A Taxonomic Review of the Clavicipitaceous Anamorphs Parasitizing Nematodes and Other Microinvertebrates

Full text

Clavici
pita
I
ea
n
Fu
ng
i
Evol
u
t
i
ona
ry
Biology,
C
h
em
ist
ry
,
and
Cultural
Impacts
Biocontrol,
edited
by
Cook College-Rutgers University
New Brunswick, New Jersey, U.S.A.
James
F.
White,
Jr.
Charles
W.
Bacon
Agricultural Research Service,
US.
Department
of
Agriculture
Athens, Georgia,
U.S.A.
Nigel
L.
Hywel-Jones
National Science and Technology Development Agency
Bangkok
,
Thai land
Joseph
W.
Spatafora
Oregon State University
Corvallis, Oregon, U.S.A.
MARCEI
MARCEL
DEKKER,
INC.
DEKKER
NEW
YORK
BASEL

Although great care has been taken to provide accurate and current information, neither the
author(s) nor the publisher, nor anyone else associated with this publication, shall be liable
for any loss, damage, or liability directly or indirectly caused or alleged to be caused by
this book. The material contained herein is not intended to provide specific advice or
recommendations for any specific situation.
Trademark notice: Product or corporate names may be trademarks or registered trademarks
and are used only for identification and explanation without intent to infringe.
Library of Congress Cataloging-in-Publication Data
A catalog record for this book is available from the Library of Congress.
ISBN: 0-8247-4255-9
This book is printed on acid-free paper.
Headquarters
Marcel Dekker, Inc., 270 Madison Avenue, New York, NY 10016, U.S.A.
tel: 212-696-9000; fax: 212-685-4540
Distribution and Customer Service
Marcel Dekker, Inc., Cimarron Road, Monticello, New York 12701, U.S.A.
tel: 800-228-1160; fax: 845-796-1772
Eastern Hemisphere Distribution
Marcel Dekker AG, Hutgasse 4, Postfach 812, CH-4001 Basel, Switzerland
tel: 41-61-260-6300; fax: 41-61-260-6333
World Wide Web
http://www.dekker.com
The publisher offers discounts on this book when ordered in bulk quantities. For more
information, write to Special Sales/Professional Marketing at the headquarters address
above.
Copyright q2003 by Marcel Dekker, Inc. All Rights Reserved.
Neither this book nor any part may be reproduced or transmitted in any form or by any
means, electronic or mechanical, including photocopying, microfilming, and recording, or
by any information storage and retrieval system, without permission in writing from the
publisher.
Current printing (last digit):
10987654321
PRINTED IN THE UNITED STATES OF AMERICA

2
A Taxonomic Review of the Clavicipitaceous
Anamorphs Parasitizing Nematodes
and Other Microinvertebrates
Walter Gams
Centraalbureau voor Schimmelcultures, Utrecht, The Netherlands
Rasoul Zare
Plant Pests and Diseases Research Institute, Tehran, Iran
1. INTRODUCTION
The association between nematodes and fungi that colonize them has been the
subject of extensive mycological studies. The voluminous and meticulous work
by Charles Drechsler (1892 – 1986; see references) has laid the foundation for our
knowledge of nematophagous fungi. Numerous publications by Barron (since
about 1969, particularly 1977, 1981, 1982, and other papers cited) subsequently
augmented our knowledge about the diversity and distribution of these fungi. A
detailed understanding of their classification in a teleomorph-based fungal
system is emerging only in recent years.
The nematophagous fungi in the broadest sense (for recent reviews see,
e.g., Barron, 1977, 1981, 1982; Gray, 1988; Kerry and Jaffee, 1997) comprise
zoosporic, zygomycetous (Zoopagales), ascomycetous (anamorphic Orbiliaceae
and Clavicipitaceae), and basidiomycetous (Pleurotaceae) representatives. In a
17

first instance, they are subdivided into nematode-capturing fungi, endoparasites
of the free-living animals, and cyst and egg parasites. While nematode-capturing
capacities are found mainly in some Zoopagales, the ascomycete family
Orbiliaceae and basidiomycetes of Hohenbuehelia, the majority of nematode
endoparasites and egg parasites belong to the Clavicipitaceae and associated
anamorph taxa. Not only nematode-capturing but also endoparasitic parasites
attract nematodes (Jansson and Nordbring-Hertz, 1988). The endoparasites
colonize the animal from small propagules, produce internal mycelium, and
finally penetrate the corpse to sporulate on its surface. Endoparasites generally
employ two modes to penetrate the host animal, oral ingestion of conidia which
germinate in the intestines (mostly the esophagus or mastax), or direct
penetration of the cuticle from conidia which adhere firmly to the surface. This
adhesion occurs only in certain combinations of host and parasite taxa as
reviewed below for Drechmeria,Haptocillium, and Hirsutella. From the attached
conidium a narrow penetration tube grows through the nematode cuticle,
apparently with some mechanical pressure (Dackman et al., 1992; Dijksterhuis
et al., 1994). Egg parasites will be treated further below. Besides nematodes,
mainly terrestrial bdelloid rotifers (genus Adineta), some ciliated Protozoa, and
rarely Tardigrada have been found to be colonized by similar fungi.
The nematode body is covered by a thick proteinaceous cuticle, the eggs by
lipid and chitin layers, and an outermost protein layer characterized as vitellin
(Jatala, 1986; Bonants et al., 1995), and the cysts of certain genera by protein and
mucopolysaccharide. These recalcitrant layers provide barriers to penetration
that can be crossed only by enzymatically qualified fungi (Morgan-Jones et al.,
1983; Stirling, 1991). Strong chitinolytic activities are observed in many of the
fungi involved, which share this capacity with most entomogenous and many
fungicolous fungi. It has been hypothesized, in a phylogenetic analysis of
Cordyceps species, that a jump from entomogenous to fungicolous lifestyle is
possible within a certain common biotope (Nikoh and Fukatsu, 2000). Therefore
it is conceivable that certain endoparasites could also have jumped from insects
to nematodes or other animals. Such a jump has been suggested to have occurred
within a species of Lecanicillium (Hall, 1980) or between closely related taxa.
Some of the species to be mentioned have also been isolated as apparently
saprotrophic soil fungi.
Nematodes are subdivided into saprotrophic or bacterial feeders, plant-
pathogenic, and predatory groups. Parasites affecting the second group have most
frequently been studied in view of their possible role in biological control.
However, the plant-feeding nematodes have a specialized narrow mouth
apparatus with a stylet, which prevents ingestion of fungal propagules. These
animals can only be attacked in their free-living stage by fungi that produce
adhesive conidia. Bacterial feeder nematodes are much more prone to parasitism
by both nematode-capturing and endoparasitic fungi. They can ingest conidia of
Gams and Zare18

their parasites, such as Harposporium species, through their wider mouth
openings. For the adhesion of conidia of Drechmeria (or to a lesser extent of
Haptocillium) to specific parts of the body, chemical factors are responsible,
which are still not sufficiently elucidated (see below under Drechmeria). The
difference between bacteriophagous and phytophagous nematodes is of great
ecological significance in relation to endoparasitic and other nematophagous
fungi. The fact that we see relatively little host specificity is inconsistent with the
“host relatedness hypothesis” proposed for endoparasites (Nikoh and Fukatsu,
2000). Particularly in Harposporium and Rotiferophthora, a broad range of taxa
occurs on a rather limited array of prey animals.
In contrast with the numerous free-living nematodes, some plant-
pathogenic nematodes spend most of their life inside plant roots or on their
surface in cysts and/or root knots. These resting structures persist in the soil and
act as a selective substratum for fungal colonization by egg parasites. Certain
clavicipitaceous anamorphs, now comprised in the genus Pochonia (Zare et al.,
2001), are specialized to parasitize such resting structures (for a review see
Morgan-Jones and Rodrı
´guez-Ka
´bana, 1988). In addition, the free-living phase of
the same nematodes can be attacked by a different array of parasites, mainly of
Haptocillium and Hirsutella.
Species of Pochonia,Haptocillium, and Hirsutella are therefore among the
most promising biocontrol agents against plant-pathogenic nematodes (Dackman
et al., 1992; Kerry, 1987, 1989, 1990, and later work; Tribe, 1979, 1980; Stirling,
1988, 1991). Such endoparasitic fungi are often considered more amenable to
practical applications than nematode-capturing fungi (Persmark et al., 1996).
Biological control can manifest itself in three situations: (1) natural control,
which includes decline phenomena, is mainly depending on soil characteristics;
(2) addition of antagonists together with additional nutrient sources, which often
requires application of prohibitively high quantities of inoculum; or (3) the
stimulation of the resident population of antagonists, e.g., by organic
amendments or various cropping systems (Dackman et al., 1992; Stirling,
1989, 1991). Practical approaches to biological control therefore include the
exploitation of naturally suppressive soils, soil amendments to encourage the
activity of indigenous nematode parasites, application of selected strains of
bacteria or fungi, and the application of fungal toxins and enzymes (Jansson et al.,
1997; Kerry, 1990; Stirling, 1989, 1991). Seed treatment is considered less
feasible for these fungi because of poor reproduction in the rhizosphere (Kerry
and Jaffee, 1997). A fungus, Pochonia chlamydosporia, and a bacterium,
Pasteuria penetrans, are regarded as promising control agents in small-scale
tropical agriculture with low input of chemicals (Davies et al., 1991).
Clavicipitaceous Anamorphs Parasitizing Nematodes 19

2. METHODS OF STUDY
Methods of handling nematophagous fungi have been detailed by Duddington
(1955), Barron (1969, 1977, 1981, 1982), Gray (1984), and Bailey and Gray
(1989). The sprinkled soil plate method introduced by Drechsler (1933) and
frequently modified (e.g., Barron, 1977) retrieves all kinds of nematophagous
fungi. Particles of soil or various organic substrata are spread onto water agar or
a dilute cornmeal agar, and nematodes (particularly Panagrellus redivivus,
Turbatrix aceti, or others) are added as baits. The plates are checked at weekly
intervals for fungi growing on or out from dead nematodes. To retrieve
endoparasitic species preferentially, the Baerman funnel technique, adopted for
this purpose by Giuma and Cooke (1972), is significantly more effective (Barron,
1977, 1978, 1982; Gray, 1984). However, both methods are usually employed
together to reveal the maximum of taxa present in a soil. A differential
centrifugation technique was proposed by Barron (1969): the supernatant of a soil
suspension obtained after gentle centrifugation is supposed to contain most
conidia; when spun down at high speed, the pellet obtained from the first
supernatant can be plated on agar with nematodes. Subsequent comparative
studies by Barron (1978) and Gray (1984) showed that this technique had little
additional advantage over the Baerman funnel. Dackman et al. (1987) combined
a dilution plate method (otherwise similar to the soil sprinkling method) with
most probable number analysis to quantify populations of nematode parasites.
Persmark et al. (1996) almost exclusively obtained zoosporic parasites with this
method and therefore discontinued its use. Banck et al. (1990) compared methods
for retrieving plant-parasitic nematodes and their parasites. They retrieved
species of Harposporium and Hirsutella by Seinhorst’s elutriation and Cobb’s
sieving, decanting, and centrifugation method with either silica or sugar
solutions. To detect ovicidal fungi in soil, Fassatiova
´and Lysek (1982) buried
eggs of Ascaris lumbricoides in soil.
To recover parasites of bdelloid rotifers from samples of soil or organic
debris, Barron (1985) used cultured rotifers of the genus Adineta (originally
recovered from sprinkled-soil plates as used for nematophagous fungi) as bait.
The animals are maintained in Petri dishes in a thin film (2 mm) of water or
physiological saline layered over sucrose-free Czapek agar. Cultures of rotifers are
maintained by periodic transfer (1 – 2 weeks) to fresh Petri dishes. From 10 to 25 g of
soil or organic debris are mixed with an equal volume of sterile water in a plastic bag
and squeezed and agitated vigorously for several minutes. Then 1 –2 mL of the
slurry is transferred to a rotifer culture in a Petri dish and swirled gently. The dishes
are incubated at 18– 228C and examined at weekly intervals for 3– 6 weeks.
Infected rotifers can be spotted under a dissecting microscope by a cluster of
conidiophores arising from the floating body of a dead rotifer. These rotifers are
Gams and Zare20

then transferred with a flattened needle to a fresh rotifer culture; new crops of
infected rotifers will appear in 3 – 7 days and can be used for preparing pure cultures.
For methods developed for specific taxa, see the texts under each genus.
Mass production of inoculum for nematode control has not yet been
upscaled industrially. To assure optimal longevity and infectivity of the
conidia, the fungi are generally grown in solid-state surface cultures; see under
Pochonia chlamydosporia. For the production of Hirsutella rhossiliensis
inoculum, stirred cultures in 5-L containers were used (Patel et al., 2001).
3. ECOLOGY
The ecology of nematophagous fungi has been reviewed extensively
(Duddington, 1951; Barron, 1977, 1981; Gray, 1983a, 1984). Soil and various
organic substrata, particularly dung (Juniper, 1967; Glockling and Yamada,
1997), are suitable sources for nematophagous fungi. Extending this range,
Gray (1983b) found nematode endoparasites in deciduous and conifer litter, old
dung, moss cushions, and decaying vegetation. Addition of farmyard manure to
agricultural soil increased the population of endoparasites (Dackman et al.,
1987).
Densities of Harposporium anguillulae in a Swedish agricultural soil
reached maxima in March and June. The highest densities of nematophagous
fungi in general were found in the upper 40 cm, with Harposporium anguillulae
going down to 30 –40 cm and Hirsutella rhossiliensis strongly declining after
20 cm. During fallow periods the population of nematode parasites declined
(Persmark et al., 1996). Certain nematophagous fungi were stimulated under
plants with a strong rhizosphere effect, particularly peas; among these,
Harposporium anguillulae was found on extracted nematodes in 4 of 5 soils
examined, and Haptocillium balanoides in one rather acidic soil, both grown with
barley (Persmark and Jansson, 1997).
Nematodes are attracted to colonies of Drechmeria coniospora,
Haptocillium balanoides, and other endoparasites (Barron, 1982; Jansson,
1982a, 1982b; Jansson and Nordbring-Hertz, 1979, 1980). As a mechanism of
nematicidal action, antibiotic (antifungal) activities have been demonstrated for
Drechmeria coniospora,Harposporium anguillulae (Barron, 1977), Lecanicil-
lium (Ha
¨nssler, 1990), Paecilomyces lilacinus (Jatala, 1986), and Pochonia
(Segers et al., 1999).
Even the large eggs of Ascaris can be parasitized when exposed in soil.
Fassatiova
´and Lysek (1982) obtained Pochonia chlamydosporia,P. bulbillosa,
Paecilomyces marquandii,P. lilacinus, and P. carneus from such eggs buried in
soils in the Czech Republic, Pakistan, Afghanistan, and Cuba. Pochonia spp. and
P. lilacinus rapidly infected and killed the eggs.
Clavicipitaceous Anamorphs Parasitizing Nematodes 21

