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Lakkireddyetal. AMB Expr (2020) 10:141
https://doi.org/10.1186/s13568-020-01085-5
ORIGINAL ARTICLE
Mycoparasite Hypomyces odoratus infests
Agaricus xanthodermus fruiting bodies innature
Kiran Lakkireddy1,2†, Weeradej Khonsuntia1,2,3† and Ursula Kües1,2*
Abstract
Mycopathogens are serious threats to the crops in commercial mushroom cultivations. In contrast, little is yet known
on their occurrence and behaviour in nature. Cobweb infections by a conidiogenous Cladobotryum-type fungus iden-
tified by morphology and ITS sequences as Hypomyces odoratus were observed in the year 2015 on primordia and
young and mature fruiting bodies of Agaricus xanthodermus in the wild. Progress in development and morphologies
of fruiting bodies were affected by the infections. Infested structures aged and decayed prematurely. The mycopara-
sites tended by mycelial growth from the surroundings to infect healthy fungal structures. They entered from the
base of the stipes to grow upwards and eventually also onto lamellae and caps. Isolated H. odoratus strains from a
diseased standing mushroom, from a decaying overturned mushroom stipe and from rotting plant material infected
mushrooms of different species of the genus Agaricus while Pleurotus ostreatus fruiting bodies were largely resistant.
Growing and grown A. xanthodermus and P. ostreatus mycelium showed degrees of resistance against the mycopatho-
gen, in contrast to mycelium of Coprinopsis cinerea. Mycelial morphological characteristics (colonies, conidiophores
and conidia, chlamydospores, microsclerotia, pulvinate stroma) and variations of five different H. odoratus isolates are
presented. In pH-dependent manner, H. odoratus strains stained growth media by pigment production yellow (acidic
pH range) or pinkish-red (neutral to slightly alkaline pH range).
Keywords: Mycopathogen, Hypomyces, Agaricus, Mushrooms, Conidiation, Microsclerotia
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Introduction
Commercially cultivated mushrooms can be attacked by
distinct mycoparasites such as the edible Agaricus bispo-
rus by the ascomycetes Lecanicillium fungicola, Myco-
gone perniciosa (teleomorph Hypomyces perniciosus),
and Cladobotryum dendroides (teleomorph Hypomyces
rosellus) which cause dry bubble, wet bubble and cobweb
disease, respectively (Largeteau and Savoie 2010; Berend-
sen etal. 2010; Carrasco etal. 2017). Such infections can
result in severe crop losses, particularly in later flushes, if
hygienic standards during cultivation are not high. Infec-
tions might originate from contaminated soil or spawn
and the fungi might be introduced into mushroom cas-
ing in the form of spores or mycelium (Adie etal. 2006;
Soković and Van Griensven 2006; Szumigaj-Tarnowska
etal. 2015; Carrasco etal. 2017).
Lecanicillium fungicola not only infects the genera-
tive stage of A. bisporus but at all phases of fruiting body
development (North and Wuest 1993; Calonje et al.
2000; Bernardo etal. 2004; Largeteau etal. 2007; Nunes
etal. 2017). Depending on the developmental stage that
becomes infected, disease symptoms range from totally
undifferentiated spherical masses formed together by
mycelia of host and pathogen (“dry bubble”), over par-
tial disruption of stipe and cap tissues resulting in stipe
deformations (“stipe blowout”) to small necrotic lesions
in the cap (“spotty cap”) (North and Wuest 1993; Soler-
Rivas et al. 2000; Largeteau et al. 2007, Largeteau and
Savoie 2010; Bailey etal. 2013). Early infection of fruiting
body initials by M. perniciosa also leads to the formation
Open Access
*Correspondence: ukuees@gwdg.de
†Kiran Lakkireddy and Weeradej Khonsuntia contributed equally to the
work
1 Department of Molecular Wood Biotechnology and Technical Mycology,
Büsgen-Institute, Georg-August-University, Göttingen, Germany
Full list of author information is available at the end of the article
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Page 2 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
of undifferentiated hyphal masses (“sclerodermoid mush-
rooms”). ese “wet bubbles” are first white and spongy.
en, they turn brownish and may be covered by amber-
coloured liquid excretions. Mushroom deformations and
cap spotting result from infections at later developmen-
tal stages (Fletcher etal. 1995; Umar and Van Griensven
1999; Umar etal. 2000; Glamoclijaet al. 2008; Kouser
and Shah 2013; Zhang etal. 2017). e soil inhabiting C.
dendroides covers all stages of fruiting bodies in form of
coarse white mycelium (“cobweb”) under massive con-
idiospore production. Overgrown mushrooms eventu-
ally rot and collapse. Further symptoms linked to cobweb
disease are brown spotting on caps instigated by ger-
minating spores (Bhatt and Singh 2002; Potočnik 2006;
Parrag etal. 2014; Carrasco et al. 2017). In recent time,
other Cladobotryum species (mainly C.mycophilum,
teleomorph Hypomyces odoratus; C. varium, teleomorph
Hypomyces aurantius) have more often been reported to
cause cob-web diseases including cap spotting and patch-
ing on A. bisporus (McKay etal. 1999; Grogan and Gaze
2000; Back et al. 2010, 2012b; Lee et al. 2011; Sharma
et al. 2015; Carrasco et al. 2016, 2017; Chakwiya etal.
2019). According to McKay etal. (1999), Grogan (2006)
and Tamm and Põldmaa (2013), when H.odoratus occurs
in mushroom farms, it is quite often misidentified under
the name H. rosellus. e sexual fruiting bodies (peri-
thezia) cannot easily be differentiated morphologically
between the species unlike their conidiophores with
the asexual conidia (Rogerson and Samuels 1993, 1994).
Asexual strain features together with molecular data are
therefore used to define species (Kirschner et al. 2007;
Põldmaa 2011; Tamm and Põldmaa 2013; Gea et al.
2019).
e different mycopathogens are not restricted to A.
bisporus but may affect also other commercially culti-
vated species. Incidences of L. fungicola disease were
reported for other Agaricus species (Gea etal. 2003) and
Pleurotus ostreatus (Marlowe and Romaine 1982). M.
perniciosa is shown to also infect Pleurotus eryngii and
Pleurotus nebrodensis as well as Volvariella volvaceae,
with the result of fruiting body malformations (Sisto etal.
1997; Sharma and Kumar 2000; Carrasco et al. 2017).
Aggressive cobweb infections by Cladobotryum species
were described for cultured Calocybe indica (Sharma
etal. 2015), Coprinus comatus (Wang etal. 2015), Flam-
mulina velutipes (Kim et al. 1999; Back et al. 2012b),
Ganoderma tsugae (Kirschner et al. 2007), Hypsizygus
marmoreus (Back etal. 2012a, b, 2015), Pleurotus sajor-
caju (Sharma etal. 2015), P. eryngii (Kim etal. 1998, 2014;
Gea et al. 2011, 2016, 2017; Back et al. 2012b), and P.
ostreatus (Pérez-Silva and Guevara 1999; Gea etal. 2019).
While attention is paid on pathogen infections in
commercial mushroom cultures due to the high eco-
nomic interest, infection events observed in nature are
scattered and usually not deeply described. In nature,
an association with basidiomycete fruiting bodies and
verticillium-like anamorphs (conidiophores are verticil-
late with whorls of few to several phialides which give
rise to the phialoconidia) can help to identify potential
mycopathogens (Gray and Morgan-Jones 1980; Zare and
Gams 2008; Rogerson and Samuels 1989, 1993, 1994;
Põldmaa and Samuels 1999; Põldmaa 2003; Tamm and
Põldmaa 2013; Chakwiya etal. 2019). From the wild, L.
fungicola has been isolated from fruiting bodies of Agari-
cales (e.g. Marasmiellus ramealis, Hypholoma capnoides
and Laccaria laccata) and of decaying samples of el-
ephora terrestris from the elephorales. Lecanicillium
flaccidum from the same species complex was obtained
from basidiocarps of Coltricia perennis of the Hymeno-
chaetales and of Gomphidius glutinosus from the Bole-
tales, and of decaying samples of Russula nigricans of
the Russulales (Zare and Gams 2008). Incidences of
Hypomyces/Cladobotryum infections appear to be more
common. C. dendroides and C. mycophilum have a broad
host range and have been isolated from mushrooms of
varied species of Agaricales, Boletales, Hymenochaetales,
Polyporales, Russulales, Telephorales and others. How-
ever, there are several more mycopathogens between the
paraphyletic Hypomyces/Cladobotryum species group,
several of which are producing yellow to red-coloured
pigments and some of which have a more restricted host
range (Gray and Morgan-Jones 1980; Sohi and Upadhyay
1986; Rogerson and Samuels 1989, 1993, 1994; Helfer
1991; Põldmaa and Samuels 1999; Douhan and Rizzo
2003; Põldmaa 2003; Valdez and Douhan 2012; Tamm
and Põldmaa 2013; Marzuko etal. 2015; Wang etal. 2015;
Zare and Gams 2016). In particular, orange-red lobster
mushrooms are fruiting bodies of Russula, Lactarius and
Lactifluus species from the Russulales which are infested
by staining Hypomyces lactifluorum and are collected and
commercially marketed as culinary delicacy in Mexico
and Northern America (Laperriere etal. 2018).
In this report, we describe our observations on infes-
tations of Agaricus xanthodermus fruiting structures in
nature with strongly sporulating ascomycetous myco-
pathogens. We isolated mycopathogenic strains from
infested material and describe their morphology and
molecular identity with ITS sequences as H. odoratus/C.
mycophilum. Furthermore, we performed infection stud-
ies with vegetative mycelium and fruiting structures of
different basidiomycetous species.
