• TABLE OF CONTENTS
HIDE
 Title Page
 Acknowledgement
 Table of Contents
 List of Tables
 List of Figures
 Abstract
 Introduction
 Review of the literature
 Materials and methods
 Electron microscopy of hyphal...
 Cellulase and UDPG transferase
 The association of wall synthesis...
 Isolation of cellulase-containing...
 General discussion
 References
 Biographical sketch






Title: Isolation and characterization of vesicles involved in hyphal tip growth of Achlya /
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 Material Information
Title: Isolation and characterization of vesicles involved in hyphal tip growth of Achlya /
Physical Description: xii, 152 leaves : ill. ; 28 cm.
Language: English
Creator: Hill, Terry William, 1947-
Publication Date: 1978
Copyright Date: 1978
 Subjects
Subject: Achlya   ( lcsh )
Fungi -- Physiology   ( lcsh )
Botany thesis Ph. D
Dissertations, Academic -- Botany -- UF
Genre: bibliography   ( marcgt )
non-fiction   ( marcgt )
 Notes
Thesis: Thesis--University of Florida.
Bibliography: Bibliography: leaves 137-151.
Additional Physical Form: Also available on World Wide Web
General Note: Typescript.
General Note: Vita.
Statement of Responsibility: by Terry William Hill.
 Record Information
Bibliographic ID: UF00097469
Volume ID: VID00001
Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
Resource Identifier: alephbibnum - 000013237
oclc - 04803295
notis - AAB6258

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Table of Contents
    Title Page
        Page i
    Acknowledgement
        Page ii
    Table of Contents
        Page iii
        Page iv
    List of Tables
        Page v
        Page vi
    List of Figures
        Page vii
        Page viii
        Page ix
    Abstract
        Page x
        Page xi
        Page xii
    Introduction
        Page 1
    Review of the literature
        Page 2
        Page 3
        Page 4
        Page 5
        Page 6
        Page 7
        Page 8
        Page 9
        Page 10
        Page 11
        Page 12
        Page 13
        Page 14
        Page 15
        Page 16
        Page 17
        Page 18
        Page 19
        Page 20
    Materials and methods
        Page 21
        Page 22
        Page 23
        Page 24
        Page 25
        Page 26
        Page 27
        Page 28
        Page 29
        Page 30
        Page 31
        Page 32
        Page 33
        Page 34
    Electron microscopy of hyphal apices
        Page 35
        Page 36
        Page 37
        Page 38
        Page 39
        Page 40
        Page 41
        Page 42
        Page 43
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        Page 45
        Page 46
        Page 47
        Page 48
        Page 49
        Page 50
        Page 51
        Page 52
        Page 53
        Page 54
    Cellulase and UDPG transferase
        Page 55
        Page 56
        Page 57
        Page 58
        Page 59
        Page 60
        Page 61
        Page 62
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        Page 78
        Page 79
        Page 80
        Page 81
        Page 82
        Page 83
        Page 84
    The association of wall synthesis and enzymes with hyphal growth
        Page 85
        Page 86
        Page 87
        Page 88
        Page 89
        Page 90
        Page 91
        Page 92
        Page 93
        Page 94
        Page 95
        Page 96
        Page 97
        Page 98
        Page 99
        Page 100
    Isolation of cellulase-containing membranes
        Page 101
        Page 102
        Page 103
        Page 104
        Page 105
        Page 106
        Page 107
        Page 108
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        Page 124
        Page 125
        Page 126
        Page 127
        Page 128
        Page 129
        Page 130
    General discussion
        Page 131
        Page 132
        Page 133
        Page 134
        Page 135
        Page 136
    References
        Page 137
        Page 138
        Page 139
        Page 140
        Page 141
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        Page 145
        Page 146
        Page 147
        Page 148
        Page 149
        Page 150
        Page 151
    Biographical sketch
        Page 152
        Page 153
        Page 154
Full Text












ISOLATION AND CHARACTERIZATION OF VESICLES INVOLVED IN
HYPHAL TIP GROWTH OF ACHLYA













By

TERRY WILLIAM HILL


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




UNIVERSITY OF FLORIDA


1978














ACKNOWLEDGEMENTS


I thank Drs. H. C. Aldrich, G. E. Bowes, and J. H. Gregg for

serving as members of my supervisory committee and Dr. F. C. Davis,

who freely provided his advice and facilities throughout the research.

I especially thank Dr. J. T. Mullins, who, as chairman of my

supervisory committee, suggested the problem and supported the re-

search through his time, counsel, monies, and facilities. Dr. Mullins'

role in all these regards is sincerely appreciated.

Lastly, I thank my wife, Evalie, whose love and support in so

many ways made the successful conduct and completion of my studies

possible.














TABLE OF CONTENTS

Page
CHAPTER

ACKNOWLEDGEMENTS . . . . . . . . ... . . . ii

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

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

ABSTRACT . . . . . . . . . ......... x

INTRODUCTION . . . . . . . . . . .... 1

REVIEW OF THE LITERATURE . . . . . . . . . . 2

Achlya ambisexualis Raper . . . . . . . . 2
Apical Growth and the Fungal Cell . . . . . . 5
The Fungal Cell Wall . . . . . .. . . . 6
Cell Wall Modification During Growth . . . . . 8
Involvement of Cytoplasmic Structures in Wall
Formation. . . . . . . . ... ..... 13

MATERIALS AND METHODS. . . . . . . . . ... . 21

General Culture Methods . . . . . . . ... .21
Cell Homogenization . . . . . . . .... . 23
Centrifugations . . . . . . . .... .. .. .23
Assays . . . . . . . . . . . 24
Liquid Scintillation Counting . . . . . . . 29
Statistical Methods . . . . . . . .... . 30
Electron Microscopy . . . . . . . .... . 30
Cytochemical Tests. . . . . . . . .... . 31

ELECTRON MICROSCOPY OF HYPHAL APICES . . . . . .... .35

The Cytoplasmic Organization of Achlya Hyphal Apices. . 35
Cytochemical Localization of Enzymes and Other Materials
in Hyphal Apices . . . . . . . . ... .42
Discussion. . . . . . . . . ... ...... 49

CELLULASE AND UDPG TRANSFERASE . . . . . . .... .55

Some Properties of Mycelial Cellulase . . . .... .55
Some Properties of UDPG Transferase . . . . .... .70
Discussion. . . . . . . . . ... ...... 76


iii








Page
THE ASSOCIATION OF WALL SYNTHESIS AND ENZYMES WITH
HYPHAL GROWTH . . . . . . . . . . . 85

Enzymes in the Culture Filtrate . . . . 85
Comparisons of the Growing and Nongrowing Conditions . 86
Discussion. . . . . . . . . . . . .. 96

ISOLATION OF CELLULASE-CONTAINING MEMBRANES. . . . . ... 101

The Distribution of 280 nm-absorbing Materials and
Cellulase-containing Particles in Isopycnic
Sucrose Gradients . . . . . . . 101
Enrichment of Cellulase-containing Particles by
Sequential Differential, Velocity, and Isopycnic
Centrifugation . . . . . . . . . . 112
Discussion. . . . . . .. . . . . . 125

GENERAL DISCUSSION . . . . . . .. . . . . 131

REFERENCES . . . . . . . . . . . . 137

BIOGRAPHICAL SKETCH. . . . . . . . . . . .. 152














LIST OF TABLES


Table Page

1 The composition of Defined Liquid Medium (DLM)
(modified from Mullins and Barksdale, 1965) . . .. 22

2 Embedding medium for electron microscopy. . . .. 31

3 Cellulase activity in salt-soluble or buffer-soluble
and buffer-insoluble fractions from homogenates of
replicate 2g FW samples of A. ambisexualis mycelium
produced by the method of Thomas (1966) or by
grinding in a buffered osmoticum, respectively ... 57

4 The effect of triton X-100 on the activity of
particulate and buffer-soluble cellulases from
A. ambisexualis mycelial homogenates. . . . ... 59

5 The distribution of Cx activity between particulate
and soluble phases after treatment of A. ambisexualis
cellular particles with various concentrations of
triton X-100. . . . . . . . . ... . 60

6 The distribution of protein and Cx activity between
particulate and soluble phases after treatment of
A. ambisexualis cellular membranes with salts,
freezing, or sonication . . . . . . ... .63

7 The effect of DTT upon cellulase activity during
incubation of A. ambisexualis cellular membranes for
24 hr at room temperature . . . . . .... .64

8 The effect of temperature upon cellulase activity
during incubation of A. ambisexualis cellular mem-
branes for 24 hr in 0.5 mM DTT. .. . . . ... 65

9 The effect of temperature upon solubilization of
cellulase from A. ambisexualis cellular membranes
during incubation in the presence of 0.5 mM DTT for
24 hr . . . . . . . . .. .. . 67

10 The distribution of protein and UDPG transferase
activity between "wall" and "protoplasm" fractions of
A. ambisexualis mycelial homogenates produced by
sonication . . . . . . . . .. . 71








Table


11 The distribution of protein and UDPG transferase
activity between particulate and soluble proto-
plasmic fractions of A. ambisexualis mycelial
homogenates produced by grinding with a mortar and
pestle . . . . . . . . ... .. .. . .73

12 The distribution of radioactivity among different
extracts of the products of UDPG transferase
activity from the particulate fraction of A.
ambisexualis mycelial homogenates. ... . . . .. 75

13 The activities of enzymes in the mycelium and
culture filtrate of 48 hr old cultures of
A. ambisexualis. . . . . . . . .... . 87

14 Changes in fresh weight of 2 g FW lots of A.
ambisexualis mycelium during incubation in Defined
Liquid Medium (DLM) or 0.2% Glucose Medium (GM). ... 89

15 The incorporation of exogenous glucose into walls
of 2.0 g FW lots of A. ambisexualis mycelium during
incubation in Defined Liquid Medium (DLM) or 0.2%
Glucose Medium (GM). . . . . . . . . .. 91

16 The recovery of cellulase from media after incuba-
tion of 2.0 g FW lots of A. ambisexualis mycelium
in 50.0 ml volumes of Defined Liquid Medium (DLM) or
0.2% Glucose Medium (GM) . . . . . . . . 92

17 Protein content, carbohydrate content, and the
specific activities of mycelial enzymes in the
particulate and soluble phases of A. ambisexualis
mycelial homogenates, after incubation of 2 g FW
lots of mycelium for 3 hr in either Defined Liquid
Medium (DLM) or 0.2% Glucose Medium (GM) . . ... 94

18 Distribution of cellulase, cytochrome oxidase, and
glucose-6-phosphatase in differential centrifugation
fractions of A. ambisexualis mycelial homogenates. . 114

19 Carbohydrate content and the specific activities of
selected enzymes at each stage in the enrichment of
cellulase-rich particles from A. ambisexualis mycelial
homogenates. . . . . . . . .. .. .120


Page














LIST OF FIGURES


Figure Page

1 Longitudinal section of the subapical region of an
A. ambisexualis hypha. . . . . . . . ... 38

2 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the many cyto-
plasmic vesicles in the region . . . . .... .38

3 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows a dictyosome
surrounded by cytoplasmic vesicles . . . .... 38

4 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows a dictyosome
bearing an incipient fibrous vesicle . . . ... 38

5 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows details of the
cytoplasmic vesicles . . . . . . . ... 38

6 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows cytoplasmic
vesicles beneath the cell wall . . . . . .. 38

7 Tangential longitudinal section of the subapical
region of an A. ambisexualis hypha . . . .... 41

8 Longitudinal section of the subapical region of an
A. ambisexualis hypha. . . . . . . . ... 41

9 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows coated vesicles
in the vicinity of and attached to dictyosomes. .... .41

10 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the lack of
deposition of cellulase reaction product in the
hypha . . . . . . . . . . . . 41

11 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the distribution
of acid phosphatase-positive and acid phosphatase-
negative vesicles in the apex. . . . . . .. 41







Figure Page

12 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows the deposition of
acid phosphatase reaction product in some of the
cytoplasmic vesicles, but not in others . . . ... 41

13 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows the deposition of
acid phosphatase reaction product in a single cisterna
of a dictyosome . . . . . . . . ... 45

14 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the deposition of
IDPase reaction product in some of the cytoplasmic
vesicles, but not in others . . . . . . ... .45

15 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the distribution
of IDPase-positive and IDPase-negative vesicles in
the apex. . . . . . . . ... ....... 45

16 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows the lack of IDPase
reaction product in a dictyosome. . . . . . ... 45

17 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows the lack of
alkaline phosphatase reaction product in dictyosomes. . 45

18 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the lack of
alkaline phosphatase reaction product in cytoplasmic
vesicles. . . . . . . . . . .. . . 45

19 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the stainability of
the plasma membrane and some (but not all) of the
cytoplasmic vesicles with the PTA-Cr03 stain. . . ... 48

20 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the distribution
of PTA-CrO3-positive vesicles in the apex . . ... 48

21 Longitudinal section of the apical region of an
A. ambisexualis hypha, which shows the stainability
of cytoplasmic vesicles and the cell wall with the
PASM stain for carbohydrate . . . . . .... .48

22 Longitudinal section of the subapical region of an
A. ambisexualis hypha, which shows a dictyosome
that has been stained with the PASM stain for
carbohydrate. . . . . . . . .. .. . 48


viii







Figure Page

23 The distribution of 280 nm-absorbing materials
after centrifugation to equilibrium of the
270 x g x 10 min supernatant from an A. ambisexualis
homogenate in a linear sucrose gradient. . . . ... 105

24A The distribution of cytochrome oxidase, IDPase,
B-glucosidase, and UDPG transferase after centri-
fugation to equilibrium of the 270 x g x 10 min
supernatant from an A. ambisexualis homogenate in a
linear sucrose gradient. . . . . ... . .. 108

24B The distribution of ATPase, cellulase, glucose-6-
phosphatase, and carbohydrate after centrifugation
to equilibrium of the 270 x g x 10 min supernatant
from an A. ambisexualis homogenate in a linear
sucrose gradient . . . . . . . .... . 110

25 The distribution of cellulase and 280 nm-absorbing
materials after velocity centrifugation of the
"25 K x g" differential centrifugation fraction of
an A. ambisexualis homogenate in a 15-35% linear
sucrose gradient over a 65% sucrose cushion. . . ... 118

26 Section of a purified membrane fraction from an
A. ambisexualis mycelial homogenate, which contains
dictyosome cisternae and unidentified membrane
vesicles . . . . . . . . ... . . .124

27 Section of a purified membrane fraction from an
A. ambisexualis mycelial homogenate, which shows a
fragmented dictyosome cisterna, bearing incipient
vesicles . . . . . . . . ... . . . 124

28 Section of a purified membrane fraction from an
A. ambisexualis mycelial homogenate, which shows
ribosomes associated with isolated membranes . . .. .124

29 Section of a purified membrane fraction from an
A. ambisexualis mycelial homogenate, which shows the
stainability of dictyosome cisternae and some of the
unidentified membrane vesicles with the PASM stain
for carbohydrate . . . . . . . .... . 124

30 Section of a purified membrane fraction from an
A. ambisexualis mycelial homogenate, which shows the
lack of stainability of isolated membranes with the
PTA-Cr03 stain . . . . . . . . ... . 124

31 Section of a purified membrane fraction from an
A. ambisexualis mycelial homgenate, which has been
oxidized with periodic acid, but not stained with
PTA-Cr03 . . . . . . . . . . . .. 124







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


ISOLATION AND CHARACTERIZATION OF VESICLES INVOLVED IN
HYPHAL TIP GROWTH OF ACHLYA

By

Terry William Hill

December 1978

Chairman: J. T. Mullins
Major Department: Botany

Achlya ambisexualis Raper is a heterothallic, oomycetous fungus.

During sexual reproduction, hormone-induced male strains produce hyphal

branches, which elongate by apical growth and differentiate into anther-

idia upon contact with the female. Antheridial hyphae presumably

elongate by the same mechanism that permits vegetative growth. Theories

have been proposed to account for the pattern of apical vegetative

growth, and these involve the coordinated action of cell wall synthe-

sizing enzymes (e.g., UDPG transferase) and cell wall hydrolyzing

enzymes (e.g., cellulase) in the hyphal tip. The present study was

conducted in order to gain information about the mechanism of vegetative

hyphal tip growth in Achlya.

