Bull. Natl. Mus. Nat. Sci., Ser. B, 38(3), pp. 77–85, August 22, 2012

A Preliminary Survey on Larval Diversity in Mushroom Fruit Bodies
Kentaro Hosaka* and Kunihiko Uno
Department of Botany, National Museum of Nature and Science,
4–1–1 Amakubo, Tsukuba, Ibaraki 305–0005, Japan* E-mail: khosaka@kahaku.go.jp(Received 17 May 2012; accepted 26 June 2012)

Abstract We investigated diversity of insect larvae in mushroom fruit bodies. A total of 24 larvae from nine species of mushrooms (7 genera, 3 orders) were collected and DNA sequences wereobtained. The BLAST searches could not identify these larvae to species but they all belong toDiptera. Phylogenetic analyses revealed the presence of a total of nine presumable species of ﬂies
in mushroom fruit bodies. Each species of ﬂies was found from a single fruit body, except for themaggots of Group 2 and 3, which were collected from two species of mushrooms. Although oursampling effort is very limited for this study, the following conclusions can be drawn: (1) Different
species of mushrooms can be inhabited by the same Diptera species. (2) Different species of larvae
can be present in the same mushroom fruit bodies. (3) Same species of larvae can be present in the
different mushroom fruit bodies at different times.
Key words : COI, Diptera, DNA barcode, maggots, mycophagy.

Introduction
Mycophagy (fungal consumption) by mammals and arthropods has attracted researchers for
centuries. Among which, mycophagy by small
animals, especially by rodents and small marsupials, has been well documented as an exampleof trufﬂe-animal interaction (Claridge and May,
1994; Lehmkuhl et al., 2004; Meyer et al.,2005). Because fruit bodies of trufﬂe-like fungi
are produced below ground, the spores are notdisseminated by wind, as is the case in many
above-ground mushroom-forming species, whichhave potential of long distance spore dispersal(Thiers, 1985; Bruns et al., 1989). Instead, they
produce a unique aroma to attract small animals,
which rely on trufﬂe-like fungi as a large part oftheir diet (Thiers, 1985; Castellano et al., 1989).
Trufﬂe-like fruiting bodies are eaten by small
animals and the fungal spores are disseminatedwith the animal feces (Castellano et al., 1989).
This association between small animals and trufﬂe-like fungi can be seen in many areas of theworld (Malajczuk et al., 1987). Because of thetight interaction between mycophagous animals
and trufﬂe-like fungi, some sort of co-evolution
scenario is a possibility. For example, one could
hypothesize that the origin of mycophagous animals and hypogeous fungi occurred at the sametime.
The other well documented examples of interactions include mycophagy by arthropods, which
was reported from 1800ʼs or even earlier (Fulton,
1889). Many reports came from the order
Phallales (commonly known as stinkhorns) as
one of the prime examples of insect mycophagy(Smith, 1956; Shorrocks and Charlesworth,
1982; Tuno, 1998). Most notably, spore mass
(gleba) of Phallales becomes slimy and emits
strong odor when matured, which attracts small
insects, especially ﬂies, and their spores are dispersed by those insects (Fulton, 1889). There arealso large numbers of trufﬂe-like taxa, in addition to stinkhorn-like taxa, in the order Phallales(Hosaka et al., 2006). Interestingly, however,spores of trufﬂe-like taxa in Phallales have not