4. DISTRIBUTION
Most species of nematode endoparasites were originally described in the United
States and Canada. Reports from other countries, however, indicate an almost
cosmopolitan distribution, but a few species are either tropical or temperate.
Species lists have been compiled from Ireland (Gray, 1983b), New Zealand
(Hay, 1995), and El Salvador (Bu
´caro, 1983); and from vegetation and soils in the
maritime Antarctic (Gray, 1982; Gray et al., 1982; Gray and Lewis Smith, 1984).
Pochonia chlamydosporia is one of the most cosmopolitan species, but its
Cordyceps teleomorph is so far known only from slug eggs in the tropics
(Zare et al., 2001). Drechmeria coniospora,Haptocillium balanoides,Harpos-
porium anguillulae, and Hirsutella rhossiliensus were also encountered infre-
quently in Central America (Persmark et al., 1995).
5. MORPHOLOGY
In axenic culture, colonies are slow- to medium-fast growing (reaching 5 –40 mm
in diameter in 10 days at about 208C on common laboratory media, depending on
the taxon), white to yellowish, with some cottony aerial mycelium, often
consisting largely of fertile hyphae or conidiophores. In most taxa conidiophores
are at least partly verticillate, either erect or soon procumbent or prostrate, so that
indefinite numbers of phialides can arise from arched hyphae of the aerial
mycelium. Conidiogenesis is mostly phialidic; sometimes only solitary conidia
are formed on a conidiogenous cell, i.e., blastic conidiogenesis. Phialides are
more or less swollen in the lower part or aculeate, with a single (monophialide)
or several conidiiferous tips (polyphialides or polyblastic conidiogenous cells).
If the conidiogenous cells are phialidic, they produce usually several conidia in
slimy heads, sometimes in fascicles that aggregate at the tip in a position
perpendicular to the phialide (characteristic of Lecanicillium). Conidia are mostly
one-celled, of various shapes and sizes. Some taxa produce complex resting and
propagative structures called dictyochlamydospores, i.e., hyaline, thick-walled,
pluricellular bulbil-like structures supported by a short stalk. The development of
the characteristic dictyochlamydospores was studied by Campbell and Griffiths
(1975). These dictyochlamydospores can also be absent in some strains of species
supposed to produce them, but then some irregularly swollen, thick-walled
intercalary cells are usually present in the vegetative hyphae.
6. TAXONOMY
The taxonomy of clavicipitaceous nematode parasites has been considerably
modified in recent years, leading to the distinction of several phylogenetically
distinct genera. Drechmeria was segregated from Meria (Gams and Jansson,
1985); its unrelatedness with Meria Vuill. (Rhytismatales) and its affinity with
Gams and Zare22

the Clavicipitaceae has been proven (Gernandt and Stone, 1999). The type
species of Tolypocladium was connected with the teleomorph Cordyceps
subsessilis Petch (Hodge et al., 1996), but the nematophagous species of
Tolypocladium are not yet critically classified. Some species demonstrate a
continuum between the genera Harposporium and Hirsutella, producing two
kinds of conidia with the associated kinds of conidiogenesis (Hodge et al., 1997;
Glockling 1998b). The species of the former Verticillium sect. Prostrata W. Gams
(1971) comprised several groups of nematophagous taxa as reviewed by Gams
(1988), but molecular analyses (Zare et al., 2000; Sung et al., 2001) have shown
that members of this section are heterogeneous and must be distributed among
several genera (Gams and Zare, 2001), a classification that is adopted here and
elaborated below in reference to microinvertebrate-parasitizing taxa. Barron
(1991a) had already singled out the very slowly growing rotifer parasites in a
separate genus, Rotiferophthora Barron. The most characteristic nematophagous
verticillium-like genera are Haptocillium W. Gams & Zare, comprising species
with adhesive conidia that stick to free-living nematodes, and Pochonia Bat. &
O. M. Fonseca, formerly often called Diheterospora Kamyschko ex Barron &
Onions, species which are particularly capable of penetrating nematode cysts and
eggs. The production of dictyochlamydospores was mostly used to characterize
Diheterospora, but this is an unreliable criterion for recognizing species of this
genus, because they are absent or scanty in some species, while similar structures
also occur in species of Rotiferophthora and Haptocillium. The present
separation of several verticillium-like genera has the advantage of reflecting
correlated ecological, morphological, and phylogenetic traits.
Methods for identification can be summarized as follows. Colonies can be
grown on various media that are not too rich in nutrients, such as cornmeal agar
(Difco), a dilute oatmeal agar (OA), malt extract agar (not more than 2% sugar),
synthetic nutrient-poor agar (SNA), potato-carrot agar, or even water agar (Gams
et al., 1998a). Transfer by streak inoculation is recommended to induce good,
homogeneous sporulation from colonies developing from a conidial inoculum.
Direct observation of the undisturbed colony in the open Petri dish under the
compound microscope allows the observation of the branching system and
structures of the conidiophores and the arrangement of the conidia in situ.
Microscopic mounts are made in lactic acid (sometimes with cotton blue or a
similar stain) and recorded in camera-lucida drawings or photographs.
7. KEY TO THE GENERA THAT INCLUDE PARASITES OF
NEMATODES AND OTHER MICROSCOPIC ANIMALS
In this general survey, we illustrate representative species of each genus mainly
with drawings taken from original publications. In the special part we add some
drawings of our own.
Clavicipitaceous Anamorphs Parasitizing Nematodes 23

1. Conidiogenous cells with almost globose venter and sharply delimited
slender neck.......................................................................................... 2
10. Conidiogenous cells aculeate or with moderately inflated venter or of
reduced shape.......................................................................................3
2. Conidia bearing curved phialoconidia on one or several necks
(cylindrical in some Harposporium species); generally parasitizing
nematodes or bdelloid rotifers ...................... 5. Harposporium (Fig. 1)
FIGURE 1Harposporium, conidiophores arising from infected nematodes
and conidia: (a) H. anguillulae. (From Zopf, 1888.) (b) H. helicoides. (From
Drechsler, 1941.)
Gams and Zare24

20. Conidia bearing globose to cylindrical conidia, usually from
single necks; soilborne or entomogenous, rarely associated with
rotifers ....................................................................14. Tolypocladium
3. Intercalary conidiogenous cells (mostly phialides) with short
conidiiferous necks commonly produced .......................................... 4
30. Intercalary conidiogenous cells absent (or rarely formed in
Haptospora) ....................................................................................... 6
4. Intercalary phialides mostly produced singly, frequently in
verticillate end-branches of the conidiophore; a conspicuous oil
globule present in each conidium; parasites of rotifers; dictyo-
chlamydospores present ......................... 12. Rotiferophthora (Fig. 2)
40. Several intercalary conidiogenous cells produced below a terminal
one; dictyochlamydospores absent .................................................... 5
5. Conidiogenesis phialidic with single openings; conidia obclavate
with adhesive tip; parasitizing nematodes or ciliated protozoa ............
......................................................................... 2. Drechmeria (Fig. 3)
50. Conidiogenesis polyblastic; conidia globose, with a conspicuous
basal slime pad; parasitizing rotifers11. Pseudomeria mucosa (Fig. 4)
6. Conidiogenous cells with more or less swollen venter; discrete
dictyochlamydospores absent ............................................................ 7
60. Conidiogenous cells hardly swollen, aculeate, often in whorls;
dictyochlamydospores often present ............................................... 12
7. Conidiogenous cells phialidic, with a flaring collarette; conidia with
basal appendage .............................................. 4. Haptospora (Fig. 5)
70. Conidiogenous cells phialidic or with solitary conidia, apparently
blastic, lacking a discernable collarette; conidia lacking a basal
appendage........................................................................................... 8
8. Conidia adhering in regular chains.............. 8. Paecilomyces (Fig. 6)
80. Conidia adhering in heads or formed singly ..................................... 9
9. Conidiophores synnematous or mononematous; conidia with a
distinct, chromophilic slime layer or covered by a finely warted
epispore, often somewhat fusiform ................... 6. Hirsutella (Fig. 7)
90. Conidiophores mononematous; conidia thin- and smooth-walled..10
10. Conidiogenous cells single, with only the tips protruding from the
nematode; in vitro single swollen phialides supported by slender
stalks................................................................ 9. Plesiospora (Fig. 8)
100. Complex conidiophores appearing outside the host animal, more or
less verticillate ................................................................................. 11
11. Conidia with a distal adhesive surface, appearing as a wall
thickening; parasites of nematodes... ............ 3. Haptocillium (Fig. 9)
110. Conidia lacking an adhesive structure; parasites of rotifers ...............
................................................................ 14. Tolypocladium (Fig. 10)
Clavicipitaceous Anamorphs Parasitizing Nematodes 25

FIGURE 2Rotiferophthora globispora, conidiophores and dictyochlamy-
dospores arising from an infected rotifer. (From Barron, 1991a.)
Gams and Zare26

12. Conidiophores erect and well differentiated (stipe usually thick-
walled).................................see 10. Pochonia suchlasporia (Fig. 18)
120. Conidiophores usually prostrate, sometimes also erect, but hardly
differentiated from vegetative hyphae............................................. 13
FIGURE 3Drechmeria coniospora, conidiophores and conidia arising from
infected nematodes; conidia, some with developing adhesive knob;
nematode with conidia attached at the cephalic and anal regions: (a) from
Drechsler (1941); (b) from Barron (1977).
Clavicipitaceous Anamorphs Parasitizing Nematodes 27

13. Conidiogenesis polyblastic, with conidia either on sympodially
produced denticles of terminal conidiogenous cells or on densely
crowded, rapidly collapsing denticles laterally along intercalary cells
of prostrate fertile hyphae; dictyochlamydospores absent ..............14
130. Conidiogenesis phialidic; phialides aculeate, more or less persistent,
each producing several conidia ....................................................... 15
FIGURE 4Pseudomeria mucosa, two infected rotifers with juvenile and
mature conidiophores, detail of conidiophores, and conidia with mucous
sheath, in those at the bottom sheath distorted after attachment. (From
Barron, 1980b.)
Gams and Zare28

14. Conidiiferous denticles persistent, mostly in terminal position,
sometimes inserted next to discrete conidiogenous cells ...................
.........................see Beauveria and Microhilum (entomogenous taxa)
140. Conidiiferous denticles scattered along cells of fertile hyphae, soon
collapsing aphanophialides; colonies deeply woolly ..........................
..................................... aphanocladium-like species of Lecanicillium
15. Colonies slow-growing, reaching 5 –15 mm diam in 10 d; parasites of
free-living nematodes or rotifers; dictyochlamydospores often
present .............................................................................................. 16
150. Colonies growing faster, reaching 15 – 40 mm diam in 10 d; growing
on insects or fungi; if attacking nematodes, then parasitizing cysts or
eggs; dictyochlamydospores present or absent...............................17
16. Parasites of bdelloid rotifers; intercalary phialides with a lateral neck
normally present below terminal, flask-shaped or elongate phialides;
conidia adhering in heads; a conspicuous oil globule present in each
conidium; dictyochlamydospores commonly present, often flattened
....................................................................... see 12. Rotiferophthora
160. Parasites of free-living nematodes; conidia balanoid, campanulate
to cylindrical, subglobose to irregularly angular, mostly terminally
adhesive (visible as a wall thickening at the upper, more or less
FIGURE 5Haptospora, conidiophores arising from infected rotifer and
extra enlarged conidia: (a) H. appendiculata. (From Barron, 1991b.)
(b) H. endoparasitica. (From Barron and Szijarto, 1982b.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 29

truncated end), produced in heads or short chains or both;
sesquiphialides absent; dictyochlamydospore-like structures some-
times present................................................. 3. Haptocillium (Fig. 9)
17. Phialides exclusively solitary (if verticillate, conidia narrowly
acerose); dictyochlamydospores absent.... 13. Simplicillium (Fig. 11)
170. Phialides at least partly in whorls ................................................... 18
FIGURE 6Paecilomyces lilacinus, conidiophores and conidia from various
isolates or specimens. (From Samson, 1974.)
Gams and Zare30

FIGURE 7Hirsutella rhossiliensis, conidiophores and conidia, drawn with
and without the slime layer. (From Minter and Brady, 1980.)
FIGURE 8Plesiospora globosa, infected nematode with conidiophores and
conidia developing in agar culture. (From Drechsler, 1971.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 31

FIGURE 9
Gams and Zare32

18. Conidia subglobose to short-ellipsoidal, sometimes short-falcate,
often cyanophilic; dictyochlamydospores spherical or irregularly
shaped, often present in the aerial mycelium or in the agar; mostly
parasites of nematode cysts or saprotrophic, soilborne; crystals absent
in the medium ................................................. 10. Pochonia (Fig. 12)
180. Conidia short- or long-ellipsoid to cylindrical or falcate, not con-
spicuously cyanophilic; chlamydospores or dictyochlamydospores
absent (mostly entomogenous, fungicolous, or soilborne); crystals
abundantly produced in the medium; sporulation with aculeate
phialides predominant; denticles with blastoconidia, if present,
densely scattered along the cells of fertile hyphae; on various
substrata..................................................... 7. Lecanicillium (Fig. 13)
[If conidia 2-celled, see 1. “Cephalosporiopsis” carnivora (Fig. 14)]
8. THE GENERA
8.1. Cephalosporiopsis Peyronel, Mem. R. Accad.
Sci. Torino, Ser. 2, 66:52, 1916 (Fig. 3)
Type species: Cephalosporiopsis alpina Peyronel.
This genus is not generally recognized because the identity of the type species
is doubtful and species assigned to the genus are extremely heterogeneous.
The best-known species that was often referred to the genus is now recognized
as Plectosporium tabacinum (van Beyma) Palm et al., anamorph of
Plectosphaerella cucumerina (Lindf.) W. Gams in the Phyllachoraceae (Palm
et al., 1995) (syn. Cephalosporiopsis imperfecta C. Moreau & M. Moreau).
Cephalosporiopsis carnivora Drechsler (1969) was observed on free-living
Rhabditis sp., but is not available in culture for further study. It is characterized
by discrete, short flask-shaped phialides in moderately verticillate, generally
erect conidiophores and elongate-ellipsoid or somewhat obovoid conidia, mostly
divided by a cross-wall at a slight median constriction, mostly 3:0–4:2£
1:7–2:2mm:Frequently 1 or 2 conidia were seen adhering to the forward profile
of an actively motile eelworm. This statement suggests an adhesive mechanism
that would place the species near Haptocillium.
FIGURE 9Haptocillium balanoides, infected specimen of Acrobeloides
buetschlii with outgrowing hyphae and conidiophores, detail of conidia
attached to the head region, and three extra enlarged conidia. (From
Drechsler, 1941.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 33

8.2. Drechmeria W. Gams & H.-B. Jansson,
Mycotaxon 22:33, 1985 (Fig. 3)
Type species: Drechmeria coniospora (Drechsler) W. Gams & H.-B.
Jansson.
Diagnosis: Colonies very slow-growing; mycelium hyaline. Fertile hyphae
erect, bearing one terminal and 2 – 7 intercalary phialides, the intercalary
ones bearing acropleurogenous conidiiferous pegs below the upper
septum, forming several conidia in basipetal succession, which form
stellate clusters. Conidia of D. coniospora are conical, with rounded base
FIGURE 10 Tolypocladium lignicola, schematic drawing of an infected rotifer
with conidiophores growing out above the water surface and some
chlamydospores below, details of conidiophores and more strongly
enlarged conidia, those at the bottom after 24 h exposing to soil extract.
(From Barron, 1983).
Gams and Zare34

and almost pointed tip, one-celled, hyaline, smooth-walled, 4 –7 £
1:8–2:5mm:
Second species:D. harposporioides (Barron & Szijarto) W. Gams & H.-B.
Jansson (Meria harposporioides Barron & Szijarto, 1982a), a parasite of
sessile ciliated protozoans, which differs from D. coniospora by having
falcate conidia, 7:5–12 £1:7–2:0mm:
Conidiogenesis and penetration of nematodes were examined with light- and
electron-microscopic methods (Jansson et al., 1984; in more detail, Dijksterhuis
et al., 1991). The nematodes survived about 24 h after infection. The fungus can
produce 5000– 10,000 conidia at the expense of a single nematode. The
formation of adhesive knobs on the tip of the conidia is an autonomous process in
Drechmeria coniospora at the end of conidial maturation (van den Boogert et al.,
1992). This adhesive knob consists of radiating fibrils visible in TEM (Saikawa,
1982a), with best resolution in KMnO
4
-fixed material (Saikawa, 1982b). After
attachment, an infection vesicle is formed between the cuticle layers (TEM by
Dijksterhuis et al., 1990; Sjollema et al., 1993). Collagenase production is
induced before penetration of the cuticle (Jansson et al., 1985a). Hyphae
penetrate the nematode via the pseudocoel, without attacking internal organs.
FIGURE 11 Simplicillium lanosoniveum, phialides and conidia. (From Zare
and Gams, 2001a.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 35

FIGURE 12 Pochonia chlamydosporia var. catenulata;left,several
conidiophores with catenate conidia and a few extra enlarged conidia; right,
development of dictyochlamydospores. (From Barron and Onions, 1966.)
FIGURE 13 Lecanicillium lecanii, conidiophores and conidia. (From Gams
and Zare, 2001.)
Gams and Zare36