Some aspects of the growing and nongrowing states were examined.

Mycelial growth is reduced by about 90% when mycelia are transferred

from defined liquid medium (DLM) to 0.2% glucose medium (GM). Mycelia

growing in DLM incorporate exogenous glucose into cell walls and

secrete cellulase into the medium, but these processes are reduced by

about 90-100% when mycelia are transferred to GM. Of the enzymes

tested, only cellulase and alkaline phosphatase exhibit higher specific








activities in growing mycelia than in nongrowing mycelia; the specific

activity of UDPG transferase does not change.

The enzyme cellulase exists both as a buffer-soluble and a buffer-

insoluble form. The insoluble form is membrane-bound and can be

solubilized with 1% triton X-100 or by incubation at room temperature

in 0.5 mM DTT, with a subsequent 8- to 10-fold increase in activity.

Freezing, sonication, and 1 M salts do not solubilize this cellulase.

I conclude that this particulate cellulase is an integral membrane

protein.

Mycelial homogenates were centrifuged isopycnically in a linear

sucrose gradient, in which most of the cellulase activity equilibrates

at a density of 1.19 g/cm3. Most carbohydrate, UDPG transferase,

IDPase, and ATPase also equilibrate here. Enrichment of these activi-

ties was achieved by recovering those particles that sediment from

homogenates between 5,000 x g x 10 min and 25,000 x g x 10 min and

recentrifuging them in a 15-35% sucrose velocity gradient, before a

final isopycnic centrifugation in a linear 20-55% sucrose gradient.

Particles equilibrating at 1.19 g/cm3 consist of dictyosome cisternae

and unidentified smooth membranes. The PTA-CrO3 stain for plasma

membranes fails to stain these particles; cisternae and some of the

smooth membranes stain with the PASM stain for carbohydrate.

In order to identify cellulase-containing membranes, growing

hyphae were examined electron-microscopically, and a number of cyto-

chemical tests and ultrastructural enzyme localizations were performed.

Hyphal tips contain cytoplasmic vesicles, which are apparently produced

by dictyosomes. Vesicles are of at least two major classes, whose

sizes are about 150 nm and about 80 nm in diameter, respectively. Some







vesicles in each class contain IDPase and the 150 nm vesicles stain

with the PASM stain. Dictyosomes are IDPase-negative and PASM-positive.

The plasma membrane and some of the 80 nm vesicles stain with the PTA-

Cr03 stain.

The identical distribution of cellulase, UDPG transferase,

carbohydrate, and IDPase in isopycnic gradients indicates that trans-

ferase and cellulase are localized in the IDPase-positive, PASM-

positive cytoplasmic vesicles. This supports the theory that postu-

lates the coordinated involvement of cell wall synthesis and lysis in

apical growth of fungi. The involvement of vesicles provides a

mechanism for the simultaneous delivery of these materials to the apex.

The fact that repression of growth is accompanied by a reduction of

mycelial cellulase activity and abolition of cellulase secretion,

while UDPG transferase activity is unchanged, supports the proposal that

the rate of wall synthesis can be regulated by the availability of

polysaccharide chain "primer" ends in the wall. Membrane-bound cellu-

lase may be transferred to the cell surface by fusion of apical

vesicles with the plasma membrane, at which time the enzyme is solu-

bilized and produces cellulose "primer" ends in the wall by endo-

hydrolysis. The subsequent release of cellulase into the medium may

ensure that effective levels of activity occur only at the apex.














INTRODUCTION


Fungal hyphae are walled, filamentous cells, which extend by

localized growth at their tips. This process includes continued bio-

synthesis of new cell walls. Cytological and biochemical studies of

growing hyphae suggest that at least part of the synthetic process may

involve apical vesicles derived from the Golgi apparatus, and attempts

have been made to isolate these and other cellular structures in hopes

of determining their contents. Among the proposed contents are wall

synthesizing enzymes, wall hydrolyzing enzymes, and carbohydrate wall

precursors; as yet, their simultaneous association with purified

membranes of filamentous fungi has not been demonstrated.

The present study utilizes the fungus Achlya ambisexualis Raper in

an examination of apical growth. The choice of the organism is appro-

priate because the involvement of the enzyme cellulase with a specific

wall-related morphogenetic event in the life cycle is already well

characterized, and a model employing cytoplasmic vesicles to explain

this event has been proposed. Though the present study does not ex-

amine these events of hormone-induced branching, it is hoped that a

better understanding of the mechanism of ordinary vegetative growth in

Achlya will contribute to further studies both of hormone induction and

of fungal growth in general.














REVIEW OF THE LITERATURE


Achlya ambisexualis Raper


The fungal species Achlya ambisexualis Raper is classified in the

order Saprolegniales, class Oomycetes, division Mastigomycotina (Dick,

1973). Like other fungi, individuals exhibit an absorbtive, hetero-

trophic mode of nutrition plus a conspicuous cell wall and lack of

motility in the vegetative state. Unlike most other fungi, however,

asexual reproduction involves the production of heterokont zoospores, and

the cell wall contains cellulose instead of chitin (Sparrow, 1960).

The growth form of Achlya is typical of most fungi. The body or

thallus consists of branched cylindrical filaments called hyphae, and

the assemblage of all the hyphae of a thallus is called a mycelium

(Alexopoulos, 1962). Ordinarily, the mycelium is considered to consist

of but one cell, which contains many nuclei and is not divided by cross

walls. This is, perhaps, strictly true only of very young mycelia,

because cross walls do form as the mycelia age. They function to wall

off older, nonfunctional regions of cytoplasm and to delimit repro-

ductive structures (Johnson, 1956). Thus, the vegetative portion of the

mycelium, even in mature individuals, consists of from one to several

extensively branched coenocytic cells, each capable of independent

colonization, growth, and reproduction. In the case of water molds,

there is evidence that, nutritionally at least, there is little








communication between widely separated parts of the same mycelium, re-

enforcing the impression of functional autonomy among growing regions

(Jennings et al., 1974).

Achlya is capable of both sexual and asexual reproduction. Asexu-

ally, individuals reproduce by fragmentation, by differentiation of

hyphae into resistant gemmae, or by differentiation of vegetative apices

into clavate zoosporangia (Johnson, 1956; Sparrow, 1960). Zoosporangial

formation can be induced by depletion of nutrients (Klebs, 1899) and

results in the formation of biflagellate spores, which settle and encyst

after a period of swimming. Germination may then occur on a suitable

substrate via a germ hypha (Coker, 1923).

Achlya ambisexualis is one of three Achlya species that are het-

erothallic; but, as the specific epithet implies, members of the species

are ambivalent toward any rigid assignation of gender (Raper, 1951).

Individuals exhibit varying degrees of sexual disposition, and many can

be induced to act as either male or female, depending on the partners

with which they are paired. In response to the proper hormonal cues,

compatible vegetative thalli form appropriate sex organs de novo; their

subsequent fusion is also under hormonal control (Raper, 1951;

Barksdale, 1969). The best studied of the hormonal responses is that

elicited by the steroid antheridiol, which is secreted constitutively by

the vegetative female. Individuals capable of responding as a male ex-

hibit a series of specific molecular responses to antheridiol induction

(e.g., Kane et al., 1973), culminating in the copious production of

lateral antheridial branches (Barksdale, 1970). The branches grow toward

the female thallus, where they make contact with oogonial initials.








Delimitation of sex organs, meiosis, gametogenesis, and fertilization

follow (Raper, 1951).

It is the events between antheridiol uptake and lateral branch pro-

duction that have attracted the most interest of Achlya researchers

because the system provides a model system for investigating steroid

hormone action. However, the steps of greatest significance to fungal

biology are those of branch initiation itself, and the most relevant work

here has dealt with the events of wall modification. At the time of

branch initiation, the intracellular level of endocellulase (Cx) ac-

tivity rises and then declines as the enzyme is secreted into the medium

(Thomas and Mullins, 1967, 1969). Cellulase secretion and branch ini-

tiation are dependent on synthesis of RNA and protein and can be pre-

vented by selective inhibitors (Kane et al., 1973; Horowitz and Russell,

1974; Timberlake, 1976). This response is seen only in those Achlya

strains that produce antheridial branches and has led to the hypothesis

that induction of antheridial branches requires the delivery of cellu-

lase to specific points on the wall, where lytic wall thinning and

softening permit a turgor-driven "blow out" of the wall (Thomas and

Mullins, 1967, 1969).

Support for the above hypothesis derives from the observation that

lateral walls are thinned at points of antheridiol-induced branch ini-

tiation, and these points are subtended by accumulations of cytoplasmic

vesicles (Mullins and Ellis, 1974). In another investigation, hormone-

induced hyphae were shown to contain vesicles beneath the lateral to

subapical walls, which react positively to a cytochemical test for

cellulase (Nolan and Bal, 1974). However, the vesicles in question were

not associated with points of branch initiation, leaving open the








possibility that they were involved in some other cell process, such as

ordinary vegetative growth.

It must, at this time, be emphasized that hormone-induced branch-

ing may not differ qualitatively from similar events during vegetative

growth, inasmuch as branching is a regular part of fungal growth. In an

induced thallus, branching increases, and growth occurs both in newly

formed branches and in existing apices. Furthermore, after the initial

modification of lateral walls for branch initiation, continued growth of

antheridial hyphae presumably occurs by the same mechanism as that which

permits vegetative growth. Knowledge of the method of vegetative fungal

growth is, therefore, relevant to an understanding of reproductive growth.


Apical Growth and the Fungal Cell


Growth of filamentous fungal cells has been shown to occur by

localized extension of the hyphal tips (Smith, 1923; Robertson, 1965).

Regions more than about five micrometers from the tip do not elongate.

New tips are, of course, initiated by branching in lateral regions, but

subsequent growth is apical. Other forms of fungal growth are known,

but they serve specialized functions, such as the elevation of terminal

sporangia by intercalary growth of the sporangiophore (Castle, 1942).

Apical growth is not unique to fungi but is a characteristic growth form

of a variety of filamentous cells. Among these are algal filaments and

rhizoids (Ott and Brown, 1974; Sievers, 1967) and pollen tubes and root

hairs of higher plants (Rosen et al., 1964; Bonnett and Newcomb, 1966).

Because of its growth pattern, a fungal hypha consists of an

apical region, which is constantly growing, and an older region, which

was once the site of growth but is no longer capable of extension.







This dichotomy of growth potential is reflected in the cytoplasmic

structure and chemistry of growing and nongrowing regions. Apical and

mature regions have been shown to differ in their distributions of vari-

ous macromolecules, enzyme activities, and reducing potential (Zalokar,

1965; Turian, 1976). Older regions of the hypha are typically highly

vacuolate, and the cytoplasm is restricted to a thin, peripheral layer

between the tonoplast and plasma membrane (Bracker, 1968). The cyto-

plasm contains a variety of eucaryotic organelles, which includes

dictyosomes in the case of Oomycetes. Typically, the terminal 40-100

micrometers are nonvacuolate and are particularly rich in organelles.

At the very apex, however, the cytoplasm is particularly devoid of most

organelles and is occupied almost exclusively by a collection of apical

vesicles (McClure, et al., 1968; Girbardt, 1969; Grove and Bracker, 1970;

Grove et al., 1970). Based on this distribution, three distinct cyto-

plasmic zones are recognized, corresponding to the older vacuolate region,

the subapical organelle-rich region, and the terminal vesiculate region

(Grove et al., 1970).


The Fungal Cell Wall


In the mature cell walls of Oomycetes, typically from 80-90% of the

dry weight is composed of carbohydrate, with protein and lipid consti-

tuting the rest; similar proportions are found in walls of other fungal

groups (Bartnicki-Garcia, 1968). Analysis of Achlya cell walls reveals

that the carbohydrate fraction contains from 10% to 15% cellulose in a

weakly crystalline form (Parker et al., 1963; Aronson et al., 1967),

while the remainder consists primarily of a highly branched glucan







containing B-1,3 and B-1,6 linkages (Aronson et al., 1967). Small

amounts of nonglucose sugars are also present (Thomas, 1966; Dietrich,

1973).

In a manner resembling the walls of higher plants, these components

are organized into what is essentially a biphasic system, which consists

of a fibrillar phase enmeshed in an amorphous matrix phase (Preston,

1974). In fungi, however, the fibrillar elements are typically re-

stricted to the inner part of the wall, so that the outer portion con-

sists of amorphous materials only (Hunsley and Burnett, 1970). Recent

work in which Oomycete cell walls were disassembled with specific

enzymes has served to reveal the identity of some of the materials con-

tributing to the various wall phases.

In Pythium acanthicum Drechsler, for instance, the outer layer was

shown to be removable by laminarinase treatment, indicating a glucan

with a high proportion of B-1,3- and B-1,6-linkages. Inner fibrils re-

quired treatment with both laminarinase and cellulase for complete

dissolution, and the pattern of degradation indicated that these con-

sisted of B-1,3- and B-1,6-glucan surrounding a weakly crystalline

cellulose core (Sietsma et al., 1975). In Phytophthora parasitica Dastur,

the outer matrix was also removable by laminarinase, though the matrix

in the fibril layer required pronase treatment for removal. Micro-

fibrils were completely removable with cellulase (Hunsley and Burnett,

1970; Hunsley, 1973). Thus, even within the Oomycetes, there seems to

be room for considerable variation in cell wall architecture.







Cell Wall Modification During Growth


The importance of the cell wall in fungal morphogenesis is immense.

The form of almost every fungal cell, and thus the function it performs,

is determined by the structure of the cell wall. It is not surprising,

therefore, that changes in fungal form (i.e., morphogenesis) usually

involve cell wall modification in some way (Bartnicki-Garcia, 1968).

Growth, the fundamental expression of morphogenesis, is no exception.

It is unarguable that a growing fungal cell must increase the area

of its cell wall to accommodate the increase in volume of the cytoplasm;

and this must be accompanied by wall synthesis, if growth is to continue

indefinitely. Therefore, the prime concern of most models of fungal

cell growth is the explanation of the development of the cell wall.

As was seen to be the case with the fungal protoplast, the fungal

wall shows variations between growing and nongrowing regions of the same

hypha. Lateral walls are typically thicker and possess more easily re-

solved layers than do apical walls; walls of P. parasitica vary in thick-

ness from about 175 nm laterally to about 54 nm at the apex (Hunsley,

1973). Enzymatic disassembly reveals that there is only a small con-

tribution of the outer amorphous glucan layer to the apical wall, and

the apical fibrils are narrower and more loosely arranged than are

lateral fibrils (Hunsley and Burnett, 1970; Hunsley, 1973). Further

support for the gradation of materials is provided by reports of differ-

ential exposure of antigenic sites along a growing hypha (Fultz and

Sussman, 1966; Hunsley and Kay, 1976) and differential staining with

fluorescent brighteners (Gull and Trinci, 1974).

Because the apical wall is quite thin, one might expect this to be

one of the weakest points of the wall, and this can be demonstrated by








immersing the hypha in dilute acidic solutions, which causes the cells

to burst preferentially at the apex (Park and Robinson, 1966; Bartnicki-

Garcia and Lipmann, 1972). Theories of wall development account for the

differential osmotic stability by postulating that there exists in the

apex a delicate balance between wall synthesis and wall lysis, which

permits turgor-driven expansion of the plastic apical dome (Park and

Robinson, 1966; Bartnicki-Garcia, 1973). Lateral rigidification could

be accounted for by a predominance of synthesis in maturing regions

(Bartnicki-Garcia, 1973) or by the activation of a secondary synthetic

mechanism (Park and Robinson, 1966).

The requirement of turgor pressure for apical growth is demonstrated

by cessation of hyphal elongation in hypertonic solutions (Robertson,

1965; Park and Robinson, 1966) and prevention of hormone-induced branch-

ing in Achlya by water stress (Thomas, 1970). Although turgor may pro-

vide no more than an expansive force for deformation of the plastic

apex, contact between the plasma membrane and cell wall may also be

required for activity of wall synthesizing enzymes (Shore and Maclachlan,

1975; Maclachlan,1976) or for exocytosis (Robinson and Cummins, 1976).

For example, hormone-treated Achlya hyphae under water stress fail to

produce branches, and they accumulate intracellular cellulase, which is

not secreted (J. T. Mullins, unpublished data).