78

Kentaro Hosaka and Kunihiko Unobeen recorded from animal feces in previous
studies. This may indicate that that spore dispersal of Phallales, including trufﬂe-like taxa, isentirely dependent on arthropods. It is therefore
possible, as in the case of trufﬂe-like fungi, that
mycophagous insects and stinkhorns have coevolved.
Mycophagy by arthropods is known not only
from Phallales, but also from a wide variety ofmushrooms (Martin, 1979; Pacioni et al., 1991).
At the same time, a wide variety of arthropods,
mostly insects, mites, and springtails (Collembola)
has been reported as mycophagous (Tuno, 1999;
Sawahata et al., 2002, 2008; Takahashi et al.,2005; Yamashita and Hijii, 2007; Kadowaki,2010). It is a general trend that insect fauna of
perennial fruit bodies (polypores or conks) is
composed of a wide variety of groups, often
dominated by Coleoptera, but mushrooms withephemeral, soft fruit bodies are mostly fed byCollembola and Diptera, often dominated by thefamily Drosophilidae (Atkinson and Shorrocks,
1981; Takahashi et al., 2005; Kadowaki, 2010;
Tuno, 1999; Yamashita and Hijii, 2007).
Among mycophagous insects, the diversity
and behavior of drosophilid ﬂies have mostextensively been studied (Kimura et al., 1977;
Jeanike, 1978; Grimaldi, 1985; Worthen, 1989;
Worthen et al., 1996; Toda and Kimura, 1997;
Toda et al., 1999; Tuno et al., 2007). For example, spore dispersal of stinkhorns by ﬂies has
been documented for centuries (Fulton, 1889;
Tuno, 1998). Most studies, however, are basedon ﬁeld observation of adult ﬂies, or rearing
experiments from ﬁeld-collected mushrooms(Jaenike, 1978; Grimaldi, 1985; Toda andKimura, 1997; Yamashita and Hijii, 2007;
Kadowaki, 2010), both of which depend on theidentiﬁcation of the adult ﬂies.
We argue that those methods may signiﬁcantly
bias results because not all species of larvae canbecome adults under the experimental settings,
and we therefore may miss the true diversity oflarvae in fruit bodies. In this study, we attempted
to reveal such “hidden” diversity of larvae inmushroom fruit bodies using DNA barcode data

directly sequenced from larval stage. We focusedon collecting only larval stage of ﬂies (maggots),and the identiﬁcation of each ﬂy was entirely
dependent on DNA sequence data. By this way,
we also attempted to link the sequence data withspecies names, and to reveal the presence of
cryptic or new species of ﬂies.
Materials and Methods
Collecting Sites, Collecting Scheme, andCuration of Specimens
Fieldwork was conducted in the year of 2011
(July–September). The main collecting sites werelocated in Tsukuba Botanical Garden, Tsukuba,
Japan (36°06′N, 140°06′E). The garden occupies
ca. 14 ha. (36 acres), and has ca. 5000 taxa ofdomestic and exotic plants. It is divided into several sections simulating natural habitat, such as“Evergreen broad-leaved forest section”, “Temperate coniferous forest section”, and “Warmtemperate deciduous broad-leaved forest section”.
A general collection scheme for mushroomforming fungi was followed. At each collecting
site, fruit bodies were collected and the speciﬁc
location (section) in the Tsukuba Botanical Garden and habitat information were recorded. If
presumable ectomycorrhizal mushrooms werecollected, potential ectomycorrhizal host trees
near fruit bodies, mostly Pinaceae and Fagaceae,
were identiﬁed to species. If alternative ectomycorrhizal hosts were present near fruit bodies,those were recorded as well.
All fruit bodies were collected, photographedand dried using a food dehydrator (Snackmaster
Express FD-60, Nesco/American Harvest, WI,USA) for 24 to 48 hours. In addition to driedmaterials, small fragments of hymenial tissue
from freshly collected samples were soaked in
1.5 ml tubes with DMSO buffer (Seutin et al.,1991) with an addition of 100 mM Tris-HCl (pH8.0) and 0.1 M sodium sulﬁte (Na2SO3) under
4°C, following the procedures of Hosaka (2009)
and Hosaka and Castellano (2008). Voucher
numbers were assigned to representative speci-