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Page 3 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
Materials andmethods
Mushroom observations, collection andfungal strain
isolation
Mushrooms of A.xanthodermus growing underneath a
Pseudotsuga menzii tree on the north side next to build-
ing Büsgenweg 5 of the Faculty of Forest Sciences and
Forest Ecology (latitude 41.55933; longitude 9.95722)
on the grounds of the North Campus of the University
of Göttingen were usually observed and photographed
at noon (at about 13 to 14 o’clock). Climate data (tem-
perature and humidity) were routinely collected on the
grounds through a hygro-thermo transmitter (Adolf
ies GmbH & Co. KG, ies Clima, Göttingen, Ger-
many). Mushrooms were identified by morphology using
Breitenbach and Kränzlin (1995).
Crippled and decaying mushrooms were collected as
well as rotting grass/moss samples with obvious white
fungal mycelium. e samples were directly brought to
a classroom laboratory and photographed by an IXUS
115 HS digital camera (Canon, Krefeld, Germany). For
enlarged views, a M205 FA stereomicroscope with an
integrated CF420 camera was used and the Leica Appli-
cation Suite v3.8 software (Leica, Wetzlar, Germany).
Samples of infesting mycelium from the cap of a crip-
pled mushroom and mycelial samples of isolated cultures
were observed with an Axioplan 2 imaging microscope
(Carl Zeiss, Göttingen, Germany) equipped with a Soft
Imaging System ColorView II digital camera. Digital
photos taken were processed with the Soft Imaging Sys-
tem analySIS software (EMSIS, Münster, Germany). Size
parameters were measured with the Arbitrary Distance
function of the program and Excel (Microsoft, Redmond,
WA) was used for calculations.
To isolate the basidiomycete, small mycelial samples
were aseptically taken from the inner stipe regions of a
healthy mushroom, and tissues were transferred onto
MEA (2% malt extract, 1% agar; initial pH 5.0) plates
with added antibiotics (ampicillin 100µg/ml, kanamycin
50µg/ml, tetracycline 10µg/ml, chloramphenicol 20µg/
ml and streptomycin 100µg/ml) as described formerly in
Badalyan etal. (2011). To isolate the potential mycopath-
ogens, foreign mycelia were taken from outer infested
stipe and cap regions as well as from a grass/moss sam-
ple and transferred onto MEA plates supplemented with
antibiotics. Plates were incubated at room temperature
(RT) in the classroom. Growing mycelial samples were
transferred for strain isolation and colony observations
onto fresh MEA and YMG/T (0.4% yeast extract, 1%
malt extract, 0.4% glucose, 0.001% tryptophan, 1% agar;
Granado etal. 1997; initial pH6) for growth at RT. Plas-
tic Petri dishes (9cm in Ø) with vents were used. Yeast
extract (LP0021) and malt extract (L39) were from Oxoid
(Basingstroke, UK), agar (Nr. 11396) from Serva (Heidel-
berg, Germany).
e isolated dikaryotic mycelium of A.xanthodermus
(strain KKRL1) and the five different mycopathogen iso-
lates (AscoA1, AscoB1, AscoC1, AscoD1, AscoE1) of this
study were deposited in the DSMZ (Deutsche Sammlung
von Mikroorganismen und Zellkulturen GmbH) strain
collection in Braunschweig (Germany) under Catalog
numbers DSM 111245 (KKRL1) to DSM 111250 (AscoA1
to AscoE1), respectively.
Colony characterisation
Cultures were grown at RT if not otherwise stated. Cul-
tures were photographed with the IXUS 115 HS digital
camera. pHs of culture medium were estimated with pH
indicator strips which were dipped into squeezed agar
pieces cut out from fresh and from mycelium overgrown
medium. Mycelial samples with conidiophores, conidia
or chlamydospores were observed under a Zeiss Axi-
oplan 2 imaging microscope, digital photos were taken
and size parameters measured with the analySIS software
as described above. Diameters of chlamydospores as of
microsclerotia and dense mycelial patches were meas-
ured crosswise in two directions and averages were calcu-
lated from all data. White mycelial patches were analysed
in digital photos of complete cultures and microsclerotia
using colony views of older cultures with collapsed aer-
ial mycelium as photographed under the M205 FA ster-
eomicroscope. Conidia from fully grown whole cultures
were harvested from the culture surfaces as described in
Kertesz-Chaloupková etal. (1998), spores attached to the
lids of Petri dishes were washed off with sterile water and
added to the spores harvested from the colony surfaces
and total spores were counted using a hematocytometer.
ITS sequencing
Genomic DNA was isolated from mushroom samples
taken from outside and from mycelium in culture (Zolan
and Pukkila 1986). ITS sequences of basidiomycetes were
PCR-amplified with primers ITS1 (TCC GTA GGT GAA
CCT GCG G) and ITS4 (TCC TCC GCT TAT TGA TAT
GC) (White et al. 1990) and of ascomycetes with prim-
ers ITS-1* (TCC GTT GGT GAA CCA GCG G) (Waal-
wijk et al. 1996) and ITS4 and analyzed as described
before (Naumann et al. 2007). Gene sequences were
deposited in GenBank under the Accession numbers
KX098646-KX098654.
Mushroom infestation tests
Commercial mushrooms of A. bisporus (cap Ø 3.7 to
5.7 cm) and P. ostreatus (cap width between 2.4 and
6.7 cm) were purchased from a local supermarket. A.
bisporus fruiting bodies were longitudinally cut into
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Page 4 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
halves and transferred into sterile crystal dishes (18.5cm
in Ø, 4.5cm in height) with the cut side alternatively
positioned to the top or to the bottom of the dish. Other
A. bisporus mushrooms were used in whole in erect
condition. P. ostreatus fruiting bodies were used either
in whole or as halves in upside-top (lamellae oriented
down) and in upside-down position (lamellae oriented
to the top). Non-injured caps or cuts of caps or cut or
non-cut sides of stipes of the fruiting bodies of A.bispo-
rus and either caps or stipes of P. ostreatus were infested
with small freshly grown MEA agar pieces of mycelial
isolates, the crystal dishes were closed by their lids and
incubated at RT. Every 12 to 24 h, mushrooms were
inspected and photographed. For every isolate, at least
35 mushroom samples of A. bisporus and 25 mushroom
samples of P.ostreatus were tested in at least 4 rounds of
experiments.
Further, Agaricus mushrooms collected in Septem-
ber 2015 from the wild in other places in Göttingen-
Weende/-Nordstadt were transferred into sterile glass
jars and infested either on the cap or at the bottom of the
stipe by small MEA agar pieces with freshly grown myce-
lial samples. Mushroom identities were determined by
morphological means (Breitenbach and Kränzlin 1995)
and ITS sequencing as A. xanthodermus (KX098653) and
Agaricus sp. section Arvenses (KX098654).
Culture infestation tests
Mycelial cultures of Coprinopsis cinerea strain AmutB-
mut (A43mut, B43mut, pab1-1; Kertesz-Chaloupková
etal. 1998), P. ostreatus monokaryon Pc9 (CECT20311),
and of the isolated dikaryon KKRL1 of A. xanthodermus
were prepared by inoculating one or two small freshly
grown mycelial samples in the middle or at equal dis-
tances distributed on MEA or YMG/T plates and incu-
bating them for vegetative growth at 37°C (C.cinerea
for subsequent grown mycelial challenge tests) or room
temperature (RT, about 22°C, used for other species in
all grown mycelial challenge tests). Once a basidiomy-
cete mycelium was fully established, a culture was chal-
lenged with two ca. 1 × 1 mm small inocula of freshly
grown MEA agar pieces of a mycelial isolate to be tested
by placing them onto the already grown basidiomycete
mycelium 2 cm apart from the basidiomycete inocu-
lum. e dual cultures were further incubated at RT and
observed on daily basis for at least 20days and in some
instances for up to 2months. Plates were photographed
by an IXUS 115 HS digital camera. Five (A.xanthoder-
mus) to six repeats (others) with two to three plates each
were followed up per strain combination and MEA or
YMG/T medium. Mycelial samples were observed under
a Zeiss Axioplan 2 imaging microscope (Carl Zeiss, Göt-
tingen, Germany).
In other sets of experiments (mycelial confronta-
tion tests), basidiomycetes were inoculated on MEA or
YMG/T medium 1.5cm apart from the edge of a Petri
dish (and pregrown when needed; see “Results” section),
and the mycelial test strains 1.5cm apart from the edge
of the opposite side of the Petri dish. All plates were incu-
bated at RT and observed for about a month and more
after they were fully overgrown by the two mycelia. Five
(A.xanthodermus) to six repeats (others) with two to
three plates each were followed up per combination on
MEA medium or YMG/T medium. Plates were regularly
observed and photographed by an IXUS 115 HS digital
camera and under a Zeiss Stemi 2000-C Binocular (Carl
Zeiss, Göttingen, Germany). Presence of conidiophores
and -spores of test isolates and hyphae of basidiomycetes
were followed up by observing small mycelial samples
from confrontation zones under a Zeiss Axioplan 2 imag-
ing microscope.
Results
Mushroom development ofAgaricus xanthodermus
innature
Since 2012, we observed every year but in 2018 and
2019 as 2years with very dry hot summers that multi-
ple white fruiting bodies of an Agaricus species appeared
singly or in small loose groups variably in the months
June to November in the thick layer of needle and cone
litter underneath a P.menziesii (Douglas fir) tree and
in the nearby grass of the surrounding meadow on the
North Campus of Göttingen University (Fig. 1a). Ini-
tially, we noticed the conspicuous mushrooms either in
still closed or in already opened conditions. Later with
better attention we also saw smaller primordia (< 1cm
(See figure on next page.)