Synthetic Processes

The need for wall synthesis in apical growth is, as stated, patent.

The expectation of maximal activity in the apex stems both from poetic

necessity and from autoradiography, which confirms that maximal deposi-

tion of wall precursors is in the hyphal tip (Gooday, 1971; McMurrough







et al., 1971). Enzymes capable of synthesizing products with the prop-

erties of wall polymers have been demonstrated both in fungi and higher

plants.

For instance, a "soluble" cell fraction from Mucor rouxii (Calm.)

Wehmer has been shown to contain the enzyme chitinn synthetase" in a

zymogenic form that can be activated by a protease (Ruiz-Herrera and

Bartnicki-Garcia, 1974; Ruiz-Herrera et al., 1975). Such synthetases

have been found in several fungi of various groups (e.g., de Rousset-Hall

and Gooday, 1975; Duran et al., 1975; Mills and Cantino, 1978), but that

from M. rouxii is noteworthy because it is part of an enzyme complex

borne by minute cytoplasmic particles termed "chitosomes" (Bracker et al.,

1976). Apparently as a result of this association, not only is chitin

synthesized de novo, but the resulting chains are also assembled into

crystalline chitin microfibrils in vitro (Ruiz-Herrera and Bartnicki-

Garcia, 1974; Ruiz-Herrera et al., 1975).

This is, unfortunately, not yet the case with the so-called "cellu-

lose synthetases" and "glucan synthetases." These enzymes, though known

in fungi (Wang and Bartnicki-Garcia, 1966; Meyer et al., 1976, Fevre and

Dumas, 1977), have been best characterized from higher plants, where

they normally exhibit the ability to transfer a limited number of radio-

glucose molecules from nucleoside-diphosphoglucose to an endogenous

acceptor (for references, see Preston, 1974). Furthermore, enzyme prepa-

rations are usually crude, and the products in vitro are often hetero-

geneous and very much a function of assay conditions (Ordin and Hall,

1968). For these reasons, it may be better to refer to these enzymes as

"transferases," reserving the term "synthetases" until their roles in

vivo are more certainly known.







Lytic Processes

Evidence for the involvement of lytic forces in apical growth is

based in part on the already mentioned plasticity of the apical wall, as

demonstrated by osmotic rupture. The involvement of specific wall-

hydrolytic enzymes in distinct morphogenetic events of fungi and higher

plants is well documented and includes such phenomena as leaf abscission

(Lewis and Varner, 1970), ripening of fruits (Hall, 1964), autolysis of

Coprinus fruiting bodies (Iten and Matile, 1970), colonial growth

morphology of Neurospora (Mahadevan and Mahadkar, 1970), antheridiol-

induced branching in Achlya (Thomas and Mullins, 1967, 1969), and

fruiting in Schizophyllum (Wessels, 1966).

The involvement of specific hydrolases with ordinary vegetative

growth has been indicated in studies of both fungi and higher plants.

The enzyme cellulase is secreted by growing hyphae of A. ambisexualis

and accumulates in the medium (Thomas and Mullins, 1969). Its function

is apparently not nutritional because mycelia are incapable of using

cellulosic substrates as carbon sources (Thomas, 1966). And, in a re-

lated fungus, Saprolegnia monoica Pringsheim, the activity of intra-

cellular cellulase was found to be highest in regions of the colony

nearest the growing edge (Fevre, 1977).

A number of manipulations of growing cells elicit responses that can

be interpreted as the result of disturbances in the presumed balance

between synthesis and lysis. Exposure of pollen tubes to exogenous

hydrolases can enhance the rate of growth (Roggen and Stanley, 1969), and

Neurospora hyphae can be induced to branch by similar treatments (de Terra

and Tatum, 1961). Uptake of cholesterol by Pythium hyphae results in

decreased levels of wall hydrolases and corresponding morphological







aberrations (Sietsma and Haskins, 1968), and yeast cell walls lyse

upon exposure to 2-deoxyglucose, presumably because wall synthetases

are inhibited (Johnson, 1968).

The distribution of cellulase in the pea epicotyl also suggests a

relationship between vegetative growth and "the delicate balance." Al-

though synthetase activity is about equal in both growing and nongrowing

regions of the stem, the cellulase activity is high in actively growing

tissues and absent elsewhere (Maclachlan, 1976). Furthermore, auxin

treatment increases cellulase activity of decapitated epicotyls beyond

the pretreatment level (Fan and Machlachlan, 1966), whereas net synthetase

activity is merely maintained (Ray, 1973; Spencer et al., 1971).

Other cases in which high levels of wall hydrolases are associated

with growth include the budding stages of yeast, which exhibit high ac-

tivities of protein disulfide reductase (Nickerson and Falcone, 1959) and

-glucanase (Cortat et al., 1972).

An exact role for degradative enzymes in wall biosynthesis is not

readily apparent. In their simplest role, hydrolases might be involved

in mere wall loosening, which could permit the passive extension of the

wall by turgor. Loosening can be demonstrated by treating isolated cell

walls with exogenous polysaccharidases to increase their extensibility

(Olson et al., 1965). In higher plants, however, evidence seems to

argue against a role of polysaccharidases as regulators of extension

(Cleland, 1968; Ruesink, 1969). Instead, current interest is on the

potential for hydrogen ions to disrupt hydrogen bonds between cellulose

elementary fibrils and attached matrix polysaccharides (Keegstra et al.,

1973; Davies, 1973) or the polypeptide extension, which may represent








a covalently bonded, selectively cleavable linker between polysaccharide

chains (Lamport, 1974).

In the case of the enzyme cellulase, an indirect, though critical,

role in wall development may lie in its ability to generate free cel-

lulose chain ends by endohydrolysis. According to Maclachlan (1976),

the availability of cellulose "primer" ends is a rate limiting step in

cellulose biosynthesis, and the rate of synthesis should be increased by

pretreatment or cotreatment of walls with cellulase. The expected en-

hancement has been demonstrated using pea epicotyl segments pre-

incubated with either fungal cellulases (Maclachlan, 1976) or native pea

cellulases (Wong et al., 1977a). Proof that endogenous cellulases ac-

tually function this way in vivo is lacking, though the relative dis-

tributions of cellulase and synthetase activities in pea epicotyls is

consistent with this explanation (Maclachlan, 1976).


Involvement of Cytoplasmic Structures in Wall Formation


Morphological Evidence

Models assigning roles to cellular components in hyphal growth have

relied heavily on the evidence provided by electron microscopy of hyphal

tips. Although there are taxonomically related variations in the dis-

tribution of organelles in fungal tips, vesicles are present in hyphal

apices of all growing fungi (Grove and Bracker, 1970; Bartnicki-Garcia,

1973). Vesicles are also associated with the tips of basidial sterig-

mata (McLaughlin, 1973), buds of yeast (Moor, 1967; Sentandreu and

Northcote, 1969), germinating spores (Bracker, 1971), fungal rhizoids

(Barstow and Lovett, 1974), and apically growing structures of algae and







higher plants (Rosen et al., 1964; Bonnett and Newcomb, 1966; Ott

and Brown, 1974).

Vesicles often contain fibrous materials, which react with cyto-

chemical stains for carbohydrates (Heath et al., 1971; Dargent, 1975;

Meyer et al., 1976). Their apparent carbohydrate content and apical

location suggest that they may contribute their contents to the growing

cell wall (Grove et al., 1970). The origin and fate of wall vesicles

cannot be proven from fixed material, but there is sufficient morpho-

logical evidence to suggest that apical vesicles arise from dictyosomes

or their equivalents and secrete their contents into the cell wall

(Grove et al., 1970).

The involvement of dictyosomes in secretion is exhaustively docu-

mented in a number of plant and animal systems (e.g., Mollenhauer and

Morre, 1976; Palade, 1975). According to the theory of Palade (1975),

secretary products are sequestered within membrane-delimited spaces and

transferred through the endomembrane system (including the Golgi ap-

paratus) to the extracellular mileau by exocytosis. The current model

of apical growth as a secretary event involving exocytosis of Golgi-

derived apical vesicles is consistent with that theory. Evidence for

exocytosis is tenuous, of course, but support derives from electron

microscopical images that seem to show apical vesicles in a state of

fusion with the plasma membrane (Grove et al., 1970; Bracker, 1971);

these images resemble those seen in more rigorously documented examples

of exocytosis, such as mucocysts of Tetrahymena (Satir et al., 1973).

This would result in a release of vesicle contents into the wall and

would have the added virtue of contributing vesicle membrane to the

plasma membrane.







Theories proposed to explain the direction and motive force for

vesicle migration toward the apex have taken into account the already

mentioned gradients of enzyme activity and reductive potential observed

in hyphal tips; a result might be an electrochemical gradient sufficient

to account for vesicle migration by "electrophoresis" (Bartnicki-Garcia,

1973). An explanation with more experimental support requires the action

of contractile cytoplasmic microfilaments; cytochalasin B, which dis-

rupts microfilaments, prevents apical growth in root hairs (Franke

et al., 1972) and pollen tubes (Mascarenhas and Lafountain, 1972).


Biochemical Evidence

On morphological and theoretical grounds, vesicles associated with

a number of apically growing systems have been suggested to contain a

variety of materials that are consistent with a role in wall synthesis.

Among these are carbohydrate wall precursors and presynthesized wall

components (Larson, 1965; Seivers, 1967; McClure et al., 1968; Grove

et al., 1970; Bartnicki-Garcia, 1973), wall synthesizing enzymes (Grove

et al., 1970; Bartnicki-Garcia, 1973), and wall softening enzymes (Moor,

1967; Girbardt, 1969; Grove et al., 1970; Bartnicki-Garcia, 1973). In

addition, other enzyme activities have been indicated by in situ cyto-

chemical tests (Dargent, 1975; Meyer et al., 1976). However, electron

microscopical evidence can only be confirmed by actual isolation and

biochemical analysis of apical vesicles.


Filamentous fungi

There are reports of attempts to isolate apical vesicles from three

fungi. The first, in Gilbertella persicaria (Eddy) Hesseltine, is un-

substantiated by published data and reports the recovery of subcellular







particles that contain polysaccharides composed of sugars characteris-

tically found in the cell wall (Grove et al., 1972). In Phytophthora

palmivora Butler (Meyer et al., 1976), it was shown that "UDPG transfer-

ase" activity is associated with cell walls and with a membrane fraction

that may contain vesicles derived from the endoplasmic reticulum (ER).

Exo--1,3-glucanase is generally associated with a second membrane

fraction which may contain Golgi vesicles. In Saprolegnia monoica

(Fevre and Dumas, 1977), "glucan synthetase" activity is associated with

a crude "wall" fraction and with membrane fraction that apparently

contain both dictyosome cisternae and unidentified membranes. These

fractions also contained B-1,3-glucanase and cellulase activities (Fevre,

1977). Convincing correlations between these materials and distinct

classes of subcellular particles is lacking in reports on all three

fungi, and biochemical support for the involvement of specific organelles

in tip growth is, at this time, largely by analogy to other, better

characterized systems in yeast and higher plants. These will now be

discussed.


Pollen tubes

A higher plant cell with apical growth, which has been biochemically

investigated, is the pollen tube. Here, work has largely been re-

stricted to identification and labeling of carbohydrates in walls and

membrane fractions. Membranes obtained from Lilium pollen contain

carbohydrates characteristic of the wall matrix (Van Der Woude et al.,

1971), and the kinetics of labeling with radioactive precursors indi-

cates that this material eventually contributes to the tube wall (Morre

and Van Der Woude, 1974). Morphological evidence indicates that these








membranes are derived from dictyosomes and/or endoplasmic reticulum

(Van Der Woude and Morr6, 1968; Van Der Woude et al., 1971).

A noteworthy report is that of Engels (1973, 1974) who isolated

from petunia pollen a fraction of membranes that he identified as Golgi

vesicles using morphological criteria. On the basis of X-ray diffrac-

tion spectra, one component of the carbohydrate in these vesicles was

identified as a mixture of cellulose I and cellulose II (Engels, 1974).

This is the only report of true cellulose in cytoplasmic particles of a

higher plant. Despite numerous investigations in other organisms, the

only other report of in vivo cellulose synthesis in an intracellular

compartment is a report of the unusual wall of the alga Pleurochrysis

(Brown et al., 1970). Here the wall is composed of overlapping cellu-

losic scales, which are apparently preassembled entirely within

dictyosomes and exported in vesicles to the cell surface.


Yeast cells

In yeast, investigations have focused primarily on the assembly and

modification of the mannoprotein component of the wall matrix. Evi-

dence from the dimorphic fungus M. rouxii has implicated this glyco-

protein in the control of yeast morphogenesis, and the enzyme disulfide

reductase is postulated to soften the wall by cleaving disulfide bridges

between mannoprotein molecules (Bartnicki-Garcia and Nickerson, 1962).

The critical components of the yeast endomembrane system are the

ER, cytoplasmic vesicles, and the plasma membrane; yeasts lack dictyo-

somes, and the vesicles are believed to be produced directly by the ER

(Moor, 1967). Evidence indicates that all these compartments are in-

volved in the assembly of mannoprotein and in subsequent modification of







these and other wall components (Matile et al., 1971; Cortat et al.,

1972, 1973). Autoradiography shows initial mannose incorporation to be

intracellular (Kosinova et al., 1974), and the results of cell frac-

tionations indicate that the mannan chain is assembled sequentially,

with the first sugar incorporated in the ER and subsequent sugars in-

corporated as the complex moves through the vesicles to the plasma

membrane (Lehle et al., 1977). Other materials which may be involved in

wall metabolism have also been detected in yeast cell particles, in-

cluding B-1,3-glucanase, which is present in ER, vesicle, and plasma

membrane fractions (Matile et al., 1971; Cortat et al., 1972). Wall

fibril synthesis appears to be a function of the cell surface and is

regulated independently of matrix synthesis, which can be preferentially

uncoupled by cycloheximide (Necas, 1971).

Formation of the chitinous yeast septum appears to involve at least

two cellular compartments. Chitin synthetase is attached to the inner

side of the plasma membrane as an inactive zymogen (Duran et al., 1975).

The septum begins to form after a proteolytic activator is delivered in

vesicles to the appropriate sites on the plasma membrane (Cabib and

Farkas, 1971).


Higher plants

The contribution of internal membranes to cell wall synthesis is

perhaps best demonstrated in the auxin-stimulated pea epicotyl. The

work of Peter Ray and collaborators has demonstrated the role of the Golgi

apparatus in the synthesis of matrix materials and in their subsequent

transfer to the wall (Ray et al., 1969, 1976; Robinson and Cummins, 1976;

Robinson et al., 1976). Polysaccharides labeled in vivo with radio-

glucose are associated both with dictyosome cisternae and dictyosome








vesicles. Intracellular synthetase activity is associated with the

cisternae only (Ray et al., 1976). The in vivo labeled products con-

tain polysaccharides with linkages characteristic of the wall matrix,

and only 3% to 8% of the linkages are B-1,4-glucan. Pulse-chase ex-

periments demonstrate that labeled materials are transferred from

dictyosome cisternae to vesicles and ultimately to the wall (Robinson

et al., 1976). Further work by Maclachlan and coworkers (Shore and

Maclachlan, 1975; Shore et al., 1975) supports the concept of matrix

formation by dictyosomes, as does that by Harris and Northcote (1971),

who worked with pea roots.

The site of microfibril (cellulose) synthesis in higher plants has

not been biochemically defined, but evidence continues to indicate the

plasma membrane-cell wall interface (Preston, 1974). Synthetase ac-

tivity capable of generating B-1,4-glucan linkages can be detected in

isolated dictyosomes (Ray et al., 1969), smooth ER (Shore and Maclachlan,

1975), and plasma membrane fractions (Van Der Woude et al., 1974), but

this can account for no more than about 5% of the in vivo rate of

cellulose synthesis (Maclachlan, 1976). Furthermore, since most re-

ports indicate that, in vivo, only matrix polysaccharides are synthesized

intracellularly, cellulose synthetase activity in these membranes is

interpreted either as activity of enzymes in transit to the wall (Shore

and Maclachlan, 1975) or as activity required to produce the B-1,4

linkages found in certain matrix components (Ray et al., 1976).