A Preliminary Survey on Larval Diversity in Mushroom Fruit Bodies

mens.
When maggots were found, they were collected under dissecting microscope using clean
forceps, and stored in 1.5 ml tubes with DMSO
buffer. At least ten maggots were collected froma single fruit body, and the collection data wererecorded by assigning the voucher number
according to the mushrooms from which maggots were found. If maggots were detected frommultiple fruit bodies of the same specimens(under the same voucher numbers), they werestored in separate tubes. The exact positions ofmaggots found (e.g., cap, gills, stalk, etc) were
not recorded for this study.
Specimens collected during the ﬁeldwork were
deposited at the fungal herbarium of the National
Museum of Nature and Science, Tsukuba, Japan(TNS). All maggot samples were stored in refrigerator at the laboratory of KH.
DNA Preparation, PCR, and Sequencing
DNA of mushrooms and maggots was
extracted from the samples stored in DMSO buffer. Tissues of mushrooms were ground underliquid nitrogen using a mortar and pestle. Maggots were ﬁrst transferred to new 1.5 ml tubes
using clean forceps, and CTAB buffer wasadded. Maggots were then ground using plastic
pestle by hands. Only one maggot was used foreach extraction. DNA was extracted using themodiﬁed CTAB extraction protocol followed byglass milk puriﬁcation methods as summarizedby Hosaka (2009) and Hosaka and Castellano(2008). Brieﬂy, ground samples were incubatedin CTAB buffer at 65°C for 1 hour, and proteins
were removed using the mixture of chloroform :isoamylalcohol (24 : 1). The materials were further puriﬁed using 6 M sodium iodine buffer withglass milk, washed with ethanol/buffer solution,and ﬁnally eluted in 100 μl of TE buffer.
DNA sequence data of mushrooms wereobtained from the internal transcribed spacer
regions (ITS) of the nuclear ribosomal DNA. For
amplifying the ITS region, the primer combination of ITS5 and ITS4 (White et al., 1990) wasused. DNA sequence data of maggots were79

obtained from the cytochrome c oxidase I gene
(COI). For amplifying the COI, the combinationof LCO1490 and HCO2198 (Folmer et al., 1994;
Hebert et al., 2003) was used. PCR reactions
were carried out using 20 μl reaction volumes
each containing: 1 μl genomic DNA, 1 μl dNTP,(4 mM), 1 μl of each primer (8 μM), 0.5 units of
Taq polymerase (TaKaRa, Tokyo, Japan), 2 μl
MgCl2 (25 mM), 2 μl Bovine Serum Albumin(BSA). Cycling parameters were 1 cycle of 94°C
for 3 min, 30 cycles of 94°C for 1 min, 51°C for
30 s and 72°C for 1 min, with a ﬁnal extension at
72°C for 15 min. PCR products were electrophoresed in 1% agarose gels stained with ethidium
bromide and visualized under UV light. Whenampliﬁcation bands were conﬁrmed, PCR products were then puriﬁed using the ExoSap-IT(Millipore, Molsheim, France) and directly
sequenced using the Big Dye Terminator Cycle
Sequencing Kit (Applied Biosystems Inc., Norwalk, CT, USA), following the manufacturerʼs
instructions.
Phylogenetic Analyses
DNA sequences of the COI were aligned manually using the data editor of BioEdit ver. 7.0.1(Hall, 1999). Ambiguously aligned regions were
excluded from the analyses. The dataset was thenanalyzed by maximum parsimony (MP) andneighbor-joining (NJ) analyses. MP analyses
were conducted under the equally weighted parsimony criterion using PAUP* version 4.0b10(Swofford, 2002), with heuristic search option(with TBR and Multrees on) with MAXTREES
set to 10,000. Support for the individual nodes
was tested with bootstrap (BS) analysis under theequally-weighted parsimony criterion. BS analysis was based on 1,000 BS replicates using theheuristic search option (TBR and Multrees
options off), with ten random addition sequences.
NJ analysis was conducted using PAUP* (Swofford, 2002) with the Kimura-2-parameter model,
with a gamma shape set to 0.5. Support for individual nodes was tested by BS analysis based on
1,000 BS replicates under the same settings.

80

Kentaro Hosaka and Kunihiko Uno

Results
Collections of Mushrooms and Larvae
A total of nine species of mushrooms (7 gen-era, 3 orders) contained multiple maggots in theirfruit bodies (Table 1). We intentionally avoided
collecting rotten fruit bodies because maggots
found in such materials could simply be sapro-

Table 1. Maggots sequenced for this study with host mushrooms and collection information
Vocuhernos. ofmaggots

Host mushrooms ofmaggots (orders inparentheses)

Herbarium
nos. ofmushrooms

375-1

Tylopilus sp. (Boletales)

TNS-F-41193

July 7, 2011375-2

Tylopilus sp. (Boletales)

TNS-F-41193

July 7, 2011375-3

Tylopilus sp. (Boletales)

TNS-F-41193

July 7, 2011375-4

Tylopilus sp. (Boletales)