Fig. 1 Agaricus xanthodermus fruiting bodies. a Mushrooms (marked by arrows) underneath a Pseudotsuga menziesii tree on the 3rd of September
2015. b Drum-stick-like young mushrooms: the left one is grown to full size (the arrow points to the partial veil underneath the cap). c Mushroom
opening and d fully opened mushroom with vestiges of the partial veil at the edge of the opened cap and a skirt-like annulus around the stipe
(marked by arrows). e–l Diseased crippled young mushroom with split stipe and cap and an infested primordium partially covered by a fluffy
foreign mycelium (marked by an arrow) on 1st of September 2015 at the day of detection (e), 1 day after (f, h; the arrow points to partial veil still
attached to cap tissues), 2 days after (g, i; note the pinkish still healthy lamellae in i) and 3 days after (j), when the mushroom was harvested (k; note
the now brown colour of the lamellae and the white foreign mycelium which covers the crippled stipe and grows onto the lamellae). After harvest,
white mycelium was seen spread over the needle and cone litter layer, the decayed primordium and a cone from the Douglas fir underneath (l)
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Page 5 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
a
bcd
ef g
hi
jk l
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Page 6 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
in Ø) emerging through the soil from broken ground.
Mushroom production appeared to correlate with 1 to
3 prior days of high humidity triggered by good rain-
fall (about 80% and 95% humidity at days and nights)
when temperatures reduced with the rainfall by about
5 to highest 10° from prior day time temperature val-
ues which were between 18 and to up to 30°C at former
warmer days (the actual temperatures depended on the
time of the year). Spherical primordia were observed
aboveground 2 to 4days after the inducing days of high
humidity and reduced temperature, still closed mush-
rooms with lengthened stipes (“drum-sticks”) 4 to 8days
and opened mushrooms 10 to over 20 days after the
rainfalls, when the days following induction were sunny
and again warmer in temperature by an increase of 2 to
5° and when humidity values differed between night (ca.
80–95% humidity) and day periods (ca. 60–85% humid-
ity). Fruiting body development continued usually in a
temperature range from 15 to slightly above 20°C. How-
ever, depending on the month there were also exceptional
days encountered in later fruiting body development with
temperatures up to 30°C.
We observed round ball-like primordia (about
1.5–2 cm in Ø) on the floor and closed young white
mushrooms that had a drum-stick shape and were gen-
erated from the spherical primordia by stipe growth and
increase in cap size. Growth from a spherical primor-
dium into a full-sized drum-stick-like young mushroom
took several days, 2 to 3days at warmer days (18–22°C),
while it slowed down up to 6 to 8days at colder tempera-
ture (12–15°C). Fully grown drum-sticks were up to 10
to 12cm tall with a cap diameter of about 3 to 5cm and
a white partial veil at the underside of the cap that cov-
ered the lamellae (Fig.1b). During maturation in the fol-
lowing 2days, the white partial veil perforated with cap
extension at the edge of the pileus. e remaining con-
nections ripped apart with further cap opening and gave
the stretched pileus a gear-wheel appearance by tooth-
like vestiges (Fig. 1c, d). With the ripping, the partial
veil stayed first as a well-shaped skirt-like white annu-
lus around the stipe (about 1.0 to 1.5-cm in Ø, with the
lower base somewhat swollen) at a distance of about 2
to 2.5cm beneath the cap, but it degenerated with time
over the following days. Opened caps were about 10 to
13cm in diameter. On the upper surfaces towards the
centres of the pilei were small yellowish to light brown
scales. With cap opening, the densely arranged masses
of initially pinkish thin lamellae (over 60 full length
primary lamellae per cap with 5 to 7 secondary lamel-
lae in between) turned quickly dark brown. Within 2 to
3 days, the cap colour turned pale-greyish and, slowly
over 10 to 15days, the open matured mushrooms grew
old. e brown thick-walled smooth basidiospores
(examples can be seen in Fig.2m) measured in average
5.05 ± 0.5 × 3.89 ± 0.63µm (n = 21).
By mushroom morphology and spore sizes, our mor-
phological observations on the mushrooms concur with
the descriptions by Breitenbach and Kränzlin (1995) for
A.xanthodermus. However, strong yellow coloration
upon injury of stems as typical for the species was first
not noted; a faint yellow colour was seen on scratched
freshly harvested mushroom stipes in September 2016
and again in July and more intensively in August 2017.
e odour of healthy mushrooms of the colony was
rather a faint mushroom scent than the typical pun-
gent phenol odour of the species (Gill and Strauch 1984;
Petrova etal. 2007) which in contrast was noticed by us
for other A.xanthodermus colonies in the Göttingen-
Weende area. Lack of both parameters together initially
lead to a misidentification as Agaricus macrosporus by
its very similar mushroom shapes and sizes (Lakkireddy
etal. 2016). e species identity A.xanthodermus of the
mushroom colony underneath the P. menzii tree was
here confirmed by sequencing ITS DNA which was PCR-
amplified from genomic DNA of a stipe of a mushroom
harvested on 4th of September 2015. e established
sequence (KX098652) was 99 and 100% identical to A.
xanthodermus sequences AY484689 and DQ182529.1
from GenBank (Geml etal. 2004; Kerrigan etal. 2005).
Diseased mushrooms ofAgaricus xanthodermus innature
On 1st of September 2015, among several normal healthy
fruiting bodies, we noticed a crippled young mushroom
at the late drum-stick state that had a bended deformed
stipe and a split cap (Lakkireddy etal. 2016). A directly
neighboured primordial mushroom had dropped and was
half-covered by a mycelial white network that extended
over the stipe onto the edges of the cap of the other
crippled individual (Fig. 1e). Over the next 2days, the
still healthy parts of the cap of the crippled mushroom
extended in size to expose the pinkish lamellae while the
primordial mushroom degenerated into an amorphous
clump under actions of the foreign mycelium (Fig.1f–i).
As seen a day later, cap tissues of the crippled mushroom
quickly aged, probably accelerated through the presence
of the foreign mycelium. A thick mycelial layer of a fun-
gal infestation was present at the side of the cap that was
closer to the ground (Figs.1j, 2a, b) and as cover over
the stipe of the mushroom (Figs.1k, 2c, d) from which it
grew onto the lamellae (Fig.2d–i). e harvested infected
mushroom had an unpleasant smell. Sequencing of ITS
DNA (KX098651) PCR-amplified from mushroom tis-
sues again confirmed A.xanthodermus as the species
identity.
Conidiophores with oblong spores were obvious
in thick older mycelium grown on the upper side of
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Page 7 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
the cap, on the stipe and the lamellae (Fig.2b–g). We
microscoped mycelial samples from the lamellae and
found conidiophores and hyaline dry conidia (Fig.2j–n)
which suggested that the infestation was of the anamor-
phic genus Cladobotryum of the family of Hypocreaceae
(Hypocreales, Sordariomycetes) of the Ascomycota (Cole
and Kendrick 1971). Conidia were one to four-celled
(18.0% one-celled, 63.9% two-celled, 9.8% three-celled;
8.2% four-celled; n = 61) with the majority being two-
celled as it is typical for e.g. the mycopathogenic type
species C. varium and C. mycophilum (Hughes 1958;
Cole and Kendrick 1971; Rogerson and Samuels 1993;
Back etal. 2012b; Tamm and Põldmaa 2013). Individ-
ual colonies were isolated from mycelium covering the
ab c
def
i
klmn
hg j
Fig. 2 Diseased crippled Agaricus xanthodermus fruiting body. a, b The upper surface of the cap, c, d the surface of the bended stipe, and e, f parts
of the lamellae are overgrown by foreign mycelium with conidiophores visible as white flocks in the aerial mycelium (d). g–i For infestation of the
lamellae, the foreign mycelium grew first over their edges, with conidiophore production starting about 6 to 7 mm behind the growth front (g; see
white flocks at the right side of the photo). j–n Conidia and conidiophores in mycelial samples taken for microscopy from the lamellae. Note the
blastic generation of conidiospores at the tips of phialides (arrows in k–l) and dehiscence scars (basal hilum) at the spores (see arrow in j) and also
the small brown basidiospores of the host (m). Sizes bars correspond to 5 mm (g), 1 mm (h), 200 µm (i), 50 µm (k) and 20 µm (j, l–n)
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Page 8 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
stipe (strains AscoA1 and AscoB1) and from lamellae
(AscoC1) of the infested mushroom.
Upon aging, degenerating mushrooms in the meadow
were also visibly attacked by similarly sporulating fungi
(not further shown). Another mycelial strain (AscoE1)
was thus isolated on 11th of September 2015 from a
heavily infested rotting stipe of a formerly healthy A.
xanthodermus mushroom when it was found knocked-
down in course of aging on the meadow.
Following some heavy rainfall on 14th and 15th of
September 2015 with a drop in temperature from the
16–21 °C at previous days, a second flush of A. xan-
thodermus mushrooms was observed in the 3rd week
of September 2015, at day temperatures (noon) of 12 to
19°C. Small spherical primordia were seen first on the
16th of September. Several structures were found 5days
later to be diseased at different developmental stages of
mushroom development. Infestations started from white
mycelial patches of several cm in diameter that devel-
oped first well visible on the 18th of September in the
neighbourhoods on moss and decaying grass (Fig. 3),
needles and cones (not shown). Sometimes these patches
originated clearly from the remains of older mushrooms
(Fig. 3f) but there were also multiple patches of fluffy
white mycelium that did not obviously connect to a place
of former mushroom production (Fig.3n). Another fun-
gal colony (strain AscoD1) was isolated from a decaying
grass and moss sample from such a patch of sporulating
white mycelium.
Mushrooms of differential developmental ages became
infested by foreign mycelium, even very young pri-
mordia (Fig. 3a). Fluffy white mycelium grow onto the
lower base of another young mushroom at the begin-
ning of stipe outgrowth and, possibly as a consequence,
the stipe of the young mushroom strongly bended with
the mushroom cap laying down on the floor (Fig.3a, b).
