Hyphal tip growth

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Title:
Hyphal tip growth molecular composition of elongating and non-elongating regions of Achlya cell wall
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vi, 107 leaves : ill. ; 29 cm.
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English
Creator:
Shapiro, Alexandra, 1971-
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Subjects / Keywords:
Fungi -- Hyphae -- Growth   ( lcsh )
Achlya -- Physiology   ( lcsh )
Botany thesis, Ph.D   ( lcsh )
Dissertations, Academic -- Botany -- UF   ( lcsh )
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bibliography   ( marcgt )
theses   ( marcgt )
non-fiction   ( marcgt )

Notes

Thesis:
Thesis (Ph.D.)--University of Florida, 2000.
Bibliography:
Includes bibliographical references (leaves 94-106).
Statement of Responsibility:
by Alexandra Shapiro.
General Note:
Printout.
General Note:
Vita.

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University of Florida
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All applicable rights reserved by the source institution and holding location.
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aleph - 025872528
oclc - 47103163
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AA00013527:00001


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HYPHAL TIP GROWTH: MOLECULAR COMPOSITION OF ELONGATING AND
NON-ELONGATING REGIONS OF ACHLYA CELL WALL












By

ALEXANDRA SHAPIRO


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


2000






TABLE OF CONTENTS


Page

ACKNOWLEDGMENTS.............. ........... ...................................................... iv

ABSTRACT.. .... ... ......... .... .......... .... ...................v

CHAPTERS

1 INTRODUCTIO N...... ...... ...............................................................................................

Apical Growt............................................................. ..1
The Organism o .............................. ..............................................................3
Class Oomycetes........... ...................................................................5

2 LITERATURE REVIEW ........ ....................................................................... ..... 9

Mechanism of Apical Growth.. .....................................................................9
Structure of the Hyphal Wall ....................... ............ ....... 10
Electron Microscopic Studies of Fungal Cell Walls........... 14
Biosynthesis of the Fungal Cell Wall ............................................21
Role of Turgor in Wall Expansion ........... ....... ........... 25
Role of Cytoskeleton in Hyphal Growth............. .................... 6
Cytology of Growing Hyphal Apices .............................................2 8
Calcium Gradient ............................................................................................. 31
Ion Current s......................... .......... ............................................................. 32

3 HYPHAL GROWTH................ ....... ............................... ................. 33

Introduction .......... ..........................3............................. ................................. 3
Materials and Methods.................-........ ............. ..............3 3
Results ............................................................................................................... 35
Discussion........................................................... ................... 35

4 LOCALIZATION OF CELLULOSE IN THE CELL WALL
AS REVEALED BY ELECTRON MICROSCOPY AND
CYTOCHEMICAL TECHNIQUE S ............... .......................... ....... .......... 39

Introduction .............................................................................................................. 39
Materials and Methods ............................................................................ ........ 40
Culture Methods and Microscopy Techniques.. .....4...0.............
Electron Microscopy................... ..............................4 0
Cellulose Localization Using Enzyme-Gold
Affinity Labeling..................................... ............................................4 2
Cytochemical Contro s........................ .......... ................. ....... 43
Cellulase Enzyme Activity during Labeling .......................... 44
Detection of Gold Particles with a
Backscatter Detector ......................................................................... 45
Zymolyase Hydrolysis ....................................................... ............4 5
Treatment of Growing Colonies with
Dichlorobenzonitril e...................................... ... ..... .... 4 5
Results ........................................................................................ 46
Cellulose Localization ..................................................................... 46






Cytochemical Control s........................ ..................................4 7
Cellulase Activity during Labeling........................................4 8
Cellulose Localization on the Surface of
the Hyphae in Colonies Incubated
with Zymo 1yase... ...................... ................ ....... ........ ........ 68
Hyphal Elongation, Spores Germination and
Cellulose Localization in the Presence
of DCB .................................................................. ............. 68
Discussion..... .. ......................................... .. 69

5 LOCALIZATION OF 1,3-B-GLUCANS IN THE CELL
WALL AS REVEALED BY ELECTRON MICROSCOPY
AND CYTOCHEMICAL TECHNIQUES ............................ ............. ..... 75

Introduction n........................ ................................................ 75
Materials and Methods.......................................................... ................... 78
Culture Methods, Fixation and Microscopy
Technique s............................................. .. 78
Localization of 1,3-B-Glucans Using
Monocl onal An tibody... ........................ ..................................
Cytochemical Control s.................................................................... .............79
Results ........................... ................. ................................................................................ 80
Localization of 1,3-B-Glucans on Sections and
Hyphal Surfaces Using Monoclonal Antibodies ................ 80
Cytochemical Controls.............................................................................. 80
Discussion.. ........................................................................... 86

6 LOCALIZATION OF CHITIN IN THE CELL WALL AS
REVEALED BY ELECTRON MICROSCOPY AND
CYTOCHEMICAL TECHNIQUES ........ ......................................................... 87

Introduction ....... ................. ................................................................................................... 87
Materials and Methods ..... ........................................ .......................................... 88
Chitin Localization Using Lectin................ ....... ....... 88
Cytochemical Control s. .................................................................. 88
Results ................................................................... 89
Discussion..................... ...................... ................. 89

7 CONCLUSIONS ...................................................................................... 2

LIST OF REFERENCE S....................... .................................................... 9 4

BIOGRAPHICAL SKETCH............................................... .......... 107


iii











ACKNOWLEDGMENTS

I thank Drs. Tom Emmel, Greg Erdos and Alice Harmon for

serving as members of my supervisory committee, and for their

time and expertise.

I would like to express my appreciation to Dr. J.T.

Mullins, my supervisory chairman, for his support,

understanding, patience, interest in the research project and

his help and guidance throughout the work on my dissertation.

I also would like to thank Karen Vaughn of ICBR EM core

lab for her technical assistance. My special thanks go to

Scott Whittaker of the same lab for his help, time and

technical expertise.

The support of my family was very important for me during

these years. I am truly indebted to my parents and my

parents-in-law for their love, constant encouragement and

inspiration, help with the kids and readiness to help anytime

I needed it.

Finally, my greatest gratitude goes to my husband, Andrei

Sourakov. His love, patience and help made the completion of

my dissertation possible.














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

HYPHAL TIP GROWTH: MOLECULAR COMPOSITION OF ELONGATING
AND NON-ELONGATING REGIONS OF ACHLYA CELL WALL

By

ALEXANDRA SHAPIRO

December 2000

Chair: J. Thomas Mullins
Major Department: Botany

Although apical growth is a widespread process in the

biological world and has been known for over a hundred

years, the mechanisms that underlie this process are not

yet understood. Knowledge of these mechanisms would allow

the development of techniques for inhibiting or

stimulating growth of medically or economically

important species. I approached the problem of hyphal tip

growth by comparing the cell wall composition of

elongating and non-elongating regions of the oomycete

Achlya bisexualis. Light microscope observations were

used to determine the growth rate and to distinguish

elongating and non-elongating hyphae for further EM

studies, because non-elongating hyphae often are found

among growing mycelia. I found that hyphal growth is a







discontinuous irregular process with periods of

elongation and no elongation. The elongation rate is not

steady, but instead fluctuates with periods of fast and

slow elongation. Both transmission and scanning electron

microscopes were used with a variety of cytochemical

labels, and several fixation techniques. Cellulose, the

microfibrillar component of the Achlya wall, was

identified with cellulase enzyme-gold affinity labeling.

Elongating hyphae have cellulose in mature and subapical

regions, but not at the apex. In non-elongating hyphae,

cellulose was found in all the regions including the

apex. These results suggest that the apices of elongating

hyphae lack cellulose. This contradicts the long-standing

hypothesis that the microfibrilar component is present in

the elongating hyphal apex. The 1,3-E-glucans, the major

matrix wall components, were immuno-localized in all

regions of elongating and non-elongating hyphae. A number

of cytochemical, biochemical and physiological controls

were performed to assure the reliability of these

findings. I suggest that in elongating regions, the

matrix is synthesized first and synthesis of

microfibrilar component follows. Another explanation for

these results is that localized apical cellulose

hydrolysis by endoglucanase creates plastic wall regions

consisting mainly of 1,3-1-glucans, which expand under

turgor and/or cytoskeleton pressure. Cellulose deposition

quickly follows to prevent "blowing out" of the hypha.














CHAPTER 1
INTRODUCTION

Apical Growth

Hyphal tip growth is a hallmark of the fungi, even

though it also occurs in specialized plant cells (i.e.,

growth of pollen tube, root hair, moss protonema).

Diverse animal cells share this capacity to protrude

their cytoplasm and then move in that direction, a

process termed ameboid movement. The essential feature of

tip growth is that the tip of the hypha is protruded into

the environment from the subapical region. This protrusion

involves the synthesis and extension of the cell wall and

cytoplasm (with its contained organelles). The organism

is thus able to explore and exploit its environment.

Although apical growth is a widespread process in the

biological world and has been known for over a hundred

years, the mechanisms underlying this process are not yet

understood. Knowledge of these mechanisms would allow an

understanding of other related characteristics of fungi,

such as the influence of environmental factors on growth

and morphogenesis and the interaction between fungi and

other organisms. Ultimately, detailed knowledge of hyphal

tip growth would allow the development of techniques for








inhibiting or stimulating growth of medically or

economically important species.

Studying hyphal tip growth is a complex problem

because the apex represents only a tiny part of a hypha.

Most of the important growth events occur within 5

micrometers of the tip. On the other hand, the mature

part of the hypha is not inactive. In growing hyphae, the

wall synthesis per unit area is maximal at the tip. The

total amount of wall material synthesized subapically at

the same time is appreciable (Sietsma et al 1985). This

also contributes to the difficulty of studying tip

growth. Finally, not all of the hyphae in an actively

growing colony are growing (apically elongating) at a

given moment in time. Therefore, conventional

biochemical, autoradiographical and cytological

techniques must be adapted to the specificity of the

problem.

In this study I approached the problem of hyphal tip

growth by comparing cell wall architecture in elongating

versus non-elongating hyphal apices of an oomycete Achlya

bisexualis. Electron microscopy, both transmission and

scanning, was used with a variety of immunocytochemical

labeling of hyphae. Several fixation techniques were used

to ensure that the results were not only reproducible but

also not artifacts of the fixation procedure. The results

allowed me to propose a new hypothesis for the mechanism

of hyphal tip growth.














The Organism

Members of the genus Achlya grow as branched

coenocytic hyphae, which collectively are termed a

mycelium. Septa are formed only to delimit reproductive

structures, while vegetative growth occurs at the apex.

Achlya has both asexual and sexual cycles of

reproduction. Asexual reproduction occurs by

fragmentation, differentiation of resistant gemmae, or by

the differentiation of vegetative apices into sporangia

(Sparrow 1960). Achlya differs from related genera by the

fact that the primary zoospores immediately encyst in a

loose cluster at the orifice of the sporangium after

discharge (Johnson 1956).

Sexual reproduction occurs by gametangial contact.

The male gametes produced in an antheridium are

transported via fertilization tubes to female gametes

produced in an oogonium (Mullins 1994). Sexual

morphogenesis is initiated and sequentially controlled by

a series of diffusible steroid hormones (Raper 1939).

While most water molds are monoecious, bearing both male

and female reproductive structures on a single diploid

mycelium, some members of the genus Achlya are dioecious.

True "male" and "female" strains of dioecious species of

Achlya may exist, but the expression of mating type in a







strain depends on that of its mating partner (Raper

1939). The involvement of hormones in sexual reproduction

in this genus is very noteworthy, as species of Achlya

appear to be the most primitive eukaryotes known to

produce and respond to steroids.