Trophic hyphae contain numerous lipid droplets, often associated with
microbodies (Dijksterhuis et al., 1991). Nematodes with attached conidia were
seen in animals recovered from soil (cryo-SEM by Jansson et al., 2000).
Conidia can be ingested by nematodes but do not germinate in the intestine
(Jansson, 1994); direct penetration from adhering conidia is thus the only
mechanism of infection. The fungus attracts susceptible nematodes (Jansson,
1982a, 1982b). Based on observation of a limited number of potential hosts
(Du
¨rschner, 1983), the conidia were seen to adhere specifically in the mouth
region, where chemoreceptors are situated (Jansson and Nordbring-Hertz, 1983),
in male nematodes also in the anal region of certain species. Infected animals are
disturbed and are no longer attracted by colonies of the fungus. In more extended
studies, this localized mode of adhesion was found in bacteriophagous, a few
plant-parasitic (Meloidogyne and Aphelenchus), and animal-parasitic nematodes,
while other plant-parasitic nematodes (Pratylenchus, Ditylenchus and Cricone-
mella) became infected at any point, but were then not equally strongly
FIGURE 14 Cephalosporiopsis carnivora, infected specimens of Rhabditis sp.
with outgrowing conidiophores developing on an agar plate, and conidia,
some extra enlarged. (From Drechsler, 1969.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 37

parasitized (Jansson et al., 1987). Insect-parasitic species of Neoaplectana and
Heterorhabditis occasionally showed adhesion of conidia but normally no further
penetration (Poinar and Jansson, 1986), while plant-parasitic species of Rhabditis
were susceptible. On Acrobeloides conidia also adhered but without subsequent
penetration (Dijksterhuis et al., 1993). The observations on nematode attraction
and conidial adhesion were reviewed by Dackman et al. (1992). In several studies
before 1990, sialic acid (acetyl-neuraminic acid), localized in the head and tail
regions, was assumed to bind to a lectin located on the parasite’s conidia. Sialic
acid was the only one of 21 carbohydrates tested that inhibited adhesion. Limulin,
a lectin specifically binding sialic acid, reduced attraction to the fungus and
adhesion of conidia (Jansson and Nordbring-Hertz, 1983, 1984). A carbohydrate
receptor for chemotaxis with sialyl- and mannosyl-related residues was
demonstrated in Caenorhabditis elegans and Panagrellus redivivus. In support
of this hypothesis, adhesion was shown to be inhibited by flooding conidia with
sialic acid or by treating nematodes with sialidase, a treatment that also reduced
attraction of nematodes to the fungus (Jansson and Nordbring-Hertz, 1983, 1984;
Jansson et al., 1985a, 1987). However, no lectin could be isolated from the
conidia and no sialic acid residues were directly demonstrated in the cephalic
region of P. redivivus. Other compounds mimicking sialic acid were then
suggested as mediators of the effect. Surface receptors are still unknown in the
host, contrasting with those involved in the lectin-mediated contact between
nematode-capturing Arthrobotrys species and their prey (review by Tunlid et al.,
1992). Pronase treatment of the nematodes also prevents adhesion of these
conidia to Caenorhabditis elegans, but the nematodes regenerate the lost protein
material after 2 h in Tris buffer (Jansson, 1994). In the same paper, the adhesion is
suggested to be mediated by sensilla exudates. The fibrillar layer of the adhesive
knob is not altered during adhesion. This layer is dissolved in Pronase E. The
protease inhibitor chymostatin inhibits infection, suggesting involvement of
chymotrypsin-like proteases in the infection process (Jansson and Friman, 1999).
Motile nematodes infected with conidia of D. coniospora can also be caught by a
second nematophagous fungus. Dijksterhuis et al. (1994) found that hyphal
penetration by Arthrobotrys oligospora as a secondary invader is inhibited and its
hyphae are often dead in proximity to those of D. coniospora. Populations of
Meloidogyne incognita were reduced in 250-cm
3
pots with sterile or unsterile soil
by adding suspensions of 10
6
conidia per pot or 1000 living, infected Panagrellus
redivivus as carriers (Jansson et al., 1985b). Population densities of D. coniospora
in soil can be enhanced by increasing the organic matter content (by application
of lucerne meal or a barley root system), which first stimulates bacteriophagous
nematodes and, secondarily, the population of the fungus (van den Boogert et al.,
1994). In field soils, no positive rhizosphere effect was observed. Application of
this fungus for biological control is not considered feasible because of its narrow
host spectrum and because phytophagous nematodes are not intensely parasitized
Gams and Zare38

(van den Boogert et al., 1994; Dijksterhuis et al., 1993; Du
¨rschner, 1983; Jansson
et al., 1987).
8.3. Haptocillium W. Gams & Zare, Nova Hedwigia 72:334,
2001 (Fig. 9)
Type species:Haptocillium balanoides (Drechsler) Zare & W. Gams
(Verticillium balanoides (Drechsler) Dowsett et al., 1982).
Diagnosis: Colonies slow-growing, reaching 5– 15 mm diam on PDA after
10 days. Conidiophores erect or prostrate, bearing verticillate or solitary
phialides which are more or less swollen near the base. Conidia balanoid,
campanulate to cylindrical, subglobose to irregularly angular, mostly
terminally adhesive, produced in heads or short chains or both.
Dictyochlamydospores sometimes present. Mostly parasites of free-
living nematodes, to which the conidia become attached by means of an
adhesive apical layer. A detailed account of the genus is given by Zare
and Gams (2001b).
Gams (1988) synonymized several taxa described by Drechsler under
V. balanoides. Molecular studies have shown, however, that more species are to
be distinguished.
8.3.1. Key to the Species (modified after Zare and Gams,
2001b)
1. Conidia at least partly in chains, sometimes in heads ........................ 2
10. Conidia always in heads, never in chains ........................................... 3
2. Conidia slightly irregularly angular, small, 1:3–2:0£1:0–1:7mm;
in heads and short chains.......................................................................
H. sinense (K. G. Zhang, L. Cao & Z. Q. Liang) Zare & W. Gams
[If conidia globose, 1:3–1:8mm diam, then “Verticillium” cocco-
sporum (Drechsler) W. Gams]
20. Conidia of two types; campanulate to cylindrical, 2:7–3:0£
1:3–1:5mm;in heads, and globose to subglobose, 4:2–4:5£
3–3:5mm;often catenate .......... H. campanulatum (Glockling) Zare
& W. Gams
3. Conidia globose with mostly 5 apical adhesive buds,
3:5–5:0£2:2–2:5mm......... “Verticillium” coronatum Barron (1989)
30. Conidia cylindrical...............................................................................4
300. Conidia balanoid to elongate balanoid ................................................5
4. Conidia cylindrical to campanulate, 2:7–3 £1:3–1:5mm..................
.............................................................................see H. campanulatum
Clavicipitaceous Anamorphs Parasitizing Nematodes 39

40. Conidia cylindrical, 3:8–4:0£1:3–1:5mm.........................................
.....................................................H. rhabdosporum Zare & W. Gams
5. Conidia triangular to elongate-balanoid, with inconspicuous terminal
wall thickening, measuring 3:0–4:0£1:7–2:0mm;elongate conidia
absent...............................H. zeosporum (Drechsler) Zare & W. Gams
50. Balanoid conidia mostly with pronounced terminal wall-thickening,
elongate conidia scarce or frequent..................................................... 6
6. Balanoid to triangular conidia mostly 2:5–3:0£1:5–2:0mm;
elongate conidia ð5:0–7:5£1:5–2:0mm) frequent; phialides slender,
aculeate on OA; polyphialides absent ....................................................
........................................ H. balanoides (Drechsler) Zare & W. Gams
60. Balanoid to subglobose conidia 2:2–3:2£1:3–2:3mm;elongate
conidia scarce (if present); phialides slender or swollen on OA;
polyphialides usually absent...........H. sphaerosporum (J.B. Goodey)
Zare & W. Gams (Fig. 15)
600. Balanoid conidia 3:2–3:6£2:5–3:0mm;elongate conidia rather
frequent, 5:7–6:5£2:0–2:2mm;polyphialides frequent ....................
......................................................... H. glocklingiae Zare & W. Gams
Because of the lectotypification of H. balanoides by CBS 250.82 (Gams,
1988), an isolate that takes a somewhat isolated position among those available,
most of the isolates commonly identified as that species now have to be called
H. sphaerosporum (Fig. 15), and most of the information compiled below is
likely to refer to that species.
8.3.2. Ecology and Application
The conidia of H. balanoides attract nematodes (Jansson, 1982a, 1982b).
Drechsler (1941) originally described Cephalosporium balanoides from Plectus
parvus and Acrobeloides buetschlii. The species was then reported as an
endoparasite of Rhabditis terricola from various soils in Ontario (Barron, 1978),
in nematodes in mosses in a birchwood in England (Duddington, 1951), and from
deciduous and coniferous leaf litter, old dung, and coastal vegetation in Ireland
(Gray and Duff, 1982; Gray, 1983c). Haptocillium balanoides occurred in 50% of
agricultural soil samples taken in Westfalen, Germany (Du
¨rschner, 1983). It was
also commonly encountered in the maritime Antarctic (Gray et al., 1982; Gray
and Lewis Smith, 1984). The species was regularly found on nematodes in
Manitoba soils (Dowsett et al., 1982). The host range included all species of the
Rhabditida, Aphelenchida, and Tylenchida tested, but not the Dorylaimida
(Du
¨rschner, 1983). The fungus seems to recognize suitable host nematodes
(Du
¨rschner-Pelz and Atkinson, 1988). The temperature minimum for growth was
about 158C (Gams, 1988). Haptocillium balanoides (sensu lato) was frequently
observed in nematodes associated with needles of decaying Japanese red pine
Gams and Zare40

trees (Watanabe, 2000); it has also been reported from nematodes in various soils
of El Salvador (Bu
´caro, 1983). An over-winter decline in populations of the stem
nematode Ditylenchus dipsaci was ascribed to infections by H. balanoides
(Atkinson and Du
¨rschner-Pelz, 1995). Conidial production was assessed on
different host nematodes: the most frequent host, the stem nematode Ditylenchus
dipsaci, yields about 15,000 conidia per cadaver, Globodera rostochiensis about
11,600, and Panagrellus redivivus about 840. This could mean that the
association with phytophagous nematodes is advantageous to the parasite, while
bacteriophagous nematodes may attract too many other microbes that antagonize
the endoparasite (Atkinson and Du
¨rschner-Pelz, 1995).
FIGURE 15 Haptocillium sphaerosporum, conidiophores and conidia from
agar cultures of several isolates. (From Zare and Gams, 2001b.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 41

The great differentiation of species recently observed in this genus will
necessitate detailed ecological studies with different isolates in order to select
efficient antagonists and to determine which are the most promising for
biological control in a particular soil (Hay, 1995; Hay and Regnault, 1995).
8.4. Haptospora Barron, Can. J. Bot. 69:503, 1991 (Fig. 5)
Type species:Haptospora appendiculata Barron.
Diagnosis: Conidiophores simple or slightly branched; conidiogenous cells
(phialides) flask-shaped, solitary or in clusters, with a membranous
collarette; intercalary phialides occasionally present. Conidia one-celled,
hyaline, with a basal appendage. Parasitizing bdelloid rotifers, forming a
mycelium of swollen cells.
8.4.1. Key to the Species
1. Conidia globose, 2:5–3:2mm diam .......................................................
..................................... H. endoparasitica (Barron & Szijarto) Barron
10. Conidia bilobed.................................................................................... 2
2. Conidia T-shaped, 2:8–3:2mm long, 2:8–3:2mm broad ......................
..................................... H. tribrachispora (Barron & Szijarto) Barron
20. Conidia Y-shaped, 5:5–6:5mm long, 5:0–7:0mm broad .....................
....................................................................... H. appendiculata Barron
As in Rotiferophthora, conidia of Haptospora are ingested and germinate
from the mastax (Barron, 1991b).
8.5. Harposporium Lohde, Tagebl. Vers. Deutsch. Naturf.
A
¨rzte Breslau 47:206, 1874
Type species:Harposporium anguillulae Lohde.
Diagnosis: Colonies slow-growing, whitish. Vegetative hyphae tending to
be broad and thick-walled, particularly when submerged in the agar.
Fertile hyphae with elongate aggregates or clusters of phialides. Phialides
mostly consisting of an almost globose venter and one to several narrow
conidiiferous necks. Conidia adhering in heads (in one species, in chains),
in most species gracefully bent, in others cylindrical or more irregularly
shaped. Most species of the genus parasitize nematodes, but some recently
described ones are found in bdelloid rotifers.
Synanamorphs: Arthroconidia or blastoconidia of Hirsutella (Glockling,
1998b; Hodge et al., 1997).
Gams and Zare42

Teleomorph of H. angullulae:Atricordyceps harposporifera Samuels,
New Zealand J. Bot. 21:171, 1983 (found on an arthropod [millipede?]).
Classical descriptions: Zopf (1888) gave a detailed description, establish-
ing its classification among the hyphomycetes, and a first description of
intercalary chlamydospores; Karling (1938) frequently found the species
in the U.S.A. and gave an extended description of the development of
conidiophores and conidia.
8.5.1. Key to the Species
(Measurements are given unaltered from the original diagnoses; some authors
give the total length, others the straight line spanning the curved conidia as
length. Species parasitizing nematodes unless otherwise stated.)
1. Conidia bent in circles or spirals..........................................................2
10. Conidia of other shapes ...................................................................... 16
2. Conidia with an almost straight base and bent distal part...................3
20. Conidia consistently bent throughout................................................... 4
3. Conidia with a circular bend in the upper part, 15– 25 (as a
straight line) £0.7–1.0 mm, with a terminal 2 – 5-mm-long point
filled with wall material, and with a viscid drop at the base ..................
....................................................................... H. oxycoracum Drechsler
30. Conidia with a rather sharp bend above the middle, base 14 –
17 mm, distal arm almost straight, 9– 11 mm, broadest part
2.2 mm ..................................................................... H. angulare Barron
4. Conidia bent in one direction ............................................................... 5
40. Conidial bends changing direction .....................................................11
5. Conidia forming about half a circle ..................................................... 6
50. Conidia forming less than half a circle................................................ 9
500. Conidia forming about three-fourths of a circle or ellipse................10
6. Besides helicoid conidia blastoconidia and/or arthroconidia commonly
produced................................................................................................ 7
60. Secondary conidia unknown or all conidia very small .......................8
7. Conidia 7 –16 (straight line, total length 12.5 – 35 mm) £0.7–
2.5 mm; arthroconidia conspicuous, cylindrical, 9.5–11.5 £3–
4mm................................................................. H. arthrosporum Barron
70. Circular conidia 9– 16 (total length) £1.0–2.0mm, obovoid
blastoconidia 3.0 – 6.5 £1.5–2.0mm, arthroconidia 14 – 17 £
4mm..................................................... H. janus Shimazu & Glockling
8. Conidia 13 –17 mm long (7 –13 mm when measured in a straight line),
1–2mm wide ....................................................... H. anguillulae Lohde
Clavicipitaceous Anamorphs Parasitizing Nematodes 43

80. Conidia smaller, 4.5– 9 (straight line?) £1–1.5mm; arthroconidia
present....................................................... H. lilliputanum M. S. Dixon
9. Conidia curved in ascending spiral, distally hardly pointed, 7.0–
(9.5)– 14.0 (straight line?) £0.6 – 0.9 mm............................................
............................................................H. microspirale X. Z. Liu et al.
90. Conidia crescent-shaped, distal point filled with wall material, 10 –16
(straight line) £2.5 –3.0 mm .................... H. microsporum Glockling
(if on rotifers, see Rotiferophthora torquatispora)
10. Conidia filamentous, bent to enclose three-fourths of an ellipse, total
length 18 – 50 £0.8 – 1.0 mm............................... H. arcuatum Barron
100. Conidia widening in the distal part, with pointed end filled
with wall material for 1.5 –2 mm, total length 20– 30 £
1.3 –1.9 mm ..................................................... H. cycloides Drechsler
11. Conidia filiform, undulate, spirally bent, total length 8 – 12 £
0.5 –0.8 mm.....................................................H. leptospira Drechsler
110. Conidia broader................................................................................ 12
12. Conidia twisted in the middle, appearing V-shaped, 5 – 11 £
0.7 –1.5 mm, additional clavate phialoconidia with rounded base,
12– 15 £3–4mm ............................................... H. drechsleri Barron
120. Change of direction more gradual................................................... 13
13. Spiral conidia 12.5–20.5 £1.2 –2.0 mm, with basal slime drop;
synanamorph of broadly fusiform hirsutella-like blastoconidia,
5.6–8.7 £1.8–3.1mm, abundantly produced; arthroconidia
11.0– 23.5 £3.2 –4.7 mm .......................... H. cerberi W. Gams et al.
130. Blastoconidial synanamorph absent ................................................ 14
14. Conidia with both ends pointed, 6.0 –8.0 £1.5 –2.0 mm....................
........................................................................ H. spirosporum Barron
140. Conidia up to 1.7 mm wide, not sharply pointed ............................ 15
15. Conidia with constant width over most of the length, 20 – 48 £
0:5–1:3ð–1:7Þmm;with mucous base ..........H. helicoides Drechsler
150. Conidia with pronouncedly wider distal third, 25 – 33 £1:3–1:7mm....
......................................................................... H. cocleatum Drechsler
16. Conidia straight for the major part .................................................. 17
160. Conidia of more irregular shapes .................................................... 22
17. Conidia regularly cylindrical, mostly straight................................. 18
170. Conidia of other shapes ................................................................... 21
18. On rotifers; conidia cylindrical, slightly bent, forming irregular
chains, 9 – 11 £1:7–2:5mm............................. H. botuliforme Barron
180. On nematodes; conidia mostly straight, adhering in heads ............ 19
19. Conidia narrowly cylindrical, straight, 6– 9 £0:3–0:5mm...............
...................................................H. angustisporum Monoson & Pikul
190. Conidia mostly exceeding 1 mm in width ....................................... 20
Gams and Zare44