Erect older drum-stick-like stages with extended stipes
were also seen to be confined from the bases of the stipes
(Fig.3d–g). In some instances, heavy infestation lead to
reddish-brown to lilac decolourisation of stipes (Fig.3d,
f) and also caps, and to collapse of the young mushrooms
(Fig.3f, g). Also older structures at and after cap open-
ing were attacked by foreign mycelium (Fig.3h–m). Caps
of attacked mushrooms turned brown to blackish-brown
and shrivelled, thus quickly grew old (Fig.3i, j; l, m) and
rapidly rotted (not further shown). e reactions on
older fruiting bodies appeared to be more aggressive and
faster than reactions on younger stages.
Mycopathogens inculture
All five isolated strains formed conidiogenous mycelium
and grew well on MEA at RT (about 22°C) with increases
in colony radii of 3.7 ± 0.2, 3.8 ± 0.3, 3.6 ± 0.1, and
3.6 ± 0.1 mm/day (AscoA1, AscoB1, AscoC1, AscoE1)
and of 2.4 ± 0.1mm/day (AscoD1), respectively. On the
nutrient-rich YMG/T, the colonies increased in radius
by 4.3 ± 0.1, 4.3 ± 0.1, 4.2 ± 0.2, and 4.4 ± 0.1 mm/day
(AscoA1, AscoB1, AscoC1, AscoE1) and 2.0 ± 0.1 mm/
day (AscoD1). e odour of the fungi when grown on
MEA was pleasant faint sweet aromatic (camphor-like,
resembling Eucalyptus smell). During growth phases on
YMG/T, the odour was also first pleasant faint aromatic
to medicine-like but when cultures on YMG/T aged
and turned wine-red it became unpleasant sharp. e
mycelial scents became stronger on both media with an
increase in growth temperature to 28°C.
All five strains grow on MEA at RT as a first slightly
pigmented mycelium. Growing colonies on MEA of four
of the strains stained first light yellow, while cultures of
strain AscoD1 were stronger yellow from the begin-
ning. Comparably little aerial mycelium was produced
by all strains resulting in overall flat colony appear-
ances. Growing colonies had small white fringed borders
due to the production of multiple conidiophores with
white flocks of masses of dry hyaline conidia. Within 2
to 3days upon production, conidia separated from con-
idiophores and fell in larger aggregates onto the surface
of the yellowish colonies (Fig.4a). Per fully grown MEA
(See figure on next page.)
Fig. 3 Mycoparasitic mycelium infested Agaricus xanthodermus mushrooms at different developmental stages in a larger disease outbreak in the
3rd week of September of 2015. a, b Foreign mycelium grew from surrounding moss to a primordium and the stipe base of a young mushroom
at the stage of stipe elongation and cap growth. c 24 h later the stipe base was surrounded by a thick layer of foreign mycelium, the stipe and
cap were enlarged but the cap laid down on the floor due to strong bending of the stipe. d–m Strong white mycelium found at multiple places
in the grass and moss served as infection source of A. xanthodermus. Bases of elongating stipes of growing drum-stick-like young mushrooms
were covered by a layer of foreign mycelium (d, f) and the same structures 24 h later photographed from different angles (e, g). While the yet less
infected structure with the foreign mycelium confined only to the stipe base was still erect (e), the heavily infected structure with foreign mycelium
reaching up to the cap already collapsed (g). Infested young mushrooms at the start of partial veil rupture (h, i). 24 h later, the cap of the mushroom
shown in i coloured brownish and the rupture of the partial veil blocked. Thick white patches of the pathogen were obvious on the cap surface (see
arrow; j). Also young stages of opened caps (the arrows mark the skirt-like annulus injured by the infestation) were attacked by mycelium growing
upwards the stipe (k) and eventually also onto the lamellae (l, m). Rapid decolourization and mushroom collapse within 24 h resulted from strong
pathogen infestation (l, m). Strong white mycelium found at multiple places in the grass and moss (n)
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Page 9 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
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Page 10 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
AscoA1 AscoB1 AscoC1 AscoD1 AscoE1
a
b
c
Fig. 4 Colony growth of five isolated conidiogenous strains on MEA at RT. a After 3–4 days of incubation with white fringed colony borders (upper
row: plates from top, lower row: from reverse) and b with white pulvinate stroma in fully grown cultures after 25 days of incubation at RT (upper
row: plates from top, lower row: from reverse). c Morphologies of conidiophores (top) and conidia (bottom) taken from aerial mycelium of growing
cultures of isolated Hypomyces odoratus strains. Size bars: 50 µm
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Page 11 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
Table 1 Features ofconidiophores oftheve isolated strains grown atRT
Strains were cultivated for 13days at RT on MEA and conidiophores were taken for microscopy from the outer white growth zone characterized by ocks of conidia.
n = number of structures or cells analyzed. For counting spores per plate, ve plates per strains and per medium were inoculated and spores were counted after fully
growth of plates at RT
Cells Parameter AscoA1 AscoB1 AscoC1 AscoD1 AscoE1
Conidiophores Stem
1st order whorls 2–5 2–4 2–4 2–4 2–3
Whorls with branches 0–3 1–3 0–3 1–3 0–3
Branches per whorl 0–3 1–3 0–3 1–3 0–3
1st order branches 0–7 1–4 0–3 1–3 1–3
1st order branch
2nd order whorls 0–4 0–3 0–2 0–3 0–3
Whorls with branches 0–2 0–1 0–1 0–1 0–1
Branches per whorl 0–1 0–1 0–1 0–1 0–1
2nd order branches 2 1 1 1 1
2nd order branch
Whorls 2 1 1 1 1
Branches total 0–4 0–3 0–2 0–3 0–2
Whorls total 3–7 1–4 1–5 1–4 1–4
n 15 9 8 14 7
Ampulliform phialides Per whorl without branches 2–5 2–5 2–5 2–5 2–5
Per whorl with branches 2–4 2–3 1–3 1–2 1–2
Length (µm) 35.8 ± 7.7 35.7 ± 7.4 38.1 ± 8.3 31.6 ± 5.8 32.9 ± 5.7
Apex width (µm) 2.7 ± 0.4 3.3 ± 0.6 3.4 ± 0.5 3.8 ± 0.6 3.1 ± 0.5
Width broadest point (µm) 8.1 ± 1.1 8.3 ± 1.4 8.1 ± 0.8 8.7 ± 1.2 8.3 ± 1.2
Base width (µm) 5.2 ± 1.0 4.4 ± 0.7 5.0 ± 1.0 4.6 ± 1.1 4.6 ± 0.9
n 27 24 16 20 27
Conidia No septum
Length (µm) 15.3 ± 1.2 15.1 ± 1.4 15.5 ± 1.8 15.4 ± 1.6 16.2 ± 1.0
Width (µm) 10.7 ± 1.3 10.3 ± 1.3 9.6 ± 1.4 10.0 ± 1.0 11.0 ± 1.3
n 21 13 16 15 13
% of total 20.0 14.8 14.4 14.9 13.1
One septum
Length (µm) 20.5 ± 3.0 20.7 ± 3.1 21.2 ± 3.3 21.1 ± 4.8 20.8 ± 3.0
Width (µm) 10.8 ± 1.2 10.3 ± 1.2 10.0 ± 1.3 10.7 ± 1.2 10.4 ± 1.2
n 71 64 67 64 68
% of total 67.6 72.7 60.4 63.4 68.7
Two septa
Length (µm) 24.7 ± 3.8 23.4 ± 1.5 24.4 ± 3.7 26.2 ± 4.4 24.3 ± 2.3
Width (µm) 11.2 ± 1.4 10.5 ± 0.7 10.3 ± 0.9 11.4 ± 1.3 11.2 ± 1.3
n 12 9 17 16 16
% of total 11.4 10.2 15.3 15.8 16.2
Three septa
Length (µm) 26.4 25.3 ± 1.4 31.7 ± 3.3 30.9 ± 4.8 26.6 ± 0.1
Width (µm) 11.9 11.1 ± 0.3 10.5 ± 0.5 12.7 ± 1.3 12.8 ± 1.0
n 1 2 11 6 2
% of total 1.0 2.3 9.9 5.9 2.0
All
Length (µm) 20.0 ± 4.0 20.2 ± 3.6 22.0 ± 5.2 21.7 ± 5.8 20.9 ± 3.5
Width (µm) 10.8 ± 1.2 10.3 ± 1.2 10.0 ± 1.2 10.8 ± 1.3 10.7 ± 1.3
n 105 88 111 101 99
Conidia per plate MEA 4.4 ± 1.2 × 1078.3 ± 4.9 × 1077.9 ± 2.5 × 1071.8 ± 1.4 × 1074.4 ± 1.7 × 107
YMG/T 1.2 ± 0.6 × 1081.2 ± 0.2 × 1089.2 ± 1.3 × 1072.3 ± 0.1 × 1079.9 ± 3.9 × 107
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Page 12 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
plate, the strains produced between 2 × 107 and 8 × 107
hyaline 0- to 3-septate conidia (Table 1). With time,
fully grown colonies turned from slighter yellow to dark
yellow (after 4 to 6days of growth at RT to after about
14days; when cultured at 28°C, these processes were 1
to 3days faster) while strain AscoD1 was still darker pig-
mented as compared to the others. Cultures appeared to
be in a final stage at RT after about 25days of incubation,
for the appearance of mycelium and colony color. e
observations on plates cultivated at RT usually lasted up
to 36days as the plates became drier. Mainly the myce-
lium but also the agar was stained by the strains by yellow
pigments. In one experiment when plates were kept for
2months in very humid conditions, some plates of strains
AscoB1 and AscoD1 adopted in the end a mixed yellow-
slightly pinkish pigmentation while none of the other
strains did change the color. Further in aging MEA cul-
tures after around 3weeks of incubation, all five strains
started to produce hard white patches of dense mycelial
pulvinate stroma which increased in numbers with time
(between dozens to > 100 per plate). First they were small,
less than 0.1mm in Ø, but with time they could grow to
patches of up to 3–4mm in Ø (Fig.4b). After around
30 days of cultivation with drying out medium, round
dark brown microsclerotia (AscoA1: 0.33 ± 0.08 mm in
Ø, n = 16; AscoB1: 0.37 ± 0.08mm in Ø, n = 13; AscoC1:
0.35 ± 0.08 mm in Ø, n = 14; AscoD1: 0.36 ± 0.07 mm
in Ø, n = 17; AscoE1: 0.36 ± 0.07mm in Ø, n = 15) filled
with large round unstained cells formed in aging colonies
(not shown). In addition, masses of round chlamydo-
spores (AscoA1: 13.3 ± 1.7 µm in Ø, n = 22; A scoB1:
13.5 ± 1.7 µm in Ø, n = 26; AscoC1: 13.9 ± 1.6 µm in
Ø, n = 28; AscoD1: 13.5 ± 2.2µm in Ø, n = 22; AscoE1:
13.9 ± 1.2µm in Ø, n = 24) arose in chains from swelling
and fragmenting of vegetative hyphal cells (not shown).