In higher plants, cellulose has not been reported within vegetative

cells using biochemical criteria, which suggests its assembly at the

cell surface (but cf. Brown et al., 1970; Engels, 1974). In addition,

autoradiographic data from sycamore seedlings indicate that, under







conditions of maximal cellulose synthesis and minimal matrix synthesis,

most activity is associated with the plasma membrane-cell wall interface

(Wooding, 1968). That the interface itself is critical to synthesis is

demonstrated by the reduction of incorporation upon plasmolysis

(Maclachlan, 1976) or physical disruption (Shore and Maclachlan, 1975).

Current attention has been directed toward plasma membrane particle

complexes postulated by Preston (1974) and which may have been demon-

strated recently in corn (Mueller et al., 1976).

Cellulases, which have been implicated in wall synthesis (Wong

et al., 1977a), are also associated with subcellular compartments, in-

cluding the cell wall and endoplasmic reticulum in peas (Bal et al.,

1976) and plasma membrane in kidney beans (Koehler et al., 1976). Asso-

ciations with dictyosomes have not been reported.



In summary, models of hyphal tip growth employ the coordinated

activities of wall degradation and wall synthesis. These processes

would require specific enzymes, wall precursors, and a mechanism for

their simultaneous delivery to the apex. Morphological studies indicate

that Golgi-derived apical vesicles are involved. Furthermore, evidence

from other systems in which cytoplasmic vesicles play a role in wall

metabolism lends support to models assigning a similar role to vesicles

in hyphal growth.














MATERIALS AND METHODS


General Culture Methods


An isolate of Achlya ambisexualis Raper, strain E87, was provided

by Dr. J. T. Mullins; derivation of the strain is described by

Barksdale (1960). Stock cultures were maintained on YPSS agar slants

(Emerson, 1941) at 5 C.

Mycelia were grown in a defined liquid medium (DLM) (Mullins and

Barksdale, 1965) of the composition given in Table 1. The inoculum was

obtained by a modification of the method of Griffin and Breuker (1969).

A small amount of mycelium was transferred from the YPSS slant to a

petri plate containing "enriched medium" (Kane, 1971), which contains

the same ingredients as DLM, except monosodium-Z-glutamate, 3.0 mM;

D-glucose, 77.7 mM; casein hydrolysate, 0.15% w/v; agar, 2.5% w/v.

After two days' growth at 24 C, a plug was removed from the center of

the colony with a #10 cork borer and cut into about 9 pieces. These

were washed in 50 ml of sterile 0.5 mM CaC12 in a 250 ml flask for 2 hr

at 24 C on a shaker set at 100 rpm. The liquid was decanted and re-

placed with another 50 ml volume of sterile 0.5 mM CaC12, and the flask

was returned to the shaker. After 15 to 20 hours, the sample was re-

moved aseptically, and the zoospores and spore cysts were counted on a

hemacytometer. A volume of liquid containing 200,000 spores was trans-

ferred to 200 ml of DLM in a 500 ml Erlenmeyer flask. This was in-

cubated under the same conditions as those used to obtain the spore














TABLE 1. The composition of Defined Liquid Medium (DLM) (modified
from Mullins and Barksdale, 1965)



Monosodium-z-glutamate 2.4 mM

D-glucose 11.1 mM

Tris-SO4 buffer, pH 6.9 10.0 mM

k-methionine 0.1 mM

KC1 2.0 mM

MgSO4 0.5 mM

CaC12 0.5 mM

HEDTA 72.0 pM

KH2P04 1.5 WM

Fe(NH4)2(S04)2 36.0 iM

ZnSO4 15.0 pM

MnSO4 9.0 pM

Sulfosalicylic acid 46.0 pM








inoculum. After a 24 hr lag, mycelial fresh weight (FW) increased to

about 8 g per flask at about 72 hr. Accordingly, cultures were har-

vested at about 48 hr, which represented the midpoint in the growth

curve and yielded about 5 g FW per flask. Harvest was accomplished by

pouring the contents of each flask into a funnel lined with miracloth

(Chicopee Mills, Inc.).

Cell Homogenization


Disruption was accomplished by grinding mycelia in a mortar with

5 g of acid-washed sea sand. The amount of mycelium and the composi-

tion of the homogenizing solution varied with certain experiments, and

these details will be specified with the descriptions of the individual

experiments. Generally, the homogenizing solution contained 0.03 M

tris.HC1 buffer (pH 7.6 measured at RT) and 30% w/w sucrose. In certain

experiments, 15 mM dithiothreitol (DTT) and/or 0.3% bovine serum albumin

(BSA) were included. Mycelia were ground for 30 sec at 5 C, and the

homogenate was filtered through miracloth. The retained material was

rehomogenized for 15 sec in a small volume of homogenizing solution

which had been diluted until the sucrose concentration was 10% w/w. The

second homogenate was refiltered on the same piece of miracloth, and the

retained material was discarded.

Centrifugations


Isolation of cell particles was affected using either of two

centrifuges. The Sorvall RC-2B high speed centrifuge with an SS-34

rotor was used for all differential centrifugations of homogenates. The

Beckman L2-65B preparative ultracentrifuge with an SW 27 or SW 27.1







horizontal rotor was used for all gradient centrifugations, and sedi-

mentation of gradient fractions was achieved with the 65 rotor.

In addition, the Sorvall GSA rotor was used in the recovery of

ethanol-insoluble cellulase from culture filtrates.


Assays


Unless otherwise stated, all substrate biochemicals were obtained

from Sigma Chemical Co., St. Louis, Mo. All pH's were measured at room

temperature with a "tris" electrode (Sigma Chemical Co.). All spectro-

photometry was performed with a Gilford model 240 spectrophotometer.


Bio-Rad Assay for Protein (Bio-Rad Technical Bulletin, 1977)

Five milliliters of Bio-Rad dye reagent were added to 50-200 pl

of protein sample and mixed. After 10 min, absorbance at 595 nm

was read. Standards were made with Bovine Serum Albumin (BSA).


Anthrone Test for Carbohydrate (Herbert et al., 1971)

Anthrone reagent:

anthrone 200.0 mg

absolute ethanol 5.0 ml

75% v/v H2SO4 95.0 ml


Five milliliters of cold (4 C) anthrone reagent were added to

1.0 ml of cold sample. The mixture was heated at 100 C for

10 min, and absorbance was read at 625 nm. Standards were made

with glucose.








Phenol Test for Carbohydrate (Herbert et al., 1971)

Reagents:

5% w/v phenol

concentrated H2SO4


One milliliter of phenol and 5.0 ml of H2SO4 were added to 1.0 ml

of aqueous sample. After cooling, absorbance was read at

488 nm. Standards were made with glucose.


Ferrous Sulfate-Ammonium Molybdate Assay for Inorganic Phosphorus
(modified from Taussky and Shorr, 1953)

Reagent:

FeSO4*7H20 5.0 g

(NH4)6MoO24.4H20 (10 mg/ml in 10 N H2S04) 10.0 ml

distilled water 85.0 ml


Two milliliters of reagent were added to 250 pl of sample. Absorb-

ance at 710 nm was read after 10 min. Standards were made with

KH2PO4.

Acid Phosphatase (EC 3.2.3.2) (modified from Ray et al., 1969)

Substrate:
MgC12 2.2 vmol

p-nitrophenyl phosphate 0.8 mg

80 mM citrate buffer, pH 5.0 1.0 ml

Fifty to one hundred microliters of sample were added to 50 pl of
substrate and incubated for 30 min at 37 C. The reaction was

terminated by adding 2.0 ml of 0.2 M Na2C03, and absorbance was
read at 410 nm. A molar extinction coefficient for p-nitrophenol
of 18,000 was assumed (Meyer, 1976).








Adenosine Triphosphatase (ATPase) (EC 3.6.1.3) (modified from
Marriott, 1975)

Substrate:

tris-ATP 1.5 pmol

MgC12 5.0 Pmol

16.7 mM tris-HCl buffer, pH 7.2 1.0 ml


Fifty to one hundred microliters of sample were added to 500 ul of

substrate. Two hundred fifty microliters were removed and

assayed for P.. The remainder was incubated at 37 C for 4 hr,

then 250 pl were removed and assayed for P. released.
1

Alkaline Phosphatase (EC 3.1.3.1)

Substrate:

MgC12 2.0 pmol

p-nitrophenyl phosphate 0.8 mg

50 mM tris-HCl buffer, pH 9.0 1.0 ml


One hundred microliters of sample were added to 500 pl of sub-

strate. The mixture was incubated 4 hr at 37 C, and 2.0 ml of

0.2 M Na2CO3 were added to dilute the mixture to a volume readable

in the spectrophotometer. Absorbance was read at 410 nm.


Cellulase (EC 3.2.1.4) (modified from Bell et al., 1955)

Substrate:

carboxymethyl cellulose (type 7Mf, Hercules 12.0 g
Powder Co.)

merthiolate 0.5 g

0.018 M sodium citrate buffer, pH 5.0 1.0 1








One milliliter of sample was added to a size "300" Ostwald-Fenske

viscometer tube containing 5.0 ml of substrate, which had

equilibrated at 30 C. After mixing, the flow time was measured

at TO and after various intervals, and the difference in flow

time was determined.


Cytochrome Oxidase (EC 1.9.3.1) (modified from Hodges and Leonard, 1974)

Substrate:

cytochrome c (Sigma type III, from horse heart, 11.0 mg
oxidized form)

50 mM potassium phosphate buffer, pH 7.5 60.0 ml

Cytochrome c solution is reduced chemically with sodium dithionite.


Twenty to one hundred microliters of sample were added to 2.0 ml of

substrate in a spectrophotometer cuvette. The change in absorbance

at 550 nm was monitored during the phase of linear change. A molar

extinction coefficient of 18,500 was assumed for cytochrome c.


Glucose-6-phosphatase (EC 3.1.3.9) (modified from Hubscher and West, 1965)

Substrate:

EDTA 4.0 pmol

KF 2.0 pmol

glucose-6-phosphate 6.8 mg

0.4 M sodium "PIPES" buffer, pH 6.5 1.0 ml


Fifty to one hundred microliters of sample were added to 500 pl of

substrate, and 250 pl were removed for assay of P.. The remainder

was incubated for 4 hr at 37 C, and 250 pl were removed for assay

of P. released.
1







B-glucosidase (EC 3.2.1.21) (modified from Parish, 1975)

Substrate:

p-nitrophenyl-6-D-glucoside 0.5 mg

0.1 M sodium citrate buffer, pH 5.0 1.0 ml


Fifty to one hundred microliters of sample were added to 500 pl of

substrate. The mixture was incubated for 30 min at 37 C, and the

reaction was terminated with 2.0 ml of 0.2 M Na2CO3. Absorbance

was read at 410 nm, and a molar extinction coefficient for p-

nitrophenol of 18,000 was assumed.


Inosine Diphosphatase (IDPase) (modified from Ray et al., 1969; Shore
and Maclachlan, 1975)

Substrate:

MgC12 1.0 pmol

inosine diphosphate 1.4 mg

0.1 M tris.HC1 buffer, pH 7.5 1.0 ml


One hundred microliters of sample were added to 500 p1 of sub-

strate, and 250 pl were removed for assay of P.. After incubation

of the remainder for 4 hr at 37 C, 250 p1 were removed for assay

of P. released.


UDPG transferase (EC 2.4.1.12) (modified from Ray et al., 1969; Shore
and Maclachlan, 1975)

Substrate:

Solution I: UDPG 925 pg

1C-UDPG (250 pCi/pmol in 1.0 ml, 100 1l
New England Nuclear)

distilled water 2.0 ml








Solution II: MgC12*6H20 68 mg

cellobiose 51 mg

dithiothreitol (DTT) 8.0 mg

0.2 M sodium phosphate buffer, pH 5.8 10.0 mi


One hundred microliters of solution I, 100 pl of solution II, and

100 pl of sample were added to a 15 ml conical glass centrifuge

tube. (The final concentration of reagents was UDPG, 0.242 mM;

cellobiose, 5 mM; MgC12, 11 mM, DTT, 1.7 mM; buffer, 67 mM. The

reaction vessel contained 7.27 x 10-8 moles of UDPG with 119 nCi

of radioactivity.) After incubation at room temperature for 20 min,

the reaction was terminated by adding 5.0 ml of 70%.v/v ethanol.

About 30 mg of powdered Whatman cellulose were added, and the tube

was centrifuged in an IEC clinical centrifuge at about 1,000 x g

for 5 min. The supernatant was discarded, and the sediment was

washed three more times in 70% ethanol (15.0 ml total volume).

Excess ethanol was permitted to evaporate from the final pellet,

which was then resuspended in 3.0 ml of distilled water for liquid

scintillation counting.


Liquid Scintillation Counting


Radioactive materials in 3.0 ml of water were transferred to a glass

scintillation vial, and 5.0 ml of PCS scintillation fluid (Amersham

Searle Co.) were added. The sample was shaken to form a gel and to

disperse any solid materials, and radioactivity in the vials was meas-

ured with a Packard Tri-Carb Liquid Scintillation Spectrometer, model

3385. Results in counts-per-minute were converted to decays-per-minute







(dpm) using the channels ratio method, and pCi of radioactivity present

was calculated, assuming 2.22 x 106 dpm/pCi. For some experiments,

substrate incorporation (in moles) was calculated by isotope dilution.


Statistical Methods


When sufficient data permitted, values were reported with the as-

sociated standard deviation, which was calculated as the square root of

the sample variance. When possible, comparisons between two samples

are accompanied by a figure representing the degree of confidence in

their statistical difference. This was calculated by a "two-tail"

Student's "t" test. (All methods are from Runyon and Haber, 1971).


Electron Microscopy


Hyphae or subcellular fractions (obtained by methods to be de-

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

glutaraldehyde in 0.05 M sodium cacodylate buffer, pH 7.2. After rins-

ing in several changes of buffer, the material was postfixed in 1% w/v

Os04 in 0.05 M cacodylate buffer, pH 7.2, for 30 min at room tempera-

ture. Samples were again washed several times in buffer and dehydrated

in a graded series of ethanol washes, terminating in absolute acetone.

Material was infiltrated with an epoxy embedding medium (Table 2) and

polymerized at 60 C for 24 to 48 hr in a flat embedding mold. Embedded

samples were mounted on metal microtome stubs and sectioned on a Sorvall

Porter-Blum MT-2 ultramicrotome. Thin sections were poststained for

5 min with lead citrate (Reynolds, 1963) or with 1% BaMnO3 before ex-

amination with a Hitachi HU-11C or HU-11E electron microscope.

On some samples, cytochemical tests were performed, and this

necessitated various modifications of the above scheme.








TABLE 2. Embedding medium for electron microscopy


Reagent Manufacturer Amount

Epon 812 Polyscience, Inc. 12.5 g

Araldite 6005 R. P. Cargille Lab., Inc. 11.5 g

DDSA Tousimis Research Corp. 25.5 g

BDMA Ladd Research Ind., Inc. 0.14 ml/10 g*


* BDMA is added just before use.



Cytochemical Tests


Phosphotungstic Acid-Chromic Acid (PTA-Cr03) Stain (Roland et al., 1972)


Staining reagent:

phosphotungstic acid

Cr03

distilled water


0.1 g

1.0 g

10.0 ml


Thin sections carried in polyethylene rings were oxidized by

flotation on 1% w/w periodic acid for 30 min at room temperature.

After washing by flotation on distilled water, sections were

floated on PTA-Cr03 staining reagent for 5 min at RT and rinsed on

distilled water. Sections were mounted on grids for viewing

without poststain.







Periodic Acid-Silver Methenamine (PASM) Stain (Martino and Zamboni, 1967)

Staining reagent:

hexamethylene tetramine 90 mg

AgNO3 10 mg

4 mM sodium borate buffer, pH 9.0 10 mg


Thin sections in polyethylene rings were preoxidized in acidic

H202 (15% H202 in 2% HC1) for 30-60 min at RT to remove osmium

stain. After rinsing, sections were further oxidized in 1% w/w

periodic acid for 15-30 min at RT and rinsed. Staining was

performed by incubating oxidized sections for up to 2 hr in

staining reagent at 60 C, followed by a distilled water rinse.

After incubating in 1% v/v Kodak photographic fixer for 5 min at

RT, sections were rinsed and mounted on grids for viewing.