20. Conidia cylindrical, straight, 2:5–5:0£0:7–1:5mm..........................
.....................................................................H. baculiforme Drechsler
200. Conidia long cylindrical with rounded ends, 22 –27 £1:7–2:5mm;
mostly straight........................................... H. cylindrosporum Barron
21. Conidia cylindrical to allantoid, 3 – 5 £0:9–1:2mm...........................
.......................................................................... H. sicyodes Drechsler
210. Conidia slender obclavate, tapering distally, with bent tip and
adhesive apical spur, 12 –26 £1:0–1:8mmH. subuliforme Drechsler
22. Conidia reniform or triangular.........................................................22
220. Conidia with a laterally displaced basal apiculus and various shapes
of the distal part ............................................................................... 24
23. On nematodes; conidia reniform, 3.2– 4 mm long ...............................
...................................................... H. reniforme S. D. Patil & Pendse
230. On rotifers, conidia with bilobed apex, thus appearing regularly
triangular, with rounded ends, 7:2–9:0£6:3–7:2mm (up to 11 mm
in culture) ................................ H. trigonosporum Barron & Szijarto
(If conidia with a basal appendage, see Haptospora appendiculata)
24. Conidia rather slender, apex rounded or with a simple point.........25
240. Conidia with bulbous base, apex with upwards directed beak or with a
double beak ...................................................................................... 26
25. Conidial apex rounded, conidia 4:5–6:5£0:8–2:1mm......................
...............................................................H. bysmatosporum Drechsler
250. Conidial apex pointed in axial direction, conidia 3:5–4:0£
1:0–1:5mm .............................. ...................... H. diceraeum Drechsler
26. Conidial apex with a sharp point in distal direction, conidia 4 – 5 £
4–5mm...................................................... H. rhynchosporum Barron
260. Conidial apex with two unequal beaks, conidia 3 – 9 £2–3mm.........
..................................................................... H. dicorymbum Drechsler
Electron-microscopic observations were concerned with H. anguillulae
(Saikawa, 1982c), H. subuliforme (Saikawa and Morikawa, 1985), and
H. oxycoracum (Saikawa et al., 1983).
8.5.2. Ecology
Harposporium anguillulae is the most widely distributed species. Most reports of
the genus are from temperate regions, but a few species have also been reported
from El Salvador (Bu
´caro, 1983) and Central America (Persmark et al., 1995).
The biology of H. anguillulae was studied in detail by Aschner and Kohn (1958),
who showed that species of this genus could be easily grown in culture.
Harposporium lilliputanum and H. cycloides were grown in culture by Glockling
and Shimazu (1997). The conidia of H. anguillulae and other species are ingested
orally and lodge in the esophagus of the prey, whence they colonize the body
Clavicipitaceous Anamorphs Parasitizing Nematodes 45

(see also TEM study by Saikawa et al., 1983), but H. subuliforme can also adhere
to a nematode externally due to an adhesive apical bud, which produces an
adhesive substance after contact with a nematode (Saikawa and Morikawa,
1985). Because the conidia can be ingested only by saprophagous
(bacteriophagous) nematodes, species of Harposporium have no effect on
phytophagous nematodes. The seasonality and distribution in agricultural soils
were studied by Persmark et al. (1996a) as described in the general part. Living
mycelium (but not conidia) of H. anguillulae attracts nematodes (Jansson and
Nordbring-Hertz, 1979), in particular species of Panagrellus,Ditylenchus, and
Aphelenchoides, but not of Pratylenchus (Jansson and Nordbring-Hertz, 1980).
In Brazil, infective larvae of Haemonchus contortus (Trichostrongylid nematode
parasites of sheep) were eliminated for 99.5% by adding 300,000 conidia of H.
anguillulae to 1 g of feces (Charles et al., 1996).
8.6. Hirsutella Patouillard, Rev. Mycol. Toulouse 14:69, 1892
(Fig. 7)
Type species:Hirsutella entomophila Pat.
Diagnosis: Colonies medium-fast-growing. Sporulation synnematous or
mononematous. Conidiogenous cells flask-shaped, tapering in the
middle or the distal part into one or several conidiiferous necks;
conidiogenesis mostly phialidic, with several conidia agglutinated in
slimy heads at each opening, sometimes solitary conidia produced with
apparently blastic conidiogenesis. Conidia mostly somewhat fusiform,
hyaline, smooth-walled, surrounded by adhesive slime, sometimes
globose and roughened.
Synanamorphs:Harposporium (Glockling, 1998b; Hodge et al., 1997).
Most species of the genus are entomogenous, and many are synnematous. No
comprehensive revision of the species has so far been published. A common
nematode parasite is the mononematous H. rhossiliensis Minter & B. L. Brady
(Minter and Brady, 1980), a name that predates the synonymous H. heteroderae
Sturhan & R. Schneider 1980 by a few months. Phialides solitary, 18– 33 £
3–5mm;tapering to 0.5–0.7 mm; conidia formed singly or in pairs on phialides,
ellipsoidal with a more or less apiculate base and a voluminous persistent slime
layer, 7 – 11 £4:8–7:5mm (measured including the slime).
8.6.1. Ecology of H. rhossiliensis
The species has been isolated from Criconemella xenoplax,Heterodera
avenae,Meloidogyne javanica, and, as H. heteroderae, from Heterodera
humuli (Sturhan and Schneider, 1980). It successfully infected several
other species of Heterodera,Ditylenchus destructor,Meloidogyne hapla,
Gams and Zare46

Pratylenchus penetrans,Anaplectus granulosus, and even larvae of Globodera
rostochiensis, but not members of the Tylenchidae (Sturhan and Schneider,
1980). A Petri dish population of Ditylenchus dipsaci is killed in vitro in
4 days, and one of M. incognita juveniles in 2 days (Cayrol et al., 1986).
Conidia adhere to the nematode body and penetrate it; in Acrobeloides a
preferential adhesion in the head and tail regions was observed (Venette et al.,
1997). Hirsutella rhossiliensis is considered responsible for rapid fluctuations
of C. xenoplax populations in peach orchards (Stirling, 1988, 1991; Zehr, 1985).
Without nematodes, the population of H. rhossiliensis in soil dies out (Jaffee,
1991). In legume rhizospheres the population of the bacteriophagous nematodes
of the genus Acrobeloides increased, followed by an increase in the population of
H. rhossiliensis, but these nematodes were much less vigorously attacked by the
fungus than were populations of Heterodera schachtii (Venette et al., 1997).
Densities of H. rhossiliensis in a Swedish agricultural soil peaked in September –
November (Persmark et al., 1996). The population in the soil follows that of its
host, Heterodera glycines in soybean fields, with midseason maxima, declining in
alternating maize crops (Chen and Reese, 1999).
The conidia of the fungus are infective only as long as they are attached to
the phialide (McInnis and Jaffee, 1989), and therefore the species seems less
suited for biological control than species of Haptocillium (Hay and Bateson,
1997). Moreover, conidial germination can be greatly affected by soil fungistasis
(Jaffee and Zehr, 1985; Stirling, 1988). The fungus exerted little population
control unless host densities were high (Kerry and Jaffee, 1997). On an agar pH
gradient, no growth was observed below pH 5 (Lo
´pez-Llorca et al., 1994).
Isolates obtained from different nematodes and origins showed considerable
genetic variation but had uniform characters of nematode pathogenicity; only
isolates originating from Hoplolaimidae grew more slowly, had larger conidia,
and were less pathogenic toward nematodes than isolates from other hosts
(Tedford et al., 1994). When growing on J2 larvae of Meloidogyne hapla,the
fungus produced 78 – 124 conidia from a single individual. Addition of 1.9
vegetative colonies/cm
3
soil caused a 50% decrease in J2 penetration of lettuce
roots, while lettuce weight, root galling, or egg production were not affected
(Viaene and Abawi, 2000). When applied in combination with Pochonia
chlamydosporia,H. rhossiliensis could not be detected from lettuce roots and
control was not improved by the combination; an inundative release of the fungus
would be necessary at every lettuce planting, as the fungus did not survive over
long periods in the soil (Viaene and Abawi, 2000). Patel et al. (2001) attempted
production of inoculum in liquid culture.
A second nematophagous species, Hirsutella minnesotensis S. Chen et al.,
was found as a pathogen of second-stage juveniles of the soybean cyst
nematode, Heterodera glycines (Chen et al., 2000). This species is morpholo-
gically very similar to the mite parasite H. thompsonii F. E. Fisher, with equally
Clavicipitaceous Anamorphs Parasitizing Nematodes 47

roughened globose solitary conidia, but it has a more strongly swollen base of
the conidiogenous cells and larger conidia, 4– 6 mm in diameter. Other,
entomogenous species of Hirsutella did not attach to nematodes with their
conidia and had no controlling effect (Cayrol et al., 1986).
8.7. Lecanicillium W. Gams & Zare, Nova Hedwigia 72:50,
2001
Type species:Lecanicillium lecanii (Zimmerm.) Zare & W. Gams.
Diagnosis: Colonies rather fast-growing, reaching 15 – 30 mm diam in 10 d
at 208C on PDA or MEA, white or yellowish. Conidiophores little
differentiated from the subtending hyphae, commonly arising from aerial
hyphae, initially erect with one or two whorls of phialides, then usually
prostrate and bearing large numbers of phialide whorls or single phialides.
Phialides aculeate, bearing often fasciculate groups of conidia, often
positioned at a right angle with the phialide tips, in some taxa forming
chains; some taxa also forming short, basally swollen, rapidly collapsing
“aphanophialides” which bear single conidia. Conidia short- to long-
ellipsoidal to falcate with pointed ends. Chlamydospores, dictyochlamy-
dospores, or swollen hyphal portions absent. Octahedral (sometimes also
prismatic) crystals commonly present in the agar medium. Most species
are entomogenous or fungicolous (Zare and Gams, 2001a).
Teleomorphs:Torrubiella,Cordyceps
Lecanicillium psalliotae (Treschow) Zare & W. Gams (once found in a cyst of
Globodera rostochiensis) and “Verticillium”leptobactrum W. Gams (mainly in
Heterodera eggs) were occasionally isolated from nematodes (Gams, 1988).
Godoy et al. (1982) and Gintis et al. (1983) mentioned the rare observation of
V. lecanii (Zimm.) Vie
´gas [probably now L. muscarium (Petch) Zare & W. Gams
(Fig. 16a)], V. lamellicola (F. E. V. Smith) W. Gams (Fig. 19a), and
V. leptobactrum (Fig. 19b) as parasites of cysts and eggs of Heterodera and
Meloidogyne species. For the latter two species see Simplicillium. Uziel and
Sikora (1992) deliberately applied isolates of “Verticillium lecanii” originating
from insects to control cyst nematodes, Globodera pallida, under artificial
conditions in water agar cultures. Among 14 isolates tested, 8 [most of them
probably L. muscarium and one L. longisporum (Petch) Zare & W. Gams
(Fig. 16b)] successfully parasitized eggs after 2 months. Meyer and Wergin
(1998) observed colonization of cysts and females of soybean cyst nematodes
when L. lecanii was added to monoxenic nematode cultures, with fungal
multiplication in the gelatinous matrix but relatively little penetration of the eggs.
Observed antagonistic effects by this fungus reducing viability of cyst nematode
eggs (Ha
¨nssler, 1990) might be attributed to chemical action.
Gams and Zare48

8.8. Paecilomyces Bain., Bull. Trimest. Soc. Mycol. Fr.
213:26, 1907
Type species:P. variotii Bain.
Diagnosis: Colonies medium-fast-growing, powdery due to long chains of
dry conidia. Conidiophores erect, with terminal and intercalary whorls of
phialides. Phialides consisting of a swollen venter that tapers strongly in
the upper part into a slender condiiferous neck. Conidia one-celled,
hyaline, dry, adhering in long chains.
FIGURE 16 (a) Lecanicillium muscarium; (b) L. longisporum, conidiophores
and conidia (From Zare and Gams, 2001a.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 49

The type species of the genus belongs to the Eurotiales. Several members of
Clavicipitaceae have been included in the genus (Samson, 1974). The best known
of these, P. farinosus (Holm : Fr.) A. H. S. Brown & G. Smith, teleom. Cordyceps
memorabilis Ces., is a common entomogenous species, for which the genus Isaria
Pers. : Fr. is being reintroduced (K. T. Hodge et al., in preparation). Paecilomyces
lilacinus (Thom) Samson (Fig. 6) also belongs to the Clavicipitaceae, although it
is unrelated to P. farinosus (Sung et al., 2001). This is a rather common pathogen
of nematode eggs in soil (Dackman and Nordbring-Hertz, 1985; Dackman et al.,
1985; Stirling, 1998, 1991). Numerous experiments have been carried out in view
of its application in biological control (Kerry and Evans, 1996; Stirling, 1991).
However, most experiments carried out so far with the species for biological
control lacked adequate controls to ascertain the specific fungal effects (Kerry,
1989, 1990). Its potential human pathogenicity (ocular and cutaneous infections,
onychomycosis, sinusitis, and deep infections in immunocompromized patients,
de Hoog et al., 2000) seems to preclude its practical application, but some genetic
difference was found between human-pathogenic isolates and the nematode
parasites (R. A. Samson, personal communication, 2002). Different isolates vary
greatly in their pathogenicity to nematodes (Stirling, 1991). The nematophagous
ability of the isolates tested was somewhat correlated with their UV resistance; a
similar grouping of isolates could also be achieved by means of random amplified
polymorphic DNA (RAPD) (Gunasekera et al., 2000). The species was found to
efficiently parasitize eggs of Meloidogyne incognita and Globodera pallida in
Peru (Jatala et al., 1979). Some other sensitive nematode species were listed in a
comprehensive review by Jatala (1986). The fungus not only attacks the egg
shell, it also has toxic effects (Jatala, 1986). Production of the antibiotics
leucinostatin and lilacin and chitinolytic enzymes have been documented
(Morgan-Jones and Rodrı
´guez-Ka
´bana, 1988). Production of a basic serine
protease is induced in P. lilacinus by cultivation on chitin or vitellin; this enzyme
destroys eggs of Meloidogyne hapla and is regarded as a crucial component
in pathogenesis (Bonants et al., 1995; Segers et al., 1999). Colonization of
Meloidogyne eggs with appressorium formation was illustrated in SEM
photographs by Segers et al. (1996). Increased populations of the fungus were
also associated with decline of Rotylenchulus reniformis in tomato in India,
giving a control comparable to that by carbofuran (Reddy and Khan, 1988). In
some Meloidogyne-suppressive soils in California, the natural populations of
Pochonia chlamydosporia and P. lilacinus seemed to play a minor role in
regulating the nematode population (Gaspard et al., 1990a). Population densities
of P.lilacinus were positively correlated with those of P. chlamydosporia in
tomato field soils in California, but not with those of Meloidogyne incognita
(Gaspard et al., 1990b). Application of 10 or 20 g of fungus-infested wheat
kernels per microplot (0.76 m diam) at planting time (even better with an
additional treatment 10 days before planting) gave good protection against
Gams and Zare50

