Content uploaded by Danushka S. Tennakoon
Author content
All content in this area was uploaded by Danushka S. Tennakoon on Aug 17, 2022
Content may be subject to copyright.
Vol.:(0123456789)
1 3
Fungal Diversity (2022) 115:73–103
https://doi.org/10.1007/s13225-022-00508-x
Fungal community succession ondecomposing leaf litter acrossfive
phylogenetically related tree species inasubtropical forest
DanushkaS.Tennakoon1,2,3,4· Chang‑HsinKuo5· WitoonPurahong6· EleniGentekaki1,2· ChayakornPumas4·
ItthayakornPromputtha4,8· KevinD.Hyde1,2,3,4,7,8
Received: 20 March 2022 / Accepted: 1 July 2022 / Published online: 1 August 2022
© The Author(s) under exclusive licence to Mushroom Research Foundation 2022
Abstract
Fungi are an essential component of the ecosystem. They play an integral role in the decomposition of leaf litter and return
nutrients to the ecosystem through nutrient cycling. They are considered as the “key players” in leaf litter decomposition,
because of their ability to produce a wide range of extracellular enzymes. Time-related changes of fungal communities during
leaf litter decomposition have been relatively well-investigated. However, it has not been established how the tree species,
tree phylogeny, and leaf litter chemistry influence fungal communities during decomposition. Using direct observations
and a culturing approach, this study compiles fungi found in freshly collected leaf litter from five phylogenetically related,
native tree species in Taiwan: Celtis formosana (CF), Ficus ampelas (FA), Ficus septica (FS), Macaranga tanarius (MT),
and Morus australis (MA). We investigated (i) the effects of tree species (including tree phylogeny) and leaf litter chemistry
on fungal community succession, and (ii) specific patterns of fungal succession (including diversity and taxonomic com-
munity assembly) on decomposing leaf litter across the selected tree species. We hypothesized that host species and leaf
litter chemistry significantly affect fungal community succession. A total of 1325 leaves (CF: 275, FA: 275, FS: 275, MT:
275 and MA: 225) were collected and 236 fungal taxa were recorded (CF: 48, FA: 46, FS: 64, MT: 42 and MA: 36). Tree
species relationships had variable associations on the fungal communities, as even closely related tree species had strongly
differing communities during decomposition. A high number of species were unique to a single tree species and may indicate
‘host-specificity’ to a particular leaf litter. The overlap of microfungal species in pair wise comparisons of tree species was
low (7–16%), and only 1–2% of microfungal species were observed in leaves of all tree species. The percentage of occur-
rences of fungal communities using Hierarchical Cluster Analyses (HCA) showed that there were at least four succession
stages in each tree species during decomposition. Fungal diversity increased at the beginning of each tree species leaf decay,
reached peaks, and declined at the final stages. Overall, our findings demonstrate that tree species and leaf litter chemistry
are important variables in determining fungal diversity and community composition in leaf litter. Referring to the establish-
ment of fungal discoveries from this experimental design, two new families, two new genera, 40 new species and 56 new
host records were reported. This study provides a host-fungus database for future studies on these hosts and increases the
knowledge of fungal diversity in leaf litter.
Keywords Fungal diversity· Ecology· Leaf litter decomposition· Percentage of occurrences· Taxonomy
Introduction
Leaf litter is a vital component of forest ecosystems and
constitutes a major source of soil organic matter (Berg and
Laskowski 2006; Purahong etal. 2016; Osono 2017; Bani
etal. 2018; Tennakoon etal. 2021a). Fungal communities
are regarded as "key players" in leaf litter decomposition
as they produce a wide spectrum of extracellular enzymes
(Pointing etal. 2005; Promputtha etal. 2010; Purahong
etal. 2016; Tennakoon etal. 2021a). These enzymes aid
Handling Editor: Chayanard Phukhamsakda
* Kevin D. Hyde
kdhyde3@gmail.com
Extended author information available on the last page of the article
74 Fungal Diversity (2022) 115:73–103
1 3
in the breakdown of the layers of leaf litter, particularly
lignocellulose (Berg and McClaugherty 2003; Promputtha
etal. 2010; Qu etal. 2019; Osono 2020; Tennakoon etal.
2021a). Many studies have looked at the fungal communities
that emerge during the breakdown of leaf litter. These have
revealed that fungal populations tend to alter both numeri-
cally and qualitatively during decomposition (Koide etal.
2005; Tang etal. 2005; Duong etal. 2008; Shirouzu etal.
2009; Voříšková and Baldrian 2013; Promputtha etal. 2017).
The time-related change of fungal species during decom-
position has been termed as “fungal succession” (Dix and
Webster 1995; Fryar 2002; Promputtha etal. 2017; Tenna-
koon etal. 2021a). Fungal succession on decomposing leaf
litter has previously been investigated in temperate (Cooke
and Rayner 1984; Pascoal etal. 2005; Kara etal. 2014;
Voříšková and Baldrian 2013; Purahong etal. 2016) and
tropical environments (Promputtha etal. 2002, 2017; Paulus
etal. 2006; Duong etal. 2008; Osono etal. 2020).
Several factors have an effect on the fungal succession
or composition of fungal communities in leaf litter (Dix
and Webster 1995; Photita etal. 2001; Promputtha etal.
2002; Duong etal. 2008; Purahong etal. 2016; Saitta etal.
2018; Angst etal. 2019). In particular, both biotic and abi-
otic variables influence the makeup of fungal communities
(Purahong etal. 2016; Saitta etal. 2018; Angst etal. 2019).
Knowing the variables that affect leaf litter fungal commu-
nities is especially important for understanding how eco-
systems function and their susceptibility to environmental
disturbances like climate change, biodiversity loss, and bio-
logical invasions (Glassman etal. 2018; Dossa etal. 2021).
Biotic factors, such as heterogeneity of tree species (tree spe-
cies effect), colonization priority effects, interspecific com-
petition or relationships between fungal species and inter-
kingdom relationships (i.e. bacteria and fungi), play vital
roles during leaf litter decomposition (Lodge and Cantrell
1995; Promputtha etal. 2002, 2017; Paulus etal. 2003;
Duong etal. 2008; Saitta etal. 2018; Angst etal. 2019).
Furthermore, the chemical content and physical quality of
leaves vary according to tree species and may have a signifi-
cant impact on the makeup of fungal communities (Kubar-
tová etal. 2009; Purahong etal. 2016; Huang etal. 2017).
Another important biotic factor is the priority effect of
fungal species during fungal succession. Most succession
investigations have identified comparable patterns of fungal
presence across the course of decomposition, namely early,
intermediate, and late colonizers (or alternative names: Pio-
neer, Mature and Impoverished communities) (Garrett 1963;
Hudson 1968; Tsui etal. 2000; Promputtha etal. 2002, 2017;
Tang etal. 2005). The general view is that early coloniz-
ers are non-cellulolytic and rely on easily available sugars,
therefore they dominate the early stages of decomposi-
tion. Fungi that degrade cellulose and other hydrolysable
plant polymers are prominent in the second stage, whereas
lignin-degraders are dominant in the later stages (Garrett
1963; Hudson 1968; Kjøller and Struwe 2002). Early colo-
nists impact subsequent community dynamics through either
interspecific competition or facilitating the growth of other
species (Voříšková and Baldrian 2013; Lin etal. 2015; Veen
etal. 2019). This is referred to as priority effect and is an
important biotic factor relating to the colonization of fun-
gal species in leaf litter (Boddy and Hiscox 2017). Specifi-
cally, it has been hypothesized that the priority effect may
be responsible for certain fungal species switching from
an endophytic to a saprotrophic lifestyle (Promputtha etal.
2007, 2010; Purahong and Hyde 2011). Fungal species that
are associated with living leaves have also been identified in
leaf litter (Hyde etal. 2007; Promputtha etal. 2007, 2010;
Purahong and Hyde 2011; Persoh etal. 2018). Phylogenetic
analyses showing that fungi isolated from living leaves and
decaying litter may indeed belong to the same taxa have
further strengthened this hypothesis (Promputtha etal. 2007,
2010; de Silva etal. 2019). For example, Promputtha etal.
(2007) found that isolates of Colletotrichum gloeospori-
oides, Fusarium oxysporum, Nectria haematococca, Phyl-
losticta mangiferae which are common plant endophytes,
exhibit significant sequence similarity and phylogenetic
affinity to their saprotrophic counterparts in Magnolia liliif-
era. In addition, Lasiodiplodia pseudotheobromae has been
recorded as both as endophyte and saprobe in Magnolia can-
dolii (de Silva etal. 2019).
During the succession process, interactions between fun-
gal species, such as interspecific competition, are determin-
ing factors as to whether fungi are successful in colonizing
and for how long they retain that territory (Jones and Hyde
2002; Boddy and Hiscox 2017). Early colonizers may pos-
sess adaptations for substrate colonization, but later colo-
nizers might be better suited for interspecific competition
and thus invasion of existing communities (McNaughton
and Wolf 1973; Boddy and Hiscox 2017). Some taxa may
also contribute to the excretion of compounds that directly
affect the growth and development of other fungal species
(Shearer 1995; Duong etal. 2008). For instance, metabolites
such as glycol, chlorohydrins and bromohydrin can affect
the development of fungal communities (Paulus etal. 2003;
Duong etal. 2008). Several studies have noted an increase
in bacterial species (i.e. Actinobacteria, Bacteroidetes, and
Proteobacteria) during the early and later stages of leaf litter
decomposition, but the specific functions of bacterial-fungal
interactions throughout this process is unknown (Romaní
etal. 2006; Purahong etal. 2016; Zhao etal. 2020). In gen-
eral, bacterial and fungal communities may compete or
cooperate for the same substrates (Lynd etal. 2002; Johnston
etal. 2016; Purahong etal. 2016). Kirby (2005) revealed that
some bacterial species (i.e. Bradyzhorium, Burkholderia,
Streptomyces) contribute to the production of extra cellu-
lar degrading enzymes at the later stages of decomposition.
75Fungal Diversity (2022) 115:73–103
1 3
Purahong etal. (2016) suggested that nitrogen-fixing bac-
teria contribute to increased availability of nitrogen in leaf
litter thus supporting other decomposers (i.e. fungi). High
throughput studies have revealed that lignocellulose decom-
posers (i.e. Clitocybe spp., Mycena spp.) co-exist with nitro-
gen-fixing bacterial OTUs (i.e. Bradyrhizobium, Mesorhizo-
bium, Pseudomonas and Rhizobium) suggesting synergistic
interactions between the two communities (Purahong etal.
2016; Zhao etal. 2020).
Abiotic factors, such as leaf litter quality, temperature,
precipitation, seasonal fluctuations, soil properties, and vari-
ous plot-related parameters have a strong impact on leaf lit-
ter decomposition, because they change the circumstances
for decomposers to act and convert litter organic matter into
plant-friendly forms (Bothwell etal. 2014; Ge etal. 2017;
Osono 2017). The physical nature and biochemical compo-
sition of the leaves comprise the sole determinants of leaf
litter quality (Hou etal. 2005; Purahong etal. 2016; Osono
2017). Physical qualities of leaves, such as hardness, particle
size, and leaf surface features, can impact the accessibility of
litter to soil organisms, altering rates of colonization (Prom-
puttha etal, 2002; Duong etal. 2008; Berg 2014). Some of
the elements that determine the biochemical composition of
leaf litter and thus its quality include initial C or N content,
C/N ratio, soluble sugars, polyphenols, waxes, cellulose,
hemicellulose, and lignin content concentration. These ele-
ments contribute to nutrient mineralization and immobili-
zation dynamics, which may have an impact on microbial
decomposers (Tian etal. 1997; Cotrufo etal. 1998; Perez-
Harguindeguy etal. 2000; Hou etal. 2005; Purahong etal.
2014; Osono 2017; Tennakoon etal. 2021a).
Numerous informative studies have investigated the influ-
ence of biotic and abiotic factors on fungal communities
during leaf litter decay, however knowledge gaps remain.