TNS-F-41193

July 7, 2011

535-1

Boletus sp. (Boletales)

TNS-F-41353

August 11, 2011535-2

Boletus sp. (Boletales)

TNS-F-41353

August 11, 2011

535-3

Boletus sp. (Boletales)

TNS-F-41353

August 11, 2011

566-1

Amanita sp. (Agaricales)

TNS-F-41384

August 12, 2011

566-2

Amanita sp. (Agaricales)

TNS-F-41384

August 12, 2011

566-3

Amanita sp. (Agaricales)

TNS-F-41384

August 12, 2011

566-4

Amanita sp. (Agaricales)

TNS-F-41384

August 12, 2011576-1

Calvatia craniiformis(Agaricales)
Russula sp. (Russulales)

TNS-F-41393

August 12, 2011

TNS-F-41397

August 12, 2011

TNS-F-42118

September 22, 2011

TNS-F-42118

September 22, 2011

TNS-F-42118

September 22, 2011

TNS-F-41945

August 18, 2011

593-1

Amanita hemibapha(Agaricales)
Amanita hemibapha(Agaricales)
Amanita hemibapha(Agaricales)
Strobilomyces sp.
(Boletales)
Xerocomus sp. (Boletales)

TNS-F-41950

August 18, 2011593-2

Xerocomus sp. (Boletales)

TNS-F-41950

August 18, 2011593-3

Xerocomus sp. (Boletales)

TNS-F-41950

August 18, 2011

675-1

Russula sp. (Russulales)

TNS-F-42032

September 2, 2011

675-2

Russula sp. (Russulales)

TNS-F-42032

September 2, 2011

675-3

Russula sp. (Russulales)

TNS-F-42032

September 2, 2011

675-4

Russula sp. (Russulales)

TNS-F-42032

September 2, 2011

580-1
761-1
761-2
761-3
588-1

Collection date

The classiﬁcation at the ordinal level was based on Kirk et al. (2008).

Speciﬁc location in
the Tsukuba Botanical Garden
Cool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Evergreen broad-leaved forest
sectionEvergreen broad-leaved forest
sectionEvergreen broad-leaved forest
sectionCool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Cool temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Evergreen broad-leaved forest
sectionWarm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section
Warm temperate deciduous
broad-leaved forest section

A Preliminary Survey on Larval Diversity in Mushroom Fruit Bodies

81

Fig. 1. Representative fruit bodies of mushrooms from which maggots were isolated. A. Amanita hemibapha(Agaricales), TNS-F-42118. B. Calvatia craniiformis (Agaricales), TNS-F-41393. C. Tylopilus sp. (Boletales),TNS-F-41193. D. Russula sp. (Russulales), TNS-F-42032. Bars＝2 cm.

trophic not speciﬁc to mushrooms, and therefore
not necessarily mycophagous. Most of the collected fruit bodies had no apparent insect damage
judging from the outside appearance (Fig. 1).
Typically, maggots were found only when fruit
bodies were cut in half prior to drying. At leastten maggots from each fruit body were successfully collected without damaging the outer layerof their bodies, and stored in DMSO buffer for
further molecular studies.
DNA Preparation
The number of maggots sequenced variedamong samples (Table 1). Only one maggot was
extracted for DNA each from Calvatia craniiformis(TNS-F-41393; Fig. 1B), Russula sp. (TNS-F41397), and Strobilomyces sp. (TNS-F-41945).
On the other hand, three or four maggots wereextracted from Amanita hemibapha (TNS-F42118; Fig. 1A), Amanita sp. (TNS-F-41384),

Boletus sp. (TNS-F-41353), Russula sp. (TNS-F42032; Fig. 1D), Tylopilus sp. (TNS-F-41193;
Fig. 1C), and Xerocomus sp. (TNS-F-41945).
DNA was extracted from a total of 24 maggots,
and PCR and sequencing of the COI were successful for all samples. The BLAST searches of
the COI sequences only resulted in the identiﬁcation at the ordinal level (Diptera) and could notlead to the species names. All mushroom samples
were successfully extracted and sequenced, andthe identiﬁcation at the generic level was conﬁrmed by the BLAST searches.
Phylogenetic Analyses
The initial alignment resulted in 680 bp long,and the beginning and end regions were deleted
from the analyses. This resulted in the dataset of
455 characters without gaps or ambiguouslyaligned regions. Out of 455 characters, 317 wereparsimony uninformative and the remaining 138