Mycelia of all five strains on YMG/T medium were first
nearly unpigmented during the fresh growth at RT. e
cultures were characterized by loose white fluffy aerial
mycelium starting to regularly develop behind the colony
growth fronts on the 1-day-old mycelium and to produce
conidia over the following days. Spreading in the growing
colony outwards from the inoculum, substrate mycelium
with the agar began to stain yellowish 1 to 2 days after
first aerial mycelium production (at 28 °C 1 or 2 days
earlier than at RT), while the yellow colour intensified
continuously with further mycelial age in growing. e
yellow colour increased in intensity during the further
incubation also after plates were fully grown (after 5 and
in the case of AscoD1 7days of incubation) while after
about 15 and, in the case of AscoD1, 20days there was a
switch in colour to first light pinkish and later wine-red
(Fig.5). e final cultural stages also appeared to have
been reached on YMG/T plates after culturing at RT for
about 25 days, followed by only desiccation reactions
with continued incubation up to 36days.
Pink to red medium coloration has been
reported before e.g. from older PDA cultures of
Hypomyces/Cladobotryum strains (Back et al.
2010; Carrasco et al. 2016; Muhammad et al. 2019).
Hypomyces/Cladobotryum species are known to produce
aurofusarin as pigment (Rogerson and Samuels 1989;
Põldmaa 2011; Tamm and Põldmaa 2013; Carrasco etal.
AscoA1AscoB1AscoC1 AscoD1 AscoE1
Fig. 5 Colony morphology of five isolated conidiogenous strains on YMG/T at RT after 25 days of incubation. Upper row: plates from top, lower
row: reverse
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Page 13 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
2016) which changes in color from yellow to red depend-
ing on the pH (Ashley etal. 1937). We therefore checked
the pH in the medium over the time of cultivation.
When colonies on YMG/T plates were stained yellow-
ish to dark yellow, the pH in the medium did not much
change and was around 5.5 to 6. With onset of pinkish
coloration however, the pH increased to values of 6.5 to
7. With increasing colorization when the cultures turned
pink to finally wine-red, the pH rendered into the alkaline
range to values of around 7 to 7.5 and then to 7.5 to 8.
For comparison, the pH in the MEA cultures was in the
acidic range with pH 4.5 in slightly yellow cultures and
pH 4 and sometimes even pH3 in dark yellow cultures.
In 1-month-old cultures of the stronger yellow cultures
of strain AscoD1, the pH raised first slightly to pH 5. In
2-month-old plates of strain AscoD1 and also of strain
AscoB1, within a few days under color changes to mixed
yellow-pinkish and then yellowish-pink, the pH raised
further to 6 to 6.5 and then pH 7. Cultures of strains
AscoE1, AscoA1, and finally AscoC1 also increased in
pH to 6 but two to several days later, along with color
changes into yellow-pinkish.
e pigments in YMG/T cultures stained majorly the
submerged mycelial agar layer and to less part the agar
beneath. Notably, the colony surfaces remained white in
appearance due to the considerable amounts of whitish
aerial mycelium with huge amounts of hyaline conidia
produced (for spore numbers per fully grown plates see
Table 1). e dry conidia assembled into larger flocks
on the tips of the conidiophores. During colony growth,
thick aerial mycelium arose as high as up to the lids of
the Petri dishes, transferring large parts of the clumps of
spores onto the plastic surface (not shown). is thick
aerial mycelium was longer lasting. After mycelial growth
on a plate was completed, and after the change in col-
our of the substrate mycelium with agar from yellow to
wine-red and along with the evaporation of any humid-
ity from the lids of the Petri dishes (after about 25days
of incubation), the aerial mycelium in undisturbed plates
collapsed slowly throughout the colony. With opening
the lid however, the aerial mycelium collapsed imme-
diately. Eventually, the aggregated conidial clusters fall
down from aerial mycelium and lids of Petri dishes in
irregular patterns onto the surfaces of the colonies. Spore
aggregates collected from agar and from lids of Petri
dishes needed harsh forces to separate them into individ-
ual cells for counting (Table 1). Furthermore, all strains
produced on YMG/T on the surfaces of aging cultures
(after about 30days of cultivation, mainly in the outer
regions of colonies) also masses of dark brown microscle-
rotia which were much more in numbers but of similar
sizes than those on MEA (AscoA1: 0.40 ± 0.08mm in Ø,
n = 15; AscoB1: 0.39 ± 0.05 mm in Ø, n = 20; AscoC1:
0.37 ± 0.05mm in Ø, n = 16; AscoD1: 0.35 ± 0.03mm in
Ø, n = 20; AscoE1: 0.32 ± 0.03mm in Ø, n = 22). Aging
cultures on YMG/T did not produce white stromas but
they gave rise to some chlamydospores resulting in chains
from swellings and fragmenting of hyphal cells (AscoA1:
13.3 ± 1.4µm in Ø, n = 22; AscoB1: 13.5 ± 1.5µm in Ø,
n = 21; AscoC1: 13.2 ± 1.8 µm in Ø, n = 20; AscoD1:
12.9 ± 1.6µm in Ø, n = 24; AscoE1: 13.4 ± 1.7µm in Ø,
n = 23).
Species identication
Conidiophores with conidia were analyzed in more detail
from the strains grown on MEA (Table1). Conidiophores
with conidia on mycelia of all five strains were verticillate
as typical for the Hypomyces/Cladobotryum genus. Con-
idiophores were separated over their length into several
cells. ey had stems with 2 to 5 whorls with up to 5 phi-
alides each and they were usually irregularly branched,
with 1st order sidebranches arising in numbers between
1 and 3 among some phialides at the lower whorls of
the stem and with some 2ndorder sidebranches arising
at the lower whorls of 1st order sidebranches (Fig.4c;
Table1). e up to 5 phialides per whorl were succes-
sively produced (Fig.4c and see also Fig.2j–n) and grew
into lengths of > 30µm (Table1). e ampulliform phial-
ides tapered from broader regions (width > 8µm) shortly
above their bases (width ca. 5µm) to slim blunt apexes
of widths of around 3µm. Conidiospores were produced
at the simple tips of the ampulliform phialides in mono-
blastic mode (Fig.4). First, the young blastospores were
equally swelling but with increase in size, they often
buckled with further growth to the lateral side (Fig. 4c
and see also Fig. 2j–n). Released conidia were hyaline,
oblong in shape with rounded edges, had sometimes
visibly a hilum at the basal ends and different numbers
of septa (Fig.4c). e majority of conidia of all strains
(60–72%) were two-celled. However, strain AscoA1 had
more non-septated spores (20%, comparably to the myce-
lium grown on the infected cap from which AscoA1 was
isolated, please see above) than the other strains. Strains
AscoC1, AscoD1 and AscoE1 had higher numbers of
spores with two or also three septa (in sum 18.2 to 25.2%;
Table 1). Between the strains, there were some meas-
ured minor size variations of the spores (Table1). Spore
lengths ranged from 13.2 to 33.0µm (AscoA1), 13.4 to
28.8µm (AscoB1), 11.7 to 36.5µm (AscoC1) and 13.6
to 37.4µm (AscoD1), 13.8 to 28.2µm (AscoE1). In ten-
dency, spore lengths and widths increased with numbers
of septa (Table1). e strains AscoC1 and AscoD1 with
higher percentages of both 3- and 4-celled spores had
thus a bit more of the longer spores as compared to the
other three strains.
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Page 14 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
e general morphological parameters of conidio-
phores and conidia of the five strains matched descrip-
tions of H. odoratus/C. mycophilum in the literature
(Arnold 1963; Gams and Hoozemans 1970; Cole and
Kendrick 1971; Gray and Morgan-Jones 1980; Back etal.
2012b; Tamm and Põldmaa 2013; Gea etal. 2014). e
occurrence of microsclerotia and presence of round
chlamydospores in the aged mycelium and yellow to
red stained colonies with a strong smell on nutrient-
rich YMG/T medium also concur with descriptions of
H. odoratus/C. mycophilum (Helfer 1991; McKay etal.
1999; Grogan 2006; Gea etal. 2014; Carrasco etal. 2017).
In other instances reported in the literature, no pecu-
liar stronger smell was noted by isolates of H. odoratus
(McKay etal. 1999; Gea etal. 2019), similar as in this
study when growing the five strains on MEA plates.
We amplified and sequenced the 530 bp long ITS
rDNA regions of all five isolates (KX098646-KX098650).
e sequences of the strains are identical to each other
and 99–100% identical to the ITS sequences of H. odo-
ratus (FN859435; Põldmaa 2011) and C.mycophilum
strains (JF693809, JF505112, AB527074, JQ004737,
Y17094, Y17095, KP267826) shown to infect mushrooms
in culture (McKay etal. 1999; Back etal. 2010; Kim etal.