Cytochemical Localization of IDPase (modified from Novikoff and
Goldfischer, 1961)

Substrate:

IDP 1.0 mg

MnC12 1.0 mg

PbNO3 12.0 mg

0.4 M tris.HCl buffer, pH 7.2 10.0 ml


Hyphae were fixed by the standard method and washed in cacodylate

buffer, followed by 0.5 M tris-HCl buffer, pH 7.2. The material

was then incubated in substrate solution for 60 min at 37 C and

rinsed in tris buffer. Samples were postfixed and embedded as

before, and thin sections were stained with lead citrate before

viewing.








Cytochemical Localization of Alkaline Phosphatase (Hugon and Borgers,
1968)

Substrate:

Na-B-glycerophosphate 25.0 mg

PbNO3 13.0 mg

0.04 M tris-maleate buffer, pH 9.0 10.0 ml


Hyphae were fixed and rinsed in cacodylate buffer, followed by

0.05 M tris-maleate buffer, pH 9.0. Incubation in substrate solu-

tion followed for 30-60 min at 37 C; and hyphae were washed, post-

fixed, and dehydrated in the standard manner. Thin sections were

viewed with or without poststaining.


Cytochemical Localization of Cellulase (modified from Bal, 1972)

Substrate:

carboxymethyl cellulose (type 7 MF) 1.0 mg

0.018 M citrate buffer, pH 5.0 10.0 ml


Hyphae were fixed for 1 hr on ice and washed with cold buffer

overnight. They were next transferred directly to substrate

solution for 10 min to 2 hr at RT. After incubation, hyphae were

transferred to 80 C Benedict's solution (Bauer et al., 1968) for

5 min, washed in distilled water, and postfixed in osmium tetroxide

as described earlier. Subsequent treatment adhered to standard

techniques.


Cytochemical Localization of Acid Phosphatase (Gomori, 1952)

Substrate:

Na-B-glycerophosphate 30.0 mg

0.05 M acetate buffer, pH 5.0 11.0 ml








12% w/v lead nitrate 0.1 ml

sucrose 0.8 g


Hyphae were fixed by the standard method; however, 7.5% sucrose

was included in the fixation medium and in all buffer washes

because hyphal tips tend to burst in dilute acidic solutions

(Park and Robinson, 1966). After fixation, hyphae were washed

in cacodylate buffer, followed by 0.05 M acetate buffer, pH 5.0,

and incubated in substrate solution for 45 min at 37 C. Follow-

ing a rinse in acetate buffer, samples were postfixed in OsO4

and embedded as described previously. Thin sections were ob-

served after poststaining with lead citrate.














ELECTRON MICROSCOPY OF HYPHAL APICES


It is the aim of this research to study the location of the enzyme

cellulase (Cx) within hyphae of A. ambisexualis and to determine what

evidence, if any, exists to implicate this enzyme in wall morpho-

genesis during vegetative growth. In this section, growing hyphae will

be examined with emphasis being placed on the involvement of subcellular

structures in wall formation.


The Cytoplasmic Organization of Achlya Hyphal Apices


Growing fungal hyphae extend at their tips, and this process is

reflected in the organization of the apical region (Grove and Bracker,

1970). Hyphal apices of A. ambisexualis were examined electron micro-

scopically, in order to compare the apical organization of this fungus

with that of other fungi.


Methods

Fungal hyphae were obtained in either of two ways. In the first

method, #2 cork borer plugs were removed from the edge of a 48 hr old

Enriched Agar Medium colony and placed in a 5 cm petri plate which con-

tained about a 5 mm deep layer of DLM. The culture was incubated at

RT for 8-12 hr, after which time hyphae had grown out from the agar

plugs to a distance of 2-5 mm. Agar plugs bearing hyphae were then

processed for electron microscopy according to standard methods.







In the second method, a small piece of mycelium was transferred

from a YPSS agar slant to a 5 cm petri plate containing about a 2 mm

deep layer of Enriched Agar Medium, and the culture was incubated at RT

for 12-24 hr. The entire layer of agar with submerged hyphae was then

processed for electron microscopy, as previously described.

These two methods produced robust, straight hyphae, which are

vastly more suitable for electron microscopy than are the narrow, con-

torted hyphae that are obtained from shaken liquid cultures. As a result,

hyphae can be conveniently manipulated with minimal risk of damage, and

embedded specimens can be easily oriented for longitudinal sectioning.


Observations

The apical regions of Achlya hyphae, like those of other fungi, can

be divided into the extensive subapical zone, which contains an abundance

of organelles and no central vacuole (Fig. 1), and the apex-proper,

which is characterized by its population of small apical vesicles

(Fig. 2). Walls in thin sections taken from both zones are not con-

trasted by poststaining with lead citrate (e.g., Figs. 1,6), but post-

staining with BaMnO4 reveals their presence (e.g., Figs. 2,8). While the

width of the wall varies greatly in each region, measurements indicate

a width of 178 100 nm for older, lateral walls and a width of 59

20 nm at the very apex. This agrees very well with measurements of

the cell walls of Phytophthora hyphae, which are 175 nm wide laterally

and 54 nm wide at the apex (Hunsley, 1973).

Organelles in the subapical zone include nuclei, mitochondria,

endoplasmic reticulum (ER), dictyosomes (Golgi apparatus), and numerous

cytoplasmic vesicles. These vesicles are of various sizes, shapes, and



















Fig. 1. Longitudinal section of the subapical region of an A.
ambisexualis hypha. M, mitochondrion; N, nucleus. x 5,500.
All magnifications are approximate.

Fig. 2. Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the many cytoplasmic vesicles (V) in the
region. The section was poststained with BaMnO4 to reveal the
cell wall (CW). M, mitochondrion. x 22,000.

Fig. 3. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows a dictyosome (0) surrounded by
cytoplasmic vesicles (V). N, nucleus. x 48,000.

Fig. 4. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows a dictyosome (D) bearing an
incipient fibrous vesicle (arrow). M, mitochondrion; N,
nucleus; V, vesicle. x 43,000.

Fig. 5. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows details of the cytoplasmic
vesicles (V). Arrows indicate the central fiber-free zone of
two vesicles. x 90,000.

Fig. 6. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows cytoplasmic vesicles (V)
beneath the cell wall (CW). The wall is not contrasted with
the lead citrate poststain. x 75,000.







N1 A5 4 : ".
. t t .."V ,




, : . A t IFt Nk1",A ', i

44.
1 .. I ... '


N M


EE. N .... .
, -- ,. ,- ,





- %.E.' ..u .
* .... r ,i
^ 1<'^ ,, .... ..,,. f" l ...





" I iS^^ N







contents; and many are located near dictyosomes (Figs. 3,4,9). Indeed,

dictyosomes bear what appear to be incipient vesicles and may be one

source of the many cytoplasmic vesicles (Figs. 4,9). There is a virtual

continuum of vesicle sizes, with diameters commonly extending from about

40 nm to about 160 nm. The largest of these vesicles (those with diame-

ters above about 120 nm) exhibit a characteristic morphology in thin

section, which consists of a fibrous matrix found mostly in the peri-

pheral region of the vesicle's interior; the innermost region often

appears free of fibrous material (Fig. 5).

In the subapical region, vesicles are commonly found just beneath

the cell wall (Fig. 6). These vesicless," however, are not always the

roughly spherical structures implied by the term. Tangential sections

of hyphae reveal that some of these structures are quite elongated and

may be better described as submural tubules (Fig. 7). These correspond

in morphology to the large vesicles with fibrous contents (Fig. 5).

Most such structures, though, appear to be legitimate vesicles, and the

possibility exists that submural vesicles and submural tubules are dis-

tinct, but related, structures. Perhaps, one gives rise to the other;

structures intermediate between tubules and vesicles (Fig. 8) may

represent fusion of vesicles or vesicle production. Some vesicles that

are attached to dictyosomes (Fig. 4) have a peripheral fibrous matrix

like that of large fibrous cytoplasmic vesicles, suggesting that this may

be their true origin. But the resolution of questions like this is very

difficult using thin-sectioned material.

The smaller vesicles (those with a diameter less than about 120 nm)

may have contents which appear either fibrous or featureless (Figs. 5,6),

but in most, the fibrous nature of the contents is difficult to discern.




















Fig. 7. Tangential longitudinal section of the subapical region of
an A. ambisexualis hypha. Arrows indicate elongated con-
figurations of "submural tubules." The cell wall (CW) is
not contrasted by the lead citrate poststain. x 24,000.

Fig. 8. Longitudinal section of the subapical region of an A.
ambisexualis hypha. The arrow indicates a structure that may
be intermediate between a tubule and vesicles. The cell wall
(CW) is contrasted with BaMnO4. M, mitochondrion. x 30,000

Fig. 9. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows coated vesicles (CVT in the
vicinity of and attached to dictyosomes (D). M, mitochondrion;
N, nucleus. x 48,000.

Fig. 10. Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the lack of deposition of cellulase reaction
product in the hypha. CW, cell wall; M, mitochondrion; V,
vesicle. x 24,000.

Fig. 11. Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the distribution of acid phosphatase-
positive and acid phosphatase-negative vesicles in the apex.
CW, cell wall; M, mitochondrion; V. vesicle, x 22,000.

Fig. 12. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows the deposition of acid
phosphatase reaction product in some of the cytoplasmic
vesicles, but not in others. M, mitochondrion; V, vesicle.
x 55,000.




41




cw I ',A w
rA


Al e


t.. I-















1 .A |
,4






S e... . , .. .
.CW



ab. *. .) S M
r .I

M






* // .hV

' ..
rCm
IL1


--CW
,. :::i ::.


'Id4







If the contents are fibrous, an inner "fiber-free" zone is usually not

present. Another kind of vesicle that is present is the coated vesicle,

which has a diameter of 85 5 nm; this is apparently produced by

dictyosomes (Fig. 9). These are the same size as the coated vesicles

in the cytoplasm of radish root hairs (Bonnett and Newcomb, 1966).

The apical region is populated for the most part by vesicles,

though mitochondria intrude almost to the apical wall itself (Fig. 2).

These vesicles seem to be identical to the vesicles already described,

though elongated profiles reminiscent of "submural tubules" are not

commonly seen in the apex, and most apical vesicles would seem to be

true spheres. No sections revealed images that could be interpreted

as representing stages in the fusion between vesicles or the fusion

between a vesicle and the plasma membrane.


Cytochemical Localization of Enzymes and Other
Materials in Hyphal Apices


Methods

Cytochemical tests were performed using the techniques already

described under Materials and Methods.


Results

Cellulase. The attempt to use this technique was apparently

unsuccessful; no reaction product was observed in any part of the hyphae

examined (Fig. 10).

Acid phosphatase. The reaction product of the acid phosphatase

reaction is located in some 20-30% of the apical vesicles present in

the terminal 3 Pm of the apex (Fig. 11), and the average size of the

reactive vesicles is 143 19 nm. The proportion of reactive vesicles








in the overall vesicle population increases with distance from the

apex. The reaction product is preferentially associated with the

fibrous contents of those vesicles that stain, but nearby vesicles of

similar morphology may not stain at all (Fig. 12).

Stain in dictysomes can also be seen (Fig. 13), and it is re-

stricted to a single cisterna in those dictyosomes that react; reaction

product is also found in nearby small vesicles.


IDPase. Like acid phosphatase, IDPase activity is found in associa-

tion with the fibrous material of large cytoplasmic vesicles, which

measure 142 18 nm in diameter (Fig. 14). Similarly, not all such

structures react; but unlike acid phosphatase, IDPase is found in a

much higher proportion of apical vesicles (Fig. 15).

What may be an as yet unrecognized class of apical vesicles is

represented by an intensely reactive vesicle in which the reaction

product is deposited both within and without the membrane (Fig. 14).

The outside diameter of these structures is 89 16 nm. In size and

the presence of material outside the vesicle membrane, these IDPase-

positive vesicles resemble the dictyosome-derived coated vesicles

(Fig. 9).

Dictyosomes are unreactive, as are attached incipient vesicles

(Fig. 16).


Alkaline phosphatase. Alkaline phosphatase reaction product is not

associated with any cellular structure, including dictyosomes (Fig. 17)

and cytoplasmic vesicles (Fig. 18). The fine electron-dense deposits

associated with the cytoplasmic vesicles and dictyosomes are also

present in control material from which B-glycerophosphate was omitted
















Fig. 13. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows the deposition of acid
phosphatase reaction product in a single cisterna of a
dictyosome (D). M, mitochondrion; N, nucleus. x 54,000.

Fig. 14. Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the deposition of IDPase reaction product
in some of the cytoplasmic vesicles, but not in others. Ar-
rows indicate smaller vesicles, in which the reaction product
is deposited both inside and outside the membrane. V,
vesicle. x 49,000.

Fig. 15. Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the distribution of IDPase-positive and
IDPase-negative vesicles in the apex. M, mitochondrion; V,
vesicle, x 19,000.

Fig. 16. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows the lack of IDPase reaction
product in a dictyosome (D). V, vesicle, x 43,000.

Fig. 17. Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows the lack of alkaline phos-
phatase reaction product in dictyosomes (D). The finely
granular deposit in dictyosomes and the coarse cytoplasmic
deposit are nonspecific products, which are also present in
control incubations. M, mitochondrion. x 45,000.

Fig. 18. Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the lack of alkaline phosphatase reaction
product in cytoplasmic vesicles (V). The finely granular
deposit in vesicles and the coarse cytoplasmic deposit are
nonspecific products, which are also present in control
incubations. CW, cell wall; M, mitochondrion. x 25,000.











- -i



M) 0p3 F..r




... .... ...
M *t -'I .-





4p,
'''. ... 1




p1-.


.100
-c w







040




S...' J- .;*.
* I







F ~~ r. r~4~
4. a Ug'








(not shown), as are the coarser deposits that are scattered throughout

the cytoplasm.


PTA-Cr03 stain. Only two types of membranes stain with the PTA-

Cr03 stain: the plasma membrane (Fig. 19) and some of the cytoplasmic

vesicles (Figs. 19, 20). These vesicles are 86 18 nm in diameter;

the membranes of the larger fibrous vesicles do not stain (Fig. 19).

In some of the reactive vesicles, only part of the membrane is stained

(Fig. 19), though in most, the staining is complete.


PASM stain. PASM, a cytochemical stain for polysaccharide, stains

most cell structures to some degree and has the highest "background"

reactivity of the techniques employed. This is also true of control

sections from which periodate oxidation has been omitted (not shown).

Sections oxidized by periodic acid show enhanced deposition of silver

grains primarily in three cellular locations: the cell wall (Fig. 21),

dictyosomes (Fig. 22), and fibrous cytoplasmic vesicles (Fig. 21). In

addition, the plasma membrane may react, but if it does, the reaction

is masked by the heavy silver deposition in the cell wall.

The size of the reactive cytoplasmic vesicles is 152 24 nm (the

same size as the large IDPase-positive and acid phosphatase-positive

vesicles), and virtually all vesicles of this size react. The silver

reaction product is preferentially deposited in the region of the

fibrous matrix (Fig. 21), and the inner fiber-free zone is unstained.

In many of the dictyosomes, cisternae at one pole are stained more

intensely than cisternae at the opposite pole (Fig. 22).

























Fig. 19. Longitudinal section of the apical region ofanA. ambisexualis
hypha, which shows the stainability of the plasma membrane
(PM) and some (but not all) of the cytoplasmic vesicles with
the PTA-Cr03 stain. The arrow indicates a large fibrous
vesicle. CW, cell wall; M, mitochondrion; V, vesicle.
x 68,000.

Fig. 20. Longitudinal section of theapical region of an A. ambisexualis
hypha, which shows the distribution of PTA-Cr03-positive
vesicles in the apex. CW, cell wall; M, mitochondrion.
x 23,000.


Fig. 21.




Fig. 22.


Longitudinal section of the apical region of an A. ambisexualis
hypha, which shows the stainability of cytoplasmic vesicles
(V) and the cell wall (CW) with the PASM stain for carbo-
hydrate. x 79,000.

Longitudinal section of the subapical region of an A.
ambisexualis hypha, which shows a dictyosome (D) that has
been stained with the PASM stain for carbohydrate. x 80,000.

