Specifically, little is known about factors, such as tree spe-
cies effect, tree phylogeny and leaf litter chemistry. Some
studies have found positive relationships between tree
species heterogeneity and fungal diversity, but still more
experimental clarifications are needed (Saitta etal. 2018;
Angst etal. 2019). This study provides a compilation of
fungal communities during the decay of leaf litter from five
native host tree species. Celtis formosana (CF), Ficus ampe-
las (FA), F. septica (FS), Macaranga tanarius (MT) and
Morus australis (MA) were selected from relatively natural
forests in Taiwan and studied using direct observations cou-
pled with a culture-based approach. Specifically, we inves-
tigated (i) the relationship of tree species (including tree
phylogeny) and leaf litter chemistry on fungal community
succession and (ii) specific patterns of fungal succession
(including diversity and taxonomic community assembly)
on decomposing leaf litter across the five phylogenetically
related tree species in a subtropical forest. We hypothesized
that (i) tree species heterogeneity and leaf litter chemistry
are significantly related to fungal community succession,
(ii) tree species heterogeneity explains more of the observed
variation in fungal community composition rather than leaf
litter chemistry, and (iii) fungal community composition is
more similar in phylogenetically closely related tree species
rather than phylogenetically distant tree species.
Materials andmethods
Study site andhost species selection
The study site is located in the Alishan Mountain, Chi-
ayi, in the Southwestern part of Taiwan (23°27.582'N
120°36.285'E). The mean annual temperature is approxi-
mately 18–24°C and 2502mm precipitation according to
weather observation data from central weather bureau (Chi-
ayi, Taiwan). Alishan Mountain has a rich biodiversity and
most parts are covered by various types of subtropical and
temperate forests (hardwood, coniferous and grassland) (Wu
etal. 2004; Chou and Tang 2016). These climatic conditions
account for the luxuriant vegetation and great diversity of
species in this area (1050 endemic plant species, 5700 fungi
species and 19,000 animal species) (Chou and Tang 2016).
The experiment was conducted during the rainy season
(July–September 2018). Five phylogenetically related host
species were selected as follows (Fig.1): two host species
were congeners of Ficus; one belongs to the same family as
Ficus (i.e. Moraceae), but is a different genus (i.e. Morus
australis); one host species belongs to a different family (i.e.
Cannabaceae), but is in the same order; and one belongs to
a different order (Fig.1). Leaf litter samples were collected
from Ficus ampelas, F. septica, Morus australis, Celtis
formosana and Macaranga tanarius (Figs.1, 2). All host
species were growing in the same altitude at a considerable
distance from each other. Five replicates (five trees per each
host) were selected.
Litterbag design andsampling
Initially, 125 senescent leaves were collected from each
host species (Total number of leaves = 125 × 5 replicates × 5
hosts = 3125 leaves). Selected senescent leaves had just
fallen, and were yellow-green with a fresh abscission scar.
Of them, five leaves from each tree were randomly col-
lected and represented day 0 of the experiment (Initial col-
lection, day 0 = 5 × 5 replicates × 5 hosts = 125 leaves). The
rest 120 leaves put in nylon mesh bags (size = 25 × 34cm
with 2mm pores/ 10 leaves per bag) (Total number of
bags = (120 ÷ 10) × 5 replicates × 5 hosts = 300 bags). The
prepared nylon mesh bags were placed under each tree
species. Leaves were collected at 11 time points. At each
sampling time point, five leaves were randomly collected
76 Fungal Diversity (2022) 115:73–103
1 3
from the placed nylon mesh bags beneath each tree species
at days 3, 7, 14, 21, 28, 35, 42, 49, 56 and 63 (late decom-
position stage) (Number of collected leaves per day = 5 × 5
replicates × 5 hosts = 125 leaves). After 63days, leaves were
highly skeletonized comprising vascular tissue with attached
remnants of non-vascular tissue. Therefore, samples were
not collected after 63days. Collected samples were placed
in separate plastic bags in the forest and taken back to the
laboratory. Samples collected on day 0 were examined for
the presence of fungi on the day of collection. All other
samples were incubated in the laboratory in separate plastic
Fig. 1 Experimental design and selected host species
Fig. 2 a, b. Experimental sites
(Alishan Mountain, Taiwan). c.
Ficus ampelas. d. Ficus septica.
e. Morus australis. f. Celtis for-
mosana. g. Macaranga tanarius
77Fungal Diversity (2022) 115:73–103
1 3
boxes containing tissue paper moistened with sterile distilled
water for five days.
Morphological studies andisolation
The incubated leaves were observed periodically for up to
a week using the AXIOSKOP 2 PLUS compound micro-
scope for the presence of microfungi. Fruiting structures
were mounted in water, agitated gently and observed using
the microscope and photographed with an AXIOCAM
506 COLOR digital camera fitted to the microscope. For
ascomycetes and coelomycetes, free hand sections of the
fruiting body structures were mounted in water on slides
for microscopic studies and photomicrography. For hypho-
mycetes, fungal structures were picked and added to water
on a slide using a needle. Permanent slides were preserved
in lactoglycerol and sealed by applying nail-polish around
the margins of the cover slip. All measurements were made
with ZEN2 (blue edition) and the images used for figures
were processed with Adobe Photoshop CS3 Extended ver-
sion 10.0 software (Adobe Systems, USA). Single spore
isolation was carried out following the spore suspension
method described in Chomnunti etal. (2014). Germinated
spores were individually transferred to potato dextrose agar
(PDA) plates and grown at room temperature (20–26°C) in
the daylight.
Species identification, preservation, DNA extraction
andPCR amplification
The aim of the fungal observation was to identify the fungal
communities occurring on leaf litter at the time of collection
using culture based approches. The collected fungal species
were subjected to morphological and molecular analysis as
previously described by Tennakoon etal. (2019a, b, 2020,
2021b). DNA was extracted directly from fruiting bodies
using the DNA extraction kit E.Z.N.A. ® Forensic DNA kit
(Omega Bio-Tek) and from mycelium using the E.Z.N.A
Fungal DNA Mini Kit (Omega Bio-Tek) following the
manufacturer’s instructions. The DNA extracts were kept at
4°C until DNA amplification and maintained at -20°C for
long-term storage. Polymerase chain reactions (PCR) were
performed as described in Tennakoon etal. (2021b). Her-
barium specimens were prepared by drying a section of the
leaf on which fungal colonies occurred with silica gel. Type
specimens of new taxa were deposited at the Mae Fah Luang
University herbarium (MFLU) and National Chiayi Univer-
sity Herbarium (NCYU). Living cultures were deposited at
Mae Fah Luang University Culture Collection (MFLUCC)
and National Chiayi University Culture Collection (NCY-
UCC). The taxonomy-related results have been published by
Tennakoon etal. (2019a, b, 2020, 2021b) and are used for
comparison here. Leaf litter chemistry (total N, C, lignin,
C/N ratio and lignin/N ratio) was analyzed in the Central
Laboratory, Faculty of Agriculture, Chiang Mai University,
Thailand. Total C and N content were determined by flash
combustion of micro-samples (5mg) by a CN-elemental
analyzer (Flash EA2000 Thermo). Lignin contents were
analyzed using the method described by Gessner (2005).
C/N and lignin/N ratios for each litter type were calculated.
Statistical analysis
The results of this study are presented in terms of the per-
centage occurrence (POC) of fungi. Fungal taxa with an
overall occurrence equal to or higher than 10% are regarded
as common species.
Fungal species diversity was measured using the Shan-
non diversity index (Shannon and Weaver 1949) and species
richness. Shannon diversity index was calculated by the fol-
lowing formula.
where Pi is the proportion of ith species (number of leaves
which ith fungal species collected in a day / total number
of leaves which fungal species collected in a day). Species
richness was the number of fungal species in each host spe-
cies leaf. Fungal diversity patterns were examined using R
Project for Statistical Computing version 4.0.2 supported
by CRAN (R Core Team, 2017). Diversity succession was
grouped based on the average percentage occurrences of
fungal communities using Hierarchical Cluster Analyses
(HCA) and carried out with h cluster function of R using
Average distance. Venn diagram was generated by the Inter-
actiVenn web-based tool (Heberle etal. 2015). To visualize
fungal community succession and community composition
in different tree species, we performed non-metric multidi-
mensional scaling (NMDS) and cluster analysis based on
the relative abundance data and Bray–Curtis distance meas-
ure using PAST software (Hammer etal. 2001). Degrees
of separation of fungal community composition (R) within
selected tree species were analysed using analysis of similar-
ity (ANOSIM) in PAST software based on the relative abun-
dance data and Bray–Curtis distance measure (R = 0–0.24,
no separation to barely separated; R⩾0.25–0.75, separation
with different degrees of overlap; R > 0.75–1, well separated
to complete separation; significant P-values (P < 0.05) are
based on 9999 permutations and Bonferroni corrections
were applied in all cases. To analyse the effect of tree
species, tree family, phylogeny, leaf litter chemistry, and
Percentage of occurrence
=Number of leaves which fungus was detected
Total number of leaf samples examined
×
100
Shannon index
(H)=
∑
Piln P
i
78 Fungal Diversity (2022) 115:73–103
1 3
time, NMDS was performed based on abundance data and
Bray–Curtis distance measure using the function metaMDS
in the “vegan package” with default settings (Oksanen etal.
2013). Goodness-of-fit statistics (r2) were calculated for dif-
ferent factors fit to the NMDS coordination with P values
based on 999 permutations (Oksanen etal. 2013). Variance
partitioning analysis was performed using the varpart func-
tion in the vegan package in R to take into account the influ-
ence of tree species heterogeneity, leaf litter chemistry, and
time on overall fungal community composition (Oksanen
etal. 2013). The graph of overlap between fungal commu-
nities occurring on pairs of tree species was made using
Microsoft Excel (2010).
Results
Fungal species diversity andoccurrence
A total of 1325 leaves (CF: 275, FA: 275, FS: 275, MT: 275
and MA: 225) were collected and 236 fungal taxa recorded
(CF: 48, FA: 46, FS: 64, MT: 42 and MA: 36). One hun-
dred and fifteen taxa (CF: 27, FA: 27, FS: 28, MT: 19 and
MA: 14) were isolated into culture and identified to species
level, using both morphology and phylogeny (Tennakoon
etal. 2019a, b, 2020, 2021b). The percentage occurrences
of fungal species and the Shannon diversity index (H) for
each host species are presented in Tables1–6. The time
for fungal communities to reach the peak of species diver-
sity was 28days for Ficus ampelas (H = 2.89), F. septica
(H = 3.33), Macaranga tanarius (H = 3.17) and Morus aus-
tralis (H = 2.82), but 35days for Celtis formosana (H = 3.27)
(Table1). For each host species, the Shannon diversity index
ranged from 0.5 to 2.01 at the beginning of decomposition,
but the index steadily increased and reached the peak of
about 2.8–3.3 at 28–35days (Table1). After that, fungal
diversity and the total number of fungal species started to
decline. The average diversity of fungi on each host species
in the experimental period ranged between 1.51 and 2.61,
viz. Celtis formosana (H = 2.38), Ficus ampelas (H = 2.27),
F. septica (H = 2.61), Macaranga tanarius (H = 2.28) and
Morus australis (H = 1.52). Species richness also increased
from the start of decomposition (1–10 taxa), reaching a max-
imum at middle stages (25–35 taxa) and steadily declined
in the final stages (5–15 taxa) (Fig.3). All identified species
belonged to Ascomycota. No Basidiomycota species were
observed during the decomposition process in this experi-
ment. The NMDS analysis indicated that the fungal com-
munity composition was significantly different among the
selected tree species (Fig.4).
Fungal community composition inFicus ampelas
A total of 275 Ficus ampelas leaves were examined dur-
ing the decay process and the percentage occurrences were
calculated (Table2). On day 63, leaves were highly skel-
etonized comprising vascular tissues with attached rem-
nants of non-vascular tissues. Fourty-six fungal taxa were
recorded and 27 species were successfully isolated into cul-
tures and identified to species level (Tennakoon etal. 2019a,
b, 2020, 2021b). This included a new genus (Longihyalos-
pora), nine new species (Acrocalymma ampeli, Cercophora
fici, Colletotrichum fici, Longihyalospora ampeli, Micro-
peltis fici, M. ficinae, Neoanthostomella fici, Neofusicoccum
moracearum and Phaeosphaeria ampeli) (Tennakoon etal.