Achlya has been proposed as a eukaryotic model

system for studying basic mechanisms of growth and

development. Species of Achlya have been used to

investigate the regulatory mechanisms of

steroid-hormone-induced and regulated sexual

differentiation (Thomas and Mullins 1967, Mullins and

Ellis 1974, Horgen 1977, Riehl and Toft 1984, Mullins

1994). They also served in studies on: (i) the

differentiation of vegetative hyphae into asexual

sporangia (Griffin and Breuker 1969, Thomas et al. 1974,

LeJohn et al. 1977, Kropf et al. 1983, Cottingham and

Mullins 1985); (ii) the mechanism of nutrient transport

in fungi (Cameron and LeJohn 1972, Manavathu and Thomas

1982, Kropf et al. 1984); (iii) the tropic responses to

nutrients and other chemoattractants (Musgrave et al

1977, Manavathu and Thomas 1985); (iv) the role of turgor

in hyphal tip growth (Money and Harold 1992, 1993); and

(v) ionic and electrical currents (Harold 1994). In this

study, I used Achlya bisexualis Coker and A. Couch (ATCC

accession number 14524) to investigate the mechanisms of

hyphal tip growth.





5


Class Oomycetes

The genus Achlya is classified in the family

Saprolegniaceae, order Saprolegniales, class Oomycetes,

subdivision Mastigomycotina, division Eumycota of the

kingdom Fungi (Carlile and Watkinson 1994). The

subdivision Mastigomycotina contains organisms that

produce motile spores zoosporess). The subdivision is

divided into three classes, based on the morphology of

zoospores: Chytridiomycetes, Oomycetes, and

Hyphochytriomycetes. The first class is similar to other

Eumycota, while the latter two show similarities to some

protests rather than to fungi. In fact, the morphological

divergence of the Oomycetes has long been recognized

based on their morphology (Gaumann and Dodge 1928). Their

biochemical properties, such as L-lysine biosynthesis

(Vogel 1964), cell wall chemistry (Bartnicki-Garcia

1968), and tryptophan-pathway enzyme organization (Hutter

and DeMoss 1967) strongly support this view. More recent

ultrastructural (Beakes 1987) and molecular studies

(Lovett and Haselby 1971, Ohja et al. 1975, Kwok et al.

1986, Forster and Coffey 1990, Forster et al. 1990) also

confirmed the divergence of the Oomycetes. According to

Bartnicki-Garcia (1996), these biochemical and

morphological differences indicate that the Oomycetes and

the higher fungi probably arose from different ancestors.

However, the same author disagrees with the idea of

breaking up the kingdom Fungi based on these phylogenetic








considerations. In the past, the classes

Chitridiyceteomycee, mycetes, and Hyphochytriomycetes

often have been grouped with nonfungal organisms with

which they have very little in common, either on a

physiological, morphological, or ecological basis. For

example, the Oomycetes were lumped with heterokont algae

in the kingdom Chromista (Cavalier-Smith 1983,

Moore-Landecker 1996), or were placed with all zoosporic

fungi, protozoa and algae in kingdom Protoctista

(Margulis et al. 1990). An admittedly polyphyletic

kingdom Fungi is a more rational taxonomical solution

than the ones listed above. This solution allows us to

assemble and study the collection of organisms that share

key morphological, physiological and ecological

properties (Bartnicki-Garcia 1996).

Though my work does not concern systematics, an

understanding of the phylogenetic position of Achlya is

relevant to the problem of hyphal tip growth. Because the

Oomycetes could have evolved independently, their

mechanism of hyphal tip growth, despite its superficial

similarity to one of true fungi, could prove to be

different.

There are about 600 species of the Oomycetes. The

sexual phase of the Oomycetes has a clear differentiation

into large female and small male structures, termed

oogonia and antheridia. These are the sites of meiosis

and gametogenesis. Each oospore produced after








fertilization has a single diploid nucleus. When the

oospore germinates, it gives rise to a diploid vegetative

mycelium, in contrast to the haploid mycelium of most

fungi. Other characters of the Oomycetes that distinguish

them from the Eumycota are the biflagellate zoospore;

mitochondria with tubular cristae; Golgi bodies

consisting of multiple flattened cisternae; cellulose as

a microfibrillar component of the cell wall; the presence

of the amino acid hydroxyproline in cell wall

glycoproteins; and various other biochemical and

molecular characteristics (Carlile and Watkinson 1994).

True fungi have mitochondria with platelike cristae and

produce Golgi bodies that are very simple in structure,

often consisting of only a single cisternal element. Cell

walls of true fungi have chitin as the microfibrillar

component and do not contain hydroxyproline (Alexopoulos

et al. 1996).

The class Oomycetes consists of 5 orders:

Saprolegniales, Lagenidiales, Peronosporales, Rhipidiales

and Leptomitales (Alexopoulos et al. 1996). The order

Saprolegniales contains a single family Saprolegniaceae.

Usually these fungi occur in fresh water and in soil as

saprotrophs and play an important role in decomposition

and recycling of materials in aquatic ecosystems. Some,

however, are obligate parasites of plants, animals, or

other fungi. For example, some species of Saprolegnia,

Achlya, and Aphanomyces attack fish and their eggs





8


(Alexopoulos et al. 1996). The members of Saprolegniaceae

are often called water molds, are distributed

universally, and are among the easiest fungi to isolate

and cultivate in the laboratory.















CHAPTER 2
LITERATURE REVIEW

Mechanism of Apical Growth

The phenomenon of hyphal tip growth has been known

for over a hundred years (Reinhardt 1892). Its mechanism,

though, is not yet understood. Several theories of hyphal

tip growth dominate the literature. They are (1) the

delicate balance theory (Park and Robinson 1966,

Bartnicki-Garcia 1973); (2) the steady-state theory

(Wessels 1990); and more recently a combination of the

first two, (3) the hybrid theory (Johnson 1996). All

three imply that the wall of the apex is plastic, while

that of the subapical nongrowing area is rigid. They also

assume that the driving force for cell elongation is

turgor pressure and/or cytoskeleton.

The delicate balance theory assumes that the

plasticity of the hyphal apex is achieved by a constant

delicate balance between biosynthesis and hydrolysis of

wall components.

The steady-state theory suggests that the plastic

region at the tip contains a mixture of nonlinked wall

polymers that are being constantly synthesized, and the

rigid condition of the wall is established by chemical

crosslinking that is initiated at or near the tip and








continues progressively further back in the hyphal wall.

Presoftening of the apical wall is catalyzed by endolytic

enzymes that briefly initiate growth but do not sustain

it.

The hybrid model retains from the steady-state model

the constant exocytosis of a plastic mixture of wall

polymers at the tip and its rigidification via

crosslinking. Among the concepts retained from the

delicate balance model is continuous endoglycanolytic

activity expressed in proportion to the rate of tip

extension (Johnson 1996).

Thus these models suggest different mechanisms to

explain the events of wall growth, while agreeing on

other aspects such as the role of turgor and the

cytoskeleton.

Structure of the Hyphal Wall

Fungal cell walls have essential roles in the life

of the fungal cell, i.e., maintenance of cell shape,

plasticity, protection against unfavorable environmental

conditions, cellular recognition, immune response, and

host-parasite interaction (Rosenberger 1976, Wessels and

Sietsma 1979). The general organization of hyphal cell

walls comprises an inner layer of microfibrillar

polysaccharides overlaid by an outer layer of amorphous

polysaccharides (Burnett 1979).

In Oomycetes, these polysaccharides are,

respectively, cellulose and 1,3-3-glucans containing some








1,6-B branches. Cellulose usually represents about 20%

(w/w), 1,3-B-glucans about 80% (w/w) of the total wall

carbohydrates (Sietsma 1969, Burnett 1979).

In the hyphal wall of A. ambisexualis Raper

(Reiskind and Mullins 1981a), acid-soluble 1,3-B-glucans

with single 1,6-8-linked residues as branches represents

40% (w/w) of the dry wall. An alkali-soluble glucan, a

polymer of 1,3-B and 1,4-B linkages with occasional 1,6-B

glucosyl residues as side chains, represents 7% (w/w) of

the wall. Cellulose represents 21% (w/w) of the wall. An

insoluble residuum with a linkage pattern similar to the

alkali-soluble fraction is present at 6% (w/w). An

insoluble component consisting of glucosamine represents

3% (w/w) of the wall. This insoluble fraction probably

represents chitin (Mullins et al 1984). Protein

containing hydroxyproline residue comprises 10% (w/w).

There is also a small amount of phosphorus.

In the study on the ultrastructural organization of

the hyphal wall of A. ambisexualis (Reiskind and Mullins

1981b) a model of the various layers in the wall was

proposed. The method used in this study of the hyphal

wall consisted of the sequential chemical or enzymatic

removal of the various fractions, followed by analysis

(with electron microscopy) of carbon-platinum replicas.

The model shows (a) a surface layer of amorphous

1,3-B-glucan hydrolyzed by acid or the enzyme

laminarinase; (b) another 1,3-B-glucan layer containing








some 1,4-B and 1,6-B linkages hydrolyzed by alkali or

laminarinase; (c) microfibrillar cellulose, removed by

cadoxen or the enzymes cellulase plus protease; and (d)

an innermost layer of insoluble residuum, faintly

microfibrillar.

The most abundant and most thoroughly studied

glucans from the fungal cell walls are B-glucans. These

1,3-B-glucans are variable in degree of 1,6-B branching

and in the length of the branches.

Cellulose is a linear polysaccharide made of

glucosyl moieties joined through 1,4-B linkages. The

glucan chains in this polysaccharide associate through

hydrogen bonding to form microfibrils. According to chain

orientation, different crystalline structures exist. The

most prevalent form is Cellulose I, where glucose chains

are arranged in parallel fashion (the free reducing

groups are in the same end of the microfibrils, and the

nonreducing ends are in the opposite one). In this sense,

as demonstrated by X-ray diffraction analysis (Reiskind

and Mullins 1981a), and also apparently in size, fungal

cellulose is similar to the polysaccharide found in

plants (Ruiz-Herrera 1991).

Chitin is an unbranched polysaccharide containing

exclusively N-acetylglucosamine residues linked 1,4-B.

Three crystalline isoforms of the polysaccharide exist in

nature, according to the arrangement of the chains. These

forms can be recognized by X-ray diffraction. In fungi,








only alpha-chitin, characterized by the antiparallel

arrangement of the chains, has been detected (Sentandreu

et al. 1994).

Structural proteins present in the cell wall of

fungi are glycoproteins. They display a basic common

structure, consisting of protein with covalently bound

carbohydrate chains. In fungi, they are usually called

mannoproteins because the carbohydrate moiety mainly

consists of mannose units, although small amounts of

other sugars and phosphodiester groups are found (Peberdy

1990, Ruiz-Herera 1991). Hydroxyproline is reported as a

constituent of cell wall proteins in the Oomycetes

(Webster 1980, Reiskind and Mullins 1981a, Ruiz-Herrera

1991). The carbohydrate moieties are attached to the

protein through two types of linkages. One type is

O-glycosidic linkage between mannose or small

oligosaccharide chains and the hydroxy-amino-acids

(Nakajima and Ballou 1974, Sentadreu and Northcote 1969,

Tanner and Lehle 1987). The second type of linkage

(N-glycosidic) connects high molecular weight, highly

branched, mannan tufts to asparagine residues of the

protein, through diacetylchitobiose (Byrd et al. 1982,

Cohen and Ballou 1981, Tanner and Lehle 1987).

Studies of the structure of fungal cell walls by

cast-shadowing or replica techniques have demonstrated

that their outer and inner surfaces appear different. The

outer surface is usually amorphous or finely granular,








whereas the inner face shows intertwining microfibrils of

different size, width and orientation. However, there is

evidence that some components such as microfibrils may

escape observation because they are masked by the

presence of the large amounts of matrix compounds. From a

structural point of view, the fungal cell wall has been

compared to such manmade composites as reinforced

concrete or fiber-reinforced plastics which are formed by

two distinct elements: an elastic one, which in the cell

wall would be microfibrils of the structural

polysaccharides, and a plastic one, which would

correspond to the rest of the wall components, generally

referred as amorphous or cementing (Ruiz-Herrera 1991).

