerospora from Dr. Kimbrough, and GS-l 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.
TABLE OF CONTENTS
ACKNOW LEDGM ENTS ..................................................................................... ................. ii
LIST O F TABLES ........................................................................................................... vi
LIST O F FIGURES ........................................................................................................ vii
ABSTRACT .................................................................................................................... ix
1 INTRODUCTIO N ........................................... ................ .............................. 1
Ascospores .................................................................................................... 1
Ascosporogenesis .............................................................................................. 3
Chem istry of the Ascospore W all ............................................. ........... .... 13
Chem istry of the Hyphal W all ................................... .............. .............. 14
Study Proposal ............................................................................................ 29
Conclusions ................................................................................................. 31
2 STRATEGIES FOR TISSUE PREPARATION AND EMBEDDING ............... 33
Literature Review ........................................................................................ 33
Tissue Preparation Strategies ................................................. ............. 40
3 DEVELO PM ENT O F ANTIBO DIES ........................................... ........... ... 44
Introduction ................................................................................................. 44
M materials and M ethods ................................................... ........................ 50
Results ........................................................................................................ 54
Discussion ................................................................................................... 58
4 IM M UNOCYTOCHEM ISTRY ................................................................... 61
Introduction ................................................................................................. 61
M materials and M ethods ................................................... ........................ 63
Results ........................................................................................................ 68
Discussion ................................................................................................... 86
5 LECTIN CYTOCH.EMISTRY ....................................................................... 90
Introduction ................................................................................................. 90
Binding Specificities ........................................................ ......................... 91
Materials and Methods ................................................... ........................ 96
Results ........................................................................................................ 98
Discussion ..................................................................................................... 116
6 CONCLUSIONS ............................................................................................ 119
Evaluation of Experimental Methods .......................................................... 119
Ascospore W all Chemistry ............................................................................. 121
Precursor Tracking ................................................................................... 122
Maturation of the Ascospore W all ............................................................... 122
APPENDICES ................................................................................................................... 124
A FUNGAL CULTURE .................. ..................................................... ... 124
B ISOLATION OF ASCOSPORE W ALLS ....................................................... 125
C PROTOCOLS FOR HYBRIDOMA CONSTRUCTION AND CLONING ....... 127
D FREEZING AND THAWING HYBRIDOMA OR SP2-O CELLS ................. 130
E LIGHT BREAK OF ASCOSPORES ............................................................. 132
F FIXATION PROTOCOL ............................................................................. 134
G ANTIBODY LABELING .................................................................................. 135
LITERATURE CITED ........................................................................................................ 137
BIOGRAPHICAL SKETCH ............................................................................................... 153
LIST OF TABLES
Table 3.1. Mean optical densities for buffer wash, substrate, and secondary
............... ......... .................... ........ ......................... ................ ... ....... ...... 56
Table 3.2. Least square means comparison of washing vs substrate and
antibody / substrate treatment.