2019a,b, 2020, 2021b) and eleven new host records (Beltra-
nia rhombica, Ceramothyrium longivolcaniforme, Coniella
quercicola, Diaporthe limonicola, D. pseudophoenicicola,
Discosia querci, Lasiodiplodia thailandica, Pseudopestalo-
tiopsis camelliae-sinensis, Torula fici, Wiesneriomyces lau-
rinus and Yunnanomyces pandanicola) (Tennakoon etal.
2019b, 2020, 2021b). An additional 21 species were identi-
fied to genus level, and belonged to Aspergillus, Backusella,
Table 1 Shannon Diversity Index (H) in each host species leaves dur-
ing succession process
Days Shannon Diversity Index (H)
CF FA FS MT MA
0 0.53 2.11 1.56 1.55 1.27
3 1.63 2.41 2.11 2.05 1.55
7 1.99 2.44 2.68 2.23 1.37
14 2.55 2.33 2.77 2.43 1.88
21 2.81 2.39 3.03 2.88 2.81
28 3.17 2.89 3.33 3.17 2.82
35 3.27 2.36 3.32 2.72 2.74
42 3.09 2.09 2.89 2.48 2.23
49 2.53 2.18 2.37 2.05 0.00
56 2.63 2.25 2.30 1.92 0.00
63 2.03 1.50 2.37 1.66 0.00
Average 2.38 2.27 2.61 2.28 1.52
Fig. 3 Fungal species richness (mean ± SE) variation with decompo-
sition process of selected host species leaves
79Fungal Diversity (2022) 115:73–103
1 3
Fig. 4 Non-metric multidimensional scaling (NMDS) analysis of fungal community composition in selected tree species
80 Fungal Diversity (2022) 115:73–103
1 3
Cylindrocladium, Fusarium, Mortierella, Mycosphaerella,
Penicillium, Periconia, Phoma, Phyllosticta, Rhizopus,
Volutella and Zygosporium (Table2). Five taxa were uniden-
tified Ascomycete species (due to lack of morphology data).
Table 2 Percentage occurrence
of fungal species which
occurred on Ficus ampelas
leaves during succession
process
Species/days 0 3 7 14 21 28 35 42 49 56 63
Acrocalymma ampeli 0 0 72 92 80 48 80 24 8 0 0
Aspergillus sp. 1 0 0 0 0 8 20 4 4 4 0 0
Aspergillus sp. 2 0 12 12 0 4 0 0 0 0 0 4
Aspergillus sp. 3 0 0 0 0 0 0 0 4 12 16 0
Asteridiella sp. 20 4 0 0 0 0 0 0 0 0 0
Backusella sp. 0 0 0 0 0 0 0 0 8 20 8
Beltrania rhombica 0 0 0 56 48 76 20 4 0 0 0
Ceramothyrium longivolcaniforme 888028120000000
Cercophora fici 0 0 0 0 0 0 0 0 24 16 28
Colletotrichum fici 20402080000000
Coniella quercicola 0 0 0 16 8 20 4 0 0 0 0
Cylindrocladium sp. 1 0 12 20 4 0 0 0 0 0 0 0
Cylindrocladium sp. 2 0 0 0 20 0 0 0 0 0 0 0
Diaporthe limonicola 100 96 96 60 20 20 24 4 20 0 0
Diaporthe pseudophoenicicola 0 0 0 0 4 52 68 4 0 0 0
Discosia querci 204420204840000
Fusarium sp. 3 0 12 8 0 4 0 0 0 0 4 32
Fusarium sp. 4 0 0 0 0 0 16 12 20 0 20 0
Lasiodiplodia thailandica 0 00003280000
Longihyalospora ampeli 100 96 76 20 36 4 0 0 0 0 0
Micropeltis fici 100 100 92 60 40 8 0 0 0 0 0
Micropeltis ficinae 20681200000000
Mortierella sp. 0 0 0 0 0 0 0 8 20 4 32
Mycosphaerella sp. 0 00004040000
Neoanthostomella fici 20 20 96 92 88 60 0 20 20 0 0
Neofusicoccum moracearum 0 0 0 4 20 64 60 20 0 0 0
Penicillium sp. 1 0 20 8 0 0 0 0 0 0 36 20
Penicillium sp. 2 0 0 0 0 0 0 0 0 8 16 4
Periconia sp. 1 0 0 0 0 0 28 12 0 0 0 0
Phaeosphaeria ampeli 04024120400000
Phoma sp. 1 0 0 0 0 0 32 8 0 0 0 0
Phoma sp. 2 0 0 0 0 16 20 8 0 0 0 0
Phyllosticta sp. 24 4000400000
Pseudopestalotiopsis camelliae-sinensis 0 0 0 0 20 64 40 20 20 20 0
Rhizopus stolonifer 2412000000000
Torula fici 0 20 40 0 4 56 20 4 20 0 0
Unidentified Ascomycete 1 0 0 0 0 0 20 4 0 0 0 0
Unidentified Ascomycete 2 0 0 0 4 16 4 0 0 0 0 0
Unidentified Ascomycete 3 0 0 0 0 0 4 0 0 0 20 4
Unidentified Ascomycete 4 0 0 0 0 20 4 0 0 0 0 0
Unidentified Ascomycete 5 0 0 0 0 8 20 8 0 0 0 0
Volutella sp.1 0 0 0 0 0 16 16 0 0 0 0
Wiesneriomyces laurinus 0 0 0 0 4 40 20 20 20 0 0
Yunnanomyces pandanicola 0 0 20 32 36 8 0 0 0 20 0
Zygosporium sp. 1 0 0 20 0 0 20 8 0 0 20 0
Zygosporium sp. 2 0 0 0 20 8 0 0 0 0 0 0
81Fungal Diversity (2022) 115:73–103
1 3
The most dominant species (POC > 10%) on Ficus ampe-
las leaves were Diaporthe limonicola (40%), followed by
Neoanthostomella fici (37.82%), Acrocalymma ampeli
(36.73%), Micropeltis fici (36.36%), Longihyalospora ampeli
(30.18%), Ceramothyrium longivolcaniforme (18.91%), Bel-
trania rhombica (18.55%), Pseudopestalotiopsis camelliae-
sinensis (16.73%), Neofusicoccum moracearum (15.27%),
Torula fici (14.91%), Diaporthe pseudophoenicicola
(11.64%), Discosia brasiliensis (10.91%) and Yunnanomyces
pandanicola (10.55%) (Fig.5).
The percentage occurrences of fungal communities using
Hierarchical Cluster Analyses (HCA), showed presence of
at least four distinct communities during succession: Stage
I (day 0–3), Stage II (day 7–21), Stage III (day 28–35) and
Stage IV (day 42–63) (Fig.5). In Stage I (0–7days) the
species number was low with low percentage occurrences.
The dominant species at this stage were Ceramothyrium
longivolcaniforme, Diaporthe limonicola, Longihyalos-
pora ampeli, Micropeltis fici and M. ficinae. The dominant
species of Stage II (7–21days) were Acrocalymma ampeli,
Beltrania rhombica, Diaporthe limonicola, Longihyalospora
ampeli, Micropeltis fici, Neoanthostomella fici and Yunnano-
myces pandanicola. The highest species diversity was noted
during Stage III (28–35days) and the dominant species
were Acrocalymma ampeli, Beltrania rhombica, Diaporthe
limonicola, D. pseudophoenicicola, Mycosphaerella sp.,
Neoanthostomella fici, Neofusicoccum moracearum, Pseu-
dopestalotiopsis camelliae-sinensis, Torula fici and Wiesne-
riomyces laurinus. In Stage IV (42–63days), the species
diversity and number of species declined. The community
was dominated by only a few species with relatively high
percentage occurrence. Dominant species were Cercophora
fici, Fusarium sp. 4, Mortierella sp., Neoanthostomella fici,
Penicillium sp. 1, Pseudopestalotiopsis camelliae-sinensis
and Wiesneriomyces laurinus.
The highest species diversity as indicated by Shannon
diversity index was at day 28 (H = 2.89) and lowest at day 63
(H = 1.50). The average diversity of fungi on Ficus ampelas
leaves during the experimental period was 2.27. The species
diversity changes during the experimental period are shown
in Table1.
Fungal community composition inFicus septica
A total of 275 Ficus septica leaves were examined during the
decay process and the percentage occurrences were calcu-
lated (Table3). On day 63, leaves were highly skeletonized
comprising vascular tissues with attached remnants of non-
vascular tissues. Sixty-four species were recorded and 28
species were successfully isolated into cultures and identi-
fied to species level. These include a new family (Cylin-
drihyalosporaceae), a new genus (Cylindrihyalospora), 16
new species (Bertiella fici, Colletotrichum fici-septicae,
Conidiocarpus fici-septicae, Coniella fici, Cylindrihyalo-
spora fici, Diaporthe fici-septicae, Diplodia fici-septicae,
Discosia ficeae, Leptodiscella sexualis, Microthyrium fici-
septicae, Muyocopron ficina, Mycoleptodiscus alishanense,
Neophyllachora fici, Ophioceras ficina, Pseudocercospora
fici-septicae and Stictis fici) and twelve new host records
(Arthrinium malaysianum, Conidiocarpus betle, Lasiodip-
lodia thailandica, L. theobromae, Neopestalotiopsis phangn-
gaensis, Parawiesneriomyces chiayiense, Periconia alis-
hanica, Pestalotiopsis portugallica, Robillarda roystoneae,
Sirastachys castanedae, Stachybotrys microspore and Torula
fici) (Tennakoon etal. 2021b). Another 27 species were
identified to genus level, which comprised Acremonium,
Appendiculella, Arthrinium, Aspergillus, Camarographium,
Colletotrichum, Cylindrocladium, Fusarium, Mucor, Peni-
cillium, Phoma, Phyllosticta, Pleurostoma, Pseudocercos-
pora, Rhizopus, Syncephalastrum and Torula (Table3). Two
species were identified as Botryosphaeriaceae species and
three species as coelomycete species. Four taxa were uni-
dentified Ascomycetes (due to lack of morphology data).
The dominant species (POC > 10%) on Ficus septica
leaves were Coniella fici (36.36%), followed by Colle-
totrichum fici-septicae (36%), Lasiodiplodia thailandica
(35.27%), Diaporthe fici-septicae (29.45%), Diplodia fici-
septicae (28.36%), Conidiocarpus fici-septicae (22.18%),
Leptodiscella sexualis (22.18%), Sirastachys castanedae
(21.82%), Conidiocarpus betle (21.81%), Neophyllachora
fici (20%), Ophioceras ficina (20%), Lasiodiplodia theo-
bromae (17.09%), Bertiella fici (16.36%), Pestalotiopsis
portugallica (16%), Parawiesneriomyces chiayiense (16%),
Arthrinium malaysianum (15.63%), Microthyrium fici-sep-
ticae (14.54%), Muyocopron ficina (14.18%), Stachybotrys
microspora (13.82%), Aspergillus sp. (13.45%), Cylindrihy-
alospora fici (12.73%) and Robillarda roystoneae (12.73%)
(Fig.6).
The percentage occurrences of fungal communities using
Hierarchical Cluster Analyses (HCA), showed presence of
at least four distinct communities during succession: Stage
I (day 0–7), Stage II (day 14–21), Stage III (day 35–42) and
Stage IV (day 49–63) (Fig.6). During Stage I (Day 0–7),
fungal communities comprised low species number with
low percentage occurrence. The dominant species at this
stage were Appendiculella sp., Colletotrichum fici-septicae,
Conidiocarpus betle, C. fici-septicae, Cylindrocladium sp.