2012; Carrasco etal. 2016; Gea etal. 2016). In contrast,
they were only 98% identical to H.rosellus (FN859440,
FN859442; Põldmaa 2011) and C. dendroides ITS
sequences (Y17090, Y17092; McKay et al. 1999). ree
subgroups of ITS fragments of H. odoratus/C.mycophi-
lum strains are distinguished (McKay etal. 1999; Tamm
and Põldmaa 2013; Gea etal. 2016) by a 1 base pair dif-
ference in the ITS1 sequence (base A at position 80 in
subgroup 1/2 versus G in subgroup 3) and 1 or 2 base
pair differences in the ITS2 region (base T at position
390 in subgroup 1/2 versus C in subgroup 3; base C at
position 507 in subgroup 1 versus T in subgroups 2/3).
Our sequences fall into H. odoratus/C. mycophilum sub-
group 3 together with strains from Ireland, Estonia, Rus-
sia and the USA (McKay etal. 1999; Tamm and Põldmaa
2013).
Fruiting body infection tests
All five isolated H. odoratus strains were tested on com-
plete or halved commercial mushrooms of A. bisporus.
All five strains regularly infected all commercial A. bispo-
rus mushrooms, regardless of whether the inoculum was
placed onto a non-injured stipe or cap or onto cuts of
stipes and caps of sliced mushrooms (Fig.6). After plac-
ing fresh mycelial MEA agar blocks of the ascomycetes
onto a stipe or a cap region of A.bisporus, the hyphae
started to grow (1st day). When intact pilei were inocu-
lated, the surrounding A. bisporus cap region in conse-
quence caved in with a growing pathogen, resulting in a
visible dent with the inoculum in the center (2nd day).
Later on, regardless of place of inoculation, the hyphae
spread over all the mushrooms (3rd day) and produced
huge white-coloured masses of conidia. During this time,
the pathogens were very aggressive and appeared to
absorb nutrients from the mushrooms, while the mush-
rooms reduced in sizes and weights, changed in color
from white to light brownish, and became watery-rotten
(5th to 6th day). All infectious strains, AscoA1 to AscoE1
gave rise to black microsclerotia on the overgrown sur-
face of the A. bisporus samples (not further shown).
In contrast to fruiting bodies of A. bisporus, strains
AscoA1, AscoB1 and AscoD1 did not grow much on
commercial P. ostreatus fruiting bodies, neither when
inoculated on the cap surface nor on the lamellae nor
on the stipes. P. ostreatus thus showed resistance to the
ascomycetes. At most, the H. odoratus hyphae spread
Day 1
Day 5
AscoA1 AscoB1 AscoC1 AscoD1 AscoE1
Control
Fig. 6 Infection test with commercial A. bisporus fruiting bodies. Top row: day of inoculation, bottom: mushrooms after 5 days of cultivation at RT
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Page 15 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
from the inocula only over very small areas of the
mushrooms without obvious symptoms of disease but
not over the complete mushrooms. Importantly, when
placing a mycelial agar block at the centre of mushroom
caps, the H. odoratus mycelia started to grow out more
likely on the side of the agar blocks towards the stipe
region than upwards of the cap region. Only strains
AscoC1 and AscoE1 showed in exceptional cases some
infection by mycelial growth on the base of P. ostrea-
tus stipes (noticed on each 2 of 25 in total tested fruit-
ing bodies; Fig. 7). Still, also in these rare cases the
aggressiveness towards P. ostreatus was comparatively
low with few amounts of conidia formed by the grow-
ing mycelium. As a further interesting observation, P.
ostreatus tissue growth (growing hyphae had clamps)
occurred at the stipe margins, cap margins and the
lamellae of inoculated mushrooms and also of unin-
fected controls. Such growth probably strengthened
the mushrooms and helped in resistance against the
ascomycetes.
We also tested in similar manner A. xanthodermus
mushrooms collected from the wild (KX098653) with
H. odoratus strains AscoA1, AscoB1 and AscoC1. Young
mushrooms with still closed caps were infected by agar
pieces with the ascomycetes positioned either at the
stipes or the caps. Within 6days at RT, mycelium from
the inocula of the stipes grow onto the darkened gills
of the matured mushrooms while stipes with the annuli
degenerated whereas the caps still remained in good
shape (Fig.8). In contrast upon inoculation of pilei, tis-
sues overgrown by the pathogens shrivelled under
appearance of liquid yellow–brown droplets on the cap
surface and the caps degenerated quickly (not shown).
Similar observations were made, when A. arvensis mush-
rooms from the wild (KX098654) were inoculated with
strain AscoC1 (not further shown).
Infection tests ofgrowing mycelial cultures
e infection potential of the five mycopathogenic strains
was further tested against mycelial cultures of A. xan-
thodermus, P. ostreatus and C. cinerea, respectively. For
Day 5
AscoC1 AscoE1
Day 1
Control
Fig. 7 Infection test with commercial P. ostreatus fruiting bodies. Top:
day of inoculation, bottom: mushrooms after 5 days of cultivation
at RT. Arrows mark infested stipe regions. Other mycelial outgrowth
noticed came from P. ostreatus
AscoA1AscoB1
AscoC1
Day 1Day 4Day 3Day 2Day 5Day 6
Fig. 8 Young A. xanthodermus fruiting bodies inoculated with H. odoratus strains. Upon stipe inoculation of mushrooms kept within sterile glass jars,
the progress in mushroom development and pathogen infestation at RT was photographed once per day
Content courtesy of Springer Nature, terms of use apply. Rights reserved.
Page 16 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
mycelial confrontation tests, we inoculated mycelial agar
plugs from a test fungus and a respective mycopatho-
gen onto MEA plates at opposite edges of Petri dishes
(Fig.9a).
e dikaryotic A. xanthodermus isolate KKLR1 grew
slowly on MEA at RT (about 1.4 ± 0.1 mm/day) with a
flat cottony-dense white-colored mycelium growing in a
loosely organized dense fan-strand pattern. erefore for
the mycelial confrontation tests, the species was inocu-
lated at the edges of Petri plates and cultivated 15days
prior to the inoculation of the mycopathogens at the
edges of the opposite side of the plates (see examples
of AscoC1 and AscoD1 confrontation tests in Fig. 9a).
Once the mycelial growth fronts of the two species
reached each other in further incubation (after about 12
to 18days of incubation), both H.odoratus as A. xantho-
dermus colonies were stopped in further growth at the
confrontation zones with some combat reactions at the
colony borders. e flat dense white mycelium of A.xan-
thodermus showed some resistance against overgrowth
by the mycopathogens. However, bunches of sporulating
H.odoratus aerialhyphae were observed to grow over the
A.xanthodermus colonies. e mycopathogens produced
huge amounts of dry aerial conidia which landed in
clumps also on A.xanthodermus mycelium but without
recognisable germination. Unevenly distributed, ribbon
or thread-like mycelial aggregates appearedas deforma-
tions in the A.xanthodermus colonies (Fig.9a) and, with
longer incubation time, on their surfaces also faint zones
of yellowish-stained aerial H.odoratus mycelium over-
laying the basidiomycete. Further in older cultures, A.
xanthodermus appeared to produce a new thicker white
aerial mycelium at the colony borderlines which grow
a few mm to over 1cm into the zones of the opponent
colonies and covered the edges of the H.odoratus colo-
nies (Fig.9a). A. xanthodermus in single culture on MEA
plates rarely produced clamp cells at its hyphae. However,
some clamp cells werealso observed on growth fronts of
the new white mycelium overgrowing the H.odoratus
mycelium, supporting that the basidiomycete survived
and revived in the dual cultures.
Observation with A.xanthodermus strain KKRL1 on
YMG/T plates were in parts similar. A.xanthodermus
colonies were for 4weeks pregrown into colonies of ca.
3 to 3.5cm in diameter prior to inoculation of the myco-
pathogens. e overgrowth of A.xanthodermus colo-
nies by aerial mycelium of H.odoratus strains was then
stronger with fast growing thick bunches of conidiog-
enous hyphae attracted to and overlaying densely the
A.xanthodermus mycelium. Masses of white clumps of
conidia were produced on the plate and fell over the cov-
ered A.xanthodermus mycelium. e areas on the plates
with the grown H.odoratum colonies turned lilac-red
unlike the unstained agar underneath the covered A.xan-
thodermus mycelium. On the reverse of the cultures,
dense assemblies of many submerged brown microscle-
rotia filled with large round cells appeared underneath
the overgrown A.xanthodermus colonies and often also
underneath in the agar zone around the H. odoratum
inocula. Microsclerotia were also observed above the
overgrown A.xanthodermus mycelium. Mycelial samples
from A.xanthodermus colonies overgrown by H. odo-
ratus strains revealed under the microscope single-celled
chlamydospores and many conidia of the ascomycete. No
new outgrowth of mycelium was observed on plates from
the A.xanthodermus colonies which might not have been
strong enoughfor suchactivity if still alive.
In mycelial confrontation experiments with P. ostrea-
tus monokaryon Pc9 on MEA, the mycopathogens grow
also faster than strain Pc9 and the yellow stained colonies
produced huge amounts of conidia (Fig.9a). Where the
growing species met, combat reactions resulted, leading
to a margin of denser white mycelium formed by the Pc9
strain as delineation from the yellowish mycopathogens.
AscoC1AscoD1AscoC1AscoD1
A. xanthodermus
KKRL1
P. ostreatus
Pc9
C. cinerea
AmutBmut
a
b
Fig. 9 Mycelial growth confrontation tests (top rows) and grown
mycelium challenge tests (bottom rows) of A. xanthodermus KKRL1,
P. ostreatus Pc9 and C. cinerea AmutBmut by H. odoratus isolates
on MEA, photographed after 20 days of incubation at RT after
inoculation of the pathogen
Content courtesy of Springer Nature, terms of use apply. Rights reserved.