................................ ....... ........... ............................. .... .............. .. ..... 5 7
Table 3.3. Least square means comparison of interaction of buffer type with
..................................... .............. ...... ............................................................. 5 7
Table 3.4. Mean optical densities for buffer control, immune mouse and test
.............. .... ... ....... .............................. .. ........................................................... 5 8
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
..............................................................................................................................1 1 7
Table 6.1. Comparison of wall labeling patterns. ....................................................... 121
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. 8F11 culture supernatant labeling on A. sphaerospora ........................... 72
Figure 4.4. Collage of 8F11 positive labeling. ........................................... ........... .... 74
Figure 4.5. Determinant characterization for 8F11. ........................................................... 75
Figure 4.6. Developmental sequence with 8F11 labeling ............................................... 76
Figure 4.7. Antibody 41-1.1 labeling. .................. ................ ..........................................79
Figure 4.8. Pronase pretreatment with antibodies 12-2 and 41-1.1. ........................... 80
Figure 4.9. Antibody 12-2 labeling .................................................. ......................... 83
Figure 4.10. Anti-A. sphaerospora ascospore wall antibody lableing on P. nigrella. .... 85
Figure 5.1. WGA labeling on A. sphaerospora. .......................... .......... ........... .. 98
Figure 5.2. GS-II labeling on A. sphaerosora .................................................................. 100
Figure 5.3. WGA labeling on P. nirella. ..................................................................... 101
Figure 5.4. GS-II labeling on P. nigrella. ...................................................................... 102
Figure 5.5. WGA labeling with sugar control. .................................................................. 103
Figure 5.6. LFA labeling on A. sphaerospora .................................................................. 104
Figure 5.7. LFA labeling on or around spent cells of A. sphaerospora ....................... 106
Figure 5.8. Con A labeling on A. sphaerospora ............................................................... 109
Figure 5.9. Con A labeling on P. nirella. ...................................................................... 111
Figure 5.10. Con A labeling with a-mannosidase and/or pronase pretreatments...... 113
Figure 5.11. Collage of lectin labelings on A. sphaerospora........................................ 115
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
DEMARIS E. LUSK
Chairman: Dr. Henry C. 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
uncloned preparation, 8F11, 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.
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
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 (1897) "free-cell formation" has been
shown via ultrastructural studies to be correct. Studies of mature ascospore
morphology via light microscopy (e.g., LeGal, 1947,1951) have been a starting point for
electron microscopy studies, as well as having provided invaluable data for taxonomic
application. In the area of cell biology, ascospore ontogeny (ascosporogenesis) has
interested researchers because it represents an unusual type of cytokinesis and cell
Ultrastructural studies of ascosporogenesis came into vogue around the early
1960's, apparently culminated in Beckett's (1981) 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
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
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
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
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.
In general, for ascomycetes, the process of meiotic spore ascosporee)
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 initial 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
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, 1973; Hohl & Streit, 1975). 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.
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
summarized the research to that date on the origin of these membranes. Structures
implicated in the formation of SDMs include: the ascus plasma membrane, mesosomes,
myelin figures, cisternae of endoplasmic reticulum, endomembrane vesicles, and the
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, 1967). Niyo and coworkers (1986) noted the presence
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 Geopyxis 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 (1967) and Rosing (1982)
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 (1976) suggested that spore plasma membrane plays a role in the synthesis of
primary wall material. These suggestions are supported by Beckett's (1981) 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 (1967) found few lomasomes in
Pyronema 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,
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, 1988b). 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 proceed use of these data for taxonomy.
The Secondary 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
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, 1976). Beckett (1981) 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 (1976) outlines seven different developmental groups
within the Pezizales alone.
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
formation and that the investing membrane might have an active role. Based on
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 (1991 a, 1991b) 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
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 (1988a) 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
materials (in H. macropus and H. elastica, Gibson & Kimbrough, 1988a; in Gyromitra
esculenta, Gibson & Kimbrough, 1988b; Dyby & Kimbrough, 1987; in Geopyxis
carbonaria, Kimbrough & Gibson, 1990; Kimbrough et al., 1990; in Ascobolus immersus,
and A. stictoideus, Wu & Kimbrough, 1991 a & 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, 1988a; in Ascobolus immersus, and A. stictoideus, Wu & Kimbrough 1991a
& 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.
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
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 al., 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 appearance 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).
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., 1990; Kimbrough & Gibson, 1990) are evident in
developing ornaments and wall thickenings.
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)
differential staining of the secondary wall, and 4) a change in the permeability of the
Chemistry of Ascospore Wall
Very little information on the chemical composition, or nature, of ascospore wall
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 primary walls of Gyromitra esculenta (1988b) and
Helvella spp. (1988a) 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, 1987). They
speculated that the secondary wall ornaments consisted of lipids, protein, glycoprotein,
and possibly chitin. In Geopyxis carbonaria (Kimbrough & Gibson, 1990) and Gyromitra
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
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 Hyphal 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, 1968).