PM


dJ^f ^*


..... .!.


-



M' r ll. -
Iini=


M~


VO


...." .
. ',
V '' "*l. "


S ., ".l\ % *. .-": '.





V


I. -
F. *. *
b* ~ ~ .. -
~- .q -


W


.4. *S. '~k .* .


CW


V


C r
I


* I


" i~"


.. O








Discussion


On the whole, the apical organization of Achlya hyphae agrees very

well with the standard Oomycete pattern that was described by Grove and

Bracker (1970). In particular, the apical zone is occupied largely by

cytoplasmic vesicles, though the intrusion of mitochondria into this

region may represent a slight deviation from the formal model. Another

point of agreement is the morphological evidence that suggests that

some or all of the cytoplasmic vesicles may be derived from the Golgi

apparatus, and many of these vesicles contain polysaccharide. Poly-

saccharide has been detected cytochemically in cytoplasmic vesicles of

several fungi, including Achlya (Dargent, 1975), Phytophthora (Meyer

et al., 1976), and Saprolegnia (Heath et al., 1971) among the Oomycetes.

The stainability of dictyosomes with PASM is also common, and the

fact that the more mature cisternae are the first sites of stainability

in the endomembrane system is often taken as evidence that poly-

saccharide synthesis is initiated in this organelle (Pickett-Heaps,

1968).

From their morphological appearance and cytochemical behavior, it

would appear that at least two types of apical (cytoplasmic) vesicles

exist. The first consists of the large (about 150 nm) vesicles with

PASM-positive fibrous matrices. These are also uniformly PTA-CrO3-

negative. Furthermore, most (but not all) vesicles of this type are

IDPase-positive, while a smaller number contain acid phosphatase. It

was not determined whether these two phosphatase activities are mutually

exclusive or whether both can be found together in the same vesicles.

If this latter case is true, then yet a third subclass must be recognized







in those 150 nm vesicles which contain neither activity. In either

event, the strong probability exists that all 150 nm fibrous vesicles

merely represent one or another stage of development, which is re-

flected in their variable enzyme content. If that is so, it is likely

that acquisition of IDPase activity occurs as acid phosphatase activity

is lost, because the dictyosomes are IDPase-negative and acid

phosphatase-positive; and more of the vesicles in the apex exhibit

activity of IDPase than of acid phosphatase.

Cytochemical localization of acid phosphatase in other fungi has

demonstrated activity in dictyosomes; for example, Meyer et al. (1976)

reported acid phosphatase activity in a single cisterna of Phytophthora

dictyosomes. This is the same result observed in Achlya (Fig. 13).

Acid phosphatase has traditionally been considered a lysosomal enzyme

(Wattiaux, 1969); and it is found in various fungal vesicles, which may

be lysosomes (Armentrout et al., 1976), though in many fungi it seems to

be an enzyme of larger cellular vacuoles (Matile, 1971). Neither of

these sites is comparable to the apical vesicles of Achlya; in fact,

studies of fungal lysosomes generally report a lack of acid phosphatase

activity in apical vesicles (Armentrout et al., 1976; Maxwell et al.,

1978), though Dargent and Denisse (1976) report acid phosphatase ac-

tivity in apical vesicles of Achyla bisexualis Coker.

An observation that may be of particular importance is that IDPase

is not present in dictyosomes. This has been a traditional marker

enzyme for dictyosomes, especially in plants (Dauwalder et al., 1972),

although not all plant dictyosomes necessarily react (Dauwalder et al.,

1969). In addition to its common association with plant dictyosomes,

IDPase has been linked to polysaccharide synthesis (Ray et al., 1969).








The fact that IDPase is very specific for 150 nm polysaccharide-rich

cytoplasmic vesicles may indicate a similar association with poly-

saccharide synthesis in Achyla. Additionally, this enzyme may serve as

a marker for this class of apical vesicles in biochemical studies.

The second group of cytoplasmic vesicles consists of those smaller

vesicles (about 80 nm), among which are found the PTA-Cr03-positive

vesicles, the small IDPase-positive vesicles, and the coated vesicles.

These last two are probably identical, judging from their morphology;

and the PTA-Cr03-positive vesicles may also be identical. However, this

has not been demonstrated, and the possibility must be accepted that

this class of small vesicles is in fact composed of distinct subgroups.

The exact chemical basis for the PTA-Cr03 reaction is unknown, but

it is thought that glycoproteins that are characteristic of plasma

membranes are involved (Roland et al., 1972). It is of interest, there-

fore, that certain cytoplasmic vesicles exhibit the same stainability as

that shown by the plasma membrane. This has also been observed in higher

plant cells, where it has been interpreted as the result of a chemical

change in cytoplasmic vesicles that are involved in secretion (Vian and

Roland, 1972). Presumably, as the vesicle membranes acquire the char-

acteristics of the plasma membrane, certain barriers to their eventual

fusion are overcome. In higher plants, the change is not uniform at

first but is initiated in a restricted part of the vesicle; as a result,

the membrane may display only partial stainability (Vian and Roland,

1972). Such is also true of cytoplasmic vesicles in Achlya. If this

change is indicative of those vesicles which are capable of fusing with

the plasma membrane, it is curious that none of the 150 nm fibrous







vesicles are PTA-CrO3-positve. Can it be that these do not fuse with

the plasma membrane?

It should be clear from the preceding discussions that at least

two groups of vesicles can be found in the cytoplasm: the large fibrous

vesicles and the smaller nonfibrous vesicles. In addition, each group

may be composed of further forms which may be distinct or which may

represent various stages in the development of the two major classes of

vesicles. The existence of more than one type of vesicle in association

with fungal wall formation has been frequently noted (Grove et al., 1970;

Bracker, 1971; Hemmes and Bartnicki-Garcia, 1975; Meyer et al., 1976).

Some investigators have concluded that the different vesicles may have

different origins (e.g., from dictyosomes or from ER) and different

functions (Meyer et al., 1976). Another possibility is that all vesicles

are derived from the same source and that the larger vesicles are formed

by the coalescence of the smaller ones (Grove et al., 1970).

In Achlya, no evidence was observed to justify the derivation of

apical vesicles from the ER. Nor was there any evidence that 150 nm

vesicles must be derived from the coalescence of vesicles of the 80 nm

class. Instead, each class seems to have been independently produced by

dictyosomes (Figs. 4,9). Though those fibrous vesicles which are still

attached to dictyosomes (Fig. 4) are smaller than their mature size

(ca. 90 nm vs ca. 150 nm), it is not necessary that their subsequent

increase in size result from fusion with vesicles of the 80 nm class.

Instead, the increase could result from the fusion of other small

fibrous vesicles or from simple incorporation of materials.

The nature and function of the elongated "submural tubules" is not

apparent. From their morphology, they would seem to be no more than







alternate forms of the large fibrous vesicles. As mentioned earlier,

forms intermediate between the tubular and vesicular morphology may

indicate that one is derived from the other (Fig. 8). Their location in

the cell is well removed from the apex, where the bulk of incorporation

of materials into the wall occurs. Instead, these tubules and vesicles

are found as much as 20 or 30 Pm behind the apex. A direct role in

apical growth, thus, seems unlikely.

The cytochemical localizations of alkaline phosphatase and cellu-

lase were unable to demonstrate activity in association with subcellular

structures. Alkaline phosphatase has been reported to be associated

with fungal ER and/or dictyosomes (Meyer et al., 1976) and with apical

vesicles (Dargent, 1975). This latter report dealt with Achlya bi-

sexualis, and it is surprising that no such association was found in

A. ambisexualis apices. It must, of course, be acknowledged that this

may merely be the result of an improper application of the technique in

this study.

The greatest disappointment of this investigation must be the

failure of the cytochemical test for cellulase activity, whose localiza-

tion is the main object of the entire study. The failure is surprising,

because the technique has already been employed successfully in another

study of A. ambisexualis, strain E87 (Nolan and Bal, 1974). In that

investigation, cellulase was reported to be localized in cytoplasmic

vesicles (about 165 nm in diameter) of antheridiol-induced hyphae.

These vesicles would appear to be identical to the 150 nm fibrous

vesicles reported in this study, though the presence of cellulase in

fibrous vesicles in noninduced mycelia cannot be automatically assumed.




54


Upon contemplation, it is interesting to note that the only pub-

lished studies in which the cytochemical localization of cellulase has

been successfully reported are those in which the technique has been

performed by its originator, A. K. Bal (see Nolan and Bal, 1974; Bal

et al., 1976 and references cited therein). While it would appear

from these that the technique holds the promise of success, it may be

that the published methodology for its successful application is

incomplete.














CELLULASE AND UDPG TRANSFERASE


In this section, the existence of particle-bound cellulase (Cx)

will be demonstrated, and some properties relevant to its bound state

will be investigated. In addition, some features of UDPG transferase

will be examined.


Some Properties of Mycelial Cellulase


The Distribution of Cellulases between Particulate and Soluble
Fractions of Mycelial Homogenates


Methods

Mycelia were harvested from 48 hr old cultures and divided into lots

weighing 2 g FW. These were homogenized with 2.0 ml of a homogenizing

solution composed of 30% sucrose in 0.03 M tris.HCl buffer, pH 7.6,

according to the standard scheme. Sand and cell fragments were removed

by centrifuging the homogenate at 270 x g x 10 min, and a "total particu-

late" fraction was obtained by centrifuging the 270 x g supernatant at

37,000 x g x 90 min. This particulate fraction was resuspended in

2.0 ml of 10% sucrose in 0.01 M tris buffer, and 1.0 ml samples of the

"particulate" fractions and of the 37,000 x g supernatant "soluble" frac-

tions were assayed directly for cellulase activity.

As a comparison, another technique for extracting mycelial cellu-

lase was employed, which has already been used in studies of Achlya

(e.g., Thomas and Mullins, 1967). Two-gram lots of mycelium were








frozen on dry ice and ground with sand in a chilled mortar with 2.0 ml

of 5.0% w/v NaCI. The slurry was centrifuged at 35,000 x g x 15 min,

and the supernatant was decanted and saved. Five volumes of 95%

ethanol were added to the supernatant, which was then recentrifuged at

35,000 x g x 15 min. The pellet was resuspended in 2.0 ml of distilled

water and centrifuged at 27,000 x g x 15 min. The final supernatant

was saved for assay of cellulase.

One unit of Cx activity is defined as that amount of enzyme ac-

tivity which is sufficient to cause a 1.0% decrease in flow time of the

substrate in 1.0 hr. This definition differs from that used in previ-

ous investigations in this laboratory (e.g., Thomas and Mullins, 1967),

where a 10.0% decrease in flow time was considered to be 1.0 unit of

activity.

Results are displayed in Table 3.


Results

As Table 3 indicates, cellulase can be recovered both from particu-

late and buffer-soluble fractions extracted by grinding in a buffered

osmoticum; and the amount in each is about equal, as assayed by this

technique. (However, as the following experiments will reveal, the

level of particulate cellulase activity has been greatly underestimated

by this assay.) Precipitation of cellulase from NaCl extracts yields

cellulase levels about equal to either the buffer-soluble or particu-

late fraction alone, i.e., about one half of that in both combined.

















TABLE 3. Cellulase activity in salt-soluble or buffer-soluble and
buffer-insoluble fractions from homogenates of replicate
2g FW samples of A. ambisexualis mycelium produced by the
method of Thomas T1966) or by grinding in a buffered
osmoticum, respectively.


Sample Fraction Cx activity Average
number reaction (units/g FW) activity

1 5.2
NaC1-
2 soluble 4.4 4.9 0.4

3 5.1


4 6.6
buffer-
5buffer- 5.7 5.3 + 1.2
soluble
6 3.7


4 3.9

5 buffer- 7.0 5.4 + 1.3
insoluble
6 5.3








The Effect of Triton X-100 on the Activity of Particulate
and Soluble Cellulases from Mycelial Homogenates


Methods

Ten grams of mycelium were homogenized in 5.0 ml of homogenizing

solution, as described in the previous experiment. The homogenate

was divided into "total particulate" and "buffer-soluble" fractions by

centrifugation, and the particulate material was resuspended in 20 ml

of 10% sucrose in 0.01 M tris buffer. Separate aliquots of resuspended

particles were made 0%, 0.25%, 0.5%, 1.0%, and 2.0% w/w in triton X-

100, and aliquots of the buffer-soluble phase were made 0% and 1% in

triton. Samples (1.0 ml) were assayed for cellulase activity. Initial

mixing produced a high degree of aeration in those viscometer tubes

that contained triton, and viscosity changes were erratic during the

first 15 minutes. Therefore, readings were made at T = 15 min and at

T = 15 min + 1 hour. The results are displayed in Table 4.


Results

Treatment of cellular particles with 0.5%, 1.0%, or 2.0% triton

X-100 increases cellulase activity on the average to about 8.7 times

the activity of untreated particles. Triton at 0.25% gives a lesser

degree of activation. Triton is not itself cellulolytic, as the com-

parison between the blanks with and without triton reveals. Finally,

the activity of triton would seem not to involve an activation of the

enzyme itself, but rather a specific effect upon the bound state is

indicated. One possible mode of action is the freeing or solubiliza-

tion of the enzymes from the particles to which they are bound. To

determine whether this is involved in cellulase activation, a second

experiment was performed.
















TABLE 4. The effect of triton X-100 on the activity of particulate
and buffer-soluble cellulases from A. ambisexualis mycelial
homogenates



Percent Cellulase*
Sample triton X-100 activity

0.0 0.00 units
10% sucrose
in tris 2.0 0.02


0.0 5.41 units/g FW

0.25 37.11
Particulate 0.5 46.18
0.5 46.18 "
fraction
1.0 47.43

2.0 45.88


Buffer- 0.0 4.18 units/g FW
soluble
fraction 1.0 4.52


*Each value is the average of two measurements.







The Effect of Triton X-100 on Solubilization of
Particle-bound Cellulase


Methods

Cellular particles were obtained and resuspended in 10% sucrose in

tris buffer, as described in the preceding experiment. Separate ali-

quots of resuspended cellular particles were made 0%, 0.01%, 0.1%, and

1.0% w/w in triton X-100. Tritonated samples were kept on ice for 15

min to prevent complete triton activation and then centrifuged for 60

min in a 65 rotor at 79,000 x g. Sedimentable material was resuspended

in 10% sucrose in tris buffer, and an aliquot was made 1% in triton

X-100 at room temperature in order to activate any cellulase still bound

to the particles. Cellulase activities of the 79,000 x g sediment and

of the 79,000 x g supernatant (solubilized activity) were determined,

and the results are displayed in Table 5.


TABLE 5. The distribution of Cx activity between particulate and soluble
phases after treatment of A. ambisexualis cellular particles
with various concentrations of triton X-100


Percent Cx activity Cx activity Percent of
triton bound to cell solubilized total activity
X-100 particles (units/g FW)* solubilized
(units/g FW)*

0.0 67.7 4.1 6

0.01 79.4 9.2 10

0.1 41.4 34.7 46

1.0 13.9 69.3 83


*Each value is the average of two measurements.








Results

From Table 5, it can be concluded that bound cellulase exists in a

relatively inactive form that can be activated by triton X-100, and

this activation is accompanied by removal of cellulase from the parti-

cles. This type of behavior is typical of biological membranes.


The Effect of Various Methods for Solubilizing
Membrane-bound Cellulase

Membrane-bound proteins have been divided into two classes:

integral and peripheral (Singer, 1974). Their distinction rests upon

the tenacity with which each is bound to the membrane and upon the

types of treatments required for their removal or solubilization. To

better determine the strength with which cellulase is bound to cellular

membranes, the following experiments were performed, in which salts,

sonication, and freezing were used in an attempt to remove cellulase

from cellular membranes.


Methods

Cellular particles were obtained and resuspended in 10% sucrose in

tris buffer, as described previously. Separate 1.0 ml aliquots were

subjected to one of the following treatments, which are designed to

remove trapped or lightly bound cellulase: sonication twice for 30 sec

each at setting #5 of a Heat Systems-Ultrasonics sonifier-cell disruptor,

model W185, fitted with a standard microtip (samples were kept chilled);

freeze-thawing twice at -70 C; making solutions 1.0 M in NaC1, KC1, or

NH4C1 for 15 min at RT. Following these treatments, samples were

centrifuged for 60 min at 79,000 x g, and the sedimentable materials

were resuspended in 10% sucrose in tris buffer. Aliquots of resuspended

79,000 x g sediments were made 1.0% in triton X-100 to activate bound








cellulase, and both 79,000 x g sediment and 79,000 x g supernatant

fractions were assayed for Cx activity and protein content. Results

are displayed in Table 6.