82

Kentaro Hosaka and Kunihiko Unocharacters were used for the MP analyses. The
MP analyses produced four equally parsimonious
trees with a tree length of 271 steps, CI of
0.6052, RI of 0.8333, and RC of 0.5043. Because
the NJ analyses produced the trees with almost
identical topology, only one of the most parsimonious trees is shown (Fig. 2).
A total of nine groups, all of which were supported by 100% bootstrap values, were detected(Fig. 2). Most groups were represented by the
maggots from a single fruit body, but the group 2
and 3 were represented by the maggots from different species of mushrooms. The group 2 represents the maggots from Strobilomyces sp. (TNSF-41945) and Xerocomus sp. (TNS-F-41950),
both of which belong to the order Boletales. Furthermore, they were both collected on the sameday (August 18, 2011), but were from different
sections in the garden (Table 1). The group 3 represents the maggots from Russula spp., but one

was collected on August 12, 2011, and the otherwas collected on September 2, 2011. Those two
species of Russula were both from the same section of the garden (Warm temperate deciduous
broad-leaved forest section). Most species ofmushrooms hosted only one group of maggots,
but Russula sp. (TNS-F-42032; Fig. 1D) hosted
the maggots of groups 3 and 5, and Xerocomus
sp. (TNS-F-41950) hosted the maggots of groups
2 and 8.
Discussion
The primer combinations of the COI successfully ampliﬁed and sequenced the DNA from all
24 maggots we investigated. Those primers(LCO1490 and HCO2198) were originallydesigned for amplifying a wide variety of animals(Folmer et al., 1994; Hebert et al., 2003), but havebeen shown to be inappropriate for certain groups

Fig. 2. An unrooted network of the COI phylogeny of maggots based on parsimony analyses. One of four equallyparsimonious trees is shown with bootstrap values on branches. The numbers in the circles indicate the individual maggot corresponding to Table 1.

A Preliminary Survey on Larval Diversity in Mushroom Fruit Bodies(Lohman et al., 2009; Park et al., 2010). Our study
showed that those primers can be used for a wide
variety of mycophagous Diptera, but more studies
are needed to conclude if they can be applied toother mycophagous insects.
All maggot sequences were tentatively identiﬁed as “Diptera” by the BLAST searches, without lower taxonomic units identiﬁed. In addition,phylogenetic analysis using the COI sequencesof selected Drosophila subg. Drosophila (http://
kyotoﬂy.kit.jp/ehime/genome2007/COI_Drosophila.
html) showed that maggots sampled for thisstudy did not form a cluster with any Drosophila
species (data not shown). This indicates thatspecies of Drosophila, which includes several
mycophagous species, were not collected during
this study. Other genera with known mycophagous
species, such as Megaselia, Cordyla, Allodia,
Rondaniella, and Mycetophila should also be
included in phylogenetic analyses for better
understanding of maggot diversity.
Although we do not have species names, we
tentatively considered each group in Fig. 2(Group 1 to 9) as species because each groupwas supported by 100% of bootstrap values, andthey were separated by ten or more substitutions.
This reﬂects the current situation of DNA barcoding in insect taxonomy, clearly indicating theneed of sequencing more well-identiﬁed specimens. The problem of absence of reliable link
between species names and DNA sequence datais also critical in mushroom identiﬁcation (Brocket al., 2008; Nagy et al., 2011). The accumulation of reliable DNA data in the barcode databasetakes more time, but we have sequenced the barcoding regions both from insects (as maggots)
and mushroom hosts (as mature fruit bodies), andwe keep the voucher specimens for all of them.
These data and specimens can therefore be used
for future studies.
We focused on collecting the larval stage of
ﬂies because they clearly use mushroom fruit
bodies as food source. The adult ﬂies and beetles
are also known to feed on mushroom spores andhyphae (Tuno, 1998, 1999; Takahashi et al.,2005; Kadowaki, 2010), but it is often difﬁcult to83