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
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, 1986). The many functional aspects and dynamic nature
of cell walls have prompted some researchers to recognize cell walls as organelles
Hyphal walls have been reported to be 80%-90% polysaccharide (Farkas, 1979;
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 (1968) stated that "at least 11 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 (B(1-3) linked homopolymer; Farkas, 1979). It is
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 B(1-3)
and 1(1-6) linkages are the principal protein-associated substances) in fungi. Glucans
of these same linkages, although with a higher degree of B(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
heteropolysaccharides, glyco-conjugates, proteins, and lipids or lipo-conjugates.
Survey of Methods
Current knowledge about the architecture and chemistry of hyphal walls is
founded in three basic research methods. These are 1) degradation (extraction,
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
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
impurities, Johnston, 1965), chemical extractions begin with hot water or/then mild alkali
(e.g., 5% KOH), followed by acid hydrolysis of the soluble fraction(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 B(1-4) and 1(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 1(1-3)
and B(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 B-(1-3)-B-(1-6) glucans have longer (1-6)-B-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
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 a -D-Man(p) and B-D-Gal(f)." These differences are primarily number of units per
Mol and Wessels (1987) 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 & Tatum, 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 (1971) used 2%
hydrochloric acid at 100C 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% H2SO, (v/v) at 40C 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, 1971). 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
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 B(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, 1971; thin-layer and HPLC, Briza et al., 1986;
gas-chromatography, Stagg & Feather, 1973; SDS-PAGE, Hearn et al., 1989),
colorimetric/spectrophotometric (e.g., Mol & Wessels, 1987; Novaes-Ledieu & Mendoza,
1981; Zonneveld, 1971, 1972; Mahadevan & Tatum, 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., 1989; NMR, Briza et al., 1986,1988; C-n.m.r., Gorin & lacomini, 1984) 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
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., 1988), and fluorescent-labelled conjugates (e.g., Briza et al., 1988) 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-
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.
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.
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,
1986). It accounts for a significant portion of the wall in some fungi (e.g., about 10% in
Neurospora crassa, Burnett, 1979; and 9-13% in Aspergillus niger, 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-, 1-, and I-) are known
(Rudall, 1969). The 1- form is made up of chains piled in parallel orientation to one
another while the a- form is of antiparallel orientation, and the '- form (Fig. 8) has both
parallel and antiparallel polymer components. The a form is the most stable (Rudall,
1969), and the form present in fungal chitin (Rudall, 1969; Wessels, 1986).
Rudall (1969) describes fungal chitin as spirallyy 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 B-
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
Chitosan has been found in the walls of Zygomycetes (Bartnicki-Garcia, 1968),
non-reproductive and non-lamellae fruit-body cells of the Basidiomycete species
Agaricus bisporus (brunnescens) and A. campestris (Novaes-Ledieu & Mendoza, 1981),
Sacchromyces cerevisiae cells in early stationary growth phase (Mol & Wessels, 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 B(1-3), B(1-6), B(1-3)-B(1-6), a (1-3), a (1-3)-a (1-4) linked, and possibly a (1-4)
Although the presence of cellulose (B(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 1(1-3) linked glucans provide
the mechanical support for fungal cells that cellulose does for higher plant walls.