Results

Freeze-thawing and monovalent cations are ineffective in dislodging

cellulase from membranes. The salts used are quite effective in remov-

ing proteins, however (about a 25% increase in soluble protein), so

cellulase must be more tightly bound than at least 25% of the other

membrane proteins. While sonication results in the loss of some 10% of

the cellulase from the particulate fraction, this is accompanied by the

loss of a comparable amount of total protein, and may, therefore, rep-

resent membrane fragmentation, rather than solubilization. In any

event, the release of cellulase by sonication is not of the same mag-

nitude as the release achieved by triton treatment (Tables 4 and 5).


The Effect of Room Temperature Incubation on Activity
of Membrane-bound Cellulase

During the course of these experiments, it was observed that on

some, but not all, occasions, samples of resuspended cellular particles

that had been left "unattended" at room temperature for a number of

hours displayed unexpectedly high cellulase activity. Much further

investigation revealed that these "activated" samples were among those

obtained from mycelium which had been homogenized in a medium containing

dithiothreitol (DTT). Though the membranes had been resuspended in media

lacking DTT, it was possible that minute amounts were carried over in

some samples and were somehow affecting enzyme activity. Experiments

were conducted to determine the effect of DTT and room temperature

"aging" on the activity of membrane-bound cellulase.






















s.-

4--)
4-

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Methods

Ten grams of mycelium were homogenized, as previously described,

and the total particulate fraction was resuspended in 20 ml of dis-

tilled water. Samples were made 0 mM, 0.5 mM, and 5.0 mM in DTT and

incubated at room temperature. Periodically, 1.0 ml aliquots were re-

moved and assayed for cellulase activity, without triton activation.

Results are displayed in Table 7.


TABLE 7. The effect of DTT upon cellulase activity during incubation
of A. ambisexualis cellular membranes for 24 hr at room
temperature


Concentration Cx activity* Cx activity*
of DTT at T = 0 hr at T = 24 hr
(units/g FW) (units/g FW)

0.0 mM 4.9 11.3

0.5 mM 6.0 45.4

5.0 mM 7.9 45.4


*Each value is the average of two measurements.




Results

In the absence of DTT, cellulase activity increases about 2.3

times, whereas in the presence of DTT, activity increases by about 6

to 8 times the original value. This confirms the suspicion that the

activity of membrane-bound cellulase can increase at RT and that DTT

enhances the process.








The Effect of Temperature on Cellulase Activity during
Incubation of Cytoplasmic Particles


Methods

Membrane samples were obtained as previously described and re-

suspended in distilled water made 0.5 mM in DTT. Portions were incu-

bated at 4 C, 25 C (RT), and 37 C, and aliquots were removed at T =

0 hr and T = 24 hr for assay of cellulase activity without triton

activation. Samples were also assayed for protein content. Results

are displayed in Table 8.


TABLE 8. The effect of temperature upon cellulase activity during
incubation of A. ambisexualis cellular membranes for 24 hr
in 0.5 mM DTT


Cx activity (units/g FW)t mg protein/g FWt
Temperature
T = 0 hr* T = 24 hr T = 0 hr* T = 24 hr

4 C 5.1 9.0 2.43 2.48

RT 5.1 42.9 2.43 1.76

37 C 5.1 9.0 2.43 1.44


*Since all three T = 0 samples are identical, only one representative
value was obtained.
Each value is the average of two measurements.




Results

Cellulase activity increased 8.5-fold in 24 hours at RT, as ex-

pected. Activation is prevented at temperatures of 4 C and 37 C. One

explanation may be that activation requires a temperature sensitive

step (enzymatic?) with an optimum around 25 C. This may be proteolytic,







since protein content during "aging" decreases at room temperature,

while no proteolysis is detected at 4 C. However, the greatest

proteolysis occurs at 37 C, which shows no activation. Thus, pro-

teolysis and activation may not be related. An alternative explanation

might require that mild or selective proteolysis may occur at RT,

causing activation, but that extensive proteolysis occurs at 37 C, and

cellulase is degraded.


The Effect of Incubation at Different Temperatures on the
Solubilization of Membrane-bound Cellulase

Activation of membrane-bound cellulase by triton has already been

shown to be accompanied by solubilization of the enzyme (Table 5). How-

ever, a requirement of solubilization for activation was not demon-

strated. To determine whether activation of cellulase by an apparently

unrelated method also is accompanied by solubilization, the following

experiment was performed with "aged" material.


Methods

Membrane samples resuspended in distilled water were obtained by

the methods used in the preceding experiments. Samples of cellular

particles, made 0.05 mM in DTT, were incubated at 4 C, RT, and 37 C

for 24 hr, and aliquots were removed and centrifuged at 79,000 x g x

60 min. The resulting sediment and supernatant fractions were assayed

for Cx activity directly, and a sample of the sediment fraction was also

assayed in the presence of 1.0% triton X-100 to activate bound cellulase.

Results are displayed in Table 9.


Results

As expected, directly assayable cellulase activity (i.e., the sum

of soluble and particulate activities without added triton) is higher




















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after room temperature incubation than after incubation at 4 C or 37 C.

The bulk of this increase is clearly in the soluble phase. Compared to

the material which has been incubated at 4 C, a smaller amount of triton-

releasable cellulase activity is still associated with the membranes

after RT incubation. The simplest explanation is that, with room

temperature "aging," inactive membrane-bound cellulase becomes both

active and soluble, and this process is retarded by low temperature. As

evidence, notice that the maximum available cellulase activity in the

4 C and RT samples is approximately equal (94.6 vs 97.9), indicating

that one effect of room temperature incubation is to shift Cx from the

particulate phase to the soluble. The fact that less membrane-bound,

triton-releasable cellulase remains after incubation at RT than after

incubation at 4 C also indicates that both triton solubilization and

"aging" act on the same pool of membrane-bound cellulase. Finally, this

comparison between triton activation and room temperature activation

suggests strongly that solubilization is a requirement for maximal Cx

activity and is not just a secondary effect of activation.

The distribution of activities at 37 C, however, holds an unex-

pected result, because the triton-activatable particulate component is

missing. Thus, unlike the lack of activation observed at 4 C, the lack

of activation at 37 C is not merely due to inhibition of the room tem-

perature effect by unfavorable temperature. That is, the membrane-

bound cellulase activity has not been left in place, as it would have

been at 4 C.

A number of possible explanations come to mind, not all of which

were treated by experimentation. It may be that cellulase is denatured

by a temperature of 37 C. This is not likely, since a hallmark of








fungal cellulases is their remarkable temperature stability (Whitaker,

1963, 1971). In addition, Thomas (1966) demonstrated that salt-

extractable cellulases from Achlya exhibit a considerable degree of

stability even at 100 C.

Another possibility is that temperature may exert some effect on

the membrane itself, rendering the bound cellulase somehow unavailable,

though not actually denaturing it. Or it may be that the cellulase ac-

tually is solubilized at 37 C, but that it is destroyed (perhaps by

proteolysis) after solubilization. Extensive proteolysis has already

been shown to occur at 37 C (Table 8).

To determine whether a cellulase from Achlya would be denatured at

37 C, samples of buffer-soluble cellulase were incubated at room tem-

perature and at 37 C for 24 hr, and cellulase activity was measured.

From an initial value of 4.53 0.21 units/g FW at TO, activity changed

to 4.92 0.08 units/g FW after 24 hr at room temperature and to 4.31

0.01 units/g FW after 24 hr at 37 C.

Buffer-soluble cellulase, then, like other fungal cellulases, is

not affected by 37 C to a degree sufficient to dismiss 37 C inactiva-

tion of particulate cellulase as mere denaturation. This explanation

cannot be rigidly excluded, however, because the temperature stabilities

of buffer-soluble and membrane-derived cellulases have not been compared.

Thus, though there is reason to doubt an effect of 37 C directly on

cellulase, the reason for the apparent destruction of membrane-bound

cellulase activity is not known.







Some Properties of UDPG Transferase


The Distribution of UDPG Transferase Activity between Protoplasm
and Wall Fractions of Mycelial Homogenates

UDPG transferases, if indeed they are involved in cell wall metabo-

lism, would be expected to be located at points where wall components

are assembled. Evidence already cited from other workers has indicated

that the wall may be assembled in part within the protoplast and in part

outside the plasma membrane. Accordingly, UDPG transferase activity

was looked for in "wall" and "protoplasm" fractions of disrupted Achlya

mycelium.


Methods

Two-gram FW samples of mycelium were harvested from a 48 hr old

liquid culture and transferred to 15 ml conical glass centrifuge tubes.

Four milliliters of homogenizing solution, consisting of 20% sucrose,

10 mM DTT, and 0.02 M tris.HC1 buffer, pH 7.6, were added to each tube,

and the tubes were chilled in a salted ice bath. When the temperature

reached 0 C, the contents were sonicated for 15 sec on the Heat Systems-

Ultrasonics sonifier at a setting of 5. The temperature, which had

reached 5-10 C during sonication, was reduced by rechilling, and the

contents were sonicated for another 15 sec. Fragmented hyphae were

sedimented by centrifugation at about 1,000 x g x 10 min, and the super-

natants were decanted and saved. Fragments were resuspended in 2.0 ml

of half-strength homogenizing solution, sonicated, and centrifuged as

before. This procedure was followed twice more, and the four 1,000 x g

supernatants from each sample were pooled.

Wall fragments were resuspended in 2.0 ml of half-strength homoge-

nizing solution, and a sample was examined under the phase contrast








microscope, where little cytoplasmic contamination was evident. UDPG

transferase activity was assayed using samples of both wall and super-

natant fractions, and protein content was determined. Results are

displayed in Table 10.


TABLE 10. The distribution of protein and UDPG transferase activity
between "wall" and "protoplasm" fractions of A. ambisexualis
mycelial homogenates produced by sonication


UDPG Protein
Fraction transferase content Specific
activity*t (mg/g FW)t activity*

Wall 1.53 0.09 0.59 0.02 2593

Protoplasm 2.12 0.13 3.70 0.06 573

E 3.65 4.29 851

*UDPG transferase activity is expressed as nmoles of glucose
incorporated per min per g FW.
tEach value is the average of 3-4 measurements.
**Specific activity is expressed as pmoles of glucose incorporated
per min per mg protein.



Results

About 42% of the total mycelial transferase activity is found in

the wall fraction, whereas only about 14% of the protein assayed is

found here. Accordingly, the specific activity of wall-bound transferase

is almost five times higher than that in the protoplasm. It cannot be

said with certainty that the high activity in the walls results from a

large number of enzyme molecules in the wall, because the enzyme assay

does not present transferase enzymes with exogenous glucose-acceptors

but relies on their demonstrated ability to transfer glucose to endogenous








acceptors. High activity of wall transferases might be due to the

nature of the acceptor available (i.e., wall), which may represent a

more "ideal" acceptor than that available to cytoplasmic samples. Be

that as it may, it is apparent that transferases are found both in cell

walls and in cell protoplasm.


The Distribution of UDPG Transferase between Particulate and
Soluble Fractions of Mycelial Homogenates


Methods

Mycelium was harvested from 48 hr old cultures and divided into

lots weighing 2 g FW. These were homogenized by grinding according to

the standard scheme in 2.0 ml of homogenizing solution consisting of

30% sucrose, 15 mM DTT, and 0.03 M tris.HC1 buffer, pH 7.6. The "total

particulate" fraction was obtained and resuspended in 2.0 ml of one-

third-strength homogenizing solution; and 100 pl fractions of the

37,000 x g supernatant "soluble" fraction and the resuspended "particu-

late" fraction were assayed for UDPG transferase activity and for

protein content. The results are displayed in Table 11.


Results

Comparing the results in Table 10 with those in Table 11, it can

be seen that grinding in a mortar yields about 38% less cytoplasmic

protein than does sonication (3.70 vs 2.30 mg/g FW), though the overall

specific activities of the cytoplasmic fractions in Tables 10 and 11 are

almost identical (573 vs 523 nmoles-min- *mg- ). The particulate

fraction holds nine times the transferase activity at about eleven

times the specific activity of that in the soluble phase. The ratio of

specific activities in wall, particulate, and soluble phases is 28:11:1.








TABLE 11. The distribution of protein and UDPG transferase activity
between particulate and soluble protoplasmic fractions of
A. ambisexualis mycelial homogenates produced by grinding
with a mortar and pestle


UDPG Protein Specific
Fraction transferase content S ii
activity*t (mg/g FW)t activity*

Particulate 1.08 0.02 1.03 0.06 1050

Soluble 0.12 0.01 1.27 0.09 93

S1.20 2.30 523


*UDPG transferase activity is expressed as nmoles of glucose
incorporated per min per g FW.
tEach value is the average of 3-4 measurements.
**Specific activity is expressed as pmoles of glucose incorporated
per min per mg protein.




Although no specific experiment was performed for this purpose,

a comparison of the results of Tables 10 and 11 can serve as a "mixing"

experiment to reveal the presence or absence of soluble cytoplasmic

inhibitors of transferase activity. In Table 10, the specific activity

of the cytoplasmic fraction is calculated from the protein content and

enzyme activity of the combined particulate and soluble phases. In

Table 11, the same value is calculated from separate assays of those

two fractions. The two values differ by only about 9%, a discrepancy

too slight to require the involvement of activators or inhibitors of

transferase.







The Solubility of Radioactive Products of UDPG
Transferase Activity

Biochemical characterization of the products of the UDPG trans-

ferase reaction was not attempted in this research. However, their

solubility in various solvents was investigated to enable some compari-

son to be made with UDPG transferases reported from other systems.


Methods

Mycelia were homogenized, as in the preceding experiment. The

"total particulate" fraction was resuspended in 2.0 ml of 0.01 M

tris-HC1 buffer with 5 mM DTT, and transferase activity was assayed

using three times the volumes of reagents and sample normally employed.

Ethanol-insoluble products were fractionated by the following procedure,

which is a modification of that used by Van Der Woude et al. (1974):

1. Two washes with 2:1 v/v chloroform:methanol (combine washes)

2. Two washes with 85 C distilled water (combine washes)

3. Two washes with 85 C 1.0 N NaOH (combine washes)

4. One wash with RT water (combine with alkali washes)

5. Two washes with RT water (discard washes)

Each wash was terminated by centrifugation at about 1,000 x g x 5 min

in a conical 15 ml centrifuge tube. The chloroform:methanol washes were

evaporated, and the residue was resuspended in distilled water and

transferred to a scintillation vial. Alkali washes were neutralized

with HC1, and samples of these and the hot H20 washes were transferred

to separate scintillation vials. Finally, the alkali-insoluble residue

was transferred in distilled water to a scintillation vial, and all

samples were counted by standard procedures. The percent of radio-

activity in each extract is displayed in Table 12.








TABLE 12. The distribution of radioactivity among different extracts
of the products of UDPG transferase activity from the
particulate fraction of A. ambisexualis mycelial homogenates


Extraction Percent of radioactivity*

Chloroform/methanol 4.9 2.5

Hot H20 27.1 2.1

Hot NaOH 56.8 3.4

Insoluble residue 11.3 2.9

*Each value is the average of 3-4 measurements.



Results

The fractionation scheme employed is derived from higher plant cell

wall methodology, where walls can be similarly fractionated to yield

the chloroform/methanol-soluble wall lipids and glycolipids, the hot-

water-soluble pectins and calloses, the hot-alkali-soluble hemicellu-

loses, and the alkali-insoluble a-cellulose (Siegel, 1968; Preston,

1974). Also, it is assumed that the products of transferases can be

fractionated into similar classes of compounds. The wisdom of this as-

sumption will be discussed later. At this point it will only be men-

tioned that, if the above assumption is taken literally, only about 10%

of the products would qualify as cellulose on the basis of solubility;

this is, perhaps only coincidentally, about equal to the proportion of

cellulose in Achlya cell walls (Parker et al., 1963).