Electron Microscopic Studies of Fungal Cell Walls

In thin sections, fixed and stained by the usual

standard method including glutaraldehyde and osmium

tetroxide, fungal cell walls appear multilayered. At

least two layers are observed in most walls: an outer one

which is electron dense; and an inner layer, thicker and

electron transparent. However, appearance of the cell

wall in sections may depend on the technique used for

fixation (Ruiz-Herrera 1991).

Variability in composition of the cell wall of fungi

does not allow the proposal of a single model of the wall

structure. In general, evidence suggests that fibrillar

polysaccharides are accumulated mostly in the inner

layers of the cell walls, while amorphous components are








more abundant in the external layers. The description of

wall structure observed in different genera of fungi

analyzed by various techniques may be more useful in

providing a general overview of fungal wall architecture

(Ruiz-Herrera 1991).

Early studies on the chemical characterization of

fungal cell wall layers were conducted by Hunsley and

Burnett (1970). They studied the wall structure of

Neurospora crassa, Schizophyllum commune and Phytophtora

parasitica after treatment with several hydrolytic

enzymes. The outer surface of N. crassa in shadow-cast

samples appeared amorphous. Laminarinase treatment

removed the amorphous coat revealing a layer of coarse

strands whose interstices were filled with amorphous

material, whereas treatment with both laminarinase and

pronase enhanced the reticular appearance. The

microfibrils were sensitive to chitinase. The authors

concluded that the external coat was made of amorphous

beta-glucans placed over a reticulum of glycoproteins.

More internally, it was suggested, a protein layer

followed in which chitin microfibrils were embedded. In

contrast to Neurospora, the cell wall of S. commune was

resistant to laminarinase, pronase and chitinase,

apparently due to the presence of superficially located

1,3-alpha-glucan which prevented the access of the lytic

enzymes. After removal of this glucan layer by KOH,

laminarinase and pronase treatment gave rise to the







appearance of a fibrillar structure sensitive to

chitinase, suggesting that inner wall layers had a

chemical composition and organization similar to N.

crassa. Appearance of the wall from P. parasitica was not

affected by pronase, but laminarinase unmasked a

fibrillar layer sensitive to cellulase treatment. These

results were interpreted as suggesting the presence of

two layers rich in amorphous beta-glucans and cellulose,

respectively, in the wall of this fungus.

The cell of yeast and mycelial cells of Candida

albicans reveals four wall layers when treated by a

standard gluteraldehyde-osmium technique (Yamaguchi

1974). When stained by Thiery's technique, eight

different layers can be observed, depending on the

intensity of staining and their electron density. The

four outermost layers are PATAg positive, whereas layers

5 and 7 appear electron transparent and PATAg negative

(Poulain et al. 1978). The authors concluded that the

inner layers must be rich in chitin and 1,3-B-glucan,

which are both electron transparent and PATAg negative.

Other outer layers must be rich in glucans and mannans.

The existence of mannans on the surface of the cell was

confirmed by Horisberger et al. (1975) who observed

binding of colloidal gold-tagged concanavalin A (ConA-Au)

by intact cells of the fungus. The presence of mannan in

two continuous layers at the periphery of blastospores

was demonstrated by staining ultrathin sections with








Concanavalin A-horseradish peroxidase-3,3'diamino

benzidine and HO (Tronchin et al. 1979). In this

technique, the lectin binds to the mannose residues of

the glycoprotein and it is recognized by peroxidase. The

peroxidase forms a dark product by the catalytic

decomposition of HO in the presence of an oxygen

acceptor. A similar method, which included treatment with

wheat germ lectin followed by chitibiosyl-horse radish

peroxidase or chitobiosyl-ferritin, was used to conclude

that chitin was located mostly in the inner layers of the

wall of C. albicans.

In related species Candida utilis, sections were

stained with ConA-Au and gold-labeled antimannan

antibodies. These techniques demonstrated that

mannoproteins were denser in the cell periphery although

labeling also was observed close to the plasmalemma

(Horisberger and Vonlanthen 1977). Similar results were

obtained with Saccharomyces cerevisiae by the same

authors (Horisberger and Vonlanthen 1977).

Lectins bound to fluorescein isothiocyanate (FITC)

were used to detect superficial polysaccharides in the

different yeasts by Barkai-Golan and Sharon (1978). The

authors observed that S. cerevisiae, S. bayanus and

Candida mycodema bound ConA only, suggesting the presence

of mannoproteins on the surface of the cells. On the

other hand, Schizosaccharomyces pombe did not bind ConA;

but it bound peanut lectin, which recognizes D-galactose,








indicating that the cell surface of fission yeast is

covered by a galactomannan, not by mannoproteins. Candida

rugosa and Sporobolomyces roseus bound both lectins. This

result may indicate the presence of both galactomannan

and mannoproteins on the surface of these cells.

Treatment of the cells with KOH resulted in a strong

reaction with wheat germ lectin which recognizes GlcNAc,

suggesting that chitin is located internally and is

covered by alkali-soluble mannoproteins. The presence of

galactomannan on the surface of S. pombe was confirmed by

use of the lectin from Bandeiraea simplicifolia bound to

colloidal gold (Horisberger and Rosset 1977). This

lectin, which recognized alpha-galactopyranosyl residues

bound to the outer layer of the wall, and in minor

amounts was distributed evenly over the whole thickness

of the cell, including the fission scars. In a further

report, these authors demonstrated differential

distribution of galactomannan depending on the growth

stage of the cells (Horisberger et al. 1978).

Galactomannan appeared in the form of two layers of the

wall: one close to the plasmalemma, and another on the

surface of the cell. Labeling by the lectin occurred at

the cell periphery and at the growing end, but not on the

wall, formed after cell division. These results were

interpreted as meaning that the polysaccharide was

synthesized during cell extension, but not during septum

formation.








Four layers in the cell wall of Dictyostelium

discoideum spores were observed by freeze-etching and

replica (Hemmes et al. 1972). The innermost layer, which

appeared amorphous or slightly fibrillar, could be

eliminated by successive treatment with cellulase and

pronase, suggesting that it was constituted by a mixture

of cellulose and proteins. The middle layers (both

fibrillar) were removed by cellulase treatment alone,

indicating the cellulosic nature of the microfibrils. The

most superficial layer was resistant to both pronase and

cellulase treatment. Hydrolysis resulted in release of

galactose, suggesting that this is a major component of

the acidic polysaccharide present in the walls.

In sections of Agaricus bisporus spores treated with

the standard method, three layers could be recognized.

The authors concluded that the middle layer contained

protein because treatment with pronase increased the

fibrillar appearance of this layer. These fibrils

corresponded to 1,3-B-glucans and chitin, as they were

removed by B-glucanase and chitinase treatment. The outer

layer was composed of melanins and 1,3-alpha-glucans,

which was deduced by chemical analyses and electron

microscopic observations. The thin inner layer was poorly

characterized, but the authors suggested it was of

mucilagenous nature (Rast and Hollenstain 1977).

The structure of the mycelial wall of the same

fungus was different (Michalenko et al. 1976). The outer








layer, which appeared amorphous in replicas, was made of

mucilage. The thin middle layer was made of amorphous

glucans. The innermost layer, which in replicas appeared

fibrillar, is probably made of a mixture of B-glucans

covering fibrillar chitin, since chitinase by itself

could not remove it, whereas the combined action of

B-glucanase and chitinase solubilized the layer. Staining

with silver examine suggested that proteins were present

in all layers of the cell wall of the fungus.

A similar approach was followed in the

characterization of the architecture of the wall from

microconidia of Trichophyton mentagrophytes. Three layers

were recognized in sections. The outer layer appeared

electron dense, and the innermost one appeared electron

transparent. The material extracted from the outer layer

contained a single glycoprotein. The median layer was

made of proteinaceous rodlets. The inner layer apparently

was composed of amorphous glucans and microfibrillar

chitin (Wu-Yuan and Hashimoto 1977).

Structure of the cell wall from Trichoderma

pseudokoningii was studied by treatment of intact cells

with different lytic enzymes (Jeenah et al. 1982).

Accordingly, the authors concluded that the outer layer

contained 3-glucans, whereas the internal layer was

composed of chitin embedded in a protein matrix.








Biogenesis of the Fungal Cell Wall

Cell wall biosynthesis takes place in three sites:

cytoplasm, plasma membrane and the wall itself.

Structural polymers such as chitin and 1,3-B- and

1,4-B-linked glucans are synthesized vectorially at the

plasma membrane, by transmembrane synthases accepting

nucleotide sugar precursors from the cytosol and

extruding the polymerized chain into the wall (Cabib et

al. 1983, Shematek et al. 1980, Girard and Fevre 1984,

Jabri et al. 1991, Cabib et al. 1991, Hromova et al.

1989). Matrix polymers such as glycoproteins are

synthesized in the cytoplasmic secretary pathway of

endoplasmic reticulum through Golgi vesicles to secretary

vesicles. Wall assembly, involving activities such as

covalent crosslinking of polymers and modifications such

as deacetylation of chitin, takes place in the wall

itself (Gooday 1995).

Fungal wall 1,3-B-glucans are biosynthesized via the

nucleotide sugar, UDP-glucose. The glucan synthases are

intrisic proteins of the plasma membrane. Preparations of

membranes from Saprolegnia monoica, when provided with

UDP-glucose, produce polymers containing varying amounts

of 1,4-B and 1,3-B links (Girard and Fevre 1984). The

vectorial synthesis of 1,3-B-glucan chains allows only

linear molecules to be made and thus any 1,6-B branches

must be added in the wall (Gooday 1995). These








1,3-B-glucan synthases are stimulated by the presence of

trypsin but inhibited by other proteases. Stimulation

occurs from the beginning of incubation in the presence

of the protease but prolonged action of trypsin leads to

inactivation of the glycosyl transferases. The

1,3-B-glucan synthases, therefore, must exist in an

inactive state that can be activated by moderate

proteolysis. Such regulation, which also appears to

modulate plant glycosyl transferases (Girard and

Maclachlan 1987), characterizes the chitin synthase

system of various fungi (Cabib 1981).

The 1,4-B-glucan synthases, like 1,3-B-glucan

synthases, have a transmembrane orientation in the

plasmalemma, leading to a vectorial synthesis of

cellulose from UDP-glucose. These enzymes may have a

common structure or organization, as revealed by

preliminary immunological studies, but they are different

systems and can be separated by glycerol-gradient

centrifugation (Fevre et al 1990). Cellulose synthases

from Saprolegnia are inactivated by trypsin, but

stimulated in the presence of certain nucleotides. Fungal

cellulose synthase enzymatic complex may resemble the

plant plasma membrane rosettes involved in cellulose

synthesis (Mullins and Ellis 1974, Muller and Brown 1980,

Montezinos 1982). Some proteins, sensitive to proteases

or capable of reacting with nucleotides, would be

involved in the regulatory processes. Other proteins








would be involved in UDP-glucose binding. Such an enzyme

seems to exist in plants (Delmer 1999). Cellulose

synthases may have a more complicated organization than

1,3-B-glucan synthases. Cellulose synthase activities of

cell free extracts are always much lower than

1,3-B-glucan synthase activities. It is possible that

cellulose synthases require a specific factor that is

lost in the course of isolation. The opposite behavior of

the synthases towards protease and nucleotides, and the

presence of a membrane bound activator of 1,3-B-glucan

synthase, may indicate a difference in the regulation of

their activities. This would have implications in the

cell wall assembly where the deposition of the different

polysaccharides during apical growth is coordinated in

time and space (Fevre et al 1990).

Chitin synthases, like glucan synthases, are

intristic proteins of plasma membrane. These enzymes

catalyze glycosidic bond formation from the nucleotide

sugar substrate, UDP-N-acetylglucosamine. Most chitin

synthase preparations are zymogenic, i.e., produced as

proenzymes requiring activation by specific proteases.