Pure B(1-3) glucan, or those polymers with infrequent 8(1-6) linkages, can
crystallize into microfibrils (Burnett, 1979). The extent to which B(1-3)-13(1-6) glucans can
crystallize seems to be dependent on the frequency of 8(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
(Mol & Wessels, 1987), associated with chitin (Wessels, 1986; Rudall, 1969) and in
Aspergillus niger (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. niger 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
Schizophyllum 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, 1987),
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
glucan now goes by the name nigeran. Johnston (1965) found this glucan in the hyphal
walls of A. niger. Based on Johnston's data, Gorin (1968) reported this component to
represent 26-42% of the total wall, Aronson (1981) reported 4-6%, and by this author's
rough calculation from that data, 10%. For A. nidulans Zonneveld (1971) reported that
few, if any a (1-4) glycosidic linkages exist. This large discrepancy between species may
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 (1965) written opinion, that pseudonigeran was the
glucan present in A. niger rather than both nigeran and pseudonigeran because neither
were soluble in hot water. Pseudonigeran is thought to be more widespread
taxonomically than nigeran (Aronson, 1981). Zonneveld (1972, 1973) found a (1-4)
linked glucose residues in the alkali-insoluble fraction along with B(1-3), 8(1-6),
mannose-galactose polymers, and chitin. Horikoshi and lida (1964) reported a glucan
of aX (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., 8-glucan-galactomannorhamnan in Fusicoccum
amygdali) "are unquesionably significant" as they knit various polysaccharides into larger
wall complexes. No reports of a consecutively X (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
described as having an a-D-(1-6) backbone, and a-D-(1-3) and a-D-(1-2) branches
(Gorin, 1968) of two to five residues (Reiss, 1986). Farkas (1979) described yeast
mannan as a glycoprotein with 2 distinct carbohydrate moieties; one with a -D-(1-6)
backbone and a0-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,
1968). In Candida albicans cell wall mannans with a -D-(1 -2) and a -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 (9-10 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, 1968).
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
10-15% 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
of sealing in unbound wall materials (Farkas, 1979). 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
two major categories; the proteinases and 8(1-3)glucanases (Reiss, 1986). Other
enzymes known to occur murally are invertase, acid phosphatase (Farkas, 1979; Reiss,
1990) and 8(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 oryzae 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).
Bartnicki-Garcia (1968) presented evidence supporting lipid(s) as a bonafide wall
component. Cell walls of Aspergillus niger have been reported to be 2-7% lipid
(Johnston, 1965). No further information was found on its relationships) with other wall
components, conjugate partnerss, or roles within the wall.
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
Brown's (1957) description of chitin synthesis in fungi. The bulk of chitin synthetase has
since been found to be attached to the plasma membrane (Duran et al., 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 remained 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 8(1-3)glucans. B(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., 1989; in Mucor rouxii, Fevre et al., 1990; Peberdy,
1990). Activity of 8(1-3)glucan synthase has also been found in association with both
endoplasmic reticulum and plasma membrane fractions (in, Saprolegnia monoica, Fevre,
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
Cryptococcus 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 secretary vesicles. Mannoprotein
formation in yeasts has long been thought to be a process involving much or all of the
endomembrane system (Farkas, 1979).
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
in comparison to hyphae. Yet, this type of research would add greatly to our
understanding of the biology of this group of organisms.
The most fundamental questions are that of chemistry and biosynthesis of
ascospore walls and their constituent layers. Toward answering such questions some
researchers have published a limited amount of cytochemical data. Those experiments
have provided information 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 B(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
specific components 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
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.
Materials and Methods
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;
carbohydrate, lectin, enzyme, and immuno-, cytochemistry; and use of secondary
probes) was outlined by Aldrich and Todd (1986). More specific information on these
techniques was also readily available and is reviewed in chapters 2, 4, and 5.
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
proposed as a third line of techniques research.
Production of anti-spore wall antibodies was seen as necessary for successful
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
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
of ascosporogenesis. The procedures developed would be applicable to other fungi
and had the potential for addressing other biological questions. Thus, as proposed, it
was felt that this work had great potential for provision of a foundation of data for future
research on the biology of Ascomycete fungi.
TISSUE PREPARATION AND EMBEDDING
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
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,
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
these molecules specifically prior to the actual tissue processing and cytochemical
Tissue processing involves fixation of the material, sometimes a secondary
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
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
develop fixation and dehydration protocols and chose an embedding medium optimal
for the cytochemical probe, and more specifically for its target molecule.