distinguish truly mycophagous insects from
opportunistic visitors. Furthermore, fresh fruit
bodies without apparent insect damage (Fig. 1)
are often inhabited by numerous maggots. This
indicates that a variety of insects may exist inmushroom fruit bodies from earlier developmental stages, as shown by Yamashita and Hijii(2007).
There are extensive studies on mycophagous
insect fauna based on ﬁeld observation and/or
rearing experiments from ﬁeld-collected mushrooms (Jaenike, 1978; Grimaldi, 1985; Toda andKimura, 1997; Yamashita and Hijii, 2007; Kadowaki, 2010), both of which depend on the identiﬁcation of the adult ﬂies. Because competitive
coexistence, instead of competitive exclusion,
was proposed as a mechanism of mycophagous
ﬂies in mushroom fruit bodies (Grimaldi, 1985),
previous studies may already have revealed avast majority of insect diversity in mushroom
fruit bodies. However, we argue that true diversity of mycophagous insects cannot be understood by collecting adults alone. This is somewhat analogous to the study of mushroom
diversity where true mushroom diversity cannot
be understood by collecting fruit bodies alone.
Our sampling for this study is clearly limited,
and more data are necessary both for mushrooms
and insects. Based on such limited sampling, however, the following conclusions can be drawn:(1) Different species of mushrooms can beinhabited by the same Diptera species. This
clearly indicates that the interactions between
mycophagous ﬂies and mushrooms are not species speciﬁc. At this stage, however, we cannot
conclude if the maggot groups found only from asingle mushroom species (Groups 1, 4–9; Fig. 2)
have speciﬁc interactions.
(2) Different species of larvae can be present in
the same mushroom fruit bodies. This study
showed only two species of mushrooms hosted
multiple species of ﬂies. However, we have notsequenced all maggots we sampled. We believe
more mushrooms actually host multiple speciesof ﬂies in a single fruit body, which can only beanswered with increasing sampling and sequenc-

84

Kentaro Hosaka and Kunihiko Uno

ing effort of maggots and mushrooms.
(3) Same species of larvae can be present in the
different mushroom fruit bodies at different
times. It is important information to understandthe biology of ﬂies as well as mushrooms. There
was a three-week gap between the ﬁrst (voucher
number 580-1; Fig. 2) and last (voucher number
675-3, 4) collections of maggots of the same species (Group 3; Fig. 2). This may indicate maggots sampled at different times represent different developmental stages (the ones from
September were more mature), or the ﬂy species
lay eggs over a three-week span.
Some of basic questions remain to beanswered: What are the Diptera species found in
this study? What is the true diversity of larvae inmushrooms? Are there any host preferences? Are
fungal spores dispersed by larvae? Are fungal
spores stimulated to germinate by larval consumption? These questions can only be answeredby the combination of increasing sampling effort
of both adult and larval stages, developing experimental designs, including rearing experiments,
and a good taxonomy based on morphology andDNA barcoding data.
Acknowledgments
The authors thank Yoshiaki Muramatsu,
Kazuo Nishibori, Megumi Ohtsuka, and Kazuya
Tsukagoshi for assisting our curatorial work. I
also thank Kyung-ok Nam for assisting ourmolecular work. Thanks also go to Dr. Nobuko
Tuno for stimulating discussion during the Ecological Society of Japan meeting in 2012. This
research was ﬁnanced in part by a research grant(2009–2011) of the Institute of Fermentation,
Osaka, Japan, JSPS grants-in-aid for young scientists (B), and the Center for Molecular Biodiversity Research at the National Museum of
Nature and Science to KH.
References
Atkinson, W. D. and Shorrocks, B. 1981. Competition ona divided and ephemeral resource: a simulation model.
Journal of Animal Ecology 50: 461–471.