This proteolytic activation presumably plays a role in

the temporal and spatial regulation of the enzyme, by

locally activating it in the membrane when and where its

activity is required (Gooday 1995). As well as being in

zymogenic and active forms in the plasma membrane,

zymogenic chitin synthase also occurs in fungal cells as








chitosomes, which are membrane-bound microvesicles about

70 nm in diameter (Bartnicki-Garcia et al 1979, Kamada et

al 1991). After purification by differential

centrifugation, chitosomes can be activated by treatment

with proteolytic enzymes, and then produce chitin

microfibrils if incubated with UDP-GlcNAc (Gooday 1995).

Wall glycoproteins are biosynthesized in the

secretary pathway: endoplasmic reticulum > Golgi bodies >

secretary vesicles > release at the plasma membrane.

Carbohydrate material detected in apical vesicles could

be the carbohydrate portion of glycoproteins. The

transmembrane stages in glycoproteins biosynthesis

involve sugar precursors linked to polyprenol dolichol,

the "lipid intermediates" (Lehle 1981, Cabib et al.

1988). In the O-linked chains, the first mannose unit is

linked to the protein via the precursor

dolichol-phosphomannose, in the endoplasmic reticulum.

The other mannose units are added via the nucleotide

sugar guanosine diphosphomannose, GDP-Man, in the Golgi

bodies. The N-linked chains are assembled by a more

complex scheme, giving a lipid intermediate

dolichol-diphospho-(GlnNAc)2-Man9-Glc3 which is N-linked

to asparagine in the protein in the endoplasmic

reticulum, with the release of the terminal four sugar

units, Man-Glc3. The outer chain of many mannose units is

added by several linkage-specific mannosyl transferases,

with Glc-Man as substrate, in the Golgi bodies (Gooday








1995). Some secreted enzymes, notably invertase, acid

phophatase and chitinase, are also mannoproteins,

synthesized and secreted in a similar fashion (Kuranda

and Robbins 1991).

Once the cell wall components are synthesized and

secreted, they must be converted into an integrated

structure. This process includes covalent crosslinking,

hydrogen bonding, hydrophobic and electrostatic

interactions between different macromolecules

(Ruiz-Herrere 1991).

Role of Turgor in Wall Expansion

The difference in hydrostatic pressure between a

cell and its surroundings is called turgor pressure. This

actual pressure is thought to provide the driving force

for hyphal extension. Several observations and

measurements suggest that it is necessary for the apical

growth process (Robertson 1958, Park and Robinson 1966,

Robertson and Rizvi 1968). Osmometry has been used to

demonstrate a correlation between hyphal extension rates

and turgor pressure in many fungal species (Eamus and

Jennings 1986, Luard and Griffin 1981, Woods and Duniway

1986). Experiments have shown that filamentous fungi

respond to increases in external osmotic pressure by

accumulating compatible solutes, including potassium

ions, glycerol, mannitol, erythritol and arabitol (Luard

1982a,b,c; Pfyffer and Rast 1988, 1989).








The most detailed analyses of the relationship

between hyphal extension and turgor pressure have been

carried out on hyphae of Achlya bisexualis and

Saprolegnia ferax and these studies suggest that growth

can occur without significant turgor (Money and Harold

1992, Kaminskyj et al. 1992). The rate of growth under

these conditions is about half of the maximum rate.

Achlya continues to grow even after the turgor is

undetectable; however, its morphology is radically

altered. On solid medium it shows plasmodial-like growth.

In liquid medium of the same composition, it exhibited a

yeast-like morphology. Saprolegnia has a different

response to the absence of turgor, since it continues to

grow in the hyphal form. Both Achlya and Saprolegnia

appear not to respond to changes in external osmotic

pressure by controlling the concentration of internal

compatible solutes (regulation of turgor); instead, the

plasticity of the wall is modulated to balance the force

applied against it.

Role of Cvtoskeleton in Hvphal Growth

It seems very unlikely that the thin wall covering

the apices of extending cells has sufficient mechanical

strength to contain the turgor pressure of the cytoplasm.

It was suggested that other cellular components may play

a role in the regulation of tip expansion. The possible

existence of other factors is suggested by: (1) the

ability of mutants with abnormal cell wall composition to





27


generate relatively normal hyphae (Katz and Rosenberger

1970); (2) the poor correlation between growth rates and

turgor pressure (Kaminskyj et al. 1992); and (3) the

ability of some species to produce hyphae in the absence

of measurable turgor pressure (Money and Harold 1993).

The apex might be stabilized by a fibrillar

cytoplasmic network similar to that found in other

cellular systems (amoebae, slime molds). The main

structural components of such a cytoskeleton are actin

filaments (F-actin), microtubules, and intermediate

filaments. Each of these are elongated polymers composed

primarily of globular proteins known as actions, tubulins,

and other unrelated units respectively (Heath 1994). The

presence of an array of F-actin was shown in growing tips

of Saprolegnia (Heath 1987). It was always present in

growing tips, but absent in nongrowing tips. None of

these observations proves a morphogenic role for F-actin,

because it is not possible to differentiate between

direct and indirect effects in the complex system of

hyphal tips, but they do suggest that F-actin has some

role (Heath 1994).

Another explanation of the presence of actin plaques

at the growing apices is vesicle and organelle traffic

control. There is evidence for the involvement of both

microtubules and F-actin in wall vesicle transport (Heath

1994).








Cytology of Growing Hvphal Apices

A growing hypha consists of an apical region where

the extension takes place, a nonelongating subapical, and

mature regions, which were the sites of earlier growth.

This is reflected both in the structure of the wall and

the cytoplasm.

Older regions of the hyphal wall are rigid and

thick. The cytoplasm in these regions is restricted to a

thin layer between a large tonoplast and the plasma

membrane. This cytoplasm contains the usual variety of

eukaryotic organelles. The tonoplast in mature regions is

represented by a central vacuole whereas younger regions

contain many vacuoles of smaller size. The subapical

region has a thinner wall. The cytoplasm is nonvacuolated

and is particularly rich in organelles. At the very apex,

the hyphal wall is thin and the associated cytoplasm

lacks the usual organelles, containing only small

cytoplasmic vesicles of differing size (Shapiro 1995).

Based on the organelle distribution, three cytoplasmic

zones or regions are recognized: (1) an older, highly

vacuolated region; (2) a subapical organelle-rich region;

and (3) a terminal vesiculate region (Grove et al 1970).

Since one of the differences between the growing tip

region and the basal parts of the hypha is the abundance

of cytoplasmic vesicles, they are usually assumed to be

involved in the synthesis of the new wall. One

possibility would be that they carry wall polymers ready








for insertion in the growing wall. Intracellular

synthesis of wall polymers and their delivery to the wall

by vesicles occurs with pectin, hemicellulose, and

hydroxyproline-rich glycoproteins in plants (Northcote

1984), and to wall mannoproteins in yeast (Zlotnik et al.

1984). For filamentous fungi, there is no convincing

evidence for a similar process. Cytochemical staining

does detect polysaccharide material in some apical

vesicles (Grove 1978, Hill and Mullins 1980), but this

material may represent glycoproteins destined for

secretion. More likely, these vesicles contain precursors

of the cell wall, their membrane probably contributes to

the extending plasmalemma, and they may contain wall

synthase enzymes for insertion into the plasma membrane

(Heath 1994).

Growing and nongrowing hyphae differ in the type of

wall material that covers their apices (Wessels 1986).

The absence of alkali-insoluble beta-glucans at the very

apex of growing hyphae in Schizophyllum commune has been

demonstrated by light microscopic autoradiography

(Wessels et al. 1983). A subsequent study using electron

microscopic autoradiography on shadowed preparations

revealed that chitin in growing apices, though alkali

insoluble, is in a conformation state quite different

from that in nongrowing apices and subapical parts. In

contrast to the chitin in these older parts, the newly

synthesized chitin at apices appeared nonfibrillar, very








susceptible to chitinase degradation and partly soluble

in hot dilute mineral acid. Earlier observations had

indicated discontinuities in the presence of microfibrils

at hyphal apices (Strunk 1968). These have been

contradicted by other workers who showed a continuous

network of chitin microfibrils over the apex after

chemical treatments which removed a "matrix substance"

(Aronson and Preston 1960, Hunsley and Burnett 1970,

Bartnicki-Garcia 1973, Schneider and Wardrop 1979,

Burnett 1979, Aronson 1981). Wessels (1990), however,

suggested that these images showing apical microfibrils

probably represent nongrowing apices, which are known to

occur abundantly among growing hyphae.

There is a number of light microscopic studies using

fluorescently labeled probes which also suggest that the

wall covering the growing apex is different from that

covering a nongrowing apex or that of subapical regions.

In these studies fluorescently labeled antibodies (Fultz

and Sussman 1966, Marchant and Smith 1968, Hunsley and

Kay 1976), fluorescent brighteners such as calcofluor

(Gull and Trinci 1974), and fluorescently labeled wheat

germ agglutinin were used. This differential staining at

growing tips could result from the absence of outer wall

materials, or to a difference in the conformation of the

polymers that bind these probes.








Calcium Gradient

There is a tip-high calcium gradient in apically

growing cells. Free cytoplasmic calcium in the oomycete

Saprolegnia ferax is highest at the tip as demonstrated

using fluorescent dyes such as Indo-1 or Fluo-3 (Yuan and

Heath 1991, Jackson and Heath 1993, Garrill et al. 1993).

Studies using patch-clamp techniques suggest that the

tip-high gradient reflects a spatial organization of

calcium channels in the cell membrane. Using patch-clamp

electrophysiology, two types of channels were identified

in Saprolegnia ferax: (a) calcium-activated potassium

channels that were thought to be involved in turgor

regulation, but were not obligatory for growth; and (b)

stretch-activated calcium channels that were activated by

potassium ions and which may be essential for apical

extension (Garrill et al. 1992, 1993). The

stretch-activated channels were concentrated at the

hyphal apex and were blocked by Gd3 which also inhibited

hyphal extension and dissipated the tip-high calcium

gradient revealed by Indo-1 (Garrill et al. 1993). In

contrast to the stretch-activated channels, the

calcium-activated potassium channels were uniformly

distributed along the hyphal cell membrane. These could

be inhibited by tetraethylammonium, which only caused a

transient effect on growth. Stretch-activated calcium

channels have also been identified in the germ tubes

apices of the plant pathogen Uromyces appendiculatus








(Hoch et al. 1987, Zhou et al. 1991). These data suggest

that the tip-high calcium gradient is important for

polarized hyphal extension and is generated by a locally

high concentration of stretch-activated calcium channels

in the hyphal apex. It is presumed that the channels are

delivered to the surface in microvesicles. They may be

maintained there by anchoring them to the cytoskeleton or

by membrane recycling (Gow 1995).

Ion Currents

The net flow of electrical current carried by the

circulating ions can be detected with an ultrasensitive

voltmeter called the vibrating microelectrode (Jaffe and

Nuccitelli 1974). In Achlya bisexualis and filamentous

fungi in general, a positive proton-carried current

normally enters the growing apex. Inward current was

shown to be due to amino acid-proton co-transport

(symport) localized at the tip and the outward current

was due to electrogenic proton efflux via a plasma

membrane ATPase (Kropf et al. 1984, Gow et al. 1984, Gow

1984, Schreurs and Harold 1988). The proton current also

established an extracellular pH gradient around the

hypha, with the medium adjacent to the tip relatively

alkaline (Gow 1984). On the other hand, there are many

examples where there is no correlation between the

direction or magnitude of the current and the process of

tip growth (Gow 1995).













CHAPTER 3
HYPHAL GROWTH

Introduction

Until recently, hyphae were assumed to extend at a

constant linear rate when environmental factors are

favorable and stable, and nutrients are ample. Detailed

analysis of hyphal growth, however, reveals oscillating

elongation rates (Lopez-Franco et al. 1994).