1, Microthyrium fici-septicae, Mycoleptodiscus alishan-
ense, Neophyllachora fici, Phyllosticta sp., Rhizopus sp.
and Stachybotrys microspora (Fig.6). The dominant species
of Stage II (14–21days) were Colletotrichum fici-septicae,
Conidiocarpus betle, C. fici-septicae, Coniella fici, Cylindri-
hyalospora fici, Diaporthe fici-septicae, Diplodia fici-septi-
cae, Discosia ficeae, Leptodiscella sexualis, Lasiodiplodia
thailandica, Muyocopron ficina, Pseudocercospora fici-sep-
ticae and Torula fici (Fig.6). The highest species diversity
82 Fungal Diversity (2022) 115:73–103
1 3
Fig. 5 Heat-map of microbial community composition in Ficus ampelas leaves with cluster analysis. The color intensity shows the percentage of
occurrence (POC), referring to color key at the top
83Fungal Diversity (2022) 115:73–103
1 3
Table 3 Percentage occurrence
of fungal species which
occurred on Ficus septica leaves
during succession process
Species/days 0 3 7 14 21 28 35 42 49 56 63
Acremonium sp. 0 0 0 12 24 16 4 0 0 0 0
Appendiculella sp. 32 16 4 0 0 0 0 0 0 0 0
Arthrinium malaysianum 0 0 4 4 12 48 48 24 28 4 0
Arthrinium sp. 1 0 0 0 0 12 20 4 0 0 0 0
Aspergillus brasiliensis 0 4 0 0 16 28 32 40 4 4 20
Aspergillus sp. 1 0 0 0 0 8 16 20 0 0 0 20
Aspergillus sp. 2 0 0 0 0 0 0 0 4 8 20 4
Aspergillus sp. 3 0 0 0 0 0 4 8 16 16 4 0
Bertiella fici 0 0 0 0 0 20 20 60 40 20 20
Botryosphaeriaceae sp. 1 0 0 0 0 8 20 4 0 0 0 0
Botryosphaeriaceae sp. 2 0 0 4 0 0 8 16 0 0 0 0
Camarographium sp. 0 0 0 16 24 8 4 0 0 0 0
Coelomycete sp. 1 0 0 12 0 4 12 4 0 0 0 0
Coelomycete sp. 2 0 0 0 0 16 12 4 0 0 0 0
Coelomycete sp. 3 0 0 16 0 16 12 20 0 0 0 0
Colletotrichum fici-septicae 0 96 80 76 84 52 8 0 0 0 0
Colletotrichum sp. 1 0 0 4 16 20 20 0 0 0 0 0
Conidiocarpus betle 0 8 56 40 52 40 40 4 0 0 0
Conidiocarpus fici-septicae 0 100 68 56 16 0 4 0 0 0 0
Coniella fici (sexual) 0 0 4 96 24 16 4 0 0 0 0
Coniella fici (asexual) 0 0 4 80 20 24 4 0 0 0 0
Cylindrihyalospora fici 0 0 0 4 44 52 32 8 0 0 0
Cylindrocladium sp. 1 0 16 16 8 0 4 0 0 0 0 0
Diaporthe fici-septicae 0 0 4 40 80 68 64 52 12 4 0
Diplodia fici-septicae 0 0 0 40 72 72 76 44 8 0 0
Discosia ficeae 0 0 0 4 40 20 8 0 0 0 0
Fusarium sp. 1 0 0 16 0 0 8 0 4 20 20 20
Fusarium sp. 2 0 0 0 0 0 20 8 0 0 0 40
Lasiodiplodia thailandica 0 0 28 72 76 52 56 56 8 20 20
Lasiodiplodia theobromae 0 0 0 4 20 8 36 4 56 60 0
Leptodiscella sexualis 0 4 20 92 64 40 20 4 0 0 0
Microthyrium fici-septicae 0 56 64 20 12 4 4 0 0 0 0
Mucor sp. 1 0 0 0 0 0 0 0 4 24 20 20
Mucor sp. 2 0 0 0 0 0 4 4 20 28 0 0
Muyocopron ficina 0 0 0 56 52 28 12 4 0 0 4
Mycoleptodiscus alishanense 4052800400000
Neopestalotiopsis phangngaensis 0 0 0 0 0 20 12 0 0 0 0
Neophyllachora fici 88 88 24 12 0 0 8 0 0 0 0
Ophioceras ficina 0 0 0 4 24 64 68 40 0 20 0
Parawiesneriomyces chiayiense 0 0 0 0 0 4 16 4 52 40 60
Penicillium sp. 1 0 0 20 0 0 4 4 0 0 40 4
Periconia alishanica 0 0 0 0 4 8 68 20 0 0 0
Pestalotiopsis portugallica 0 0 0 0 20 52 64 40 0 0 0
Phoma sp. 1 0 20 0 8 4 4 20 4 0 0 0
Phoma sp. 2 0 0 0 20 4 8 4 0 0 0 0
Phyllosticta sp. 24 20 20 0 0 0 4 0 0 0 0
Pleurostoma sp. 0 0 0 0 0 4 20 4 0 0 0
Pseudocercospora fici-septicae 0 0 20 48 20 4 0 4 0 0 0
Pseudocercospora sp. 1 0 0 0 0 4 0 20 0 4 0 0
Rhizopus sp. 1 24 20 8 0 0 0 0 0 0 0 4
Rhizopus sp. 2 20 0 0 0 0 0 0 0 0 0 0
84 Fungal Diversity (2022) 115:73–103
1 3
was present during Stage III (35–42days) and dominant
species were Arthrinium malaysianum, Aspergillus sp.,
Bertiella fici, Colletotrichum fici-septicae, Conidiocarpus
betle, Coniella fici, Cylindrihyalospora fici, Diaporthe fici-
septicae, Diplodia fici-septicae, Lasiodiplodia thailandica,
L. theobromae, Leptodiscella sexualis, Muyocopron ficina,
Ophioceras ficina, Periconia alishanica, Pestalotiopsis
portugallica, Robillarda roystoneae, Sirastachys castane-
dae, Stachybotrys microspora and Stictis fici (Fig.6). At
Stage IV (49–63days), the species diversity and number of
species declined. The community was dominated by only
a few species with relatively high percentage occurrence.
Dominant species were Arthrinium malaysianum, Bertiella
fici, Fusarium sp., Lasiodiplodia thailandica, L. theobromae,
Mucor sp., Parawiesneriomyces chiayiense, Penicillium sp.
and Syncephalastrum sp. (Fig.6).
Shannon diversity indices showed that the species diver-
sity was highest at day 28 (H = 3.33) and lowest at day 0
(H = 1.56) (Table1). The average diversity of fungi on Ficus
septica leaves in the experimental period was 2.61. The
species diversity change during the experimental period is
shown in Table1.
Fungal community composition inCeltis formosana
A total of 275 Celtis formosana leaves were examined
during the decay process and the percentage occurrences
were calculated (Table4). On day 63, leaves were highly
skeletonized comprising vascular tissues with attached
remnants of non-vascular tissues. Fourty-eight fungal taxa
were recorded and 27 species were successfully isolated
into culture and identified up to species level. These include
eight new species (Arxiella celtidis, Colletotrichum celtidis,
Diaporthe celtidis, Discosia celtidis, Memnoniella celtidis,
Muyocopron celtidis, Periconia celtidis and Pseudoneottios-
pora cannabina) and 19 new host records (Arthrinium hydei,
Bartalinia robillardoides, Coniella quercicola, Dematiocla-
dium celtidicola, Diaporthe millettiae, Dimorphiseta acuta,
Dinemasporium parastrigosum, Lasiodiplodia theobromae,
Muyocopron dipterocarpi, M. lithocarpi, Neopestalotiopsis
asiatica, Ophioceras chiangdaoense, Pestalotiopsis dra-
caena, P. papuana, P. trachycarpicola, Phyllosticta capi-
talensis, Pseudorobillarda phragmitis, Sirastachys pan-
danicola, Strigula multiformis) (Tennakoon etal. 2021b).
Another 15 taxa were identified to genus level and belong
to Appendiculella, Aspergillus, Cladosporium, Cylindrocla-
dium, Fusarium, Idriella, Mucor, Phoma and Zygosporium
(Table4). Two taxa were unidentified Ascomycete species
(due to lack of morphology data) and another two were
Coelomycetes.
The dominant species (POC > 10%) on Celtis formosana
leaves were Muyocopron celtidis (37.82%), followed by
Strigula multiformis (37.45%) Arxiella celtidis (34.91%),
Coniella quercicola (33.45%), Pestalotiopsis dracaenea
(24%), Sirastachys pandanicola (23.64%), Dimorphiseta
acuta (22.91%), Discosia celtidis (21.09%), Muyocopron
dipterocarpi (17.09%), Memnoniella celtidis (16.73%),
Lasiodiplodia theobromae (16.36%), Phyllosticta capital-
ensis (16.36%), Periconia celtidis (16%), Dinemasporium
parastrigosum (15.27%), Pestalotiopsis trachycarpicola
(15.27%), Diaporthe millettiae (14.91%), Coniella querci-
cola (14.91%), Bartalinia robillardoides (14.55%), Dema-
tiocladium celtidicola (14.18%), Pseudorobillarda phragmi-
tis (14.18%), Pestalotiopsis papuana (13.45%), Diaporthe
celtidis (12.73%) and Arthrinium hydei (10.18%) (Fig.7).