The goal of fixation is to kill and stabilize cell structures. This should be done
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
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
(1981) 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, 1981, 1986). Glutaraldehyde reacts with the
C-amino group of lysine, N-terminal amino groups, a-amino groups of free amino acids,
protein associated DNA, and the 1 amino groups of ethanolamine containing
phospholipids (Hayat, 1986; Sternberger, 1986). Most lipids (other than phospholipids),
myelin, and glycoproteins are not fixed by glutaraldehyde (Hayat, 1986). 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 40C or room temperature is usually adequate. Low concentrations of
glutaraldehyde are recommended for immunocytochemistry (especially with monoclonal
antibodies, Beesley, 1989) and enzyme cytochemistry (Bendayan, 1989a) since retention
of biological configuration can be altered by this fixative. Loss of antigenicity or receptor
integrity during dehydration and infiltration may be reduced by glutaraldehyde (Craig &
Goodchild, 1982). DeWaele and coworkers (1983) reported that some glutaraldehyde
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
components are at least partly reversible. It penetrates tissue rapidly and in that respect
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
of lipids (Hayat, 1986). If only this fixative is used, lipids may be extracted during
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, 1986). 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, 1986). Osmium tetroxide is said to denature the a -helix regions
of membrane proteins (Lenard & Singer, 1968).
Additives to Primary Fix
Additives to the primary fixative solution such as picric acid (Stefanini et al., 1967;
Dae et al., 1982) periodate-lysine (Hixson et al., 1981; McLean and Nakane, 1974:
Pollard et al., 1987), and tannic acid (Stirling, 1989) have been recommended to improve
morphology without loss of antigenic or binding site receptivity (Stirling, 1990).
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, 1987). 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 100% followed by acetone washes.
Acetone may also be used with the acrylic Lowicryl resins. Kellenberger and coworkers
(1987) 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 1962 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)
preservation of binding site receptivity (antigenicity) within the tissue and 2) steric
hindrance of the probe (Causton, 1984). 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-
linked systems or cross-linked hydrophilic acrylics for best results and greatest flexibility
Another potential problem discussed by Newman and Hobot (1987) 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, 1987). 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
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 (RCH3)
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
hydrogen peroxide, periodic acid (periodate) or potassium permanganate produces
hydrophilic groups, thus destroying the hydrophobic barrier. These treatments may also
oxidize target molecules and therefore are best avoided (Causton, 1984; Newman &
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 likely 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-linking.
They further postulate, as Kellenberger and coworkers (1987) 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
done at low temperatures. Although ultrastructural preservation is improved by low
temperature methods, Newman (1987) 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 (500 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 (1987) 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
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
contribution to our knowledge of ultrastructural chemistry their experiments could
provide. Similar acceptance of poor morphology was proposed as a starting place for
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
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, 1983; Bendayan 1984a,
1984b, 1989a, 1989b). This treatment, in turn, could damage some carbohydrates and
oxidize alkanes. Stirling (1990) recommended preparing tissue with a number of
different fixations. Following the recommendations of Erdos (personal communication)
and Stirling (1990), the following preparations were proposed as an adequate start-up
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.
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
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
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 12% 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 (1987) 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.
DEVELOPMENT OF 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., 1974). 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
Antiserum developed against an antigen typically contains antibodies to a
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
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).
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
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, 1989). Reiss (1986) 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
Telletia sp., Banowetz et al., 1984; for Ophiostoma ulma, Benhamou & Ouellette, 1986;
for Phytophthora cinnamomi, Hardham et al., 1985, 1986; 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
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
Aspergillus 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-o -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, 1986).
Information and discussion on the production of monoclonal antibodies is
abundant and easily found in immunology text books (e.g., McMichael & Fabre, 1982;
Goding, 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 demonstrated by Mishell and coworkers (1967). Basically, these techniques differ
from the in vivo techniques by immunization of non-immune B-lymphocytes (B-cells) in
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 al., 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 106 cells or parts per immunization is recommended
(Bastin et al., 1982; Mason et al., 1983). 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, 1975), 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 (1983) 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
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 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,
antigen) can be immobilized on a carrier surface while retaining its activity or capacity
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,
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
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
Hybridoma culture supernatant contains a sufficient quantity of antibody (5-
251. g/ml; Edwards, 1981) to not only allow ELISA screening, but also be a useful source
Ascites fluid is produced by planting hybridoma cells in the intra-peritoneal cavity
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., 1983). Based on the Mason's argument, ascites will not be
used in this project.