Materials and Methods

An isolate of Achlya bisexualis Coker & A. Couch

(ATCC 14524) was used in this study. Stock cultures were

maintained on corn meal agar. Mycelia were grown on corn

meal agar (CMA), prepared from 17 g of Difco corn meal

agar, 10 g of purified grade agar (Fisher Scientific) and

1 L of distilled water. Small plugs (approximately 1

millimeter) were removed from the edge of a 24-hour-old

CMA colony. The plugs were then placed in a 250 mL flask

containing 100 mL of liquid peptone-yeast extract-glucose

(PYG) medium, pH 6.8 (Cantino and Horenstein 1953). The

PYG medium was prepared by combining 1.25 g of

bacto-peptone, 1.25 g of yeast extract (Sigma) and 3 g of

D-glucose with 1 L of distilled water. The culture was

incubated for 10 to 12 h until the hyphae had grown out








from the agar plugs to a distance of 2 to 5 mm and then

studied.

To obtain actively growing colonies, the 10 to 20 h

old cultures incubated in PYG were used. Based on dry

weight accumulation, colony size increase, glucose

incorporation, and cellulase secretion, these colonies

are in the midexponential stage of growth (Hill and

Mullins 1979).

Non-growing conditions were obtained by transferring

growing colonies from PYG to 0.2% glucose solution, by

incubation in glucose for about 48 h to cease elongation.

Procedure made it possible to find colonies with no

elongating hyphae. Such colonies were fixed and used for

studying non-elongating hyphae. Screening for no

elongation is necessary, because there are hyphae in some

colonies that are still elongating at a slow rate.

Hyphal elongation was monitored with an Olympus BH-2

light microscope. The colonies were kept on small

depression slides with cover slips. Digital images of the

hyphal tips were taken every 10 min with a Pixera 120C

digital camera. The microscope light was turned off

between the measurements to avoid heating. Average

elongation rates were calculated using a stage

micrometer. One hundred hyphae from different growing

colonies and about 50 hyphae from non-growing colonies

were studied.








Results

These light microscopic observations suggested that

in colonies growing in PYG, the majority of the hyphae

are elongating and a small number of the hyphae is not.

When individual hyphae are monitored over a long period

of time (5 to 6 h), they go through alternating periods

of elongation and non-elongation. The rate of elongation

is not steady, but fluctuates between periods of fast and

slow rates. The average rate is 3.6 um/min, but it

fluctuates from 2 to 6 um/min. The fastest rates are in

the middle of an elongating cycle, with lower rates at

the beginning and the end, resulting in a bell shaped

curve (Fig. 1). Elongating hyphae have sharp apices

(Fig. 2).

The majority of the hyphae in the colonies incubated

in glucose-only medium are not elongating and a small

portion of the hyphae (about 5%) is elongating with an

average rate of 1 unm/min. In some colonies there are no

elongating hyphae at all. All the hyphae have rounded

apices (Fig. 3).

Discussion

Light microscopic observations suggest that hyphal

growth is a discontinuous, irregular process with periods

of elongation and no elongation (Fig. 1). The elongation

rate is not constant, but instead fluctuates with periods

of fast and slow elongation. During the elongation period

the higher rates are in the middle and the rate changes








in a bell shaped curve mode. A similar irregular mode of

hyphal tip growth was demonstrated by Lopez-Franco et al.

(1994). Growing hyphal tips were recorded with

video-enhanced phase-contrast microscopy at high

magnification, and digital images were measured at very

short time intervals (1 TO 5 s). The study was conducted

using fungi from several major taxonomic groups

(Oomycetes, Pythium aphanidermatum and Saprolegnia ferax;

Zygomycetes, Gilbeltella persicaria; Deuteromycetes,

Trichoderma viride; Ascomycetes, Neurospora crassa and

Fusarium culmorum; Basidiomycetes, Rhizoctonia solani).

In all fungi, apparent steady growth of hyphal tips

revealed patterns of pulsed hyphal elongation. It was

shown that the hyphae do not grow continuously with a

steady rate but instead this rate fluctuates, with

alternating periods of fast and slow elongation. This

results in irregular pulses of growth. Pulsed growth was

observed in fungi differing in cell diameter, overall

growth rate, taxonomic position, and presence and pattern

of Spitzenkorper organization, thus suggesting that it is

a general phenomenon. The basis of these pulses was not

determined, it was proposed that their origin could be in

the pulsating mode of intracellular processes, especially

the secretary vesicle delivery/ discharge system.










60


50 .
4



I I



E .4
0 4
o 5



101 A




0 50 100 150 200 250 300
time (min)



Fig. 1. Elongation measurements of three individual
Achlya bisexualis hyphae growing in PYG medium.

















































Figs. 2-3. Scanning electron micrographs of hyphae of
Achlya bisexualis. 2. Elongating hypha from a colony
incubated in PYG. 3. Non-growing colony incubated in
glucose-only medium showing the hyphae with rounded
apices. Bars: 2 = 16.7pm; 3 = 27.3 jim.















CHAPTER 4
LOCALIZATION OF CELLULOSE IN THE CELL WALL AS REVEALED
BY ELECTRON MICROSCOPY AND CYTOCHEMICAL TECHNIQUES

Introduction

Cellulose, a crystalline 1,4-B-glucan, is the most

abundant biopolymer in nature. Its biomass makes it a

global carbon sink and renewable energy source, and its

crystallinity provides mechanical properties to cellulose

containing cell walls (Arioli et al. 1998). The

understanding of cellulose properties and metabolism is

important for understanding morphogenesis in plants and

certain fungi.

Traditionally, cell wall components have been

identified cytochemically by using indirect, extractive

methods. This approach can lead to problems such as

incomplete extraction and unseen effects of the

extraction procedure on cell wall ultrastructure.

Enzyme-linked colloidal gold labeling is a

nondestructive, direct labeling technique and can be used

to localize cellulose on thin sections (Berg et al.

1988). This labeling technique can be also used to

localize cellulose on the surface of cells.

Previously cellulose was identified in Achlya cell

walls with cellulase-gold affinity labeling. Thin








sections of growing hyphae revealed labeling for

cellulose in mature and subapical regions, but not at the

apex (Shapiro 1995).

The present study was done to observe the

distribution of cellulose along the elongating and

non-elonging hyphae, as part of an overall examination of

hyphal tip growth.

Materials and Methods

Culture Methods and Microscopy Techniques

The general culture methods are described in Chapter

3. To induce sporulation, ten discs from the edge of a 48

h old colony growing on CMA plates were punched out with

a small cork border. The discs were incubated at room

temperature in 100 mL PYG liquid medium in a 250 mL flask

for 15 h with shaking at 110 rpm. Then they were washed

several times with 0.5 mM calcium chloride. At ths point,

they were left in fresh calcium chloride for 6 to 7 h to

induce sporulation.

Electron microscopy

For chemical fixation, the agar plugs bearing hyphae

were fixed for 30 min at room temperature with 4% (v/v)

glutaraldehyde in 0.05 M sodium cacodylate buffer, pH

7.2. After rinsing in 3 changes of buffer, the material

was postfixed in 1 % (w/v) osmium tetroxide in the same

buffer, for 30 min. Samples were again washed several

times in buffer, followed by dehydration in an ethanol

series, terminating in absolute acetone. For








freeze-substitution fixation colonies were frozen in

acetone at -80 C, then substituted with methanol at -80 C

for 72 h. The samples were warmed over 2 h at room

temperature and transferred into absolute acetone for TEM

or rehydrated in a methanol/water series to be labeled

with cellulase-gold complex and processed for SEM

(modified from Bourett et al. 1998). For methanol

fixation colonies were frozen in methanol at -80 C, then

warmed at room temperature over 2 h and transferred into

absolute acetone for TEM or rehydrated in an

methanol/water series. These samples were labeled with

cellulase-gold complex and processed for SEM. Material

from absolute acetone was infiltrated with an epoxy

embedding medium and polymerized at 60 C for 48 h in a

flat embedding mold. Epon 812 embedding medium was

prepared by combining 55 g of Epon 812, 35 g of DDSA and

21 g of NMA. The accelerator DMP-30 (0.2 mL per 10 mL of

the medium) was added right before embedding. Embedded

samples were sectioned on a Reichert Ultracut R (Leica).

Thin sections (75 to 80 nm) were collected on formvar

coated nickel grids and labeled with the cellulose-gold

complex.

For scanning electron microscopy the colonies were

fixed with 4% glutaraldehyde in 0.05 M sodium cacodylate

buffer and washed several times in buffer (osmium

tetroxide fixation was omitted). Then the colonies were

processed for cellulase-gold labeling, silver enhanced,








dehydrated in an ethanol series finally critically point

dried. For silver enhancement the colonies were placed in

a non-diluted mixture (1:1) of reagents from the Aurion

Silver Enhancement Kit for 5 min and washed several times

with water to stop the reaction (Scopsi et al. 1986).

Cellulose Localization Using Enzyme-Gold Affinity

Labeling

Colloidal gold of approximately 15 nm diameter was

made via reduction of chloroauric acid by sodium citrate

as described by Frens (1973). The enzyme cellulase was

purchased from Worthington Biochemical Corporation,

catalogue No. LS02601. This is chromatographically

"purified" cellulase isolated from cultures of a selected

strain of Tricoderma reesei. A second enzyme was also

used, endocellulase III (provided by Dr. Tim Fowler,

Genencore International, Inc). The solutions used for

conjugation with this enzyme were 5.5 rather than 4.5 for

the commercial cellulase. To coat the gold with

cellulase, the pH of 10 mL of 15 nm colloidal gold was

adjusted to 4.5 and then 1 mg of cellulase dissolved in

0.1 mL distilled water was added with stirring. After 5

minutes the enzyme-gold complex was further stabilized by

the addition of 0.5 mg/mL polyethylene glycol (molecular

weight 20,000). Then the solution was poured into a

centrifugation tube and 1.5 mLof 20% glycerol (in citrate

buffer pH 4.5) was carefully placed on the bottom of the

tube (glycerol was added for long-term storage at -80 C).








The enzyme-gold complex was pelleted at 12,100 rpm for 1

h. Successful coating was evident by a mobile pellet,

which was resuspended in 0.75 mL of 20% glycerol.

Sections, on grids, were preabsorbed for 5 min by

floating them face down on citrate buffer containing 0.5%

gelatin as a blocking agent. The labeling solution was a

1:10 dilution of the enzyme-gold stock with citrate

buffer. To label, grids with sections were floated on the

labeling solution for 30 min. The grids were then floated

on citrate buffer alone for 5 min and rinsed twice for 5

min in distilled water.

The colonies destined for SEM observations were

treated with the same series of solutions, but were

completely submerged, rather than floated.

Cytochemical controls

A number of cytochemical controls were performed to

prove the specificity of the label. (1) Substrate

competition: as a control to determine that the

enzyme-gold probe was binding to cellulose the

cellulase-gold complex was incubated with 1 mg/mL

carboxy-methylcellulose (CMC) (sodium salt, medium

viscosity, Hercules CMC 7MF) for 30 min before the

labeling of sections or colonies. (2) Labeling with

nonenzymatic protein: any nonspecific protein binding

sites were determined by incubating the sections and

colonies with 18 nm Colloidal Gold-AffiniPure Goat








Anti-Mouse IgG (H+L) (Jackson ImmunoResearch

Laboratories, Inc.). (3) Substrate specificity check: two

substates similar to cellulose and present in the cell

wall of Achlya were tested to see if cellulase-gold bound

nonspecifically to them: the cellulase-gold complex was

incubated with 10 mg/mL laminarin (from Laminaria

digitata, Sigma) and 11.8 mg/mL re-acetylated glycol

chitosan (Sigma). (4) Cellulase pretreatment: to

determine the effect of predigestion by free cellulase

the sections and colonies were incubated with 1 mg/mL

cellulase in incubation buffer for 30 minutes prior to

labeling with cellulose-gold complex.