Meloidogyne incognita and increased tomato yield significantly, particularly at
temperatures between 24 and 288C (Cabanillas and Barker, 1989; Cabanillas
et al., 1989). Under field conditions in Peru, application of the fungus in potato
soils gave a lower galling index due to Meloidogyne incognita than nematicide
treatments (Jatala et al., 1980), and a single application appeared sufficient to
establish the fungus in the soil (Jatala et al., 1981). Selective media for
monitoring the species have been devised by Mitchell et al. (1987: PDA with, per
liter, 10 g NaCl, 50 mg pentachloronitrobenzene, 50 mg benomyl, 1 mL Tergitol
NP10, and antibacterial antibiotics), Cabanillas and Barker (1989: PDA with
dichloran and oxgall together with antibacterial antibiotics), and Gaspard et al.
(1990a: chitin-rose bengal agar with 50 mg/L iprodione). Fassatiova
´and Lysek
(1982) obtained this species and the similar P. marquandii (Massee) S. Hughes
from eggs of Ascaris lumbricoides exposed in soils in the Czech Republic,
Pakistan, and Cuba. Paecilomyces lilacinus (less than P. marquandii) also
infected and killed eggs of the canine roundworm, Toxocara canis, penetrating
them by means of appressoria (Basualdo et al., 2000). Therefore P. lilacinus
might also be used as a biological control agent against animal helminths in vivo.
8.9. Plesiospora Drechsler, Sydowia 24 (1970):173 – 176,
1971 (Fig. 8)
Type and only species:Plesiospora globosa Drechsler.
Diagnosis: Phialides produced singly, tolypocladium-like inflated,
remaining inside the cuticle of the infected nematode, protruding only
with the opening. Conidia globose, hyaline, smooth-walled, 1.8 – 2.5 mm
in diameter.
The species is obviously very similar to Tolypocladium and Haptocillium, but
differs from them in vitro by forming single, swollen phialides supported by
slender stalks. Plesiospora globosa was found in nematodes in forest detritus
in Wisconsin. The fungus was also observed in Canada by Barron (1978), in
Apsheron, Azerbaidzhan, by Shilova (1987) and in Japan and Illinois by S. L.
Glocking (personal communication, 2002).
8.10. Pochonia Bat. & O. M. Fonseca, Publ. Inst. Micol.
Recife 462:4, 1965 5Diheterospora Kamyschko
ex Barron & Onions, Can. J. Bot. 44:866, 1966
Type species:Pochonia humicola Bat.&O.M.Fonseca¼
P. chlamydosporia (Goddard) Zare & W. Gams (Zare et al., 2001)
(Verticillium chlamydosporium Goddard 1913).
Clavicipitaceous Anamorphs Parasitizing Nematodes 51

Diagnosis: Colonies rather fast-growing (15 –40 mm diam in 10 days).
Conidiophores usually prostrate and little differentiated from vegetative
hyphae, sometimes erect. Conidiogenous cells phialides, verticillate or
solitary, aculeate. Conidia subglobose, ellipsoidal to rod-shaped,
isodiametric-polyhedral, or falcate with blunt ends, adhering in globose
heads. Dictyochlamydospores often produced. Crystals absent.
Teleomorph:Cordyceps
Barron and Onions (1966) distinguished Diheterospora from Verticillium
because of the presence of dictyochlamydospores. Gams (1971, 1988) did not
recognize this criterion as having generic value, but molecular studies by Zare
et al. (2000) and Sung et al. (2001) showed the phylogenetic distinctness of these
parasites of nematode cysts and eggs. In addition to the common presence of
dictyochlamydospores or at least swollen hyphal cells, the chromophilic behavior
of the conidia and the absence of crystals can be taken as morphological criteria
to distinguish the genus from Lecanicillium. The genus was revised by Zare et al.
(2001).
8.10.1. Key to the Species (modified from Zare et al., 2001)
1. At least part of the conidia crescent-shaped or falcate ...........................
............................................P. bulbillosa (W. Gams) Zare & W. Gams
10. Conidia not crescent-shaped or falcate ................................................ 2
2. Conidia isodiametric-polyhedric; dictyochlamydospores present,
usually on the agar surface P. gonioides (Drechsler) Zare & W. Gams
20. Conidia rod-shaped, smooth, with truncate ends, 2:0–2:5£
0:8–1:0mm;dictyochlamydospores sparse, submerged in the agar;
so far known only from rotifers in pine litter in Japan .........................
................................................. P. microbactrospora Zare & W. Gams
200. Conidia of other shapes, oval, subglobose to subcylindrical, smooth;
dictyochlamydospores above or in the agar ........................................ 3
3. Dictyochlamydospores, at least in fresh isolates, abundant, particularly
in the aerial mycelium; conidiophores typically prostrate .................. 4
30. Dictyochlamydospores, if present, mostly submerged in the agar;
conidiophores prostrate or erect ........................................................... 5
4. Conidia only in heads, never in chains, ð1:8–Þ2:5–4:5£1:0–Þ
1:2–2:2mm ............... P. chlamydosporia (Goddard) Zare & W. Gams
................................................................ var. chlamydosporia (Fig. 17)
40. Conidia mostly in chains, ð1:5–Þ2:0–3:5£1:5–3:0mm;some heads
may be present .... P. chlamydosporia var. catenulata (Kamyschko ex
Barron & Onions) Zare & W. Gams (Fig. 12)
Gams and Zare52

5. Colony reverse developing red shades on PDA; conidiophores
prostrate, verticillate; conidia globose to subglobose, 2:5–3:5£
2:0–3:0mm;dictyochlamydospores scanty or absent .........................
........................................................................... P. rubescens Zare et al.
50. Colony reverse yellow to cream (not red) on PDA; conidiophores partly
erect, richly verticillate; dictyochlamydospores partly submerged in the
agar........................................................................................................ 6
6. Conidia only in heads, never in chains, measuring 2:3–4:0£
1:5–2:5mm .......................... P. suchlasporia (W. Gams & Dackman)
Zare & W. Gams var. suchlasporia (Fig. 18)
60. Conidia mostly in chains, some heads may be present, measuring
2:0–3:7£1:7–2:3mm.......................... P. suchlasporia var. catenata
(W. Gams & Dackman) Zare & W. Gams
Most studies deal with P. chlamydosporia var. chlamydosporia (Fig. 17).
Its teleomorph, Cordyceps chlamydosporia H. C. Evans (in Zare et al., 2001),
has been found on slug eggs in tropical countries. Diheterospora catenulata
Kamyschko ex Barron & Onions, which differs from P. chlamydosporia var.
FIGURE 17 Pochonia chlamydosporia var. chlamydosporia, conidiophores
and conidia and dictyochlamydospores. (From Zare et al., 2001.)
Clavicipitaceous Anamorphs Parasitizing Nematodes 53

chlamydosporia only by catenate phialoconidia, was relegated to varietal rank by
Gams (1988), and this position is confirmed by Zare et al. (2001), in contrast
to different statements by Carder et al. (1993). A similar, so far unnamed,
teleomorph was found for this variety by Evans on a beetle larva in Ecuador.
A second common species in central and northern Europe is
P. suchlasporia (Fig. 18), which has more richly verticillate, erect
conidiophores and mostly submerged dictyochlamydospores. Even after
P. suchlasporia was segregated from the V. chlamydosporium complex,
P. chlamydosporia is found to be rather heterogeneous in molecular analyses
and in their ecological qualities (Arora et al., 1996; Kerry et al., 1986, 1993). This
species is not very closely related to the remaining taxa of the genus (Sung et al.,
2001).
FIGURE 18 Pochonia suchlasporia var. suchlasporia, conidiophores and
conidia and dictyochlamydospores. (From Zare et al., 2001.)
Gams and Zare54

Pochonia gonioides was originally observed by Drechsler (1942) on a
species of Bunonema, but Drechsler could not establish the mode of entry into
the nematode. Recently only two isolates of this species have been available,
and these were originally not directly associated with nematodes. Pochonia
bulbillosa is commonly isolated in conifer soils but its association with
nematodes is not ascertained, apart from its isolation as an ovicidal species
from Ascaris eggs in Pakistan and Afghanistan by Fassatiova
´and Lysek
(1982).
8.10.2. Ecology and Application
P. chlamydosporia is the species most frequently cited in studies on parasites of
nematode cysts. The species has many times been associated with nematode cysts
of various Heterodera species (summarized by Stirling, 1988, 1991), and it is the
major egg pathogen of Heterodera species in all European and American
countries examined (Bursnall and Tribe, 1974; Stirling, 1988, 1991). Lo
´pez-
Llorca and Duncan (1988) illustrated the colonization of Heterodera avenae by
species of Pochonia using SEM. The species is also found as an efficient parasite
of Meloidogyne root-knot nematodes (Godoy et al., 1981; Kerry, 2001; Morgan-
Jones et al., 1983; Morgan-Jones and Rodrı
´guez-Ka
´bana, 1988). In peanut fields,
Meloidogyne arenaria was more frequently parasitized than Heterodera glycines
(Morgan-Jones et al., 1981). The fungus usually does not attack potato cyst
eelworms of the genus Globodera (Kerry and Crumpp, 1977). Barron and Onions
(1966) and Tribe (1977) found the species also on slug eggs. Its ecology has been
repeatedly studied (Juhl, 1982; Dackman and Nordbring-Hertz, 1985; Dackman
et al., 1989, 1992; Gaspard et al., 1990a, b; Kerry et al., 1993). Quantification of
diseased eggs of Heterodera species was described by Kerry and Crump (1977).
Media for the selective isolation and quantification of P. chlamydosporia were
devised by Gaspard et al. (1990a, b): a chitin-rose bengal agar with 50 mg/L
benomyl); de Leij and Kerry (1991) and Kerry et al. (1993) recommend cornmeal
agar (Oxoid) with 37.5 mg carbendazim, 37.5 mg thiabendazole, 75 mg rose
bengal, 17.5 mg NaCl, 3 mL Triton X-100, and antibacterial antibiotics. Crump
and Kerry (1981) extracted and enumerated the dictyochlamydospores from soil.
Nicolay and Sikora (1989) quantified egg parasites by a new, standardizable
technique: the cysts present in a soil are extracted and crushed, and the contents
are reincorporated into the original soil sample. Parasitic activity on newly
formed eggs is then assessed. Bourne et al. (1994) devised several methods to
quantify the capacity of the fungus to grow in sterile and unsterile plant
rhizospheres. To detect P. chlamydosporia on infected plant roots with
polymerase chain reaction (PCR), Hirsch et al. (2000) developed specific primers
from a cloned fragment of the
b
-tubulin gene.
Clavicipitaceous Anamorphs Parasitizing Nematodes 55

The proteinase VCP1 of P. chlamydosporia was found to hydrolyze egg
shell proteins of Meloidogyne species but not those of Globodera (Segers et al.,
1996). The egg shells of Globodera are also rendered resistant by being twice as
thick as those of Meloidogyne (Lo
´pez-Llorca and Robertson, 1992). Pochonia
chlamydosporia also produces a chymoelastase-like protease which hydrolyses
host nematode proteins in situ (Segers et al., 1994). One of its major classes of
extracellular proteases is subtilisin-like proteins, of which one to four isoforms
were found in different isolates; these enzymes digest the protein component of
nematodes and are important determinants of pathogenicity (Segers et al., 1999).
The enzyme is similar to that produced by Metarhizium anisopliae (Segers et al.,
1995). The fungus is a factor in natural decline of nematode populations (Kerry
et al., 1982); partial sterilization of the soil with 38% formaldehyde destroyed its
population and the nematode-decline effect (Kerry and Jaffee, 1997). The fungus
colonizes living and, somewhat more efficiently, dead eggs of Heterodera, with a
preference for young stages, before the embryo development is completed (Irving
and Kerry, 1986). Pochonia chlamydosporia is also ovicidal to the large
roundworm, Ascaris lumbricoides (Lysek and Krajci, 1987).
First attempts to apply conidial suspensions against nematodes failed
(Willcox and Tribe, 1974), but numerous subsequent trials of the potential use of
P. chlamydosporia in biological control of Heterodera and Meloidogyne species
were more successful (de Leij and Kerry, 1991; Kerry and Evans, 1996; Morgan-
Jones et al., 1983; Stirling, 1991; Tribe, 1980; Kerry, 2001). Kerry (1995, 2001)
reviewed the biology of P. chlamydosporia in view of its potential application
against cyst and root-knot nematodes. The fungus proliferates in calcareous
loams and organic soil in England and survives for at least 3 months after
application, but isolates differed greatly in their capacity to survive and
proliferate in different soils (Kerry, 1989; Kerry et al., 1993; Viaene and Abawi,
2000) and also in virulence (Irving and Kerry, 1986). Along an agar pH gradient
in Petri dishes, optimal growth occurred around pH 6, and some growth even at
pH 3 (Lo
´pez-Llorca et al., 1994). This species can affect the multiplication of
cyst nematodes by multiple means, not only by egg colonization (Kerry, 1990).
“Control of H. schachtii by different isolates was related to the proportion of
young females infected but not to the numbers of cysts colonized; infection
resulted in few eggs being produced and many of those were parasitized” (Kerry,
1990). In peaty sand a better establishment was observed than in loamy sand or
sand in tomato plots with M. incognita; but in microplots with sandy loam a 90%
control of M. hapla could be achieved, provided the temperature did not exceed
258C (de Leij et al., 1993). The species was found in 13 of 20 Californian tomato
field soils examined, and its densities were positively correlated with those of
Meloidogyne incognita and Paecilomyces lilacinus (Gaspard et al., 1990b).
Addition of Meloidogyne species to the soil increased the population of
P. chlamydosporia. In tomato soils, changes in population densities of
Gams and Zare56

M. incognita and P. chlamydosporia followed each other. De Leij and Kerry
(1991) observed that application of dictyochlamydospores and hyphal fragments
without additional food base gave the best establishment, while Kerry et al.
(1993) found addition of wheat bran to alginate pellets essential for the estab-
lishment of the fungus from granular applications. Application of chlamydo-
spores (concentrations comparable to those observed in soils naturally
suppressive to cyst nematodes, about 10
3
–10
4
CFU/g soil) is regarded as more
efficient than alginate-bran pellets (Davies et al., 1991). Addition of 5000
dictyochlamydospores/cm
3
of soil caused up to 43% colonization of egg masses
of M. hapla, without causing any effects on lettuce weight, root galling, or egg
production (Viaene and Abawi, 2000); a control could be achieved only up to a
concentration of 8 eggs/cm
3
soil. At high galling rates no successful biological
control is possible (de Leij et al., 1992). Dictyochlamydospores can be produced
on a sand-barley bran mixture in amounts of 5 £10
6
g medium in 3 weeks at
208C (Kerry and Jaffee, 1997). A small inoculum of the fungus showed strongest
multiplication in soil, provided additional nutrients were available (de Leij et al.,
1992). In maize and tomato soils, Bourne and Kerry (1999) observed a stronger
effect than in kale soils. Added dictyochlamydospores gave an acceptable control
in maize, kale, and beans, but reduced nematode infestation slightly in tomato,
where significant numbers of eggs remained protected from infection inside the
roots. Pochonia chlamydosporia cannot colonize the root cortex, and egg masses
developing inside large galls are therefore protected from fungal infection (de
Leij and Kerry, 1991). The capacity of certain isolates of P. chlamydosporia to
multiply in the rhizosphere of suitable host plants without adverse effects on the
plant (Bourne and Kerry, 1999; Bourne et al., 1994; Davies et al., 1991; de Leij
and Kerry, 1991) is particularly relevant. The understanding of the tritrophic
system is important for a successful application (Bourne and Kerry, 1999). The
rhizosphere of lettuce is not easily colonized by P. chlamydosporia, possibly
because of competition by other microorganisms (Viaene and Abawi, 2000).
Inoculation of “poor host plants” (i.e., not attractive to Heterodera species or
causing only small galls) preceding a more susceptible crop may help the fungus
to build up population levels and reduce nematode levels, thus improving
effectiveness of nematode control before a nematode-susceptible crop is planted.
The fungus “alone is unlikely to give adequate control of pest problems, but
integrated with other measures, V. chlamydosporium may provide a useful
additional approach to nematode management” (Davies et al., 1991; Kerry, 1995,
2001).
The application of P. chlamydosporia can be successfully combined with
the nematicide Aldicarb. This compound prevents initial nematode damage,
while the fungus subsequently confers a long-term protection (de Leij et al.,
1993). Application of the fungus was also successfully combined with that of an
arbuscular mycorrhizal symbiont, Glomus desertorum. A combined treatment
Clavicipitaceous Anamorphs Parasitizing Nematodes 57

gave optimal control on tomato nursery seedlings: fewer galls, fewer egg
masses, and more parasitized eggs (68%, compared with 52% with
P. chlamydosporia alone) (Rao et al., 1997). Chopped leaves of Azadirachta
indica (neem) more than those of Calotropis also acted synergistically with
P. chlamydosporia in reducing both the nematode population and galling in
tomato in pot experiments, while increasing plant growth and egg parasitism
(Reddy et al., 1999).
Pochonia suchlasporia (Fig. 18) has a lower temperature minimum
and optimum for growth and therefore certain ecological advantages over
P. chlamydosporia. Temperature effects on the development of these species
were studied by Dackman and Ba
˚a
˚th (1989). Pochonia suchlasporia is the
dominating species in Heterodera cysts in Denmark, Sweden, and
the Netherlands (Juhl, 1982; Dackman and Nordbring-Hertz, 1985; Dackman
et al., 1989; Gams, 1988), while P. chlamydosporia is more restricted to young
cysts in these countries. The former species was particularly successful in
colonizing eggs and showed high chitinase and protease activities (Dackman
and Nordbring-Hertz, 1985; Dackman et al., 1989). The most virulent isolate of
“P. chlamydosporia” and “V. chlamydosporium” tested by Irving and Kerry
(1986) was also infectious at 58C and should probably be identified as
P. suchlasporia. The similar P. rubescens showed optimal growth at pH 6, but
pigment production occurred mainly in the acidic range (Lo
´pez-Llorca et al.,
1994). In vitro, this fungus (then identified as “V. suchlasporium”) attacked eggs
of Heterodera and Globodera species, forming appressoria and penetration
hyphae with an internal infection bulb as demonstrated in TEM photographs by
Lo
´pez-Llorca and Robertson (1992).
8.11. Pseudomeria Barron, Can. J. Bot. 58:443, 1980 (Fig. 4)
Type and only species:Pseudomeria mucosa Barron.
Diagnosis: Conidiophores ascending from the attacked animal,
unbranched, repeatedly septate; from each cell a narrow conidiogenous
neck arises that develops a succession of almost beauveria-like denticles
bearing single conidia; conidia globose, 3.5 –4.5 mm diam, with a con-
spicuous basal slime pad.
Parasitizing rotifers of the genus Adineta (Barron, 1980b).
8.12. Rotiferophthora Barron, Can. J. Bot. 69:495, 1991
(Fig. 2)
Type species:R. globispora (Barron) Barron.
Gams and Zare58