The percentage occurrences of fungal communities using
Hierarchical Cluster Analyses (HCA), showed presence of
at least four distinct communities during succession: Stage
I (day 0–7), Stage II (day 7–14), Stage III (day 21–42)
and Stage IV (day 49–63) (Fig.7). The Stage I (0–7days)
communities were low in species number and had a low
percentage occurrence. The dominant species at this stage
Table 3 (continued) Species/days 0 3 7 14 21 28 35 42 49 56 63
Robillarda roystoneae 0 0 0 0 8 44 48 40 0 0 0
Sirastachys castanedae 0 0 0 20 4 64 96 56 0 0 0
Stachybotrys microspora 0 0 40 0 4 32 52 20 4 0 0
Stictis fici 0 0 0 0 8 20 36 8 0 0 0
Syncephalastrum sp. 1 0 0 0 4 0 0 4 0 20 0 20
Syncephalastrum sp. 2 0 0 0 0 0 4 8 20 0 20 20
Torula fici 0 020404000000
Torula sp. 1 0 0 0 4 0 0 20 0 4 0 0
Torula sp. 2 0 0 0 0 8 20 4 0 0 0 0
Unidentified Ascomycete 1 0 0 0 0 4 0 0 20 0 0 0
Unidentified Ascomycete 2 0 0 0 0 0 4 20 0 0 0 0
Unidentified Ascomycete 3 0 0 0 0 4 0 20 4 0 0 0
Unidentified Ascomycete 4 0 0 12 0 0 4 4 20 4 0 20
85Fungal Diversity (2022) 115:73–103
1 3
Fig. 6 Heat-map of microbial community composition in Ficus septica leaves with cluster analysis. The color intensity shows the percentage of
occurrence (POC), referring to color key at the top
86 Fungal Diversity (2022) 115:73–103
1 3
Table 4 Percentage occurrence
of fungal species which
occurred on Celtis formosana
leaves during succession
process
Species/days 0 3 7 14 21 28 35 42 49 56 63
Appendiculella sp. 24 16 8 4 0 0 0 0 0 0 0
Arthrinium hydei 0 0 0 0 12 40 40 20 0 0 0
Arxiella celtidis 0 0 40 52 76 88 80 48 0 0 0
Neopestalotiopsis asiatica 0 0 20 12 4 0 0 0 20 20 20
Aspergillus sp. 1 0 0 0 0 8 20 4 0 0 0 0
Aspergillus sp. 2 0 0 0 0 4 12 24 0 0 0 0
Bartalinia robillardoides 0 0 0 0 12 44 24 60 0 20 0
Cladosporium sp. 1 0 0 0 0 20 16 0 0 0 0 20
Coelomycete sp. 1 0 0 0 0 4 8 16 4 0 0 0
Coelomycete sp. 2 0 0 0 0 4 8 20 0 0 0 0
Colletotrichum celtidis 0 32 20 12 0 8 16 4 0 0 0
Coniella quercicola asexual 0 0 0 40 64 64 96 72 32 0 0
Coniella quercicola sexual 0 0 0 100 32 28 4 0 0 0 0
Cylindrocladium sp. 1 0 20 12 0 0 0 0 0 0 0 0
Dematiocladium celtidicola 0 0 0 0 0 20 76 60 0 0 0
Diaporthe celtidis 0 56 20 12 0 8 40 4 0 0 0
Diaporthe millettiae 0 0 0 44 20 56 40 4 0 0 0
Dimorphiseta acuta 0 0 0 20 44 60 72 56 0 0 0
Dinemasporium parastrigosum 0 0 0 0 0 0 16 40 52 20 40
Discosia celtidis 0 0 0 0 40 52 60 40 20 20 0
Fusarium sp. 1 0 0 16 16 0 0 0 0 0 20 20
Fusarium sp. 2 0 0 0 0 0 0 12 20 0 0 0
Idriella sp. 1 0 0 16 16 0 0 0 0 0 0 0
Lasiodiplodia theobromae 0 0 0 0 4 12 48 36 40 20 20
Memnoniella celtidis 0 0 0 0 0 8 20 36 60 20 40
Mucor racemosus 0 0 0 0 0 4 12 20 0 20 0
Mucor sp. 1 0 0 0 0 0 0 0 0 0 12 20
Mucor sp. 2 0 0 0 0 0 4 0 0 0 0 0
Muyocopron celtidis 0 0 0 0 40 64 96 80 84 32 20
Muyocopron dipterocarpi 0 0 0 0 4 52 20 40 32 40 0
Muyocopron lithocarpi 0 0 0 0 8 36 20 8 0 0 0
Ophioceras chiangdaoense 0004020000000
Periconia celtidis 0 0 0 0 0 16 20 60 40 40 0
Pestalotiopsis dracaenea 0 0 0 68 40 56 60 20 20 0 0
Pestalotiopsis papuana 0 0 0 0 20 28 40 20 20 20 0
Pestalotiopsis trachycarpicola 0 0 0 40 20 24 24 40 20 0 0
Phoma sp. 1 0 0 0 0 0 8 20 4 0 0 0
Phoma sp. 2 0 0 0 0 0 20 16 0 0 0 0
Phyllosticta capitalensis 0 60 40 48 20 8 4 0 0 0 0
Pseudoneottiospora cannabina 0 0 0 0 0 8 0 0 40 20 0
Pseudorobillarda phragmitis 0 0 4 36 36 20 40 20 0 0 0
Sirastachys pandanicola 0 0 0 0 84 60 56 40 20 0 0
Strigula multiformis 80 80 88 56 20 28 20 20 20 0 0
Trichoderma sp. 1 0 0 0 0 0 0 8 0 0 20 0
Unidentified Ascomycete 1 0 0 0 0 0 20 16 0 0 0 0
Unidentified Ascomycete 2 0 0 0 0 4 4 8 16 0 0 0
Zygosporium sp. 1 0 0 0 0 4 8 16 20 0 0 0
Zygosporium sp. 2 0 0 0 4 12 20 16 20 0 0 0
87Fungal Diversity (2022) 115:73–103
1 3
Fig. 7 Heat-map of microbial community composition in Celtis formosana leaves with cluster analysis. The color intensity shows the percentage
of occurrence (POC), referring to color key at the top
88 Fungal Diversity (2022) 115:73–103
1 3
were Appendiculella sp., Arxiella celtidis, Aspergillus sp.,
Colletotrichum celtidis, Cylindrocladium sp. 1, Diaporthe
celtidis, Phyllosticta capitalensis and Strigula multiformis
(Fig.7). The dominant species of Stage II (7–14days) were
Arxiella celtidis, Coniella quercicola, Diaporthe millettiae,
Dimorphiseta acuta, Fusarium sp. 1, Ophioceras chiang-
daoense, Pestalotiopsis dracaenea, P. trachycarpicola,
Phyllosticta capitalensis, Pseudorobillarda phragmitis and
Strigula multiformis (Fig.7). The highest species diversity
was present during Stage III (21–42days), dominant spe-
cies were Arthrinium hydei, Arxiella celtidis, Bartalinia
robillardoides, Coniella quercicola, Dematiocladium celti-
dicola, Diaporthe millettiae, Dimorphiseta acuta, Discosia
celtidis, Lasiodiplodia theobromae, Muyocopron celtidis,
M. dipterocarpi, M. lithocarpi, Periconia celtidis, Pestalo-
tiopsis dracaenea, Pestalotiopsis papuana, Pestalotiopsis
trachycarpicola, Pseudorobillarda phragmitis, Sirastachys
pandanicola and Strigula multiformis (Fig.7). In Stage IV
(49–63days), the species diversity and number of species
declined. The community was dominated by a few species
with relatively high percentage occurrence. Dominant spe-
cies were Aspergillus sp., Dinemasporium parastrigosum,
Discosia celtidis, Fusarium sp., Lasiodiplodia theobromae,
Memnoniella celtidis, Muyocopron celtidis, Muyocopron
dipterocarpi, Periconia celtidis and Pseudoneottiospora
cannabina (Fig.7).
Shannon diversity indices showed that the species diver-
sity was highest at day 35 (H = 3.27) and lowest at day 0
(H = 0.53). The average diversity of fungi on Celtis for-
mosana leaves in the experimental period was 2.38. The
species diversity change during the experimental period is
shown in Table1.
Fungal community composition inMorus australis
A total of 225 Morus australis leaves were examined during
the decay process and the percentage occurrences are listed
in Table5. On day 49, leaves were highly skeletonized com-
prising vascular tissues with attached remnants of non-vas-
cular tissues. Thirty-six fungal species were recorded and 14
species were successfully isolated into cultures and identi-
fied to species level. These include six new species (Arthrin-
ium mori, Memnoniella alishanensis, M. mori, Periconia
alishanica, Phaeodothis mori, Pseudopithomyces mori) and
11 new host records (Alternaria burnsii, Arthrinium para-
phaeospermum, A. rasikravindrae, Cladosporium tenuissi-
mum, Gilmaniella bambusae, Pestalotiopsis formosana, P.
neolitseae, P. parva, Pseudopithomyces sacchari, Pseudoro-
billarda phragmitis, Spegazzinia musae) (Tennakoon etal.
2020, 2021b). Another 15 species were identified to genus
level and belong to Appendiculella, Aspergillus, Cercos-
pora, Cladosporium, Fusarium, Mucor, Neopestalotiopsis,
Penicillium, Rhizopus and Syncephalastrum (Table5). Two
taxa were unidentified Ascomycetes (due to lack of morphol-
ogy data).
The dominant species (POC > 10%) of Morus australis
leaves were Arthrinium mori (42.22%), followed by Pestalo-
tiopsis formosana (36.89%), Spegazzinia musae (28.89%),
Phaeodothis mori (26.67%), Pseudopithomyces mori
(18.67%), Gilmaniella bambusae (17.33%), Pestalotiopsis
neolitseae (14.67%), P. parva (13.33%), Memnoniella mori
(12.89%) and Alternaria burnsii (11.56%) (Fig.8).
The percentage occurrences of fungal communities using
Hierarchical Cluster Analyses (HCA) showed presence of
at least four distinct communities during succession: Stage
I (day 0–7), Stage II (day 7–14), Stage III (day 21–42) and
Stage IV (day 42–49) (Fig.8). The Stage I (0–7days), fungal
communities were low in number and had a low percentage
occurrence. The dominant species at this stage were Alter-
naria burnsii, Appendiculella sp., Cercospora sp., Pestalo-
tiopsis formosana, P. parva and Pseudopithomyces mori
(Fig.8). The dominant species of Stage II (Day 14) were
Arthrinium mori, Cercospora sp., Fusarium sp., Pestalo-
tiopsis formosana, Phaeodothis mori and Spegazzinia
musae (Fig.8). The highest species diversity was present
during Stage III (21–42days) and dominant species were
Arthrinium mori, A. paraphaeospermum, A. rasikravindrae,
Cladosporium sp. 1, C. tenuissimum, Fusarium sp. 1, Gilm-
aniella sp., Memnoniella mori, Penicillium sp., Pestalotiop-
sis formosana, P. neolitseae, P. parva, Phaeodothis mori,
Pseudopithomyces mori, Pseudorobillarda phragmitis and
Spegazzinia musae (Fig.8). In Stage IV (Day 49), the spe-
cies diversity and number of species declined. The commu-
nity was dominated by a few species with relatively high per-
centage occurrence. The dominant species were Arthrinium
mori (Fig.8).
Shannon diversity indices showed that the species diver-
sity was highest at day 28 (H = 2.82) and lowest at day 0
(H = 1.27) (Fig. 4). The average of diversity of fungi on
Morus australis leaves in the experimental period was 1.52.
The species diversity change during the experimental period
is shown in Table1.
Fungal community composition inMacaranga
tanarius
A total of 275 Macaranga tanarius leaves were examined
during the decay process and the percentage occurrences
were calculated (Table6). On day 63, leaves were highly
skeletonized comprising vascular tissues with attached
remnants of non-vascular tissues. Fourty-two fungal spe-
cies were recorded and 19 species were successfully iso-
lated into cultures and identified to species level. These
include one new family (Oblongihyalosporaceae), two new
genera (Neodictyosporium and Oblongihyalospora), eight
89Fungal Diversity (2022) 115:73–103
1 3
new species (Diaporthosporella macarangae, Leptospora
macarangae, Oblongihyalospora macarange, Parawiesne-
riomyces chiayiense, Periconia alishanica, Memnoniella
alishanensis, Neodictyosporium macarangae and Nigros-
pora macarangae) and eleven new host records (Alter-
naria burnsii, A. pseudoeichhorniae, Arthrinium sacchari,
Cladosporium tenuissimum, Dictyocheirospora garethjo-
nesii, Hermatomyces biconisporus, Memnoniella echi-
nata, Periconia byssoides, P. celtidis, Pseudopithomyces
chartarum and Stachybotrys aloeticola) (Tennakoon etal.
2021b). Another 17 species were identified to genus level
and belong to Aspergillus, Asteridiella, Cladosporium,
Hermatomyces, Idriella, Meliola, Penicillium, Phoma and
Phyllosticta (Table6). Six taxa were unidentified Ascomy-
cetes (due to lack of morphology data).