Cellulase Enzyme Activity during Labeling

To determine if the cellulase enzyme, when

conjugated to gold used for labeling, retained enzyme

activity the following experiment was done. The samples

were combined with 1 mL of gold-cellulase complex (1:10

dilution) and incubated at room temperature. A control

for each sample was prepared with substrate and 0.05 M

sodium citrate buffer pH 4.5. The citrate buffer alone

was used as a blank. After 30 min (the usual time of

labeling) and 3 h, the enzyme activity was checked by

Nelson-Somogyi method, using a standard curve obtained by

plotting optical density (at 520 nm) measured on Du-64

Spectrophotometer (Beckman) against known concentrations

of glucose. The samples included: 1 mg CMC, 1 mg of the

whole Achlya wall, small colony and 1 mL of








cellulase-gold complex alone.

Detection of Gold Particles with a Backscatter Detector

To assure that the particles observed on the hyphal

surfaces were actually the gold particles, the regular

secondary images were compared with the backscattering

images of the same regions. The samples were carbon

coated, instead of the usual gold coating. Backscatter

detector (GW Electronics, USA) was used to detect the

backscattering signal.

Zymolyase Hydrolysis

Chemically fixed or live colonies were incubated at

room temperature with 0.05 mg/mL zymolyase 100 T

(Arthrobacter luteus) (Seikagaku Corporation) in 66 mM

sodium phosphate buffer pH 7.5. The hydrolysis was

monitored with light microscope. After 24 h the colonies

were fixed, labeled with cellulase-gold complex and

processed for SEM observations.

Treatment of Growing Colonies with Dichlorobenzonitrile

Small colonies were grown in 500 mL flasks

containing 250 mL of PYG. Each flask contained 10 small

colonies. After 12 h of incubation, dichlorobenzonitrile

(DCB) was added to the flasks. DCB was previously

dissolved in 100% DMSO. The concentrations of DCB were

10, 20, 30, 40, 50, 60 100, and 200 pM. The colonies were

incubated in DCB-containing medium and their growth was

monitored with light microscopy. After 36 h of

incubation, the colonies were chemically fixed, labeled








with cellulase-gold complex and processed for SEM

observations. The colonies grown in PYG only medium

served as a control in this experiment. The ability of

spores to germinate in the medium containing DCB was also

checked. For this 5 mL of fresh spore suspension were

added to PYG medium with and without DCB.

Results

Cellulose Localization

In growing colonies (incubation in PYG) cellulose is

found on the surface of mature and subapical regions on

the hyphae. In apical regions three patterns of labeling

are found in a single colony. Some of the hyphae

(approximately 5%) are not labeled at the apex and show a

sharp border between labeled and non-labeled regions. The

unlabeled area is approximately 2 to 4 urn in diameter

(Figs. 4-11). In the majority of the hyphae, there is a

gradual decrease of the labeling towards the apex (Figs.

12-19). Some of the hyphae (about 5%) are labeled at the

apex as intensively as in the mature regions (Figs.

20-27). Fifty colonies from different batches were

examined and all of them had this pattern and ratio of

labeling.

In contrast, the labeling of hyphae from colonies

incubated in glucose-only medium gave a different pattern

of cellulose labeling. In these colonies, all the hyphae

were labeled at the apices and the labeling was as

intensive as in mature regions (Figs. 28-35). Ten








colonies from different batches were analyzed. Light

microscopic analysis prior to fixation ensured that they

did not contain elongating hyphae.

The surface labeling of hyphae in the

freeze-substituted and methanol-fixed colonies has the

same patterns and ratio as in chemically fixed ones (data

not shown).

Cellulase-gold affinity labeling of cross sections

localizes cellulose in the cell wall exclusively (Fig.

36). The distribution of the gold particles in the wall

is even. The level of nonspecific labeling is very low.

On the longitudinal sections of elongating hyphae the

label is present in mature and subapical regions but is

very low or absent in apical regions (Fig. 37).

There is no labeling on the cross sections (Fig. 38)

or the hyphal surface (data not shown) when the sample of

endocellulase III from Dr. Fowler was used. The

conjugation was successful, based on the raspberry red

color of the enzyme-gold complex and the presence of the

mobile pellet. Negative staining of the enzyme-gold

complex confirmed successful conjugation (Fig. 39).

Cytochemical controls

All the cytochemical controls support the view that

the cellulase-gold complex is a specific label for

cellulose in Achlya. The preabsorbtion of the labeling

solution with CMC results in the absence of the labeling








on the hyphal surface as well as on the cross sections

(Figs. 40, 41).

The incubation of the sections and the colonies with

gold-coupled Goat Anti-Mouse IgG results in the absence

of labeling as well (Figs. 42, 43).

Cellulase pretreatment of the sections and colonies

results in the absence of the labeling (Figs. 44, 45).

The labeling pattern is regular when the

cellulase-gold complex is incubated prior to the labeling

with laminarin or chitosan (Figs. 46-49).

Detection of gold particles on the hyphal surface

with a backscatter detector gave an identical particle

distribution on secondary and backscatter images (Figs.

50-55).

Cellulase activity during labeling

Cellulase-gold complex shows no enzyme activity

during the labeling of the colonies or whole wall samples

as measured by the production of reducing sugar (Table

I). Absorbance of glucose is measured after 30 min (the

usual time of labeling with cellulase-gold complex) and 3

h. Cellulase-gold is diluted 1:10 with the buffer. Based

on glucose equivalents from a standard curve, cellulase

activity is very low in reactions with a whole wall

preparation or a colony. Thus, there are no additional

primers produced and they do not alter the results of

labeling. Enzyme activity of cellulase-gold complex

against cellulase itself is also low. Cellulase activity
























































Figs. 4-11. Scanning electron micrographs of elongating
hyphae of Achlya bisexualis showing cellulose surface
labeling with cellulase-gold. Note the unlabeled apices.
Bars: 4-9 = 2.00 iun; 10 = 2.31 pim; 11 = 3.00 pm.
























































Figs. 12-19. Scanning electron micrographs of elongating
hyphae of Achlya bisexualis showing surface labeling of
cellulose with cellulase-gold. Note the gradual decrease
of labeling towards the apices. Bars: 12 = 3.33 im;
13 = 2.73 irm; 14 = 1.67 4m; 15-18 = 2.00 um; 19 = 3.31
um.





























r .. ,,, "' .%.a. -..
*: "." .,. ,i.i ..
... fi ? .'

. 4-: y*:.-
^^*^,^' ^.' ^


Figs. 20-27. Scanning electron micrographs of surface of
non-elongating hyphae present in growing colonies of
Achlya bisexualis showing cellulose labeling with
cellulase-gold. Note the labeled apices. Bars:
20-22 = 3.00 pm; 23, 24 = 2.00 mu; 25 = 1.50 pm;
26 = 2.31 pm; 27 = 3.33 pm.





















































Figs. 28-35. Scanning electron micrographs of
non-elongating hyphae from non-growing colonies of Achlya
bisexualis, incubated in glucose-only medium showing
surface labeling of apices for cellulose with
cellulase-gold. Note the labeled apices. Bars:
28 = 2.31 jm; 29 = 4.29 im; 30 = 5.00 pm; 31 = 1.50 pm;
32, 33 = 3.75 4m; 34, 35 = 3.00 um.


















































Fig. 36. Cross section of Achlya bisexualis hypha showing
labeling of cellulose in the cell wall with
cellulase-gold complex. Bar=l jum.




54













37









4-




,' % i"





Fig. 37. Longitudinal section of the apical region of an
elongating Achlya bisexualis hypha showing labeling of
cellulose with cellulase-gold. Bar=1 pm.













































i -. "





^fe^^9. -.."...
~v-S -


Fig. 38. Cross section of Achlya bisexualis hypha labeled
for cellulose with endocellulase III-gold. Bar=0.5 pim.











- r **
** ; .' ; *


39


i


* *` L4.


i..: .


.w :. w


'pK


Fig. 39. Negative staining of endocellulase III-gold
complex. Bar=200 nm.


he






















































Figs. 40-41. Electron micrographs showing the absence of
cellulose labeling with cellulase-gold in the cell wall
of Achlya bisexualis resulting from the preabsorption of
the labeling solution with CMC. Bars: 40=1 im; 41=1.5 pm.


















































Figs. 42-43. Electron micrographs showing the absence of
gold label in the cell wall of Achlya bisexualis
resulting from the preincubation of the sections (TEM) or
colonies (SEM) with colloidal gold-affinipure goat
anti-mouse IgG. Bars: 42=1 um; 43=1.5 pm.

















































Figs. 44-45. Electron micrographs showing the absence of
cellulose labeling with cellulase-gold in the cell wall of
Achlya bisexualis resulting from the pretreatment of the
sections (TEM) or colonies (SEM) with cellulase. Bars: 44=1
im; 45=1.67 jim.

















































Figs. 46-47. Electron micrographs showing the regular
pattern of cellulose labeling with cellulase-gold in the
cell wall of Achlya bisexualis resulting from the
preincubation of the labeling solution with chitozan.
Bars: 46=1 in; 47=1.2 mun.














































Figs. 48-49. Electron micrographs showing the regular
pattern of cellulose labeling with cellulase-gold in the
cell wall of Achlya bisexualis resulting from the
preincubation of the labeling solution with laminarin.
Bars: 48=0.5 pm; 49=3 um.


















































Figs. 50-55. Scanning electron micrographs showing the
regular pattern of cellulose labeling with cellulase-gold
on the hyphal surface of Achlya bisexualis. 50, 52, 54.
Secondary images. 51, 53, 55. Backscatter images of the
same regions. Bars: 50 and 51=1.5 uim; 52 and 53=3.00 im;
53 and 55=857 nm.
























Table 1. Glucose equivalent from standard curve showing

cellulase activity during labeling



Glucose equivalent from

Sample standard curve(mg/ml)

Reaction time: Reaction time:

30 min 3 hrs

Whole wall and cellulase-gold 0. 009 0.009

Whole wall and buffer 0.005 0.006

Small colony and cellulase-gold 0.004 0.004

Small colony and buffer 0.002 0.002

Cellulase-gold 0.006 0.007

CMC and cellulase-gold 0.02 0.04





















































Figs. 56-57. Scanning electron micrographs showing
cellulose labeling with cellulase-gold of elongating
Achlya bisexualis hyphae that were hydrolyzed with
zymolyase before chemical fixation. Bars: 56=2.31 unm;
57=2.00 um.

























































Figs. 58-59. Scanning electron micrographs showing
cellulose labeling with cellulase-gold of elongating
Achlya bisexualis hyphae that were hydrolyzed with
zymolyase after chemical fixation. Bars: 58=2.31 pm;
59=1.00 unm.





















































Figs. 60-61. Scanning electron micrographs showing
cellulose labeling with cellulase-gold of an Achlya
bisexualis hypha from a non-growing colony treated with
zymolyase. Bars: 60=3.33 mun; 61=857 nm (higher
magnification of the apex).





67










S62




SAe






















Fig. 62. Transmission electron micrograph of cross
section of Achlya bisexualis hypha from a colony
incubated in 100 rM DCB. Cell wall cellulose is labeled
with cellulase-gold complex. Bar=0.5 pm.








is relatively higher when the enzyme-gold complex reacted

with CMC.

Cellulose Localization on the Surface of the Hyphae in

Colonies Incubated with Zymolyase

In the colonies, growing in PYG medium, that were

hydrolyzed with zymolyase prior to fixation, most of the

hyphal apices are intact and labeled. A small portion of

the hyphae has broken apices. In these hyphae the

remaining cell wall is labeled with cellulase-gold

complex (Fig. 56, 57).

In the growing colonies that were chemically fixed

first and then hydrolyzed with zymolyase, all the hyphae

have intact apices. Most of the apices are labeled, but

some are not (Fig. 58,59).