Diagnosis: Conidiophores verticillate or with single branches; phialides
only slightly swollen, mostly aculeate; intercalary phialides* frequently
present below the terminal phialide, with a slender conidiiferous neck.
Conidia subglobose, ellipsoidal, clavate or curved, with a characteristic
large oil drop near the apex. Dictyochlamydospores mostly conspicu-
ously present, often composed of fewer cells than in Pochonia; cells
often in a two-dimensional arrangement. Synopsis of described species:
Glockling (1998a, no key).
8.12.1. Key to the Species
(Measurements are given unaltered from the protologs.)
1. Conidia (sub-)globose...........................................................................2
10. Conidia not globose.............................................................................. 3
2. Conidia 3.3 –3.6 mm diam .................................. R. globispora
†
Barron
20. Conidia 2.0 – 2.2 mm diam .............................R. minutispora Glockling
200. Conidia 3 –4.5 £2.5 –3.0 mm..see Tolypocladium parasiticum Barron
3. Conidia with a straight longitudinal axis ............................................. 4
30. Conidia with a curved longitudinal axis ............................................ 18
4. Conidia ovoid to almost ellipsoidal ..................................................... 5
40. Conidia ellipsoidal, cylindrical or other shapes...................................9
5. Conidia 3.0 – 4.0 £2.0 –3.0 mm ............................................................6
50. Conidia larger ....................................................................................... 7
6. Phialides in pairs............................................ R. amamiensis Glockling
60. Phialides superimposed in drechmeria-like arrangement .....................
............................................................................R. gallinova Glockling
7. Conidia top-shaped, 3.5– 5.0 £3.0 –3.5 mm.........................................
.................... R. turbinispora
†
(Barron) Barron (nom. inval. Art. 37.1)
70. Conidia obovate and larger ..................................................................8
8. Conidia 4.5–5.5 £2.5–3.5 mm; dictyochlamydospores of 8 –16
cells........................................................... R. ovispora (Barron) Barron
80. Conidia 6.0 – 6.5 £5.5–6.0mm; dictyochlamydospores mostly 4-
celled.................................................... R. rotiferorum (Barron) Barron
*The term aphanophialide, introduced by Gams (1971) for ephemerous structures observed in
Aphanocladium and Lecanicillium (Zare and Gams, 2001a) that soon collapse to form inconspicuous
denticles, is not adequate to describe the structures of Rotiferophthora. Its conidiophores are more like
those of Sesquicillium W. Gams, a genus now merged with Clonostachys by Schroers (2001). No
separate term seems required to describe the structure; “intercalary phialides” suffices.
†
The spelling of the epithets was modified from the original, to bring the connecting vowel in
accordance with Art. 60.G ICBN.
Clavicipitaceous Anamorphs Parasitizing Nematodes 59

9. Conidia ellipsoidal............................................................................10
90. Conidia of other shapes....................................................................11
10. Conidia oval to broadly ellipsoidal, 2.7– 3.0 £2.4 –2.7 mm................
...........................................................................R. barronii Glockling
100. Conidia ellipsoidal, 3.5 – 4 £2.0 –2.5 mm............................................
..................................................... R. tagenophora (Drechsler) Barron
1000. Conidia ellipsoidal, 5.4 –6.9 £2.9 –3.1 mm......................................
...................................................................R. ellipsospora Glockling
11. Conidia cylindrical .................... .......................................................12
110. Conidia of other shapes....................................................................14
12. Conidia broadly cylindrical, 5.5– 7.5 £1.8 –2.3 mm...........................
.......................................................................... R. japonica Glockling
120. Conidia narrowly cylindrical ...........................................................13
13. Conidia narrowly cylindrical, in pronounced fascicles, 7.5 –
11 £2.4 –3.2 mm ........................... R. cylindrospora (Barron) Barron
130. Conidia cylindrical to slightly allantoid, 6.5– 8 £1.1 –
1.3 mm.............. ............................... R. angustispora (Barron) Barron
14. Conidia biconical with rounded ends, 6.5 –7.5 £2.2–2.5 mm..............
............................................................................... R. biconica Barron
140. Conidia with broader distal part ...................................................... 15
15. Conidia resembling a maize kernel ................................................. 16
150. Conidia more elongate ..................................................................... 17
16. Conidia 7–8 £3.2 –4.5 mm .................. R. zeispora
†
(Barron) Barron
160. Conidia 5.0 –6.5 £2.5 –3.5 mm ..........R. intermedia (Barron) Barron
17. Conidia tooth-like, clavate, slightly indented below apex, 8.4 –
10.8 £2.8 –3.6 mm ....................................... R. denticulispora Barron
170. Conidia obovoid-clavate, tearlike, 5.0 –5.5 £1.0 –1.5 mm.................
........................................................................... R. lacrima Glockling
18. Conidia comma-shaped or broadly obovate .................................... 19
180. Conidia of other shapes ...................................................................21
19. Conidia broadly obovate, apex curved, tapering toward a short,
truncate base, 3.5 –3.7 £2.9 mm ....................R. attenuata Glockling
190. Conidia comma-shaped.................................................................... 20
20. Conidia 6.0 –6.6 £2.5 –3.0 mm....... R. asymmetrica (Barron) Barron
200. Conidia 7–9 £4.5 –5.5 mm .................. R. humicola (Barron) Barron
21. Conidia bluntly kidney-shaped ........................................................ 22
210. Conidia of more slender curved shapes .......................................... 23
22. Conidia 3.0 – 5.0 £1.5– 2.0 mm .......... R. reniformis (Barron) Barron
220. Conidia 5–6 £5.0 –5.5 mm .................................. R. brevipes Barron
2200. Conidia 8–11 £5–7mm ........................... R. lunatispora
†
Glockling
23. Conidia with circular outline from one side and a slit on the other,
3.0– 3.2 £1.6 –2.2 mm ...................... R. microspora (Barron) Barron
Gams and Zare60

230. Conidia boomerang-shaped, with one arm narrower, 3.4 – 4.5 £2.0–
2.5 mm .............................................R. guttulispora
†
(Barron) Barron
2300. Conidia arcuate in about a third of a circle, 6:5–8:0ðtotal lengthÞ£
1:0–1:5mm..............................................R. torquatispora
†
Glockling
The first species of the genus was described by Drechsler (1942) as
Acrostalagmus tagenophorus from rotifers in a rich soil. Barron (1991a) is the
father of most known species. He did not generally grow these fungi in pure culture
but kept permanent slides from his material as types of the new species. The species
are very slow-growing and do not tend to form their phialidic propagules in culture,
while dictyochlamydospores are more easily obtained.
The parasitic phase is initiated by ingested conidia lodging on the wall of
the alimentary system between the mouth and the mastax. Rotiferophthora
species are among the most frequently recorded parasites of bdelloid rotifers.
They are devastating parasites, able to wipe out entire populations of rotifers in
Petri dishes in a few days (Barron, 1991a, 1991b).
8.13. Simplicillium W. Gams & Zare, Nova Hedwigia 73:39,
2001 (Figs. 11, 19)
Type species:Simplicillium lanosoniveum (van Beyma) Zare & W. Gams.
Diagnosis: Similar to Lecanicillium, but with mostly solitary phialides
arising from aerial hyphae, usually prostrate and little differentiated from
the subtending hyphae. Phialides discrete, aculeate and narrow, with a
very narrow tip in which collarette and periclinal wall thickening are not
visible. Conidia adhering in globose slimy heads or imbricate chains.
Teleomorph:Torrubiella
Species of this genus are not normally found on nematodes, but in soil, on fungi
or insects. Godoy et al. (1982) reported S. lamellicola (F. E.V. Smith) Zare &
W. Gams (Fig. 19a) and Verticillium leptobactrum W. Gams (Fig. 19b) (the latter
species may belong to Simplicillium as suggested by its solitary phialides, but
molecular study has not yet been done) from eggs of Heterodera glycines and
Meloidogyne arenaria; these two species were among the most active colonizers
of M. arenaria eggs (Morgan-Jones and Rodrı
´guz-Ka
´bana, 1988). V.
leptobactrum has also been reported from nematodes in Germany (Gams, 1988).
8.14. Tolypocladium W. Gams, Persoonia 6:185, 1971
(Fig. 10)
Type species:Tolypocladium inflatum W. Gams.
Diagnosis: Colonies rather slow-growing, pulvinate, cottony, white;
hyphae slender, mostly 1.0 –1.5 mm wide. Conidiophores scattered over
Clavicipitaceous Anamorphs Parasitizing Nematodes 61

the whole colony, short, lateral, sometimes bearing dense whorls of
phialides. Phialides consisting of a moderately swollen base and a
threadlike, often bent, neck. Conidia adhering in heads, globose to
cylindrical, hyaline, smooth-walled. Chlamydospores absent.
Teleomorph:Cordyceps
Most species of the genus are soil-borne or entomogenous.
In spite of different conidiogenesis, von Arx (1986) merged the genus with
Beauveria, but physiological (Todorova et al., 1998) and phylogenetic data
(Gams et al., 1998b) show that these genera are not closely related, although both
are associated with teleomorphs in Cordyceps (Hodge et al., 1996).
FIGURE 19 (a) Simplicillium lamellicola, conidiophores and conidia from
several isolates; (b) Verticillium leptobactrum, both original.
Gams and Zare62

Barron (1980a, 1983) described three species from bdelloid rotifers. All
have strongly inflated hyphal cells inside the prey animal. Conidia are ingested as
in Harposporium and Rotiferophthora; some conidia are digested, while others
germinate in the mastax (Barron, 1983). Bissett (1983) keyed out 11 species of
the genus, among which two were associated with rotifers. Bissett also included
Cephalosporium balanoides Drechsler in Tolypocladium; it is now classified in
Haptocillium. A few similar nematode-parasitic species with cylindrical, straight,
or slightly bent conidia are classified in Harposporium. They differ from other
species in Tolypocladium by their parasitic capacities, slower growth, and a
strong tendency to form swollen hyphae.
8.14.1. Key to the Rotifer-Parasitic Species
1. Conidia (sub)globose, 3.0– 4.5 £2.5 – 3 mm; complex stalked chlamy-
dospores present ................................................. T. parasiticum Barron
10. Conidia asymmetrical; chlamydospores unknown...............................2
2. Conidia shaped like a citrus slice, asymmetrically biconvex, 2.5 –
3.2 £1.5 –2.0 mm.....................................................T. lignicola Barron
20. Conidia symmetrically triangular with rounded ends, 2.5 –3.2 mm
diam............................................................... T. trigonosporum Barron
ACKNOWLEDGMENTS
We are grateful to Drs. R. C. Summerbell and G. F. Bills for helpful
comments on the text. We thank the authors and copyright holders of the
illustrations to grant permission for reproduction.
REFERENCES
Arora DK, Hirsch PR, Kerry B. PCR-based molecular discrimination of Verticillium
chlamydosporium isolates. Mycol Res 100:801– 809, 1996.
von Arx JA. Tolypocladium, a synonym of Beauveria. Mycotaxon 25:153 – 158, 1986.
Aschner M, Kohn S. The biology of Harposporium anguillulae. J Gen Microbiol
19:182–189, 1958.
Atkinson HJ, Du
¨rschner-Pelz U. Spore transmission and epidemiology of Verticillium
balanoides, an endozoic fungal parasite of nematodes in soil. J Invertebr Pathol
65:237–242, 1995.
Bailey F, Gray NF. The comparison of isolation techniques for nematophagous fungi from
soil. Ann Appl Biol 114:125–132, 1989.
Banck A, Persmark L, Jansson H-B, Andersson S. The influence of extraction methods on
the recovery of nematodes and nematophagous fungi (abstr). Nematologica 36:331,
1990.
Clavicipitaceous Anamorphs Parasitizing Nematodes 63

Barron GL. Isolation and maintenance of endoparasitic nematophagous Hyphomycetes.
Can J Bot 47:1899–1902, 1969.
Barron GL. The nematode-destroying fungi. Topics in Mycobiology No. 1. Guelph, Ont:
Can Biol Publ Ltd, pp 1–140, 1977.
Barron GL. Nematophagous fungi: endoparasites of Rhabditis terricola. Microb Ecol
4:157–163, 1978.
Barron GL. Fungal parasites of rotifers: a new Tolypocladium with underwater
conidiation. Can J Bot 58:439– 442, 1980a.
Barron GL. Fungal parasites of rotifers: a new genus of hyphomycetes endoparasitic in
Adineta. Can J Bot 58:443– 446, 1980b.
Barron GL. Predators and parasites of microscopic animals. In: Cole GT, Kendrick B, eds.
Biology of Conidial Fungi. Vol. 2. New York: Academic Press, 1981, pp 167– 200.
Barron GL. Nematode-destroying fungi. In: Burns RG, Slater JH, eds. Experimental
Microbiology. Oxford: Blackwell, 1982, pp 552 – 553.
Barron GL. Structure and biology of a new Tolypocladium attacking bdelloid rotifers. Can
J Bot 61:2566–2569, 1983.
Barron GL. Fungal parasites of bdelloid rotifers: Diheterospora. Can J Bot 63:211 – 222,
1985.
Barron GL. New species and new records of fungi that attack microscopic animals. Can J
Bot 67:267–271, 1989.
Barron GL. A new genus, Rotiferophthora, to accommodate the Diheterospora-like
endoparasites of rotifers. Can J Bot 69:494 – 502, 1991a.
Barron GL. A new genus of the Hyphomycetes endoparasitic in bdelloid rotifers with
conidia that lodge in the mastax. Can J Bot 69:503 – 506, 1991b.
Barron GL, Onions AHS. Verticillium chlamydosporium and its relationships to
Diheterospora,Stemphyliopsis, and Paecilomyces. Can J Bot 44:861 – 869, 1966.
Barron GL, Szijarto E. A new hyphomycete parasitic on the ciliated protozoans Vorticella
and Opercularia. Can J Bot 60:1031– 1034, 1982a.
Barron GL, Szijarto E. Structure and biology of a new hyphomycete parasitic on rotifers.
Can J Bot 60:1212–1215, 1982b.
Basualdo JA, Ciarmela ML, Sariento PL, Minvielle MC. Biological activity of
Paecilomyces genus against Toxocara canis eggs. Parasitol Res 86:854– 859, 2000.
Bissett J. Notes on Tolypocladium and related genera. Can J Bot 61:1311 – 1329, 1983.
Bonants PJM, Fitters PFL, Thijs H, den Belder E, Waalwijk C, Henfling JWDM. A basic
serine protease from Paecilomyces lilacinus with biological activity against
Meloidogyne hapla eggs. Microbiology 141:775– 784, 1995.
Boogert PHJF van den, Dijksterhuis J, Velvis H, Veenhuis M. Adhesive knob formation by
conidia of the nematophagous fungus Drechmeria coniospora. Antonie van
Leeuwenhoek 61:221– 229, 1992.
Boogert PHJF van den, Velvis H, Ettema CH, Bouwman LA. The role of organic matter in
the population dynamics of the endoparasitic nematophagous fungus Drechmeria
coniospora in microcosms. Nematologica 40:249– 257, 1994.
Bourne JM, Kerry BR. Effect of the host plant on the efficacy of Verticillium
chlamydosporium as a biological control agent of root-knot nematodes at different
nematode densities and fungal application rates. Soil Biol Biochem 31:75–84, 1999.
Gams and Zare64

