ISOLATION AND CHARACTERIZATION OF VESICLES INVOLVED IN
HYPHAL TIP GROWTH OF ACHLYA
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
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
TABLE OF CONTENTS
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
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
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
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
LIST OF FIGURES
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
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
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
Terry William Hill
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
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-
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.
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,
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).
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
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.
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
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
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).
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.
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
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.
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
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.).
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.
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.
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 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
Phenol Test for Carbohydrate (Herbert et al., 1971)
5% w/v phenol
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)
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
Acid Phosphatase (EC 188.8.131.52) (modified from Ray et al., 1969)
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 184.108.40.206) (modified from
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.
Alkaline Phosphatase (EC 220.127.116.11)
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 18.104.22.168) (modified from Bell et al., 1955)
carboxymethyl cellulose (type 7Mf, Hercules 12.0 g
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 22.214.171.124) (modified from Hodges and Leonard, 1974)
cytochrome c (Sigma type III, from horse heart, 11.0 mg
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 126.96.36.199) (modified from Hubscher and West, 1965)
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.
B-glucosidase (EC 188.8.131.52) (modified from Parish, 1975)
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)
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 184.108.40.206) (modified from Ray et al., 1969; Shore
and Maclachlan, 1975)
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
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.
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).
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.
Phosphotungstic Acid-Chromic Acid (PTA-Cr03) Stain (Roland et al., 1972)
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
Periodic Acid-Silver Methenamine (PASM) Stain (Martino and Zamboni, 1967)
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
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
Cytochemical Localization of Alkaline Phosphatase (Hugon and Borgers,
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)
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
Cytochemical Localization of Acid Phosphatase (Gomori, 1952)
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.
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.
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
1 .. I ... '
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.
cw I ',A w
1 .A |
S e... . , .. .
ab. *. .) S M
* // .hV
,. :::i ::.
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
Cytochemical tests were performed using the techniques already
described under Materials and Methods.
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
Dictyosomes are unreactive, as are attached incipient vesicles
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.
M) 0p3 F..r
... .... ...
M *t -'I .-
'''. ... 1
S...' J- .;*.
F ~~ r. r~4~
4. a Ug'
(not shown), as are the coarser deposits that are scattered throughout
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.
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.
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.
M' r ll. -
V '' "*l. "
S ., ".l\ % *. .-": '.
F. *. *
b* ~ ~ .. -
~- .q -
.4. *S. '~k .* .
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,
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
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.
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
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
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
Results are displayed in Table 3.
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
Sample Fraction Cx activity Average
number reaction (units/g FW) activity
2 soluble 4.4 4.9 0.4
5buffer- 5.7 5.3 + 1.2
5 buffer- 7.0 5.4 + 1.3
The Effect of Triton X-100 on the Activity of Particulate
and Soluble Cellulases from Mycelial Homogenates
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.
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
Sample triton X-100 activity
0.0 0.00 units
in tris 2.0 0.02
0.0 5.41 units/g FW
Particulate 0.5 46.18
0.5 46.18 "
Buffer- 0.0 4.18 units/g FW
fraction 1.0 4.52
*Each value is the average of two measurements.
The Effect of Triton X-100 on Solubilization of
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
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.
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 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.
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.
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.
U 3 -
S- r- 0
(1 0 S-
.- V) D0
r- 0- -
o S- c)
v) Q- E
c u .- 0)
u C U->
- CQ o
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
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.
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
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
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.
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.
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.
As expected, directly assayable cellulase activity (i.e., the sum
of soluble and particulate activities without added triton) is higher
4- S- .
0r V) -P
C (AI *r
5.- c --
0- > >
*r- V >
:3 C .- U
r- > X>
3 *.- C
I- > O
O *.- A 4-'
C U -
o r- *-- (U
A 0 S- *i-
4- C I *<
O *.- (A >
0) r- I
r- > 0 0
O *r- -CM +-
'- 3 4-'
o-i Lfl c~)
Ocr r- -
r- o 43
n i- C"-
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
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
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.
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
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.
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
The Solubility of Radioactive Products of UDPG
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.
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.
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).
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
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
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
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
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
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
B-1,4-glucosidic linkages. However, for the various reasons described,
this cannot be certainly stated until the Achlya products are them-
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.
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.
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,
CD, E -4-
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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
Changes in Mycelial Fresh Weight During Incubation
in DLM or GM
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.