Materials and Methods
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.
Isolation of Immunogen
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
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
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 10" parts per ml and
the second contained about 7 x 108 parts per ml. Mice were given approximately 2-3.5
x 107 parts per 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.
Only one mouse was used from the second set of 3. Its immunization schedule
was similar to that of the second mouse from the first set.
Hybridoma Production and Cloning
The complete protocol used for hybridoma production and cloning, including
materials, is given in Appendix C. A total of 3 fusions were performed.
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; Goding, 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 106 cells per ml in appropriate media and either plated out at 100.I per
well of a 96-well plate or placed in a sterile growth flask and cultured as for hybridomas.
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 37C 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 IgG/IgM specific
(Jackson Laboratory), and 5) the enzyme substrate was p-nitrophenylphosphate. Plates
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
institute Inc.) program, on an IBM PS 2 computer.
EM screening methods are given in Appendix F, and are discussed in chapter
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 600C 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 -80C. Normal
mouse serum was handled in exactly the same manner.
When small volumes of culture supernatant were harvested but not used
immediately, the supernatant was stored in microfuge tubes at either 4C, -20C, or -
80C 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 8-10 minutes). Sodium azide was added (0.1% w/v) and
then the supernatant was aliquoted, frozen and stored at -800C.
The protocol for preparation and freezing of hybridomas in given in Appendix D.
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
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.
The second and third fusions used mice that had an extended immunization
regime. These mice had greatly enlarged spleens.
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
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.
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.
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
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.
Table 3.1 Mean optical densities for buffer wash, substrate, and secondary antibody
Tab. 3.2 Tab. 3.3
Trtm. N Obs. Mean Std. Dev. LS # LS #
w/ 20 AB 8 0.5035 0.0782 1 1
w/o 20 AB 8 0.0489 0.0034 2 2
w/o 20 AB 8 0.0764 0.0018 3 3
w/20 AB 8 0.3030 0.0718 1 4
w/o 20 AB 8 0.0558 0.0079 2 5
w/o 2 AB 8 0.0896 0.0093 3 6
20 AB, secondary antibody
LS #, least square mean number
Table 3.2. Least square means comparison of washing vs substrate and antibody/
substrate treatment. (Data log transformed; x < 0.05 indicates a
Pr > ITI HO: LSMEAN (i)=LSMEAN (j)
i/j 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
Pr > ITI HO: LSMEAN (i)=LSMEAN (j)
i/j 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
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).
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
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.
Trtm. N Obs. Mean Std. Dev.
PBS 4 0.3692 0.0391
IM 4 2.4285 0.1749
NM 4 1.0942 0.3609
THST 4 0.3518 0.0350
IM 4 2.0472 0.3359
NM 4 0.7628 0.2314
IM = immune mouse
NM = normal mouse
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.
All 3 of the antibody preparations used came from the second fusion. The third fusion
was done primarily in an effort to reproduce antibody 8F11. 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 initial 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, 100p I 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. Although time has not permitted further work with the cells from
the third fusion, they are available.
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
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 (1983) and Sternberger (1986) expressed a preference for
EM screening of hybridoma culture supernatants when the final use is to be
immunocytochemistry. Mason and coworkers (1983) based their preference on the
arguments 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)
the results from immunocytochemical techniques are inherently more informative
(providing not only +/- results, but specific background and localization data). Based
on these opinions, arguments 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.
Brief History of Immunocytochemistry
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
(1966) 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
specific for particular substrates and that can demonstrate subcellular location. Causton
(1984) described immunocytochemistry as potentially being the most demanding of all
the staining techniques.
Immunolabeling 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-
Aspergillus fumigatus monoclonal antibodies. The first, MAbl, 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
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). Phytophthora cinnamon 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
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 (Goding, 1986). In any case, some idea of the nature of
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
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 monitored 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.
Ascospores were prepared for electron microscopy by an initial 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.
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.