Page 17 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
However, very long conidiogenous hyphae of the myco-
pathogens loosely overgrow the P. ostreatus colonies and
conidia were produced in small white flocks especially
at the plastic edges of Petri dishes above the P. ostreatus
colonies. With time, some mycelial patches above the Pc9
mycelium stained yellowish. In mycelial samples from
the P.ostreatus colonies under the microscope, no or
only few mostly two-celled conidia were detected. From
the reverse of plates, sometimes thinner necrotic areas
became visible in the unstained P.ostreatus colonies of
older plates (1month) while in other areas and cases the
mycelium grew denser. In mycelial confrontation tests on
YMG/T medium with much more production of aerial H.
odoratus mycelium, reactions were much stronger. Con-
idiogenous hyphae were strongly attracted in growth to
the Pc9 colonies for covering the colonies, and masses
of conidia in many very large aggregates were produced
above the P.ostreatus mycelium. On the reverse side of
cultures, production of some brown microsclerotia were
observed underneath in the unstained P. ostreatus colo-
nies, while all other parts of the medium covered with
mycelium of H. odoratum strains were stained pink to
lilac-red.
e third species tested, C. cinerea homokaryon Amut-
Bmut, in contrast was not able on MEA to defeat any of
the five mycopathogens in combat reactions. C. cinerea
was easily overgrown by all five H.odoratum strains
when the growing colonies were confronted with each
other. H.odoratum strains formed regular yellow colo-
nies with also regular conidia production over the whole
plates including the growth zones of C. cinerea (Fig.9a).
Because C.cinerea AmutBmut is a self-compatible
homokaryon by mutations in its mating type loci, it forms
clamp cells at its hyphal septa (Swamy et al. 1984). In
mycelial samples of overgrown C. cinerea colonies under-
neath the microscope, clamp cells at hyphal septa were
only exceptionally seen, suggesting that at least parts of
the existing mycelium probably came from the myco-
pathogens. White pulvinate stroma developed on top of
the C.cinerea colonies in 1 month old MEA plates. In
confrontation tests on YMG/T plates, C.cinerea colonies
of equal growth age than the mycopathogens were also
quickly overgrown by the H.odoratus strains through
outgrowth of dense fast growing hyphal fans of H. odo-
ratus mycelium being attracted to the smaller C. cinerea
colonies. Masses of conidia were produced on top of the
C. cinerea colonies while the edges of the colonies were
less sharp and, as seen on the reverse of the plates, the
pink H. odoratus staining diffused into the borders of
the C. cinerea areas. In confrontation tests with larger
pregrown C. cinerea colonies (inoculated at edges of
plates and incubated 4 days at 37°C prior to inocula-
tion of H.odoratus strains and transfer to RT), defense
was stronger with sharper colony borders against the
mycopathogens. However, the surfaces of the C. cinerea
mycelia were also quickly covered by fast growing conidi-
ophores and masses of conidia.
Infection tests ofgrown mycelial cultures
In other experimental series to challenge a grown test
fungus, mycelial agar plugs of mycopathogens were
placed at 2cm distance from inocula on the top of the
completely grown basidiomycete mycelium (Fig.9b). In
the grown mycelium challenge tests with already estab-
lished mycelium (grown with two inocula per MEA plate
for 20 days at RT), the mycelium of A. xanthodermus
could well resist the five mycopathogens. In the basidi-
omycete colonies, some white thread- or ribbon-like or in
addition also globular compact mycelial aggregates were
detected as reactions (Fig. 9b), similar as before in the
confrontations tests on the same medium. Clamp cells
were detected in the mycelium. When using slowly grow-
ing A.xanthodermus colonies on YMG/T medium for
surface inoculation with the H. odoratus strains, out-
growth of the mycopathogenic strains on thebasidiomy-
cetous colonies was impeded unlike on free agar surfaces.
In mycelial challenge tests, we also noticed on both
media little or no outgrowth of mycopathogens when
inoculated on the top of established Pc9 mycelium
(Fig.9b). Only sometimes in closer vicinity of the inocula
of H. odoratus strains, zones of some denser mycelium
or some minor necrotic reaction were observed. Like the
fruiting bodies of the species, also the vegetative myce-
lium of P.ostreatus exerts thus some but not full resist-
ance against the mycopathogens.
No much outgrowth of the mycopathogens was then
observed when inoculated on YMG/T plates that were
fully grown with dense C. cinerea mycelium (inoculated
in the middle of plates and grown for 6days at 37°C).
When inoculated on top of established but less dense
C.cinerea mycelium on fully grown MEA plates, the
H.odoratus strains however could easily overgrow the
C. cinerea myceliumand surfaces of colonies stained yel-
lowish by thepresence of the mycopathogen (Fig. 9b).
Necrotic areas became visible in the C. cinerea colonies
underneathby thinned mycelium around the inocula of
the mycopathogens in mycelial challenging tests on MEA
(Fig.9b).
Discussion
In this study, we report observations on mycoparasitic
infections of A. xanthodermus mushrooms in nature.
We have observed unimpeded developing mushrooms in
years 2012 to 2017, variably in the months June, August,
September and, in 2015, also in November, usually after
comfortably warm weather conditions. Induction of
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Page 18 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
fruiting body development of A. xanthodermus seems to
need sufficient previous rainfall possibly to both, mois-
ture the ground and create higher humidity in the air.
Consequential to the rainfall, a drop in air temperature
likely will also be favourable for induction of fruiting.
Fruiting body development proceeds from ball-like pri-
mordia over drum-stick-shaped, still closed young mush-
rooms to mature mushrooms with open umbrellas and
first pinkish and then brown lamella (Fig.1). e speed of
development from primordia to fruiting body maturation
seems to depend also on the temperature and took in our
observations between 10–13 and 6–8days at colder and
warmer temperature (around 12–15°C and 18–22°C),
respectively. Mature fruiting bodies can last further 10 to
15days.
Infections ofA. xanthodermus fruiting bodies byH.
odoratus cobweb innature
Interestingly, in a first flush of mushrooms in early Sep-
tember 2015, one split fruiting body was visibly affected
by a fungal infestation (Figs.1e–l, 2). While we do not
know whether this single mushroom was injured prior to
infestation or whether infestation resulted in the injury,
our observations from infections in the subsequent flush
of mushrooms suggest that injury is not a premise of
infection of the species in nature. Moreover, we observed
that all stages of fruiting body development were sus-
ceptible for the mycopathogen (Fig. 3). e infections
on A. xanthodermus were identified by morphological
means (conidiophores and conidia) and ITS sequencing
as H. odoratus (anamorph C.mycophilum). is fungus
is one of a group of closely related species which can
cause cobweb disease of cultivated mushrooms such as
the edible species A. bisporus, P. eryngii and P. ostreatus
(see e.g. Back etal. 2012b; Tamm and Põldmaa 2013; Gea
et al. 2014, 2016, 2019; Carrasco et al. 2016; Chakwiya
etal. 2019). e species proliferates also on mushroom
substrates (Grogan 2006; Carrasco etal. 2016; Gea etal.
2016, 2019) and has also been encountered on thepoly-
pores Ganoderma lucidum (Zuo etal. 2016) and Poly-
porus sp. in culture (Rogerson and Samuels 1994). In
commercial button mushroom cultures, any mushrooms
encountered will be engulfed by the mycopathogen with
radial outgrowth of mycelium on the substrate (Grogan
2006; Muhammad etal. 2019). We have observed similar
events in nature with the mycopathogens growing from
the surroundings (decayed fungal material, decaying
grass/moss, soil) onto the stipes of nearby developing A.
xanthodermus structures (Fig.3).
Spread ofH. odoratus cobweb clones innature
Cobweb disease can be spread by airborne conidia. In
mushroom-growing rooms, the large conidia are released
from spore clusters on the colonies into the air by physi-
cal disturbances such as by watering. When subsequently
landing and germinating on mushrooms, disease symp-
toms can be incurred (Dar 1997; Adie etal. 2006; Grogan
2006). Following initially a single infested mushroom
(Figs.1e–l, 2), we observed a larger outbreak of disease
in nature after heavy rainfalls in the 3rd week of Septem-
ber 2015 in the 2nd flush of mushrooms of A. xanthoder-
mus (Fig.3). It is thus possible that the rainfall helped to
distribute conidia from the place of the previous single
mushroom infestation over the larger area, in addition
to the general promotion of host and pathogen growth
by providing good levels of humidity through rainfall
to both. H. odoratus conidia do not survive long under
dry conditions (Lane et al. 1991) and high humidity is
needed for dispersal and germination (Carrasco et al.
2016, 2017). Attack of A.bisporus by H. odoratus in com-
mercial cultures can happen at any stage in the fruiting
body development (Carrasco etal. 2017; Chakwiya etal.
2019) while infections tend to become more severe on
the crop in later flushes at longer time of cultivation and
during autumn and winter cycles with increasing conidia
numbers (Carrasco etal. 2016, 2017). Our observations
on A.xanthodermus in nature resemble the reports
on disease on A. bisporus in commercial mushroom
production.
While H. odoratum is shown to produce perithe-
cia with ascospores in culture (Arnold 1963; Põldmaa
2011; Tamm and Põldmaa 2013), it is not known to do
so in nature. Clonality is expected to occur in nature of
asexually reproducing Hypomyces species in course of
spreading of conidia as a major mode of reproductive
distribution (McKay etal. 1999; Grogan and Gaze 2000;
Valdez and Douhan 2012; Tamm and Põldmaa 2013;
Carrasco etal. 2017; Chakwiya etal. 2019). We isolated
five H. odoratum strains from a close neighbourhood,
three of a same infested fruiting body (AscoA1, AscoB1,
AscoC1) but of different mushroom organs (from cap
and stipe, respectively). Another isolate (AscoE1) came
from a decaying stipe of a later infested mushroom. eir
properties were very similar, in measurements only dis-
tinct in some minor details. e 5th strain (AscoD1)
isolated from grass/moss was more different from the
other four such as by slower growth speed, a stronger yel-
low colony colour and by lower spore production. is
might suggest that they are not (all) clonal in relation to
each other. Larger population field studies on H. odora-
tus in nature are currently missing in order to know how
much genetic diversity exists in natural populations and
whether sexual reproduction and recombination occurs
in nature. Nearly identical clones have been isolated from
commercial A. bisporus cultivations in different Euro-
pean countries and other continents. Using worldwide
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Page 19 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
the same A. bisporus production strain and spawn and
casing soils from same sources, this could however relate
to human activities in mushroom cultivations if hygienic
conditions were not strictly kept. Further alternative
sources of primary infections in commercial mushroom
cultures were by human movements and other material
transport (Carrasco et al. 2017; Chakwiya et al. 2019).