Brock, P. M., Döring, H. and Bidartondo, M. I. 2008.
How to know unknown fungi: the role of a herbarium.
New Phytologist 181: 719–724.
Bruns, T. D., Fogel, R., White, T. J. and Palmer, J. D.
1989. Accelerated evolution of a false-trufﬂe from amushroom ancestor. Nature 339: 140–142.
Castellano, M. A., Trappe, J. M., Maser, Z. and Maser, C.
1989. Key to spores of the genera of hypogeous fungiof north temperate forests with special reference to animal mycophagy. Mad River Press, Eureka, California.
Claridge, A. W. and May, T. W. 1994. Mycophagy among
Australian mammals. Australian Journal of Ecology 19:
251–275.
Folmer, O., Black, M., Hoeh, W., Lutz, R. and Vrijenhoek, R. 1994. DNA primers for ampliﬁcation of mitochondrial cytochrome c oxidase subunit I from diverse
metazoan invertebrates. Molecular Marine Biology andBiotechnology 3: 294–299.
Fulton, T. W. 1889. The dispersal of the spores of fungiby the agency of insects, with special reference to thePhalloidei. Annals of Botany 3: 207–238.
Grimaldi, D. 1985. Niche separation and competitive
coexistence in mycophagous Drosophila (Diptera:
Drosophilidae). Proceedings of the EntomologicalSociety of Washington 87: 498–511.
Hall, T. A. 1999. BioEdit: a user-friendly biologicalsequence alignment editor and analysis program for
Windows 95/98/NT. Nucleic Acids Symposium Series
41: 95–98.
Hebert, P. D. N., Cywinska, A., Ball, S. L. and deWaard,
J. R. 2003. Biological identiﬁcations through DNA barcodes. Proceedings of the Royal Society of London, B
270: 313–321.
Hosaka, K. 2009. Phylogeography of the genus Pisolithus
revisited with some additional taxa from New Caledonia and Japan. Bulletin of the National Museum of
Nature and Science, Series B 35: 151–167.
Hosaka, K. and Castellano, M. A. 2008. Molecular phylogenetics of Geastrales with special emphasis on theposition of Sclerogaster. Bulletin of the National
Museum of Nature and Science, Series B 34: 161–173.
Hosaka, K., Bates, S. T., Beever, R. E., Castellano, M. A.,
Colgan, W., Dominguez, L. S., Geml, J., Giachini, A. J.,
Kenney, S. R., Nouhra, E. R., Simpson, N. B., Spatafora,
J. W. and Trappe, J. M. 2006. Molecular phylogeneticsof the gomphoid-phalloid fungi with an establishmentof the new subclass Phallomycetidae and two neworders. Mycologia 98: 949–959.
Jeanike, J. 1978. Resource predictability and nichebreadth in the Drosophila quinaria species group. Evolution 32: 676–678.
Kadowaki, K. 2010. Species coexistence patterns in a
mycophagous insect community inhabiting the wooddecaying bracket fungus Cryptoporus volvatus(Polyporaceae: Basidiomycota). European Journal ofA Preliminary Survey on Larval Diversity in Mushroom Fruit Bodies
Entomology 107: 89–99.
Kimura, M. T., Toda, M. J., Beppu, K. and Watabe, H.
1977. Breeding sites of drosophilid ﬂies in and near
Sapporo, northern Japan, with supplementary notes onadult feeding habits. Kontyu 45: 571–582.
Kirk, P. M., Cannon, P. F., Minter, D. W. and Stalpers, J.
A. 2008. Dictionary of the Fungi, 10th edition. CABI
Publishing, Wallingford.
Lehmkuhl, J. F., Gould, L. E., Cazares, E. and Hosford,
D. R. 2004. Trufﬂe abundance and mycophagy bynorthern ﬂying squirrels in eastern Washington forests.
Forest Ecology and Management 200: 49–65.
Lohman, D. J., Prawiradilaga, D. M. and Meier, R. 2009.
Improved COI barcoding primers for Southeast Asianperching birds (Aves: Passeriformes). Molecular Ecology Resources 9: 37–40.
Malajczuk, N., Trappe, J. M. and Molina, R. 1987. Interrelationships among some ectomycorrhizal trees, hypogeous fungi and small mammals: Western Australianand northwestern American parallels. Australian Journal of Ecology 12: 53–55.
Martin, M. M. 1979. Biochemical implications of insect
mycophagy. Biological Reviews 54: 1–21.
Meyer, M. D., North, M. P. and Kelt, D. A. 2005. Fungi inthe diets of northern ﬂying squirrels and lodgepole
chipmunks in the Sierra Nevada. Canadian Journal of
Zoology 83: 1581–1589.
Nagy, L. G., Petkovits, T., Kovács, G. M., Voigt, K.,
Vágvölgyi, C. and Papp, T. 2011. Where is the unseen
fungal diversity hidden? A study of Mortierella reveals
a large contribution of reference collections to the identiﬁcation of fungal environmental sequences. New
Phytologist 192: 592–596.
Pacioni, G., Bologna, M. A. and Laurenzi, M. 1991.
Insect attraction by Tuber: a chemical explanation.
Mycological Research 95: 1359–1363.
Park, D.-S., Suh, S.-J., Oh, H.-W. and Hebert, P. D. N.
2010. Recovery of the mitochondrial COI barcode
region in diverse Hexapoda through tRNA-based primers. BMC Genomics 11: 423–429.
Sawahata, T., Soma, K. and Ohmasa, M. 2002. The seasonal change in abundance of Hypogastrura denisana
Yoshii on agaric mushrooms in relation to its life cycle.
Edaphologia 69: 35–45.
Sawahata, T., Shimano, S. and Suzuki, M. 2008. Tricholoma matsutake 1-Octen-3-ol and methyl cinnamate
repel mycophagous Proisotoma minuta (Collembola:
Insecta). Mycorrhiza 18: 111–114.
Seutin, G., White, B. N. and Boag, P. T. 1991. Preservation of avian blood and tissue samples for DNA analyses. Canadian Journal of Zoology 69: 82–90.
Shorrocks, B. and Charlesworth, P. 1982. A ﬁeld survey