Bourne JM, Kerry BR, De Leij FAAM. Methods for the study of Verticillium
chlamydosporium in the rhizosphere. J Nematol 26:587 – 591, 1994.
Bu
´caro RD. Hongos nemato
´fagos de El Salvador. Rev Biol Trop 31:25 – 28, 1983.
Bursnall LA, Tribe HT. Fungal parasitism in cysts of Heterodera. II. Egg parasites of
H. schachtii. Trans Br Mycol Soc 62:595 – 601, 1974.
Cabanillas E, Barker KR. Impact of Paecilomyces lilacinus inoculum level and application
time on control of Meloidogyne incognita on tomato. J Nematol 21:115 – 120, 1989.
Cabanillas E, Barker KR, Nelson LA. Growth of isolates of Paecilomyces lilacinus
and their efficacy in biocontrol of Meloidogyne incognita on tomato. J Nematol
21:164–172, 1989.
Campbell WP, Griffiths DA. The development and structure of thick-walled, multicellular,
aerial spores in Diheterospora chlamydosporia (¼Verticillium chlamydosporium).
Can J Microbiol 21:963–971, 1975.
Carder JH, Segers R, Butt TM, Barbara DJ, Mende NV, Coosemans J. Taxonomy of the
nematophagous fungi Verticillium chlamydosporium and V. suchlasporium based on
secreted enzyme activities and RFLP analysis. J Invertebr Pathol 62:178 – 184, 1993.
Cayrol J-C, Castet R, Samson RA. Comparative activity of different Hirsutella species
towards three plant parasitic nematodes. Rev Ne
´matol 9:412–414, 1986.
Charles TP, Roque MVC, Santos CD. Reduction of Haemonchus contortus infective
larvae by Harposporium anguillulae in sheep faecal cultures. Int J Parasitol
26:509–510, 1996.
Chen S, Liu XZ, Chen FJ. Hirsutella minnesotensis sp nov, a new pathogen of the soybean
cyst nematode. Mycologia 92:819– 824, 2000.
Chen SY, Reese CD. Parasitism of the nematode Heterodera glycines by the fungus
Hirsutella rhossiliensis as influenced by crop sequence. J Nematol 31:437 – 444, 1999.
Crump DH, Kerry BR. A quantitative method for extracting resting spores of two nematode
parasitic fungi, Nematophthora gynophila and Verticillium chlamydosporium, from soil.
Nematologica 27:330–339, 1981.
Dackman C, Ba
˚a
˚th E. Effect of temperature and time on growth of Verticillium
chlamydosporium and Verticillium suchlasporium and infection of Heterodera avenae
eggs. Mycologia 81:161–163, 1989.
Dackman C, Chet I, Nordbring-Hertz B. Fungal parasitism of the cyst nematode
Heterodera schachtii: infection and enzymatic activity. FEMS Microbiol Ecol
62:201–208, 1989.
Dackman C, Jansson H-B, Nordbring-Hertz B. Nematophagous fungi and their activities
in soil. In: Stotzky G, Bollag J-M, eds. Soil Biochemistry 7:95 – 130, 1992.
Dackman C, Nordbring-Hertz B. Fungal parasites of the cereal cyst nematode Heterodera
avenae in Southern Sweden. J Nematol 17:50– 55, 1985.
Dackman C, Olsson S, Jansson H-B, Lundgren B, Nordbring-Hertz B. Quantification of
predatory and endoparasitic nematophagous fungi in soil. Microbial Ecol 13:89 – 93,
1987.
Davies KG, de Leij FAAM, Kerry BR. Microbial agents for the biological control of plant
parasitic nematodes in tropical agriculture. Trop Pest Manag 37:303 – 320, 1991.
Clavicipitaceous Anamorphs Parasitizing Nematodes 65

De Leij FAAM, Kerry BR. The nematophagous fungus Verticillium chlamydosporium
as a potential biological control agent for Meloidogyne arenaria. Rev Ne
´matol
14:157–164, 1991.
De Leij FAAM, Kerry BR, Dennehy JA. The effect of fungal application rate and
nematode density on the effectiveness of Verticillium chlamydosporium as a biological
control agent for Meloidogyne incognita. Nematologica 38:112– 122, 1992.
De Leij FAAM, Kerry BR, Dennehy JA. Verticillium chlamydosporium as a biological
control agent for Meloidogyne incognita and M. hapla in pot and micro-plot tests.
Nematologica 39:115–126, 1993.
Dijksterhuis J, Harder W, Wyss U, Veenhuis M. Colonization and digestion of nematodes
by the endoparasitic nematophagous fungus Drechmeria coniospora. Mycol Res
95:873–878, 1991.
Dijksterhuis J, Sjollema KA, Veenhuis M, Harder W. Competitive interactions between
two nematophagous fungi during infection and digestion of the nematode Panagrellus
redivivus. Mycol Res 98:1458– 1462, 1994.
Dijksterhuis J, Veenhuis M, Harder W. Ultrastructural study of adhesion and initial stages
of infection of nematodes by conidia of Drechmeria coniospora. Mycol Res 94:1– 8,
1990.
Dijksterhuis J, Veenhuis M, Harder W. Conidia of the nematophagous fungus Drechmeria
coniospora adhere to but barely infect Acrobeloides buetschlii. FEMS Microbiol Lett
113:183–188, 1993.
Dowsett JA, Reid J, Hopkin A. On Cephalosporium balanoides Drechsler. Mycologia
74:687–690, 1982.
Drechsler C. Morphological diversity among fungi capturing and destroying nematodes.
J Wash Acad Sci 23:138– 141, 1933.
Drechsler C. Some hyphomycetes parasitic on free-living terricolous nematodes.
Phytopathology 31:773–802, 1941.
Drechsler C. Two zoophagous species of Acrostalagmus with multicellular Desmidio-
spora-like chlamydospores. J Wash Acad Sci 32:343 – 350, 1942.
Drechsler C. A nematode-destroying species of Cephalosporiopsis. Sydowia
22(1968):194–198, 1969.
Drechsler C. A nematode-destroying parasite bearing lageniform conidiiferous branches
on endozoic hyphae. Sydowia 24(1970):173 – 176, 1971.
Duddington CL. Further records of British predacious fungi. II. Trans Br Mycol Soc
34:194–209, 1951.
Duddington CL. Notes on the technique of handling predacious fungi. Trans Br Mycol Soc
38:97–103, 1955.
Du
¨rschner U. Pilzliche Endoparasiten an beweglichen Nematodenstadien. Mitt Biol
Bundesanst Ld- u Forstw 217:1– 83, 1983.
Du
¨rschner-Pelz U, Atkinson HJ. Recognition of Ditylenchus and other nematodes by
spores of the endoparasitic fungus Verticillium balanoides. J Invertebr Pathol 51:97 –
106, 1988.
Fassatiova
´O, Lysek H. Ovicidal fungi in soil ecological system. Acta Univ Carol Biol
9:297–334, 1982.
Gams and Zare66

Gams W. Cephalosporium-artige Schimmelpilze (Hyphomycetes). Stuttgart: G. Fischer,
1971.
Gams W. A contribution to the knowledge of nematophagous species of Verticillium. Neth
J Plant Pathol 94:123– 148, 1988.
Gams W, Hoekstra ES, Aptroot A, eds. CBS Course of Mycology. Baarn: Centraalbureau
voor Schimmelcultures, 1998a, pp 1 – 165.
Gams W, Jansson H-B. The nematode parasite Meria coniospora Drechsler in pure culture
and its classification. Mycotaxon 22:33– 38, 1985.
Gams W, O’Donnell K, Schroers H-J, Christensen M. Generic classification of some more
hyphomycetes with solitary conidia borne on phialides. Can J Bot 76:1570 – 1583,
1998b.
Gams W, Zare R. A revision of Verticillium sect. Prostrata. III. Generic classification.
Nova Hedwigia 72:329–337, 2001.
Gaspard JT, Jaffee BA, Ferris H. Meloidogyne incognita survival in soil infested with
Paecilomyces lilacinus and Verticillium chlamydosporium. J Nematol 22:176– 181,
1990a.
Gaspard JT, Jaffee BA, Ferris H. Association of Verticillium chlamydosporium and
Paecilomyces lilacinus with root-knot nematode infested soil. J Nematol 22:207 – 213,
1990b.
Gernandt DS, Stone JK. Phylogenetic analysis of nuclear ribosomal DNA places the
nematode parasite, Drechmeria coniospora, in Clavicipitaceae. Mycologia 91:993 –
1000, 1999.
Gintis BO, Morgan-Jones G, Rodrı
´guez-Ka
´bana R. Fungi associated with several
developmental stages of Heterodera glycines from an Alabama soybean field soil.
Nematropica 13:181–200, 1983.
Giuma AY, Cooke RC. Some endozoic parasites on soil nematodes. Trans Br Mycol Soc
59:213–218, 1972.
Glockling SL. Two new species of rotifer-attacking fungi, Rotiferophthora, from Japan
and records of Verticillium bactrosporum in rotifer hosts. Mycol Res 102:145 – 150,
1998a.
Glockling SL. Three new species of Rotiferophthora attacking bdelloid rotifers in Japan.
Mycol Res 102:1142–1148, 1998b.
Glockling SL, Shimazu M. Culturing of three species of endoparasitic fungi infecting
nematodes. Mycol Res 101:55–60, 1997.
Glockling SL, Yamada F. A survey of fungi which kill microscopic animals in the dung of
the Amami rabbit. Mycologist 11:113– 120, 1997.
Godoy G, Rodrı
´guez-Ka
´bana R, Morgan-Jones G. Parasitism of eggs of Heterodera
glycines and Meloidogyne arenaria by fungi isolated from cysts of H. glycines.
Nematropica 12:111–119, 1982.
Gray NF. Psychro-tolerant nematophagous fungi from the maritime Antarctic. Pl Soil
64:431–435, 1982.
Gray NF. Ecology of nematophagous fungi: distribution and habitat. Ann Appl Biol
102:501–509, 1983a.
Gray NF. Further observations on predaceous fungi from Ireland. Ir Nat J 21:18 –22,
1983b.
Clavicipitaceous Anamorphs Parasitizing Nematodes 67

Gray NF. Ecology of nematophagous fungi: notes on the identification of Acrostalagmus
obovatus,Cephalosporium balanoides and Verticillium sphaerosporum. Ir Nat J
21:125–128, 1983c.
Gray NF. Ecology of nematophagous fungi: methods of collection, isolation and
maintenance of predatory and endoparasitic fungi. Mycopathologia 86:143 –153, 1984.
Gray NF. Fungi attacking vermiform nematodes. In: Poinar GO Jr, Jansson H-B, eds.
Diseases of Nematodes. Vol. II. Boca Raton, FL: CRC Press, 1988, pp 3 – 38.
Gray NF, Duff KR. Some preliminary observations on predaceous fungi from Ireland. Ir
Nat J 20:378– 380, 1982.
Gray NF, Lewis Smith RI. The distribution of nematophagous fungi in the maritime
Antarctic. Mycopathologia 85:81– 92, 1984.
Gray NF, Wyborn CHE, Lewis Smith RI. Nematophagous fungi from the maritime
Antarctic. Oikos 38:194– 201, 1982.
Gunasekera TS, Holland RJ, Gillings MR, Briscoe DA, Neethling DC, Williams KL,
Nevalainen KMH. Phenotypic and genetic characterization of Paecilomyces lilacinus
strains with biocontrol activity against root-knot nematodes. Can J Microbiol 46:775–
783, 2000.
Hall RA. Laboratory infection of insects by Verticillium lecanii strains isolated from
phytopathogenic fungi. Trans Br Mycol Soc 74:445 – 446, 1980.
Ha
¨nssler G. Verticillium lecanii als Parasit an Cysten von Heterodera schachtii.
Z PflKrankh PflSchutz 97:194–201, 1990.
Hay FS. Endoparasites infecting nematodes in New Zealand. N Z J Bot 33:401– 407, 1995.
Hay FS, Bateson L. Effect of the nematophagous fungi Hirsutella rhossiliensis and
Verticillium balanoides on stem nematode (Ditylenchus dipsaci) in white clover.
Australas Plant Pathol 26:142– 147, 1997.
Hay FS, Regnault L. Introduction of the endoparasitic nematophagous fungus Verticillium
balanoides into white clover (Trifolium repens) by its host, stem nematode
(Ditylenchus dipsaci). Australas Plant Pathol 24:249 – 251, 1995.
Hirsch PR, Mauchline TH, Mendum TA, Kerry BR. Detection of the nematophagous
fungus Verticillium chlamydosporium in nematode-infested plant roots using PCR.
Mycol Res 104:435–439, 2000.
Hodge KT, Krasnoff SB, Humber RA. Tolypocladium inflatum is the anamorph of
Cordyceps subsessilis. Mycologia 88:715– 719, 1996.
Hodge KT, Viaene NM, Gams W. Two Harposporium species with Hirsutella
synanamorphs. Mycol Res 101:1377– 1382, 1997.
Hoog GS de, Guarro J, Gene
´J, Figueras MJ. Atlas of Clinical Fungi. 2nd ed. Utrecht,
Reus: Centraalbureau voor Schimmelcultures, 2000, pp 1 – 1126.
Irving F, Kerry BR. Variation between strains of the nematophagous fungus, Verticillium
chlamydosporium. Nematologica 32:474– 485, 1986.
Jaffee BA. Population biology and biological control of nematodes. Can J Microbiol
38:359–364, 1992.
Jaffee BA, Zehr EI. Parasitic and saprophytic abilities of the nematode-attacking fungus
Hirsutella rhossiliensis. J Nematol 17:341– 345, 1985.
Jansson H-B. Attraction of nematodes to endoparasitic nematophagous fungi. Trans Br
Mycol Soc 79:25–29, 1982a.
Gams and Zare68