The non-growing colonies from glucose-only medium

were screened for the absence of elongating hyphae prior

to the treatment. Time of fixation, before or after

hydrolysis, did not make any difference in the results.

All the hyphae in these colonies are intact and are

labeled as intensively as the rest of the hyphal regions

(Fig. 60, 61).

Hyphal Elongation, Spore Germination and Cellulose

Localization in the Presence of DCB

The presence of DCB in the medium does not affect

the growth of Achlya. It does not affect the morphology

of the hyphae or the growth rate. The average hyphal

elongation rate is 3.6 um/min. Spores germinate equally








well in PYG medium with and without DCB. There is no

difference in the cellulose surface labeling of hyphae

from the colonies treated with DCB and regularly growing

colonies. The labeling patterns and their ratio is the

same (data not shown). Cross sections of hyphae from the

colonies treated with DCB have expected pattern of

cellulose labeling (Fig. 62).

Discussion

The cellulase-gold affinity labeling is specific for

cellulose in the fungus Achlya, based on its

reproducibility and a large variety of controls.

Differences in the fixation techniques do not affect the

labeling pattern. Standard chemical fixation gave the

same results as freeze-substitution and cold methanol

fixations. The results are highly reproducible and are

not artifacts of the fixation procedure. Thus this

labeling technique provides a specific and reliable

method for localizing cellulose on thin sections and

hyphal surfaces.

The fact that cellulose labeling was found on the

hyphal surface, may contradict a general assumption that

the microfibrilar component of the cell wall, cellulose

in the case of Achlya, is located next to the plasma

membrane and is covered by the matrix components of the

wall. The cell wall may not be arranged as layers of

components, but as a mixture. This would explain the

presence of some cellulose on the surface, this








explanation seems unlikely since Achlya secretes

cellulase during growth (Thomas and Mullins 1967, 1969),

and if cellulose was present on the surface, it could be

hydrolyzed. Furthermore, when cellulase is applied

exogenuously to the living Achlya cultures, it does not

destroy the integrity of the hyphae, nor does it change

their surface as revealed by shadow replicas (Reiskind

and Mullins 1981b). The unique structure of the cellulase

enzyme complex may explain the presence of cellulose

labeling on hyphal surface. The cellulolytic enzyme

complex from Trichoderma reesei used here for labeling

consists of a number of enzymes: endoglucanases (EG);

cellobiohydrolases (CBH); and cellobiase (CB); which work

synergistically. All these enzymes contain a small highly

homologous 36-residue region called the A domain,

connected to the globular enzymatically active core

domain by a threonine- and serine-rich sequence. The A

domain has no catalytic activity in CBH I and CBH II, but

it is thought to have a cellulose-binding function. The

core protein alone does not have full

cellulose-hydrolysing activity, but has normal activity

on small synthetic substrates (Rouveinen et al. 1990).

Perhaps cellulases are able to bind cellulose

microfibrils located inside the wall via the small

cellulose binding domains (CBD). CBD could penetrate the

wall and find the binding sites, while the catalytic

domains, conjugated to gold remain on the surface. The








results with EG III labeling indirectly prove the idea of

CBD penetrating the wall and leaving the catalytic

domains attached to gold on the surface. EG III provided

by Dr. Tim Fowler (Genencor International, Inc.) is a

genetically modified enzyme that does not have a

cellulose binding domain (personal communication).

Without the binding domain, this enzyme can not attach to

the cellulose microfibrils and it results in the lack of

EG III-gold affinity labeling.

The results of the experiments that measured

cellulase-gold activity during labeling also provide a

support for the idea of CBD penetrating the wall and

leaving the catalytic domain attached to gold on the

surface. The enzyme-gold does not show enzyme activity

against a whole wall sample or a colony, but is active

against the soluble cellulose derivative CMC. Perhaps, in

the case of whole wall and colony treatments the

cellulose binding domain finds the binding site by

penetrating the wall and then attaches to cellulose. The

catalytic domain conjugated to gold does not get access

to cellulose microfibrils surrounded by the matrix

material of the wall. Thus, the binding takes place

without hydrolysis. In the case of CMC the cellulose

microfibrils are not covered, they are available for the

catalytic domain. Therefore, in this case both binding

and hydrolysis take place.








According to the results of cellulase-gold labeling,

all the hyphae from non-growing colonies (glucose-only

medium) are evenly labeled in all regions, including the

apices. Light microscope screening prior to the EM

processing ensured that these hyphae are not elongating.

On the other hand, cellulase-gold labeling of the hyphae

from growing colonies (PYG medium) revealed three

patterns of cellulase-gold labeling at the apices:

labeled (small portion of hyphae), unlabeled and with the

decreasing label towards the apex. Light microscopic

observations prior to fixation revealed that a small

portion of hyphae in these colonies is not elongating,

but the majority of the hyphae are elongating. Based on

this, I propose that in non-elongating hyphae cellulose

is evenly distributed along the hypha and is present in

the apex. Elongating hyphae lack cellulose at the apices

or there is a gradual decrease in the amount of cellulose

toward the apices.

The results of the experiments with zymolyase

hydrolysis support the conclusion that in some hyphae in

growing colonies there is no cellulose in the apical cell

wall. In the colonies that were treated with zymolyase,

prior to fixation, these hyphae have broken apices. I

explain this by the fact that the wall in these apical

regions lacks cellulose and consists mainly of

1,3-8-glucans. Zymolyase has both 1,3-B-glucanase and

protease activities. It hydrolyzes not only








1,3-B-glucans, but also structural wall proteins, cell

membrane proteins and cytoplasmic proteins. Thus, the

elongating regions have hydrolyzedd" apices. In the

colonies that were chemically fixed first and then

hydrolyzed with zymolyase, the elongating hyphae have

intact unlabeled. The apices are not broken as in the

previous case because chemical fixation crosslinks

proteins so they can not be hydrolyzed. As expected, in

both experiments, non-elongating hyphae (glucose-only

medium) have intact apices with cellulose labeling as

intensive as in the other regions of the hyphae.

The experiments with DCB gave an unexpected result.

In these experiments it was the intention to use a

different approach to show the absence of cellulose in

the wall of elongation regions. DCB is a classic

inhibitor of cellulose biosynthesis in higher plants

(Delmer 1999). It was expected that the hyphae would

continue to elongate by synthesizing 1,3-B-glucans and

producing large regions of apical wall made mainly of

this component and that these hyphal regions lacking the

structural support of cellulose might not have a tubular

form. However, DCB had no inhibition effect on the growth

process of Achlya. There were no changes in hyphal

morphology as revealed by light microscopy, TEM or SEM

observations. Hyphal elongation rates in colonies

incubated with DCB were the same as in the regular growth

medium. Cellulose labeling of the cross sections and the








hyphal surface revealed no difference between the hyphae

grown in regular medium or the medium with DCB. Perhaps,

the cellulose biosynthesis system of Achlya is different

from that found in higher plants in the step that is

affected by DCB, or DCB molecules may not be able to

penetrate the Achlya wall. Similar results for the lack

of an inhibitory effect of DCB were found in the cellular

slime mold Dictyostelium (Blanton 1997, Blanton personal

communication). Actually, none of the three major

cellulose-synthesis inhibitors used in higher plants--

DCB, isoxaben, and pthoxazolin--had an effect in this

organism.













CHAPTER 5
LOCALIZATION OF 1,3-B-GLUCANS IN THE CELL WALL AS
REVEALED BY ELECTRON MICROSCOPY AND CYTOCHEMICAL
TECHNIQUES

Introduction

The term glucan applies to polysaccharides composed

of glucose units and they are divided into alpha- and

B-anomers according to their stereochemistry around the

anomeric carbon. The B-glucans include both

homopolysaccharides and heteropolysaccharides. Six

different types of B-glucans have been described in

fungi: linear 1,3-glucans; 1,3-glucans with occasional

1-6 single glucose branches, with or without phosphate;

1,3-glucans with significant amounts of 1,6-branches;

glucans containing mostly 1,6-linkages; glucans

containing 1,3-, 1,4- and 1,6- linkages (Ruiz-Herrera

1991).

These B-glucans are getting attention because of

their potential application in chemical, pharmaceutical

and food industries. Pharmaceutically, 1,3-B-glucans that

have B-glucopyranosyl units attached by 1->6 linkages as

single unit branches have been shown to enhance the

immune system. This enhancement results in antitumor,

antibacterial, antiviral, anticoagulatory and wound

healing activities (Bohn and Bemiller 1995).








The 1,3-B-glucans are important components of fungal

cell walls and they are also storage carbohydrates in

some fungi, especially in the Oomycetes and

Basidiomycota. Some B-glucans are secreted in the form of

slimy material, and may protect cells from desiccation

and other harmful environmental conditions (Ruiz-Herrera

1991). In the case of pathogenic fungi, B-glucans are

important in cellular recognition, and in eliciting

defense responses of infected plants (Ryan 1987, Dixon

and Lamb 1990, Cote and Hahn 1994).

Storage glucans accumulate intracellularly and are

used as reserve material at critical stages of growth and

reproductive development (Wang and Bartnicki-Garcia 1980,

Lee and Mullins 1994). In Phytophthora, Wang and

Bartnicki-Garcia (1973) reported a phosphorylated

1,3-B-glucan in sporangia, zoospores and cysts. This

phophorylated 1,3-B-glucan contains one or two phosphate

residues as monoester linkages at the C-6 hydroxyl groups

of some glucose units. In Achlya, a phosphorylated

cytoplasmic 1,3-B-glucan has been isolated and

characterized (Lee and Mullins 1994, Lee et al 1996),

containing 5% phosphate (w/w), and has both mono-and

diphosphoester linkages. The diester linkages are used to

form very large polymers from the smaller neutral forms.

Although the biological role of the reserve 1,3-B-glucans

is most often suggested as a source of energy or carbon








or both, in Achlya it is also an important site of

phosphate storage (Lee and Mullins 1994).

The most general role of B-glucans is a structural

one, as the major component of fungal cell walls.

Inhibition of B-glucan synthesis in yeast leads to cell

lysis and often death, resulting from a weakening of the

cell wall (Perez et al. 1983, Miyata et al. 1985). Such

inhibitors are used as important antifungal compounds

against both plant and animal pathogens.

The 1,3-B-glucans were localized on cross sections

of Achlya (Shapiro and Mullins 1997). The method used

indirect immunolabeling with a commercial polyclonal

antibody specific for 1,3-B-D-glucopyranose linkages. The

glucans occurred in the cell wall, the large vesicles in

the organelle-rich areas of the hypha, and in the large

central vacuole in more mature areas. Preabsorption of

the antibody with either purified neutral or

phosphoglucan from Achlya completely eliminated

subsequent labeling of hyphal sections. No labeling of

the large population of apical vesicles was found,

suggesting that these reserve glucans are not directly

involved in apical growth. Since the labeling occurred in

large vesicles and the central vacuole and no other

cytoplasmic sites showed conjugation, the vesicle and

vacuole membranes probably contain the synthases

responsible for the biosynthesis of the reserve glucans.

The labeling of serial sections revealed the








1,3-B-glucans in both mature and apical regions.

Additional labeling experiments have now been carried out

on hyphae that were first determined to be elongating, as

described in Chapter 3, to ensure that elongating apices

contained 1,3-B-glucans. Recall that in Chapter 4,

evidence was presented that clearly demonstrated a lack

of cellulose in elongating apices.

Materials and Methods

Culture Methods, Fixation and Microscopy Techniques

The general culture methods, fixation and microscopy

techniques are described in Chapter 3 and Chapter 4.