Discussion


Cellulase

Mycelial cellulase in Achlya is shown to exist in at least two

pools after homogenization: one which is associated with cellular

particles and another which is soluble. Attempts to dislodge particle-

bound cellulase by further physical disruption (Table 6) are unsuccess-

ful, and this indicates that the soluble cellulase is not derived from

the same population of molecules as those in the particle-bound pool.

That is not to say that the soluble cellulase may not originally have

been associated with cellular particles before homogenization. Any

number of factors during homogenization and isolation can lead to partial

or total solubilization of enzymes (Lips, 1975). Byrne et al. (1975)

reported a buffer-soluble cellulase in pea epicotyls, which eventually

proved to have been released from the endoplasmic reticulum (Bal et al.,

1976). The buffer-soluble cellulase of Achlya may also have been as-

sociated with cellular particles before disruption, but the fact that

it cannot be added to by further disruption of particles signifies a

qualitative, not quantitative, distinction between the two pools.

However, the difference may merely reflect different cellular locations

of otherwise identical molecules and not necessarily the existence of

distinct isozymes.

In previous work on Achlya, mycelial cellulase was reported as that

activity which could be extracted from frozen mycelium by grinding in a

salt solution (Thomas, 1966). While salt extraction of cellulases is a

standard method in many systems (e.g., Lewis et al., 1970), salt extrac-

tion of whole Achlya mycelium yields less cellulase than is made avail-

able by homogenization in buffer (Table 3). When one considers that








the latent, particle-bound cellulase in Table 3 has been underestimated

perhaps by a factor of 10, the proportion of mycelial cellulase re-

covered from the salt-soluble fraction becomes even less significant.

The amount of cellulase in the salt extract is about equal to the buffer-

soluble pool, and this may be the only source of salt-soluble cellulase.

Particle-bound cellulases probably do not contribute to the salt-soluble

fraction, since neither freezing nor salts dislodges cellulase from the

particles (Table 6). A contribution from wall-bound pools, as is seen

in the cases of peas (Bal et al., 1976) and beans (Reid et al., 1974),

cannot be ruled out, but the existence of similar wall-bound cellulases

in Achlya was not investigated in this research.

Whereas salts and physical disruption are ineffective in solubiliz-

ing particle-bound cellulase, solubilization can be accomplished by

triton X-100 (Table 5). Detergents act by disrupting cellular membranes

(Singer, 1974), and the effectiveness of triton in this instance is

evidence that the particles in question are cellular membranes. They

will be considered as such henceforth. Some information on the degree

of binding of cellulase to membranes is revealed in these solubilization

experiments. The failure of strictly physical disruption (freezing and

sonication) to release cellulase from membranes indicates that the

enzyme is not merely trapped inside of a vesicle as an otherwise soluble

molecule. Such a molecule would be expected to be released during the

membrane breakage which would accompany these treatments. Thus the

efficacy of triton cannot be limited to the mere rupture of membranes

but must lie in its ability to completely disrupt and disperse the

lipoidal membrane components.








The failure of salts to release cellulase indicates that the en-

zymes are not loosely bound to the membrane by weak, noncovalent bonds

and are not peripheral proteins (Singer, 1974). That peripheral pro-

teins of other types are present is attested to by the 25% increase in

soluble protein following salt treatment (Table 6). Bound cellulase,

then, would seem to fall into the class of membrane proteins termed

"integral" (Singer, 1974), i.e., they are present in the hydrophobic

interior of the membrane, or they are strongly bound to such proteins.

The degree of binding exhibited by membrane-bound cellulase in

Achlya has its counterpart in the membrane-bound cellulase of kidney

bean abscission zones (Koehler et al., 1976). This, too, is an integral

protein, which is released primarily by detergent and only to a much

smaller degree by salts or physical disruption. A similar enzyme may be

the B-glucanase in ER vesicles of yeast, which is activated by triton

X-100 (Cortat et al., 1972).

Fungal cellulases are known for their diversity (Pettersson, 1963;

Wood, 1968), and intracellular cellulases of individual higher plants

may differ markedly in their substrate specificity, extractability, and

molecular weight (Byrne et al., 1975). There is, therefore, no absolute

requirement that the secreted, the buffer-soluble, and the membrane-

bound cellulases of Achlya all be identical. However, in pea epicotyls,

a comparable trio exists in the buffer-soluble, the particulate, and

the wall-bound cellulases (Bal et al., 1976). Though at least two

isozymes are involved, the evidence indicates that these are sequen-

tially modified forms of the same enzyme (Wong et al., 1977b). Thus,

the possibility exists that in Achlya, too, the two internal pools of








cellulase represent the same enzyme in different stages of production

and secretion.

The strength with which cellulase is bound to cell membranes

raises serious questions about the way in which it is secreted or

whether membrane-bound cellulases can be secreted at all. If they are,

at some point they must become soluble. Clearly, they are not already

so, and their integral nature seems to argue against a mechanism for

their quick release. It is, therefore, of considerable interest that

membrane-bound cellulase can become soluble under conditions of adequate

temperature and concentration of DTT. Although the mechanism whereby

this occurs is not known, it seems to indicate that a strong degree of

binding to cytoplasmic membranes need not be a barrier to an enzyme's

eventual secretion.

The requirement, of course, is that cellulase release by "aging"

be reflective of a natural cellular process and not a completely arti-

ficial phenomenon, such as gross degradation of membranes. A parallel

may exist in a report by Frantz et al. (1973) in which the stainability

of isolated dictyosome vesicles by PTA-Cr03 was seen to increase with

time after isolation. This change is also part of the natural matura-

tion of the vesicles in the intact cell and apparently results from

changes of these membranes toward a more "plasmalemma-like" state (Vian

and Roland, 1972). Thus, changes in membranes in vitro can reflect the

continuation of natural developmental processes. Room temperature

activation of membrane-bound cellulase may be one such instance.

The requirement of DTT for enzyme release by "aging" raises the

possibility that some protein is critical to the process. DTT protects

protein sulfhydryl groups from oxidation (Cleland, 1964), thereby







preserving tertiary structure and/or the specificity of critical sites.

In this way, DTT may enable cellulase release to occur through the

action of a second enzyme, which may, for instance, be a protease. An

instance has been reported wherein an integral membrane protein can be

released in an active state by the activity of lysosomal enzymes; other-

wise, triton treatment is required (Spatz and Strittmatter, 1973).

Another example of autoactivation of a membrane-bound enzyme is the

chitin synthetase zymogen of M. rouxii (Ruiz-Herrera and Bartnicki-

Garcia, 1976). Activity of this enzyme increases in mixed membrane

fractions due to the action of an endogenous protease. Although

proteolytic release of this type was not demonstrated to be involved

in release of Achlya cellulase, the decrease in total protein during

incubation at RT indicates that this possibility is one of several that

bear investigation.


UDPG Transferase

The high activity of UDPG transferase in Achlya walls is also a

common observation in other fungi (Wang and Bartnicki-Garcia, 1966;

McMurrough et al., 1971; Meyer et al., 1976; Fevre and Dumas, 1977)

and in higher plants (Shore and Maclachlan, 1975). In the closely re-

lated genus Saprolegnia (Fevre and Dumas, 1977), about 45% of the ac-

tivity is wall-bound, while in Achlya, the level is 42% (Table 10).

This observation is evidence in support of two assumptions regard-

ing wall synthesis. First, it serves as an indication that these

enzymes are indeed involved in cell wall metabolism. Second, it indi-

cates that at least part of the cell's wall-synthesizing ability is

indeed extracytoplasmic, and a considerable amount of wall assembly can








occur there. This requirement had been postulated on purely theoretical

grounds,necessitated by the architectural complexity of the mature wall

(Preston, 1974).

The fact that the products of isolated transferase preparations

are often not rigorously identified (as indeed they remain in this in-

vestigation) lends itself to doubts concerning both the identity of the

natural products and their exact roles in the cell. In addition, even

well identified products produced in vitro may bear small resemblance

to the in vivo products of the same enzyme(s), because any number of

variable conditions can affect the nature of the products, and their

levels in the microcompartments of the cell cannot be known.

Among these factors are the nature and concentration of the

nucleotide-sugar donor (Ordin and Hall, 1968; Lamport, 1970; Tsai and

Hassid, 1971, 1973), the presence of metal ions (Tsai and Hassid, 1973;

Fevre and Dumas, 1977), the presence of carbohydrate or alcohol activa-

tors (Thomas et al., 1969; Spencer et al., 1971; Southworth and

Dickinson, 1975), and the presence of plant hormones (Van Der Woude

et al., 1972). The reason for much of this diversity almost surely

lies in the poorly purified nature of the enzyme preparations. Un-

doubtedly, more than one type of enzyme is present which is capable of

utilizing nucleotide sugar donors (Tsai and Hassid, 1971, 1973; Shore

and Maclachlan, 1973).

A variable which may exert fundamental control over activity in

vivo is the level of certain unidentified factors in the soluble phase

of cell homogenates. In addition to proteolytic activators (Ruiz-

Herrera and Bartnicki-Garcia, 1976), soluble inhibitors have been

reported, and these may account for the low specific activity of many








soluble transferases. One such report is that by Fevre and Dumas (1977)

in S. monoica, where the specific activity of the combined soluble and

particulate fractions was about 50% of that expected based on the ac-

tivities in the separate fractions. In Achlya, however, no evidence

of a soluble inhibitor is shown, even though the specific activity of

soluble transferase is quite low. The specific activity of the combined

soluble and particulate fractions is in fact about 9% higher than that

expected on the basis of separate assays of the two phases (cf. Tables

10 and 11). Thus, no soluble inhibitor is indicated.

In the present investigation, the identity of transferase products

was investigated only insofar as their solubility was concerned (Table

12). Most of the products are soluble either in hot water or hot

alkali. Care must be taken not to place excessive emphasis upon the

exact distribution of these products, however, because it cannot be

assumed that each fraction represents a distinct class of molecules, nor

that these solubilities correspond closely to those exhibited by ex-

tensive polymers of the types just synthesized. It is generally con-

ceded that, in vitro, transferase enzymes are successful in transferring

only a small number of sugars to the endogenous acceptor(s) (Preston,

1974). (Exceptions would be those chitin synthetases previously noted.)

Thus, the solubilities of radioactive products are determined largely

by the solubilities of the acceptors. It is to be hoped that the new

linkages are formed using an acceptor of the corresponding type, but this

has not been demonstrated.

A further complication arises, because the solubility exhibited by

long and short polymers of the same material may vary. A B-1,4-

oligoglucoside may be readily soluble in water, whereas insolubility







in most solvents is a hallmark of the longer cellulose I complexes

(Whitaker, 1971). In some systems, the hydrolysis of soluble and in-

soluble fractions yields different products (Shore and Maclachlan,

1975). However, in others, these fractions may contain identical

linkages, despite differences in solubility (Heiniger and Delmer, 1977).

Bearing these cautions in mind, it can be seen that the solubility-

distribution of Achlya transferase products (Table 12) is comparable

to that seen in S. monoica when 10 vM UDPG is used (Fevre and Dumas,

1977). The main difference is in the lipid-soluble fraction, which

constitutes a much greater proportion of the products in Saprolegnia.

One reason may be that the ethanol-insoluble products served as the

starting point for fractionation in the present research, whereas cold-

water-insoluble products were fractionated in Saprolegnia. Probably,

some lipid-soluble materials were lost in the ethanol washes.

Another source of discrepancy may lie in the 242 pM concentration

of UDPG employed in the present study, compared to the 10 uM concen-

tration used by Fevre and Dumas (1977). As already mentioned, the

concentration of the substrate is one of the many factors that can

influence the nature of the products.

An attempt was made to characterize only the alkali-insoluble

products in Saprolegnia; these yielded glucose and cellobiose upon

hydrolysis, and the alkali-insoluble materials were identified as

cellulose (Fevre and Dumas, 1977). The close taxonomic affinity between

the genera and the other similarities in the distribution and products

of their respective transferases make it probable that the alkali-

insoluble product of Achlya transferase also consists of a polymer with




84



B-1,4-glucosidic linkages. However, for the various reasons described,

this cannot be certainly stated until the Achlya products are them-

selves analyzed.














THE ASSOCIATION OF WALL SYNTHESIS
AND ENZYMES WITH HYPHAL GROWTH


Enzymes in the Culture Filtrate


Although it is probable that certain wall components are at least

partially assembled inside the cell, the architecture of the wall re-

quires that the final steps in assembly be extracellular (Preston,

1974). Enzymes involved in this assembly and others that may be present

in secretary vesicles will thus be found outside the protoplast. These

may eventually diffuse into the culture medium. To gain an indication

of the kinds of enzymes secreted during growth, the filtrate from Achlya

cultures was examined for enzyme activity. Enzyme activity in cell

homogenates was also examined to serve as a basis for comparison with

activities in the medium.


Methods

Two 48 hr old liquid cultures were harvested on miracloth, and

both mycelium and filtrate were saved and pooled separately. After

cooling the medium to 4 C, the filtrate was concentrated for 24 hr us-

ing a Millipore Immersible Molecular Separator (Millipore Corp.)

equipped with a pellicon membrane with a 10,000 nominal molecular

weight limit. When the volume had been reduced to 10 ml, samples were

assayed for enzyme activities.

Mycelium was homogenized in 2 g FW lots by standard methods.

Centrifugation (270 x g x 10 min) removed heavy material, and samples








of the homogenate were then assayed for enzyme activities. (Mycelial

cellulase was assayed in the presence of 1% triton X-100.) Results are

displayed in Table 13.


Results

Every enzyme assayed displays activity in both the mycelium and

the culture filtrate. However, the only filtrate activities which

are very high in comparison to their levels in the mycelium are those

of cellulase and ATPase. The activities of acid phosphatase, cytochrome

oxidase, and perhaps UDPG transferase are so low as to be insignificant

compared to mycelial levels.

The presence of cellulase in Achlya culture media has been reported

before (Bhargava, 1943; Thomas and Mullins, 1967, 1969). What is

demonstrated in the current experiment is that cellulase is not alone

in the medium; but, compared to its companion enzymes, cellulase ac-

tivity is perhaps the highest of all. This observation warrants our

continued investigation of the possible association between cellulase

and growth in Achlya.


Comparisons of the Growing and Nongrowing Conditions


For the growth of Oomycetes, sufficient amounts of certain min-

erals, organic nitrogen, organic sulfur, and a suitable carbon source

are required (Whiffen, 1945; Barksdale, 1962; Cantino, 1966). These

requirements are met by the Defined Liquid Medium (DLM). Removal of

nutrients will inhibit growth, and this is especially true of nitrogen

depletion (Barksdale, 1962; Griffin et al., 1974). Complete removal of

all nutrients, however, will induce sporangial development (Klebs,



















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1899), which is accompanied within a short time by its own set of

specialized physiological responses (Griffin and Breuker, 1969;

Timberlake et al., 1973; O'Day and Horgen, 1974). These might be ex-

pected to interfere with the measurements of conditions in the non-

growing state. Fortunately, sporangial formation can be avoided in a

depleted medium by maintaining the level of glucose or other carbon

source (P. A. Horgen, pers. comm.). This was confirmed by an experiment

in which sporulation was attempted by the methods already described,

but 0.5 mM CaC12 supplemented with 0.2% w/v glucose was used, instead

of calcium chloride solution alone. No spores or sporangia were found;

therefore, 0.2% glucose suppressed sporulation. Accordingly, the

medium employed to eliminate growth in the following experiments was

0.2% w/v Glucose Medium (GM), adjusted to pH 6.9 with HC1. In addi-

tion to suppression of sporulation, this medium permits the study of

radio-glucose uptake using the same concentration of glucose found in

DLM.


Changes in Mycelial Fresh Weight During Incubation
in DLM or GM


Methods

Mycelium from 48 hr old liquid cultures was harvested on miracloth

and rinsed with 250 ml of deionized water. Excess water was removed by

gentle pressure with a rubber spatula. The mycelium was divided into

2.0 g FW lots, which were resuspended in separate 250 ml Erlenmeyer

flasks containing 50 ml of either DLM or GM. Flasks were incubated at

24 C with shaking and were harvested hourly on miracloth; excess water

was removed as before. Mycelial mats were weighed, and the results are

displayed in Table 14.




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