85of the association between the stinkhorn Phallus impudicus Pers. and the British fungal-breeding Drosophila.
Biological Journal of the Linnean Society 17: 307–318.
Smith, K. G. V. 1956. On the Diptera associated with thestinkhorn (Phallus impudicus Pers.) with notes on otherinsects and invertebrates found on this fungus. Proceedings of the Royal Entomological Society 31:
49–55.
Swofford, D. L. 2002. PAUP*: phylogenetic analysis
using parsimony (*and other methods). Version 4.0.
Sunderland, Massachusetts: Sinauer Associates.
Takahashi, K. H., Tuno, N. and Kagaya, T. 2005. Abundance of mycophagous arthropods present on different
species of fungi in relation to resources abundance at
different spatial scales. European Journal of Entomology 102: 39–46.
Thiers, H. D. 1984. The secotioid syndrome. Mycologia
76: 1–8.
Toda, M. J. and Kimura, M. T. 1997. Life-history traits
related to host selection in mycophagous drosophilids.
Journal of Animal Ecology 66: 154–166.
Toda, M. J., Kimura, M. T. and Tuno, N. 1999. Coexistence mechanisms of mycophagous drosophilids on
multispecies fungal hosts: aggregation and resource
partitioning. Journal of Animal Ecology 68: 794–803.
Tuno, N. 1998. Spore dispersal of Dictyophora fungi(Phallaceae) by ﬂies. Ecological Research 13: 7–15.
Tuno, N. 1999. Insect feeding on spores of a bracket fungus,
Elfvingia applanata (Pers.) Karst. (Ganodermataceae,
Aphyllophorales). Ecological Research 14: 97–103.
Tuno, N., Takahashi, K. H., Yamashita, H., Osawa, N. andTanaka, C. 2007. Tolerance of Drosophila ﬂies to ibotenic acid poisons in mushrooms. Journal of Chemical
Ecology 33: 311–317.
White, T. J., Bruns, T., Lee, S. and Taylor, J. W. 1990.
Ampliﬁcation and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: Innis, M. A.,
Gelfand, D. H., Sninsky, J. J. and White, T. J. (eds.),
PCR protocols, pp. 315–322, Academic Press, NewYork.
Worthen, W. B. 1989. Effects of resource density on
mycophagous ﬂy dispersal and community structure.
OIKOS 54: 145–153.
Worthen, W. B., Carswell, M. L. and Kelly, K. A. 1996.
Nested subset structure of larval mycophagous ﬂyassemblages: nestedness in a non-island system. Oecologia 107: 257–264.
Yamashita, S. and Hijii, N. 2007. The role of fungal taxa
and developmental stage of mushrooms in determining
the composition of the mycophagous insect community
in a Japanese forest. European Journal of Entomology
104: 225–233.