The dominant species (POC > 10%) of Macaranga
tanarius leaves were Diaporthosporella macarangae
(77.45%), followed by Leptospora macarangae (33.09%),
Periconia byssoides (30.18%), Pseudopithomyces char-
tarum (29.46%), Stachybotrys aloeticola (22.18%), Peri-
conia alishanica (17.82%), Penicillium sp. 1 (17.09%),
Alternaria pseudoeichhorniae (14.91%), Aspergillus sp. 1
(12.73%), Alternaria burnsii (12.36%), Memnoniella alis-
hanensis (11.64%), Hermatomyces biconisporus (11.27%),
Table 5 Percentage occurrence
of fungal species which
occurred on Morus australis
leaves during succession
process
Species/days 0 3 7 14 21 28 35 42 49 56 63
Alternaria burnsii 32 24 20 8 0 0 20 0 0 0 0
Appendiculella sp. 32 24 16 4 0 0 0 0 0 0 0
Arthrinium mori 0 0 0 56 72 56 76 64 56 0 0
Arthrinium paraphaeospermum 000852440000
Arthrinium rasikravindrae 0 0 0 0 16 32 20 0 0 0 0
Aspergillus sp. 1 0 0 0 0 4 4 20 0 0 0 0
Aspergillus sp. 2 0 0 0 8 24 0 0 0 0 0 0
Cercospora sp. 1 0 0 20 16 0 0 0 0 0 0 0
Cladosporium sp. 1 0 0 0 0 4 8 20 20 0 0 0
Cladosporium sp. 2 0 0 0 4 24 0 0 0 0 0 0
Cladosporium tenuissimum 0000281600000
Fusarium sp. 1 0 0 0 0 16 16 16 0 0 0 0
Fusarium sp. 2 0 0 0 20 12 0 0 0 0 0 0
Gilmaniella bambusae 0 0 0 0 56 20 20 60 0 0 0
Memnoniella alishanensis 000082000000
Memnoniella mori 0 0 0 4 52 40 20 0 0 0 0
Mucor sp. 1 0 0 0 0 0 0 4 20 0 0 0
Mucor sp. 2 0 0 0 4 8 20 0 0 0 0 0
Neopestalotiopsis sp. 2 0 0 0 4 8 4 20 0 0 0 0
Penicillium sp. 1 0 0 0 4 8 20 20 20 0 0 0
Penicillium sp. 2 0 0 0 4 0 12 20 0 0 0 0
Periconia alishanica 000402040000
Pestalotiopsis formosana 60 44 36 20 64 80 28 0 0 0 0
Pestalotiopsis neolitseae 0 0 0 0 44 40 28 20 0 0 0
Pestalotiopsis parva 0 20 0 0 48 32 20 0 0 0 0
Phaeodothis mori 0 0 4 12 36 76 84 28 0 0 0
Pseudopithomyces mori 24 40 4 0 16 48 32 4 0 0 0
Pseudopithomyces sacchari 000042040000
Pseudorobillarda phragmitis 0004442040000
Rhizopus sp. 0 0 0 4 0 0 0 20 0 0 0
Spegazzinia musae 0 0 0 12 20 64 84 80 0 0 0
Syncephalastrum sp. 000040020000
Trichoderma harzianum 0000042020000
Trichoderma reesei 000000200000
Unidentified Ascomycete 1 0 0 0 0 20 4 0 0 0 0 0
Unidentified Ascomycete 2 0 0 0 0 0 20 0 0 0 0 0
90 Fungal Diversity (2022) 115:73–103
1 3
Fig. 8 Heat-map of microbial community composition in Morus australis leaves with cluster analysis. The color intensity shows the percentage
of occurrence (POC), referring to color key at the top
91Fungal Diversity (2022) 115:73–103
1 3
Meliola sp. 1 (11.27%), Parawiesneriomyces chiayiense
(11.27%) and Neodictyosporium macarangae (10.91%)
(Fig.9).
The percentage occurrences of fungal communities using
Hierarchical Cluster Analyses (HCA) showed presence of
at least four distinct communities during succession: Stage
I (day 0–3), Stage II (day 7–14), Stage III (day 21–42) and
Stage IV (day 56–63) (Fig.9). The Stage I (0–3days) fun-
gal communities were low in species number and had a
low percentage occurrence. The dominant species at this
stage were Diaporthosporella macarangae, Meliola spp.
and Phyllosticta sp. (Fig.9). The dominant species of Stage
II (7–14days) were Alternaria burnsii, Diaporthosporella
macarangae, Leptospora macarangae, Memnoniella
Table 6 Percentage occurrence
of fungal species which
occurred on Macaranga
tanarius leaves during
succession process
Species/days 0 3 7 14 21 28 35 42 49 56 63
Alternaria burnsii 0 4 36 32 40 16 4 0 0 4 0
Alternaria pseudoeichhorniae 0 0 0 24 44 64 4 4 24 0 0
Arthrinium sacchari 0 0 0 0 36 32 24 8 0 0 0
Aspergillus sp. 1 4 4 0 12 20 36 28 24 12 0 0
Aspergillus sp. 2 0 0 16 4 4 16 0 0 0 0 0
Asteridiella sp. 0 8 4 4 0 0 0 0 0 0 0
Cladosporium sp. 2 0 0 0 0 0 0 28 20 24 20 20
Cladosporium sp. 3 0 0 0 0 0 16 32 36 0 4 16
Cladosporium tenuissimum 0 0 0 0 4 16 20 0 0 0 0
Diaporthosporella macarangae 0 76 100 100 100 100 100 100 100 60 16
Dictyocheirospora garethjonesii 0 0 0 28 36 24 4 0 0 0 0
Hermatomyces biconisporus 0 0 0 0 40 44 24 16 0 0 0
Hermatomyces sp. 0 0 0 0 16 8 8 4 0 0 0
Idriella sp. 1 0 16 12 0 0 0 0 0 0 4 0
Idriella sp. 2 0 0 12 12 0 0 0 0 0 0 0
Leptospora macarangae 0 0 0 48 40 76 64 84 52 0 0
Meliola sp. 1 36 32 8 0 0 4 0 4 20 20 0
Meliola sp. 2 16 0 0 0 0 0 0 0 0 0 0
Meliola sp. 3 16 4 4 4 0 0 0 0 0 4 0
Memnoniella alishanensis 0 0 48 40 20 16 4 0 0 0 0
Memnoniella echinata 0 0 0 0 36 20 0 0 0 0 0
Neodictyosporium macarangae 0 0 0 4 8 44 32 32 0 0 0
Nigrospora macarangae 0 0 0 4 0 20 28 20 20 0 0
Oblongihyalospora macarange 0 0 28 20 4 16 0 0 0 0 0
Parawiesneriomyces chiayiense 0 0 0 0 0 8 44 40 28 4 0
Penicillium sp. 1 0 28 40 0 16 16 0 20 0 28 40
Penicillium sp. 2 0 0 32 0 0 28 4 0 0 28 16
Periconia alishanica 0 0 0 4 20 52 60 60 0 0 0
Periconia byssoides 0 0 20 52 56 52 60 56 36 0 0
Periconia celtidis 0 0 0 0 0 20 20 4 0 0 0
Phoma sp. 1 0 0 0 16 8 20 0 0 0 0 0
Phoma sp. 2 0 0 0 0 8 20 20 20 0 0 0
Phoma sp. 3 0 0 0 0 4 32 0 0 0 0 0
Phyllosticta sp. 28 44 4 0 4 0 0 0 0 0 0
Pseudopithomyces chartarum 0 36 52 80 8 32 4 40 44 28 0
Stachybotrys aloeticola 0 0 0 8 64 56 68 44 4 0 0
Unidentified Ascomycete 1 0 20 36 4 8 0 0 0 0 0 0
Unidentified Ascomycete 2 0 0 32 0 12 0 0 0 0 0 0
Unidentified Ascomycete 3 0 0 0 0 36 20 0 0 0 0 0
Unidentified Ascomycete 4 0 0 0 0 4 4 0 0 0 0 0
Unidentified hypomycetes sp. 1 4 4 0 36 4 0 0 0 0 20 0
Zygosporium sp. 1 0 0 0 0 0 4 16 0 0 0 4
92 Fungal Diversity (2022) 115:73–103
1 3
Fig. 9 Heat-map of microbial community composition in Macaranga tanarius leaves with cluster analysis. The color intensity shows the per-
centage of occurrence (POC), referring to color key at the top
93Fungal Diversity (2022) 115:73–103
1 3
alishanensis, Oblongihyalospora macarange, Penicillium
sp. 1, Periconia byssoides and Pseudopithomyces chartarum
(Fig.9). The highest species diversity was present during
Stage III (21–42days), dominant species were Alternaria
pseudoeichhorniae, Arthrinium sacchari, Aspergillus sp. 1,
Cladosporium spp., Diaporthosporella macarangae, Her-
matomyces biconisporus, Leptospora macarangae, Neodic-
tyosporium macarangae, Nigrospora macarangae, Parawi-
esneriomyces chiayiense, Periconia alishanica, P. byssoides,
P. celtidis, Phoma sp. 2, Pseudopithomyces chartarum and
Stachybotrys aloeticola (Fig.9). In Stage IV (56–63days),
the species diversity and number of species declined. The
community was dominated by a few species with relatively
high percentage occurrence. Dominant species were Clad-
osporium spp., Diaporthosporella macarangae, Leptospora
macarangae, Parawiesneriomyces chiayiense, Penicillium
sp. 1, Periconia alishanica, P. byssoides, Pseudopithomyces
chartarum and Stachybotrys aloeticola (Fig.9).
Shannon diversity indices showed that the species diver-
sity was highest at day 28 (H = 3.17) and lowest at day 0
(H = 1.55). The average of diversity of fungi on Macaranga
tanarius leaves in the experimental period was 2.28. The
species diversity changes during the experimental period
are shown in Table1.
Trophic modes oftaxa infungal communities
The fungi identified in this succession study were classi-
fied by trophic modes using the fungal traits ver. 1.2 online
database (https:// docs. google. com/ sprea dshee ts/d/ 1cxIm
JWMYV Tr6uI QXcTL wK1YN NzQvK JJifz zNpKC M6O0/
edit# gid= 33668 129) (Põlme etal. 2020). Eight trophic
modes were observed, viz. foliar epiphytes, endophytes,
lichenized spp., litter saprotrophs, plant pathogens, soil sap-
rotrophs, sooty molds and unspecified saprotrophs (Fig.10).
Most identified fungi were litter saprotrophs (61.1%) and
this trophic mode was recorded throughout the decompo-
sition process. Other trophic modes were soil saprotrophs
(14.4%), plant pathogens (10.4%), foliar endophytes (3.9%),
sooty molds (1.7%), foliar epiphytes (1.3%) and lichenized
species (0.8%). Fungal species with many trophic modes
(e.g. Colletotrichum spp., Pestalotiopsis spp.) were enumer-
ated in each one.
Host phylogeny andfungal composition
The findings of degree of separation (R value) of fun-
gal community composition within selected tree species
is shown in Fig.11. Statistically, a high R value implies
a strong variation in fungal composition between the
selected host species, whereas a low R value indicates a
minor difference in fungal composition. Accoding to the
results, highest R value recorded in between Celtis for-
mosana and Macaranga tanarius (R = 0.88, P < 0.001),
while lowest R value recorded in between Ficus septica
and Morus australis (R = 0.62, P < 0.001). However,
host phylogeny weakly correlated with fungal composi-
tion during the decomposition process (Fig.11). Even
Fig. 10 (a) Summary of
recorded fungal communities
and (b) fungal trophic modes
(M + P = Morphology and
Phylogeny)
94 Fungal Diversity (2022) 115:73–103
1 3
phylogenetically closely related tree species have higher
R value, than phylogenetically distant host species. For
example, phylogenetically close Ficus ampelas and F. sep-
tica (same genus) has high R value (R = 0.78, P < 0.001)
than phylogenetically distant Ficus septica and Celtis for-
mosana (R = 0.65, P < 0.001). Ficus ampelas and F. sep-
tica R value is much higher than Ficus septica and Morus
australis (R = 0.62, P < 0.001) as well. In aaddition, the
percentage of overlapping species in other tree species
pairs also indicates that host phylogeny is associated to a
lesser degree with fungal diversity in this study (Fig.12).
For example, FA is more phylogenetically close to MA
(members of the same family), than to either CF (different
family) or MT (different order). However, the number of
overlapping species between FA-MA is less than FA-CF
and FA-MT (Fig.12). In general, the expectation would
be that the number of overlapping species between phylo-
genetically close tree species should be higher than those
that are more phylogenetically distant. Another example is
that FS is phylogenetically closer to MA than CF and MT,
but result indicates that there are high number of overlap
species in FS-CF and FS-MT than FS-MA (Fig.12).
Therefore, based on our results we conclude that host phy-
logeny has a lower association with fungal composition
than expected.
Leaf litter chemistry
Total N content differed among tree species during decom-
position, but it increased in all hosts as decomposition pro-
gressed (Fig.13). The highest initial total N content was
recorded in Morus australis (2.11%) and Celtis formosana
(1.73%). The initial total N contents of the other three host
species (Ficus ampelas, F. septica and Macaranga tanarius)
varied between 1.19 and 1.27% (Fig.13). Total N content
and fungal composition of leaf litter were significantly cor-
related (r2 = 0.61, P = 0.001, Fig.14). In contrast, the total
C content of all leaf litter decreased with decomposition
(Fig.13), and there was a significant correlation with fungal
composition according to the goodness-of-fit-statistics (r2)
analysis (r2 = 0.40, P = 0.001, Fig.14). The highest initial
total C was observed in Macaranga tanarius (49.58%) leaf
litter and the lowest in Morus australis (37.86%).