In contrast, clones from mushroom farms have in some
instances been interlinked to local populations in nature
(Tamm and Põldmaa 2013). ere is thus also a possible
danger for introduction of the pathogens into mushroom
farms newly from the nature.
Outbreaks andhost range ofH. odoratum cobweb
Most of the present knowledge on the species H.
odoratus/C. mycophilum comes from cobweb outbreaks
experienced in newer time in commercial mushroom
cultivations (see “Introduction”; Grogan 2006; Tamm and
Põldmaa 2013). In essence, cobweb disease in mushroom
cultures is caused by different species and up to recently,
there was much confusion on species identities. H.
odoratus/C. mycophilum was often mistaken by H.rosel-
lus, a related species with similar disease symptoms.
H. rosellus has however distinct conidiophores with a
rachis at the apex of phialides, produces only two-celled
conidia, has a more confined host-range and appears to
be less often prevalent in the wild. In addition, the two
species differ in their ITS sequences allowing to distin-
guish the two species further by molecular data why
several misidentified strains were later reassigned to H.
odoratus/C.mycophilum (McKay etal. 1999; Tamm and
Põldmaa 2013). Our morphological and molecular data
define the five strains isolated in this study from the wild
clearly as H.odoratus.
H. odoratus has a very broad host range on mushrooms
growing in nature in temperate regions (Tamm and
Põldmaa 2013). Incidences of infections on agaric fruit-
ing bodies in the wild have sporadically been recorded
before for A. xanthodermus, Armillaria mellea, Calocybe
gambosa, Cortinarius collinitus var. mucosis, Enteloma
clypeatum, Hebeloma sp., Hygrophorus camarophyllus,
Inocybe sp., Lycoperdon pyriforme, Megacollybia platy-
phylla, Mycena galericulata, Oudemannsiella platy-
phylla, Pholiota sp., Pseudoclitocybe cyathiformis, and
Tricholoma terreum as well as occurrence on soil, leaf
litter and rotting wood (Arnold 1963; Gams and Hooze-
mans 1970; Helfer 1991; Rogerson and Samuels 1994;
Tamm and Põldmaa 2013). H. odoratus is considered to
be agaricicolous (Rogerson and Samuels 1994) whereas
other Hypomyces/Cladobotryum species are specified as
boleticolous and polyporicolous (Rogerson and Samuels
1989, 1993; Tamm and Põldmaa 2013). However, Coni-
ophora sp., Suillus aeruginascens and Suillus bovinus
from the Boletales (Arnold 1963; Rogerson and Samuels
1994; Tamm and Põldmaa 2013), Albatrellus sp., Lac-
tarius mitissimus, Lactarius deliciosus, Lactarius qui-
etus, Lactarius cf. vellereus, Russula virescens, Russula
sp., and Stereum sanguinolentum from the Russuales
(Arnold 1963; Gams and Hoozemans 1970; Helfer 1991;
Tamm and Põldmaa 2013), and Trametes versicolor from
the Polyporales (Gray and Morgan-Jones 1980) are fur-
ther named as potential hosts for H.odoratus in nature,
as well as Cantharellus cibarius from the Cantharellales,
Gloeophyllum sepiarium from the Gloeophyllales, and
Clavariadelphus truncatus from the Gomphales (Helfer
1991). Newer observations on the species in nature with
molecular identification would be helpful to unambigu-
ously confirm these claims.
Other than the many incidences in commercial mush-
room cultivations and the mostly older reports on
occasional fungal collections in the wild, little is so far
known on the ecology of necrotrophic Hypomyces spe-
cies such as H. odoratus in nature. Our observations in
nature and the infection tests in the laboratory confirm
A. xanthodermus fruiting bodies to be susceptible to H.
odoratus. e host range of the five isolated strains does
not restrict to A.xanthodermus but include further Aga-
ricus species. e strains grew on and quickly decayed
commercial fruiting bodies of A. bisporus, in accordance
with the various reports in the literature on occurrence of
the species on the white button mushroom in cultivation
(see “Introduction”; Grogan 2006; Tamm and Põldmaa
2013). e host range of the five strains extends also onto
mushrooms of an Agaricus sp. from the section Arven-
ses but not particularly to fruiting bodies of P. ostrea-
tus. Resistance against H. odoratus has been reported
from infection tests for Hypsizygus marmoreus fruiting
bodies (Back et al. 2012a, b, 2015) whereas F. velutipes
(Back etal. 2012b), G. lucidum (Zuo etal. 2016), P. eryn-
gii (Back et al. 2012b; Kim etal. 2014; Gea et al. 2011,
2014, 2016, 2017) and P. ostreatus (Pérez-Silva and Gue-
vara 1999; Gea etal. 2019) were found to be (partially)
susceptible. However, the place of inoculation can play
a role. Upper parts of intact caps of P. eryngii were thus
relatively resistant against H. odoratus infection while the
pathogen could effectively attack mushrooms of the spe-
cies through cuts (Gea etal. 2014, 2016). A recent report
on infestation of P. ostreatus by H. odoratus revealed fur-
ther that the bases of fruiting bodies of this species can
be more sensitive against infections by the pathogen (Gea
et al. 2019), similarly to our own observations on rare
events of overgrowth of mushroom stipes of P. ostreatus
(Fig.9).
Mycelial proliferation of P. eryngii is hindered by H.
odoratus and the species is attacked by the pathogen
at any cultivation stage (Kim etal. 2014). Differently to
Content courtesy of Springer Nature, terms of use apply. Rights reserved.
Page 20 of 22
Lakkireddyetal. AMB Expr (2020) 10:141
the fruiting bodies, mycelium of A. bisporus has been
reported to be resistant e.g. for the wet bubble disease
inducer H. perniciosus (Zhang etal. 2017) while the dry
bubble inducer L. fungicola has variably been found to
attack or not attack host mycelium (Dragt etal. 1996;
Calonje etal. 2000; Shamshad etal. 2009) and the cob-
web inducer Cladobotryum varium overgrow with
time cultures of the basidiomycete and caused necro-
sis (Gray and Morgan-Jones 1981). Furthermore shown
in this study, in mycelial confrontations with growing
or grown A. xanthodermus and P.ostreatus cultures,
the five H. odoratus isolates here were not or not very
aggressive with both species. In contrast, the strains
more strongly attacked mycelial C. cinerea colonies.
is latter species is a dung fungus that likes higher
temperatures around 37°C best for growth (Kües 2000)
while it is poorly growing at lower temperature ranges
such as RT (Fig.9). Strains of the temperate species H.
odoratus grow in temperature ranges of 5 to 25°C and
only very poorly at warmer temperatures up to 28°C
(Back etal. 2012b). As seen in Fig.3, strains of H. odo-
ratus proliferate in nature from soil, plant litter and
former mushroom residues onto their hosts. e two
fungi C. cinerea and H.odoratus may live under quite
different environmental circumstances and ecological
niches why a species like C. cinerea with higher tem-
perature preferences might not have developed a myce-
lial growth resistance at lower temperature towards
this particular pathogen. As also seen in this study, H.
odoratus does not generally infect all Agarics (Fig. 5)
although the mycopathogen has an apparent preference
for them. Other parasitic Hypomyces species appear
to preferentially attack polypores and boletes (Roger-
son and Samuels 1989, 1993). e broader host range
is one criterium to distinguish Hypomyces species,
temperature preferences another. H. odoratus and the
also agaricicolous H. rosellus are adapted to temperate
regions, whereas other Hypomyces species are found
on mushrooms in the tropics and subtropics (Põld-
maa 2011; Tamm and Põldmaa 2013). C. cinerea is an
edible mushroom cultivated in some tropical countries
including ailand (Kües etal. 2007) and it could be of
interest to test whether the species is better resistant
against tropical Hypomyces species.
Acknowledgements
We are very grateful to Mojtaba Zomorrodi for technical help in fungal cultiva-
tions and pH measurements and to Bernd Kopka for providing climate data.
Authors’ contributions
KL, WK and UK observed mushrooms in nature, designed experiments,
performed mycelial confrontation tests and analyzed data. KL performed
most outdoor research, mushroom infection tests and strain isolations. WK
did ITS analyses and provided most of the cell measurements. KL wrote a first
draft of the paper. UK, KL and WK revised the manuscript. All authors read and
approved the final manuscript.
Funding
Open access funding provided by Projekt DEAL. WK acknowledges a Ph.D.
student grant from the Royal Thai government.
Availability of data and materials
Not applicable.
Ethics approval and consent to participate
This article does not contain any studies with human participants or any stud-
ies with animals.
Consent for publication
Not applicable.
Competing interests
All authors declare that they have no conflict of interest associated with this
work.
Author details
1 Department of Molecular Wood Biotechnology and Technical Mycology,
Büsgen-Institute, Georg-August-University, Göttingen, Germany. 2 Center
for Molecular Biosciences (GZMB), Georg-August-University, Göttingen,
Germany. 3 Present Address: Faculty of Agricultural Technology, Rajabhat
Mahasarakham University, Mueang Maha Sarakham District, Maha Sarakham,
Thailand.
Received: 29 July 2020 Accepted: 8 August 2020
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