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
Table 4.1 demonstrates the specific variations in fixation, dehydration, and resin
used to prepare material for electron microscopy.
Pseudoplectania niirella, used for comparitive work was collected in the Oregon
coastal range in March, 1990. 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
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
Pale gold to silver (70-90 nm) sections were cut on a RMC MT6000-XL
microtome or a LKB 8800 ultramicrotome II. 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
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 8F11, 12-2, and 41-1.1. The latter two are monoclonal.
Normal-mouse and immune-mouse serums, and PBS and THST buffers were
used as controls.
Several general types of experiments were performed: screening of hybridoma
culture supernatants, testing of positive and negative serum controls, monoclonal
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
Table 4.1: Tissue preparation and embedding.
I SP 1 2 3 4 5
Spores (LB) X
AC undisturbed X X X X X
30 min. 1 fix X X X
45 min. 10 fix X
60 min. 10 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
95% EtOH X X X X X X
LR White X X X X X X
LB, lightly broken
Table 4.1 continued.
11 12 13 14 15 16 26
AC pelleted X X X X X X X
30 min. 1 fix X X
45 min. 10 fix
60 min. 10 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 / 3-4% F X X
2% G / 2% F X X X
Os04 X X
95% EtOH X X X X X X
LR White X X X X X X
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
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
most like the immunogen (I block), yet the wall structure is less disrupted and it retains
cytoplasm so that cross-reactivity could be monitored. Further, because this bolck 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, 14 and 15.
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
evaluation is sufficient for gross determination of specificity, background, and 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. 4.1A &
4.2B). Ascus and vegetative cell walls were labeled to a much lesser extent (fig.4.2A).
Additionally, sporoplasm, epiplasm (ascus cytoplasm), and vegetative cell cytoplasm
components were specifically labeled. Conversely, normal-mouse serum did not
specifically label any part of the fungus, and the background labeling was minimal (fig.
4.1C). The buffer negative control also had minimal background labeling (fig. 4.2B).
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 12 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.
Although not a monoclonal, antibody 8F11 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
SE o co -
c..E o c
o0 E (
U- c C h 0 0
II II II
0) 0.r )
S'" "'* "" .
'.** ** -
Figure 4.2. Serum labeling and buffer control on A. sphaerospora.
A) labeling of immune-mouse serum (from second fusion, diluted 1/1000);
B) buffer negative control.
Figure 4.3. 8F11 culture supernatant labeling on A. sphaerospora.
A) Labeling on the ascospore wall and sporoplasmic vesicles (pointers);
B) Labeling on the vegetative wall, including septum.
r ~ :
/'' '- 't" '' ... '
.wo, ,; .
* "- ., "'. m
Figure 4.5. Determinant characterization for 8F11.
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 sectionss. 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
0 Z3 -
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).
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 12-2 and 41-1.1.
A) antibody 41-1.1;
B) antibody 12-2;
C) buffer negative control with pronase pretreatment.
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
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.
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).
Antibodies 12-2 and 41-1.1 were tested for labeling on Pseudoplectania nigrella.
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.
4.10A). P. nigrella is the only other species these antibodies have been tested on to
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
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,
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
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
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 8F11 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
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
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
responsible for modification. An in situ modification seems the more probable of these
two suggested processes because of the suddenness of labeling and greater diversity
of ways for a modification to occur.
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
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, 1988). 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 (1984) description for epoxy resin
Antibody 8F11 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.
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
molecule before the mouse immunoglobulin response ensued. The immune system's
first response to a fungal invasion is a killer cell response and use of lytic enzymes
(Reiss, 1986). It is possible that the mouse killer response slightly altered this molecule
before the immune response proceeded to production of immunoglobulins and that
periodate and pronase pretreatments of sections in some way mimicked that alteration.
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 nigrella. 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.
Lectins are carbohydrate-binding proteins (or glycoproteins) of non-immune
origin which agglutinate cells and/or precipitate glycoconjugates (Goldstein et al., 1980).
As of 1986 no purified lectin had been shown to exhibit enzymatic activity (Goldstein &
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