As a result of N enrichment, the C/N ratio decreased in
the leaves of all host species overtime (Fig.13). The highest
initial C/N ratio was revealed in Macaranga tanarius (38.94)
leaf litter and lowest in Morus australis (17.94). The initial
C/N ratio of the other three host species (Ficus ampelas,
F. septica and Celtis formosana) varied between values of
22.87–32.75 (Fig.13). The C/N ratio and fungal compo-
sition of leaf litter were significantly correlated (r2 = 0.54,
P = 0.001, Fig.14). Lignin content also significantly corre-
sponded with fungal composition (P = 0.001, r2 = 0.25), but
it was much lower when compared to Lignin: N (r2 = 0.47,
P = 0.001) ratios. It is worthy to note that lignin content
increased with decomposition time and accumulated in the
final stages of decay (Fig.13).
Fig. 11 Degree of separation of
fungal community composition
within selected tree species
Fig. 12 Overlap between fungal communities occurring on pairs of
tree species
95Fungal Diversity (2022) 115:73–103
1 3
Host species heterogeneity, leaf litter chemistry
andsampling timesshape fungal community
succession
Variation partitioning analysis of factors demonstrated that
host species heterogeneity (31%) and leaf litter chemistry
(24%) accounted for the large variation in fungal community
composition in the five hosts. Sampling time explained a
smaller proportion of variance (5%) (Fig.14). Goodness-
of-fit-statistics (r2) also revealed significant correlations
between fungal composition and host species heterogene-
ity (r2 = 0.65, P = 0.001). However, this was considerably
lower when comparing host phylogeny (r2 = 0.21, P = 0.001)
and host families (r2 = 0.11, P = 0.001). Both variation
Fig. 13 Leaf litter chemistry variations of the selected host species during the decomposition. a Total nitrogen content (N%), b Total carbon
content (C%), c C: N ratio, d Lignin: N ratio and e Lignin content
Fig. 14 Variation partitioning analysis of factors explaining the
fungal community composition of five tree species (left) and good-
ness-of-fit statistics (r2) for factors fitted to the non-metric multidi-
mensional scaling (NMDS) ordination of the fungal community com-
position (right)
96 Fungal Diversity (2022) 115:73–103
1 3
partitioning and goodness-of-fit-statistics indicate that leaf
litter chemistry is significantly correlated to fungal com-
munity composition (Fig.14). In particular, total N shows
the strongest correlation (r2 = 0.61, P = 0.001), while lignin
content shows a much lower correlation with fungal com-
position (r2 = 0.25, P = 0.001). The significant correlation
of host species can be due to leaf litter quality, either physi-
cal or chemical characteristics of the particular tree spe-
cies leaves. These include leaf toughness, particle size, leaf
surface properties (physical characteristics) and initial C or
N content, C/N ratio, soluble sugars, polyphenols, waxes,
cellulose, hemicellulose, and lignin content (chemical char-
acteristics) are subjected to leaf litter quality of the tree
species and ultimately on fungal communities (Promputtha
etal. 2002; Bothwell etal. 2014; Purahong etal. 2016; Ge
etal. 2017; Osono 2017; Zhao etal. 2020).
Discussion
This study revealed high fungal species diversity on leaf
litter of five host tree species. The objectives of this study
were to investigate (i) the effects of tree species relationships
(including tree phylogeny) and leaf litter chemistry with fun-
gal community succession and (ii) specific patterns of fungal
succession (including diversity and taxonomic composition
of the fungal community) on decomposing leaf litter across
the five phylogenetically related tree species in a subtropi-
cal forest in Taiwan. The results support the hypothesis that
tree species heterogeneity and leaf litter chemistry signifi-
cantly correlated with the fungal communities during decay.
Our second hypothesis (leaf litter chemistry explains less
variation in fungal community composition as compared
with tree species heterogeneity) was supported by the data,
because leaf litter chemistry strongly associated with the
fungal composition, but this is less when compared with the
tree species effect (Fig.14). The third hypothesis (phyloge-
netically closely related tree species rather than phyloge-
netically distinct tree species share more fungal community
composition) is not supported here, because even the phylo-
genetically closely related tree species (Ficus ampelas and
F. septica, same genus) comprise almost totally different
fungal communities (R = 0.78, P < 0.001). Fungal taxa on
tree species from different families (Ficus septica and Celtis
formosana) were less different (R = 0.65, P < 0.001) as com-
pared to tree species of the same genus (Fig.11). This is fur-
ther confirmed by low R-value within different orders than
the same genus tree species. For instance, Ficus septica and
Macaranga tanarius have low R (R = 0.76, P < 0.001) val-
ues than Ficus ampelas and F. septica (R = 0.78, P < 0.001)
(Fig.11).
Where did thefungi onleaf litter comefrom? / Are
they prior endophytes?
The role of plant endophytes during the decomposition
process has been investigated for almost two decades. The
potential of leaf endophytes becoming early colonizers
in leaf litter has been discussed previously (Hyde 2001;
Osono 2002, Yanna etal. 2002, Promputtha etal. 2007,
2010; Hyde and Soytong 2008; de Silva etal. 2019; Hyde
etal. 2020). An experimental study on Magnolia liliifera
leaves illustrated that some endophytic fungi (e.g. Colle-
totrichum, Diaporthe, Fusarium, and Phyllosticta) switch
to the saprotrophic lifestyle (Promputtha etal. 2007). This
finding was further supported from the detection of similar
degrading enzymes in endophytic species and their sapro-
bic counterparts (Promputtha etal. 2010). The endophytes
activate these degrading enzymes once the leaf senesces
(Promputtha etal. 2010). Several other endophytes that
are morphologically and phylogenetically similar to sap-
robes have been discovered including Colletotrichum gloe-
osporioides, Colletotrichum sp. 2, Corynespora cassiicola,
Fusarium sp., Phyllosticta sp., Leptosphaeria sp. and Dia-
porthe species. De Silva etal. (2019) revealed that Lasi-
odiplodia pseudotheobromae occurred as an endophyte
and also a saprobe in Magnolia candolii (Magnoliaceae).
This suggests that leaf senescence modifies the ecological
niche for true endophytes and allows the development of
organisms that are usually better adapted to saprotrophic
life (Andrews and Hirano 1991; Promputtha etal. 2007,
2010). This would account for the differences in fungal
species in different hosts and also account for host-speci-
ficity of saprobes.
Some species found in this study are endophytes in
previous studies and according to the fungal traits ver.
1.2 database (Põlme etal. 2020). For instance, Barta-
linia robillardoides has been recorded as an endophyte in
leaves of Aegle marmelos in India (Gangadevi and Muthu-
mary 2008). The endophytic life modes of Pestalotiopsis
neolitseae (Magnolia garrettii), P. parva (Macaranga
peltata) and Phyllosticta capitalensis (Anacardium sp.,
Comocladia sp., Mangifera sp., Anthurium sp., Cinnamo-
mum sp.) have also been observed (Wikee etal. 2013;
Reddy etal. 2016; de Silva etal. 2021). If indeed sap-
robes initially have an endophytic lifestyle, then it is more
likely that they will have developed a relationship with the
host and become host-specific (Andrews and Hirano 1991;
Promputtha etal. 2007, 2010; de Silva etal. 2019; Cheth-
ana etal. 2021). Therefore, this can account for the vast
fungal numbers in host species (Fröhlich and Hyde 1999;
Wang etal. 2008; Doilom etal. 2017). This is further
confirmed in this study, where a large number of fungal
species occurred uniquely on one tree species (Fig.15).
Therefore, it would be interesting to isolate endophytes
97Fungal Diversity (2022) 115:73–103
1 3
from fresh leaves of the same host species and compare
them with the recorded saprobic species herein.
Global fungal diversity: istheratio ofsix species
offungi toone plant species (6:1) realistic?
Global fungal diversity and estimated number of fungal
species has always been controversial, as mycologists have
used various estimation criteria over time (Hawksworth
1991, 2001; Rossman 1994; Cannon 1997; Hawksworth
and Lücking 2017). For example, Hawksworth (1991) pro-
posed a ratio of about six fungal taxa per plant and provided
an estimate of 1.5 million fungal species. Various subse-
quent estimates followed, viz. 1 million (Rossman 1994),
9.9 million (Cannon 1997), 0.5 million (May 2000), 0.5–9.9
(Hawksworth 2001), 3.5–5.1 million (O’Brien etal. 2005)
and 5.1 million (Blackwell 2011). Hawksworth and Lücking
(2017) reassessed global fungal diversity upwards to 2.2–3.8
million species, taking into account cryptic species, ratio of
hosts to fungal species, unexplored niches, high throughput
sequencing surveys and the rates and patterns at which new
species are being described.
The ratio of 6:1 is still being debated, since most plants
harbor considerably more (higher than six) possibly host-
specific fungal species (Fröhlich and Hyde 1999; Wang etal.
2008; Doilom etal. 2017; Mapook etal. 2020; Tennakoon
etal. 2021b). Fröhlich and Hyde (1999) examined the fungi
of three individual palms of Licuala sp. in Brunei Darus-
salam and three individual palms of Licuala ramsayi in Aus-
tralia. The results revealed an average of 55 and 111 taxa per
plant, respectively and there were few species overlapping
between the two communities. This case clearly showcases
a high fungal diversity within a host and specificity reach-
ing host genus level. These findings suggested that a ratio
of 26:1 or even 33:1 would be more appropriate at least for
palm species in the tropics. Wang etal. (2008) conducted an
experimental study to check whether fungi are host-specific
or generalists by using five Ficus species (F. altissima, F.
virens, F. benjamina, F. fistulosa and F. semicordata) in
Thailand. The study revealed more than 24 fungal species
in each plant with few overlapping species (> 10 species),
even though they belonged to the same genus. Paulus etal.
(2006) studied the fungal species on leaf litter of Crypto-
carya mackinnoniana, Elaeocarpus angustifolius, Ficus
pleurocarpa, F. destruens, Opisthiolepis heterophylla, and
Darlingia ferruginea using direct observation of fruiting
bodies and particle filtration. A significant amount of fun-
gal diversity was revealed, with 185 species discovered from
direct observation of leaves and 419 morphotypes discov-
ered from particle filtration, respectively. The microfungal
assemblages of the tree species were distinct with 60% of
the taxa recorded only on a single host and just around 3% of
taxa shared by all tree species. Fungal diversity surveys on
single host species have also revealed more than six species,
viz. Promputtha etal. (2002) on Manglietia garrettii (22 fun-
gal species), Doilom etal. (2017) on Tectona grandis (188
fungal species), Thambugala etal. (2017) on Tamarix (24
fungal species), Phukhamsakda etal. (2020) on Clematis (88
fungal species) and Mapook etal. (2020) on Chromolaena
odorata (77 fungal species).
In this study, we also identified a considerable number
of fungal species from each tree host, viz. Celtis formosana
(48), Ficus ampelas (46), Ficus septica (64), Macaranga
tanarius (42) and Morus australis (36). The overlap of
microfungal species in pair wise comparisons of tree species
was low (7–16%), and only 1–2% of microfungal species
were observed in leaves of all tree species (Fig.15). Nev-
ertheless, the actual fungal diversity of above host species
could be much higher, if fungi were collected from other
parts of the host as well (i.e. woody litter). The results from
this study suggest that fungi appear to be host-specific, but
this needs further clarifications with additional collecting.
Therefore, the previous studies and current study results sug-
gest that the ratio of 6:1 is an underestimate and needs to be
updated (Hyde etal. 2020).
Inclusive effect oftree species heterogeneity
onfungal community composition
The quality of leaf litter (i.e. C/N ratio, lignin and cellulose
concentration, soluble sugars, polyphenols, leaf toughness,
particle size, and leaf surface properties) across tree species
varies. Consequently, this variability dictates utilization of
Fig. 15 Venn diagram analysis of fungal communities in five host
species during their decomposition process
98 Fungal Diversity (2022) 115:73–103
1 3
these substrates and may influence composition and abun-
dance of leaf litter fungal communities (Promputtha etal.
2002; Bothwell etal. 2014; Purahong etal. 2016; Ge etal.