Jansson H-B. Predacity by nematophagous fungi and its relation to the attraction of
nematodes. Microbiol Ecol 8:233–240, 1982b.
Jansson H-B. Adhesion of conidia of Drechmeria coniospora to Caenorhabditis elegans
wild type and mutants. J Nematol 26:430 – 435, 1994.
Jansson H-B, Dackman C, Zuckerman BM. Adhesion and infection of plant parasitic
nematodes by the fungus Drechmeria coniospora. Nematologica 33:480 – 487, 1987.
Jansson H-B, Friman E. Infection-related surface proteins on conidia of the
nematophagous fungus Drechmeria coniospora. Mycol Res 103:249 – 256, 1999.
Jansson H-B, Hofsten A von, Mecklenburg C von. Life cycle of the endoparasitic
nematophagous fungus Meria coniospora: a light and electron microscopy study.
Antonie van Leeuwenhoek 50:321– 327, 1984.
Jansson H-B, Jeyaprakash A, Zuckerman BM. Differential adhesion and infection of
nematodes by the endoparasitic fungus Meria coniospora (Deuteromycetes). Appl
Environ Microbiol 49:552–555, 1985a.
Jansson H-B, Jeyaprakash A, Zuckerman BM. Control of root knot nematodes on tomato
by the endoparasitic fungus Meria coniospora. J Nematol 17:327 – 329, 1985b.
Jansson H-B, Nordbring-Hertz B. Attraction of nematodes to living mycelium of
nematophagous fungi. J Gen Microbiol 112:89– 93, 1979.
Jansson H-B, Nordbring-Hertz B. Interactions between nematophagous fungi and
plant-parasitic nematodes: attraction, induction of trap formation and capture.
Nematologica 26:383–389, 1980.
Jansson H-B, Nordbring-Hertz B. The endoparasitic nematophagous fungus Meria
coniospora infects nematodes specifically at the chemosensory organs. J Gen
Microbiol 129:1121–1126, 1983.
Jansson H-B, Nordbring-Hertz B. Involvement of sialic acid in nematode chemotaxis and
infection by an endoparasitic nematophagous fungus. J Gen Microbiol 130:39 – 43,
1984.
Jansson H-B, Nordbring-Hertz B. Infection events in the fungus-nematode system.
In: Poinar GO Jr, Jansson H-B, eds. Diseases of Nematodes. Vol. II. Boca Raton, FL:
CRC Press, 1988, pp 59– 72.
Jansson HB, Persson C, Odeslius R. Growth and capture activities of nematophagous
fungi in soil visualized by low temperature scanning electron microscopy. Mycologia
92:10–15, 2000.
Jansson H-B, Tunlid A, Nordbring-Hertz B. Biological control: nematodes. In: Anke T, ed.
Fungal Biotechnology. Weinheim: Chapman & Hall, 1997, pp 38 – 50.
Jatala P. Biological control of plant-parasitic nematodes. Annu Rev Phytopathol
24:453–489, 1986.
Jatala P, Kaltenbach R, Bocangel M. Biological control of Meloidogyne incognita acrita
and Globodera pallida on potatoes (abstr). J Nematol 11:303, 1979.
Jatala P, Kaltenbach R, Bocangel M, Devaux AJ, Campos R. Field application of
Paecilomyces lilacinus for controlling Meloidogyne incognita on potatoes (abstr).
J Nematol 12:226–227, 1980.
Jatala P, Salas R, Kaltenback R, Bocangel M. Multiple application and long-term effect of
Paecilomyces lilacinus in controlling Meloidogyne incognita under field conditions
(abstr). J Nematol 13:445, 1981.
Clavicipitaceous Anamorphs Parasitizing Nematodes 69

Juhl M. Cereal cyst nematode, soil heating and fungal parasitism. EPPO Bull 12:505–511,
1982.
Juniper AJ. Dung as a source of predaceous fungi. Trans Br Mycol Soc 40:346 –348, 1967.
Karling JS. Harposporium anguillulae. Mycologia 30:512– 519, 1938.
Kerry BR. Biological control. In: Brown RH, Kerry BR, eds. Principles and Practice of
Nematode Control in Crops. London: Academic Press, 1987, pp 233 – 363.
Kerry BR. Fungi as biological control agents for plant parasitic nematodes. In: Whipps JM,
Lumsden RD, eds. Biotechnology of Fungi for Improving Plant Growth. Cambridge,
UK: Cambridge University Press, 1989, pp 153 – 170.
Kerry BR. An assessment of progress toward microbial control of plant-parasitic
nematodes. J Nematol 22(4S):621– 631, 1990.
Kerry BR. Ecological considerations for the use of the nematophagous fungus,
Verticillium chlamydosporium, to control plant parasitic nematodes. Can J Bot
73(Supp 1):S65–S70, 1995.
Kerry BR. Rhizosphere interactions and the exploitation of microbial agents for the
biological control of plant-parasitic nematodes. Annu Rev Phytopathol 38:323–441,
2000.
Kerry BR. Exploitation of the nematophagous fungus Verticillium chlamydosporium
Goddard for the biological control of root-knot nematodes (Meloidogyne spp.). In:
Butt TM, Jackson CW, Magan N, eds. Fungi as Biocontrol Agents. Progress, Problems
and Potential. Wallingford, CT: CABI Publishing, 2001, pp 155 – 167.
Kerry BR, Crump DH. Observations on fungal parasites of females and eggs of the
cereal cyst-nematode, Heterodera avenae, and other cyst nematodes. Nematologica
23:193–201, 1977.
Kerry BR, Crump DH, Mullen LA. Natural control of the cereal cyst nematode,
Heterodera avenae Woll., by soil fungi at three sites. Crop Prot 1:99–109, 1982.
Kerry BR, Evans K. New strategies for the management of plant parasitic nematodes. In:
Hall R, ed. Managing Soilborne Plant Pathogens. St. Paul, MN: APS Press, 1996,
pp 134–152.
Kerry BR, Irving F, Hornsey JC. Variation between strains of the nematophagous fungus,
Verticillium chlamydosporium Goddard. I. Factors affecting growth in vitro.
Nematologica 32:461–473, 1986.
Kerry BR, Jaffee BA. Fungi as biological control agents for plant-parasitic nematodes. In:
Wicklow DT, So
¨derstro
¨m B, eds. The Mycota IV. Environmental and Microbial
Relationships. Berlin: Springer-Verlag, 1997, pp 201 – 218.
Kerry BR, Kirkwood IA, De Leij FAAM, Barba J, Leijdens MB, Brookes PC. Growth and
survival of Verticillium chlamydosporium Goddard, a parasite of nematodes, in soil.
Biocontrol Sci Technol 3:355– 365, 1993.
Lo
´pez-Llorca LV, Duncan GH. A scanning electron microscopy study of fungal
endoparasitism of cereal cyst nematode (Heterodera avenae). Can J Microbiol
34:613–619, 1988.
Lo
´pez-Llorca LV, Moya M, Llinares A. Effect of pH on growth and pigment production of
nematophagous and entomogenous fungi. Micol Veg Mediterr 8:107 – 112, 1994.
Lo
´pez-Llorca LV, Robertson WM. Utrastructure of infection of cyst nematode eggs by the
nematophagous fungus Verticillium suchlasporium. Nematologica 39:65– 74, 1992.
Gams and Zare70

Lysek H, Krajci D. Penetration of ovicidal fungus Verticillium chlamydosporium through
the Ascaris lumbricoides egg-shells. Folia Parasitol 34:57 – 60, 1987.
McInnis TM, Jaffee BA. An assay for Hirsutella rhossiliensis spores and the importance of
phialides for nematode inoculation. J Nematol 21:229 – 234, 1989.
Meyer SLF, Wergin WP. Colonization of soybean cyst nematode females, cysts, and
gelatinous matrices by the fungus Verticillium lecanii. J Nematol 30:436 – 450, 1998.
Minter DW, Brady BL. Mononematous species of Hirsutella. Trans Br Mycol Soc
74:271–282, 1980.
Mitchell DJ, Kannwischer-Mitchell ME, Dickson DW. A semi-selective medium for the
isolation of Paecilomyces lilacinus from soil. J Nematol 19:255 – 256, 1987.
Morgan-Jones G, Godoy G, Rodrı
´guez-Ka
´bana R. Verticillium chlamydosporium fungal
parasite of Meloidogyne arenaria females. Nematropica 11:115 – 119, 1981.
Morgan-Jones G, Rodrı
´guez-Ka
´bana R. Fungi colonizing cysts and eggs. In: Poinar GO Jr,
Jansson H-B, eds. Diseases of Nematodes. Vol II. Boca Raton, FL: CRC Press, 1988,
pp 39–58.
Morgan-Jones G, White JF Jr, Rodrı
´guez-Ka
´bana R. Phytonematode pathology:
ultrastructural studies: 1. Parasitism of Meloidogyne arenaria eggs by Verticillium
chlamydosporium. Nematropica 13:245– 260, 1983.
Nicolay R, Sikora RA. Techniques to determine the activity of fungal egg parasites of
Heterodera schachtii in field soil. Rev Ne
´matol 12:97–102, 1989.
Nikoh N, Fukatsu T. Interkingdom host jumping underground: phylogenetic analysis of
entomoparasitic fungi of the genus Cordyceps. Mol Biol Evol 17:629 – 638, 2000.
Palm ME, Gams W, Nirenberg HI. Plectosporium, a new genus for Fusarium tabacinum,
the anamorph of Plectosphaerella cucumerina. Mycologia 87:397 – 406, 1995.
Patel A, Rose T, Gutberlet V, Mu
¨ller J, Vorlop KD. Der nematophage Pilz Hirsutella
rhossiliensis: Fermentation im 5-Liter Ru
¨hrreaktor und Formulierung in Kapseln und
Zuckerru
¨benpillen (abstr). Phytomedizin 31(4):59, 2001.
Persmark L, Banck A, Jansson H-B. Population dynamics of nematophagous fungi and
nematodes in an arable soil—vertical and seasonal fluctuations. Soil Biol Biochem
28:1005–1014, 1996.
Persmark L, Jansson H-B. Nematophagous fungi in the rhizosphere of agricultural crops.
FEMS Microbiol Ecol 22:303–312, 1997.
Persmark L, Marban-Mendoza N, Jansson H-B. Nematophagous fungi from agricultural
soils of central America. Nematotropica 25:117 – 124, 1995.
Poinar GO, Jansson H-B. Susceptibility of Neoaplectana spp. and Heterorhabditis
heliothidis to the endoparasitic fungus Drechmeria coniospora. J Nematol 18: 225 –
230, 1986, 1994.
Rao MS, Kerry BR, Gowen SR, Bourne JM, Parvatha Reddy P. Management of
Meloidogyne incognita in tomato nurseries by integration of Glomus deserticola with
Verticillium chlamydosporium. Z PflKrankh PflSchutz 104:419 – 422, 1997.
Reddy PP, Khan RM. Evaluation of Paecilomyces lilacinus for the biological control of
Rotylenchus reniformis infecting tomato, compared with carbofuran. Nematol Mediterr
16:113–115, 1988.
Clavicipitaceous Anamorphs Parasitizing Nematodes 71

Reddy PP, Rao MS, Nagesh M. Eco-friendly management of Meloidogyne incognita on
tomato by integration of Verticillium chlamydosporium with neem and Calotropis
leaves. Z PflKrankh PflSchutz 106:530– 533, 1999.
Saikawa M. An electron microscope study of Meria coniospora,anendozoic
nematophagous hyphomycete. Can J Bot 60:2019 – 2023, 1982a.
Saikawa M. Fixation of germinating conidia of Meria coniospora Drechsler with KMnO
4
.
J Electron Microsc 31:276– 277, 1982b.
Saikawa M. Ultrastructure of conidia of Harposporium anguillulae Lohde. J Electron
Microsc 31:90–92, 1982c.
Saikawa M, Morikawa Ch. An electron microscope study of initiation of infection by
conidia of Harposporium subuliforme, an endozoic nematophagous fungus. Trans
Mycol Soc Jpn 26:215– 219, 1985.
Saikawa M, Totsuka J, Morikawa Ch. An electron microscope study of initiation of
infection by conidia of Harposporium oxycoracum, an endozoic nematophagous
Hyphomycete. Can J Bot 61:893 – 898, 1983.
Samson RA. Paecilomyces and some allied Hyphomycetes. Stud Mycol 6:1 – 119, 1974.
Schroers H-J. A monograph of Bionectria (Ascomycota, Hypocreales, Bionectriaceae) and
its Clonostachys anamorphs. Stud Mycol 46:1– 224, 2001.
Segers R, Butt TM, Carder JH, Keen JN, Kerry BR, Peberdy JF. The subtilisins of fungal
pathogens of insects, nematodes and plants: distribution and variation. Mycol Res
103:395–402, 1999.
Segers R, Butt TM, Keen JN, Kerry BR, Peberdy JF. The subtilisins of the invertebrate
mycopathogens Verticillium chlamydosporium and Metarhizium anisopliae are
serologically and functionally related. FEMS Microbiol Lett 126:227 – 231, 1995.
Segers R, Butt TM, Kerry BR, Beckett A, Peberdy JF. The role of the proteinase VCP1
produced by the nematophagous Verticillium chlamydosporium in the infection process
of nematode eggs. Mycol Res 100:421 – 428, 1996.
Segers R, Butt TM, Kerry BR, Peberdy JF. The nematophagous fungus Verticillium
chlamydosporium produces a chymoelastase-like protease which hydrolyses host
nematode proteins in situ. Microbiology 140:2715– 2723, 1994.
Shilova TV. New parasitic for nematodes fungi from genera Acrostalagmus Drechs. and
Plesiospora Drechs. in the USSR Flora. Mikol Fitopatol 21:447 – 448, 1987.
Sjollema K, Dijksterhuis J, Veenhuis M, Harder W. An electron microscopical study of the
infection of the nematode Panagrellus redivivus by the endoparasitic fungus
Verticillium balanoides. Mycol Res 97:479– 484, 1993.
Stirling GR. Biological control of plant-parasitic nematodes. In: Poinar GO Jr, Jansson
H-B, eds. Diseases of Nematodes. Vol II. Boca Raton, FL: CRC Press, 1988,
pp 93–139.
Stirling GR. Biological Control of Plant Parasitic Nematodes. Wallingford, CT: CAB
International, 1991, pp 1– 282.
Sturhan D, Schneider R. Hirsutella heteroderae, ein neuer nematodenparasita
¨rer Pilz.
Phytopathol Z 99:105– 115, 1980.
Sung GH, Spatafora JW, Zare R, Hodge KT, Gams W. A revision of Verticillium section
Prostrata. II. Phylogenetic analyses of SSU and LSU nuclear rDNA sequences from
anamorphs and teleomorphs of the Clavicipitaceae. Nova Hedwigia 72:311 –328, 2001.
Gams and Zare72

Tedford EC, Jaffee BA, Muldoon AE. Variability among isolates of the nematophagous
fungus Hirsutella rhossiliensis. Mycol Res 98:1127– 1136, 1994.
Todorova SI, Co
ˆte
´J-C, Coderre D. Distinction between Beauveria and Tolypocladium by
carbohydrate utilization. Mycol Res 102:81– 87, 1998.
Tribe HT. Pathology of cyst-nematodes. Biol Rev Camb Phil Soc 52:477 – 507, 1977.
Tribe HT. Extent of disease in populations of Heterodera, with especial reference to
H. schachtii. Ann Appl Biol 92:61 – 72, 1979.
Tribe HT. Prospects for the biological control of plant-parasitic nematodes. Parasitology
81:619–639, 1980.
Tunlid A, Jansson H-B, Nordbring-Hertz B. Fungal attachment to nematodes. Mycol Res
96:401–412, 1992.
Uziel A, Sikora RA. Use of non-target isolates of the entomopathogen Verticillium lecanii
(Zimm.) Vie
´gas to control the potato cyst nematode, Globodera pallida (Stone).
Nematologica 38:123–130, 1992.
Viaene NM, Abawi GS. Hirsutella rhossiliensis and Verticillium chlamydosporium as
biocontrol agents of the root-knot nematode Meloidogyne hapla on lettuce. J Nematol
32:85–100, 2000.
Venette RC, Mostafa FAM, Ferris H. Trophic interactions between bacterial-feeding
nematodes in plant rhizosphere and the nematophagous fungus Hirsutella rhossiliensis
to suppress Heterodera schachtii. Plant Soil 191:213 – 223, 1997.
Watanabe T. Verticillium balanoides a nematode endoparasite associated with pine
needles of collapsing Japanese red pine trees in Tsiliba. Mycoscience 41:283 – 285,
2000.
Willcox J, Tribe HT. Fungal parasitism in cysts of Heterodera. 1. Preliminary
investigations. Trans Br Mycol Soc 62:585 – 594, 1974.
Zare R, Gams W. A revision of Verticillium section Prostrata. IV. The genera
Lecanicillium and Simplicillium gen. nov. Nova Hedwigia 73:1– 50, 2001a.
Zare R, Gams, W. A revision of Verticillium section Prostrata. VI. The genus
Haptocillium. Nova Hedwigia 73:271– 292, 2001b.
Zare R, Gams W, Culham A. A revision of Verticillium sect. Prostrata. I. Phylogenetic
studies using ITS sequences. Nova Hedwigia 71:465– 480, 2000.
Zare R, Gams W, Evans HC. A revision of Verticillium section Prostrata. V. The genus
Pochonia, with notes on Rotiferophthora. Nova Hedwigia 73:51 – 86, 2001.
Zehr EI. Evaluation of parasites and predators of plant parasitic nematodes. J Agric
Entomol 2:130–134, 1985.
Zopf W. Zur Kenntnis der Infektionskrankheiten niederer Thiere und Pflanzen. Nova Acta
Leopold Carol Dtsch Akad Naturf 52:314– 376, 1888.
Clavicipitaceous Anamorphs Parasitizing Nematodes 73