Localization of 1,3-B-Glucans Using Monoclonal Antibody

The primary antibody, raised in mouse against a

laminarin-haemocyanin conjugate, was purchased from

Biosupplies Australia PtyLtd (Parkville Victoria,

Australia), catalogue number 400-2. This antibody

recognizes linear 1,3-B-oligosaccharide segments in

1,3-B-glucans. The epitope includes at least five

1,3-B-linked glucopyranose residues. It has no cross

reactivity with 1,4-B-glucans or 1,3-B-, 1,4-B-glucans

(Meikle et al. 1991). It was diluted 1:100 in phosphate

buffered saline (PBS), pH 7.2 containing 0.5 % cold water

fish gelatin. The gold reagent, 18 nm Colloidal

Gold-Affinipure Goat Anti-Mouse IgG (H+L) was purchased

from Jackson ImmunoResearch Laboratories, Inc. (West

Grove, Pennsylvania), catalogue number 115-215-146. The

nickel grids with sections were floated on PBS containing








1% gelatin for 30 min to block non-specific labeling.

Then the grids were floated on the top of 20 ul drops of

the primary antibody solution for 1 h. After three washes

in PBS the grids were floated on a 1:20 dilution of the

gold reagent in PBS for 1 h. The solution was centrifuged

at 18 000 g in a microcentrifuge for 1 min before use.

Finally, the grids were washed: twice in PBS and twice in

water.

The colonies destined for SEM observations were

treated with the same series of solutions while

completely submerged into them, rather than floated.

Cytochemical Controls

(1) Preabsorbtion of primary antibody with

laminarin: in order to determine that the primary

antibody binds to 1,3-B-linkages it was preabsorbed with

laminarin. Laminarin from Laminaria digitata was

purchased from Sigma (St. Louis, Missouri), catalogue

number L-9634. The grids with sections were blocked

against nonspecific labeling as described above. Primary

antibody stock solution, 10 ul, was incubated with 100 mg

of laminarin in 1 ml of PBS containing 0.5 % gelatin for

1 h. The grids were floated on a drop of this solution

for 1 h. The labeling procedure as described above was

then followed. (2) Omission of the primary antibody: the

procedure was the same as during the regular labeling,

but the incubation with the primary antibody was omitted.

(3) Replacement of the primary antibody with a non-








specific primary antibody raised in mouse: the procedure

was the same as during the regular labeling but the

primary antibody was replaced by an undiluted

non-specific antibody (HL 1099) raised against

neurofilaments in mouse. HL 1099 was provided by the

Hybridoma Laboratory, Interdisciplinary Center for

Biotechnology Research (Gainesville, Florida).

Results

Localization of 1,3-B-Glucans on Sections and Hyphal

Surfaces Using Monoclonal Antibodies

On cross sections of Achlya the monoclonal antibody

detected 1,3-B-glucans in the wall, small vacuoles, and

the large central vacuole of mature regions (Figs. 63,

64). No cytoplasm-specific labeling was found.

Longitudinal sections of both elongating (Fig. 65) and

non-elongating (data not shown) hyphae show antibody

labeling in the cell wall of the apices and all along the

hyphae. No surface labeling was found using SEM (Fig.

66).

Cytochemical Controls

(1) Preabsorbtion of primary antibody with laminarin

resulted in the absence of labeling (Fig. 67). (2)

Omission of the primary antibody resulted in the absence

of the labeling (Fig. 68). (3) Replacement of the primary

antibody with a non-specific primary antibody raised in

mouse resulted in the absence of the labeling (Fig. 69).





































































Fig. 63. Transmission electron micrograph showing
localization of 1,3-B-glucans with monoclonal antibody on
cross section of Achlya bisexualis hypha. Bar=0.5 pm.
e -




















































Fig. 64. Transmission electron micrograph showing
localization of 1,3-B-glucans with monoclonal antibody on
cross section of Achlya bisexualis hypha. Bar=1.00 pm.





83













































Fg 65 T i e' m.
'. 9








hypha. Bar=1.O ii.
,* *


























Fig. 65. Transmission electron micrograph showing
localization of 1,3-B-glucans with monoclonal antibody on
longitudinal section of an elongating Achlya bisexualis
hypha. Bar=1.00 plm.














































Fig. 66. Scanning electron micrograph showing the absence
of 1,3-B-glucans labeling on the surface of Achlya
bisexualis hypha. Bar=2.31 mn.

















































Figs. 67-69. Transmission electron micrograph showing the
absence of 1,3-B-glucans labeling on the cross sections
of Achlya bisexualis hyphae resulting from: 67.
preabsorbtion of the labeling solution with laminarin.
Bar=1.00 pm; 68. omission of the primary antibody.
Bar=1.00 um; 69. replacement of the primary antibody with
a non-specific antibody raised in mouse. Bar=1.00 um.








Discussion

Based on the results of the cytochemical controls,

the monoclonal antibody used in the labeling procedure is

specific for 1,3-B-glucans. Previous results of labeling

with a polyclonal antibody (Shapiro and Mullins 1997) are

identical to the results of monoclonal antibody labeling.

Both antibodies detect 1,3-B-glucans in the cell wall and

vacuoles. Previously, serial of cross sections of apical,

subapical, and mature regions of a hypha were

immunostained with the polyclonal antibody and strong

labeling was found in the wall on all the sections

(Shapiro 1995). Based on the data presented in Chapter 3,

it can not be ascertained whether this hypha was

elongating or non-elongating. In the present study,

however, the elongating hyphae are distinguished from

non-elongating ones and the distribution of 1,3-B-glucans

is compared. It is now possible to state that

1,3-B-glucans are found in the apical wall of both

elongating and non-elongating hyphae.













CHAPTER 6
LOCALIZATION OF CHITIN IN THE CELL WALL AS REVEALED BY
ELECTRON MICROSCOPY AND CYTOCHEMICAL TECHNIQUES

Introduction

Chitin is the most characteristic polysaccharide of

the fungal cell walls. It is an unbranched polysaccharide

made of N-acetylglucosamine (GlcNAc) joined through 1,4-B

bonds. It was once thought to be absent in fungi

containing cellulose, but a number of examples from all

orders of the Oomycetes have demonstrated a least traces

of chitin (Dietrich 1973, 1975). An insoluble fraction

from the hyphal wall of Achlya radiosa Maurizio was

characterized by x-ray and infrared analyses as chitin,

and represented about 4% of the total wall (Campos-Takaki

et al. 1982). The role of chitin in Oomycete cell wall

remains unclear, and it has been suggested in Saprolegnia

that chitin does not play an important role in

morphogenesis based on results using the chitin synthase

inhibitor polyoxin D (Bulone et al. 1992). An insoluble

residue representing about 3% of the wall and containing

glucosamine was reported in Achlya (Reiskind and Mullins

1981a). This fraction was identified as chitin by x-ray

analysis (Mullins et al. 1984), but had unusual

properties in that it is more highly crystalline than the








alpha-chitin normally observed in fungi and the

characteristic lattice spacing was not readily

perceptible. Thus chitin is clearly present in those

fungi having cellulose as the major microfibrillar

component; but its role is yet to be determined.

Materials and Methods

Chitin Localization Using Lectin

Tomato (Lycopersicon esculentum) lectin conjugated

to gold was purchased from EY Laboratories, Inc. (San

Mateo, California). The tomato lectin is described by the

manufacturer as being specific for oligomers of

1,4-B-linked N-acetylglucosamine, with the binding site

being able to accommodate up to 4 carbohydrate units and

these units do not have to be consecutive. The sections,

on grids, were pretreated with phosphate buffer saline

(PBS) containing 1% bovine serum albumin (BSA) at room

temperature for 30 minutes. Then they were floated on the

labeling solution for 30 minutes. The lectin-gold complex

was a 1:9 dilution of the stock solution in PBS. The

samples then were washed with PBS three times and rinsed

twice in distilled water.

Cytochemical Control

To determine that the lectin-gold complex was

binding to chitin, the probe was pre-incubated with

re-acetylated glycol chitosan provided by Dr Michael N.

Horst (Mercer University, Macon, Georgia). The glycol

chitosan stock (0.118 g/100 mL) was diluted with PBS








(1:9). One part of the lectin-gold stock solution was

diluted with nine parts of glycol chitosan and incubated

for 30 min before labeling of the sections.

Results

Chitin is localized to the cell wall of Achlya

bisexualis with tomato lectin-gold conjugate, where it is

evenly distributed in the cross sections (Fig. 70). The

chitin labeling is absent when the labeling solution is

preincubated with glycol chitozan (Fig. 71).

Discussion

The tomato-lectin-gold conjugate appears to be a

specific label for chitin in the cell walls of Achlya,

based on the lack of labeling when the conjugate was

pre-incubated with re-acetylated glycol chitosan.

Previous studies on chitin (Campos-Takaki et al 1982,

Mullins et al 1984, Gay et al 1992) demonstrated its

presence in the cell walls of oomycetes with biochemical

and biophysical analyses. This is the first report of the

cytochemical localization and distribution of chitin in

the walls of this group. Bulone et al 1992 described

chitin as small globular particles in Saprolegnia, and

found that hyphal growth and morphology were not altered

when chitin synthesis was inhibited by polyoxin D. They

concluded that chitin did not seem to play an important

role in morphogenesis. Additional biophysical work on

Saprolegnia (Gay et al 1992) describe chitin as localized

small round granules of crystalline microfibrillar alpha





90


chitin. Chitin, however, synthesized in vitro appeared as

spindle-like particles, and was not a skeletal

polysaccharide involved in wall architecture. In

regenerating protoplast walls it might have a secondary

role in wall architecture, since it is microfibrillar.

Thus the full role of chitin is still to be determined.
















































Fig. 70. Transmission electron micrograph showing
localization of chitin with tomato lectin on cross
section of Achlya bisexualis hypha. Bar=0.5 umn.

Fig. 71. Transmission electron micrograph showing the
absence of chitin labeling on the cross sections of
Achlya bisexualis hyphae resulting from preincubation of
the labeling solution with re-acetylated glycol chitozan.
Bar=1.00 pm.















CHAPTER 7
CONCLUSIONS

The results of cellulose localization suggest that

in non-elongating hyphae, cellulose is evenly distributed

along the hypha and is present in the apex. Elongating

hyphae lack cellulose at the apices or there is a gradual

decrease of cellulose amount toward the apices. On the

other hand, the major matrix component of the wall,

1,3-3-glucan, is distributed evenly over the elongating

and non-elongating hyphae and is present in their apices.

Such distribution of these two major components of Achlya

cell wall suggests that in the elongation zone,

1,3-B-glucans are synthesized first and cellulose

deposition follows. This contradicts the idea shared by

the major theories of hyphal tip growth, that all the

wall components are present in the elongation zone.

The small diameters of the cellulose unlabeled

regions of elongating hyphae suggest that cellulose

deposition takes place almost immediately after the start

of 1,3-B-synthesis. The plastic wall that consists mainly

of 1,3-B-glucans and lacks cellulose support is stretched

under the turgor pressure and/or perhaps pressure of

cytoskeleton. The quickly following cellulose deposition

helps to maintain the tubular cell shape and prevents the








elongation regions from "blowing out" in balloon-like

structures. Cellulose is thought to provide mechanical

support for the cell wall. The initial cellulose

hydrolysis by cellulase in the growing apex is possible.

This could create new primers in existing cellulose

chains, as suggested by Maclachlan (1976) for higher

plants. It was found that the growing colonies of Achlya

secrete endocellulase (Thomas and Mullins 1967; 1969).

The authors suggested that endocellulase is important for

the wall softening since this fungus does not use

cellulose in nutrition. The recent evidence that activity

of the secreted endocellulase correlates with the tensile

strength of the apical hyphal wall support this idea

(Money and Hill 1997).

The results of cellulose and 1,3-B-glucans

distribution in the apical wall, combined with the

results of hyphal growth monitoring suggest a new aspect

of the hypothesis for hyphal tip growth. This hypothesis

would state that in the Achlya growth process, all hyphae

go through periods of elongation and no-elongation

(dormancy). The elongation is not a steady process as it

is generally assumed. It consists of alternating periods

with fast and slow growth rates. Elongation starts with

synthesis of 1,3-B-glucans, which is quickly followed by

synthesis of cellulose.













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