2017; Osono 2017). In recent years, numerous studies have
quantified how different tree species harbor distinct fungal
communities (Promputtha etal. 2002, 2017; Tang etal.
2005; Paulus etal. 2006; Duong etal. 2008; Voříšková and
Baldrian 2013; Gil-Martínez etal. 2021). These findings
have revealed the importance of interspecific variation of
tree species in shaping fungal communities. However, in
some previous experimental studies, the significance of tree
species heterogeneity has been confounded by climate and
soil variation due to large geographic separation (Purahong
etal. 2016). Therefore, our study focused on tree species
from the same geographic region and forest (similar climatic
conditions in the experiment area), which circumvented the
effect of the above-mentioned environmental factors and
showcased tree species influence more clearly.
This study brings new knowledge concerning the influ-
ence of tree species heterogeneity on fungal species diver-
sity during the decomposition process. Totally 1325 leaves
(CF: 275, FA: 275, FS: 275, MT: 275 and MA: 225) were
collected and 236 fungal species recorded (CF: 48, FA: 46,
FS: 64, MT: 42 and MA: 36). A total of 115 fungal species
(CF: 27, FA: 27, FS: 28, MT: 19 and MA: 14) were success-
fully isolated into culture and identified to species (Fig.15,
Tennakoon etal. 2019a, b, 2020, 2021b). The recorded taxa
occurring on more than two tree hosts are commonly inhab-
ited in soil, for instance Aspergillus spp., Cladosporium spp.,
Fusarium spp., Mucor spp. and Penicillium spp. (Fig.15).
In addition, several shared species belong to large and taxo-
nomically complex genera (i.e. Alternaria spp. and Phoma
spp.). The large number of fungal species, which occurred
uniquely on one tree host, indicates ‘host-specificity’ on
a particular leaf litter (Zhou and Hyde 2001; Hyde etal.
2020). However, future studies are needed to clarify the host
range of these fungi. Some recorded fungal species are new
to science (two new families, two new genera and 40 new
species) (Fig.10).
Successional patterns offungal species
duringdecomposition
Fungal communities in leaf litter change over time showing
successional patterns and are divided into early, middle and
late decomposers (i.e. Pioneer, Mature and Impoverished
communities) (Garrett 1963; Hudson 1968; Frankland 1998;
Tsui etal. 2000; Jones and Hyde 2002; Promputtha etal.
2002, 2017; Tang etal. 2005; Paulus etal. 2006). In gen-
eral, early colonizers are composed of a large number of
different species occurring at low frequency with no obvious
dominant species. Dix and Webster (1995) suggested that
fungal diversity is rich, and the number of “individuals” are
highest during the earliest stages of colonization, followed
by a period of stability. Middle fungal communities consist
of fewer species with a high-level of occurrence, while dur-
ing the late stage of decomposition, fungal diversity begins
to decline and is dominated by few species with a high level
of occurrences (Dix and Webster 1995; Promputtha etal.
2002, 2017). Then fungal diversity and total numbers of spe-
cies begin to decline.
Our findings also agreed with previous studies in showing
a higher number of fungal species with a low percentage of
occurrences during the early stages, high fungal diversity
in the middle stages, while fungal diversity declines in the
later stages (Promputtha etal. 2002, 2017; Tang etal. 2005;
Paulus etal. 2006). The percentage occurrences of fungal
communities using Hierarchical Cluster Analyses (HCA),
showed that there were at least four succession stages in
each tree species. The fungal community composition was
distinct at each stage of succession. In the initial community
stage (Days 0–3 or 0–7), fungal communities were low in
number and had a low percentage occurrence. In the initial
stages, mostly foliicolous epiphytic species were dominant
acting as “early colonizers” in each tree species (i.e. Appen-
diculella sp., Ceramothyrium longivolcaniforme, Longihya-
lospora ampeli, Meliola spp., Micropeltis fici, M. ficinae,
Mycoleptodiscus alishanense, Neophyllachora fici, Zeloas-
perisporiales sp.). Many previous studies have also empha-
sized that epiphytes can colonize as “primary saprobes”
because their growth depends on soluble carbohydrate-rich
senescent leaves (Hudson 1968; Osono etal. 2004; Osono
2006). Some epiphytic species may have already colonized
leaves even before they fall (i.e. Strigula multiformis). Some
fungi may change their endophytic lifestyles to a sapro-
trophic strategy in the initial succession stages (i.e. Bar-
talinia robillardoides, Pestalotiopsis neolitseae, P. parva,
Phyllosticta capitalensis) (Gangadevi and Muthumary 2008;
Wikee etal. 2013; Reddy etal. 2016; de Silva etal. 2021).
As decomposition progressed at 7–21days, fungal diversity
and species occurrence gradually increased and reached the
peak at 28–35days (Fig.3). After that, fungal diversity and
the total number of fungal communities started to decline in
the final stages (days 42–63).
The dynamics ofleaf litter chemistry
duringdecomposition
During leaf litter decomposition, nutrients are typically
released in three phases: (i) an initial phase where leach-
ing and nutrient release dominate; (ii) a net immobilization
(i.e. net accumulation) phase, and (iii) a net release phase,
where the nutrient mass decreases (Dutta and Agrawal
2001; Goya etal. 2008; Seta etal. 2016; Mutshekwa etal.
2020). The leaf litter nutrient release patterns of of differ-
ent plant species are diverse and related to quality, season,
99Fungal Diversity (2022) 115:73–103
1 3
and environmental factors (Semwal etal. 2003; Goya etal.
2008). In general, N concentration in leaf litter increases
through decomposition and as a result of N enrichment, the
C/N ratio decreased (Mutshekwa etal. 2020; Zhang etal.
2020). Nitrogen concentration in the leaf litter of nitrogen-
fixing tree species was higher than non-nitrogen fixing tree
species (Semwal etal. 2003; Zhou etal. 2018; Bohara etal.
2020; Zhuang etal. 2020). For example, the nitrogen fixing
genus alder (Alnus) has great actual concentrations of N (fre-
quently above 3%) than pine needle litter (frequently under
0.4%) (Berg and McClaugherty, 2003; Krishna and Mohan,
2017). In the present study, the total N concentration of the
leaf litter of all host species increased as decomposition pro-
gressed (Fig.13). The rapid increases of total N concentra-
tion observed may be attributed to its immobilization by
microbes and loss of carbon. In particular, fungal activities
have been reported to be a major source of increased N in
leaf litter (Kim etal. 2003; Bargali etal. 2006, 2015; Arslan
etal. 2010). In general, fungal mycelia contain 3–5% N on a
dry mass basis, which they can translocate from organic and
mineral soil layers during decomposition (Kim etal. 2003).
Numerous authors have proposed that N increase should be
linked to the greater abundance of fungal biomass in leaf
litter and ultimately to the decomposition process (Goya
etal. 2008; Subedi and Bhatta 2010; Voříšková etal. 2011;
Chomel etal. 2016; Angst etal. 2019). In contrast, the C/N
ratio decreased due to the loss of C with the decomposi-
tion time (Fig.13). Based on goodness-of-fit statistics (r2)
analysis, total N content, total C content and C/N ratio were
significantly correlated with the fungal composition in leaf
litter (Fig.14). Total N content was the main significant fac-
tor for the fungal composition (P = 0.001, r2 = 0.61).
The lignin concentrations in leaf litter vary widely (Koide
etal. 2005; Osono etal. 2009; Ma etal. 2020). In particu-
lar, differences between plant species are likely related to
differences in lignin structure (Rahman etal. 2013). For
instance, deciduous plant species comprise fluctuating pro-
portions of syringyl and guaiacyl forms of lignin, while
conifers generally have guaiacyl lignin (Esperschutz etal.
2013). The lignin content of plant material has been stated
as an important controlling factor in leaf litter decomposi-
tion and ultimately fungal community composition (Krishna
and Mohan 2017). Fungal communities play a central role in
leaf litter decomposition since they are primary decompos-
ers of lignin (Osono etal. 2009). In general, fungal succes-
sion in leaf litter occurs as loss of soluble components in
the initial stage, followed by holocellulose decomposition
in the second stage, and finally, lignin becomes a dominant
component in the final stages (Promputtha etal. 2002, 2017;
Tennakoon etal. 2021a). Mostly, basidiomycetes act on sub-
strates as the “lignin decomposers” in the final stages of
decay (Krishna and Mohan 2017; Tennakoon etal. 2021a).
In this study, initial lignin content was higher in the leaf litter
of Macaranga tanarius (29.83%), Ficus ampelas (26.06%)
and Celtis formosana (19.92%), but comparatively less in
F. septica (6.84%) and Morus australis (5.55%). According
to the goodness-of-fit statistics (r2) analysis in this study,
there was a significant correlation between fungal commu-
nity composition and lignin content (P = 0.001, r2 = 0.25).
High lignin accumulation was observed in the final stages of
decomposition than initial stages in all tree species (Fig.13).
Conclusions
This study reveals that the leaf litter dwelling fungal com-
munities in the selected tree species are widely diverse and
vary in species composition. Consequently, we infer that tree
species heterogeneity constitutes a major factor responsible
for fungal community composition during decomposition.
More than 35 species (CF: 48, FA: 46, FS: 64, MT: 42 and
MA: 36) were recorded on each host species and nearly 18
species (CF: 27, FA: 28, FS: 40, MT: 21 and MA: 18) were
unique to each host species. These unique species may indi-
cate ‘host-specificity’ to a particular tree species leaf litter.
This may be due to the fact that some saprobes, initially have
an endophytic lifestyle and are much more likely to be host-
specific. This is because they may have developed ways in
which to deal with the hosts defence mechanisms and may
well have evolved with the hosts (Chethana etal. 2021).
Therefore, it would be interesting to isolate endophytes from
fresh leaves of the selected host species (Celtis formosana,
Ficus ampelas, F. septica, Macaranga tanarius and Morus
australis) and compare their similarities with the recorded
saprobic species. Due to the high number of unique fungal
communities in each tree species, we believe that the ratio
of 6:1 (six species of fungi to one plant species) would be
much higher and require revision upwards in future studies.
We also found that leaf litter chemistry (i.e. total N con-
tent, C content, C/N ratio, Lignin content, Lignin/N ratio)
has a significant effect on leaf litter fungal composition,
but not as much as tree species heterogeneity. In contrast
to tree species heterogeneity and leaf litter chemistry, host
phylogeny had a smaller influence on composition of fungal
functional groups. The percentage of occurrences of fungal
communities using Hierarchical Cluster Analyses (HCA)
showed that there were at least four succession stages in each
tree species and fungal community composition was distinct
at each stage of succession. These findings provide insight
into a host-fungus database for future studies and increase
knowledge of fungal diversity, as well as novel fungal dis-
coveries, viz. two new families (Cylindrohyalosporaceae and
Oblongohyalosporaceae), two new genera (Longihyalospora
and Neodictyosporium), 40 new species and 56 new host
records (Tennakoon etal. 2019a, b, 2020, 2021b).
100 Fungal Diversity (2022) 115:73–103
1 3
Acknowledgements The Department of Plant Medicine, National
Chiayi University (NCYU) is thanked to provide facilities for DNA
molecular experiment. The authors would like to thank T.K. Goh,
Gareth Jones and Derek Peršoh for their valuable suggestions and help.
This research work was partially supported by Chiang Mai University
and K.D. Hyde thanks Chiang Mai University where he was a Visiting
Professor. He also thanks the Thailand Research Fund for the Grant No.
RDG613001, entitled “Impact of Climate Change on Fungal Diver-
sity and Biogeography in the Greater Mekong Subregion”. Danushka
Tennakoon thanks Nimali I. de Silva, T.K Goh, Derek Peršoh, Eric
McKenzie, Anuruddha Karunarathna and Milan Samarakoon for their
valuable suggestions and help. The authors would like to thank Mush-
room Research Foundation (MRF), Chiang Rai Province, Thailand for
providing research financial support and Postgraduate Scholarship.
Declarations
Conflict of interest The authors declare that there is no conflict of in-
terest.
References
Andrews JH, Hirano SS (1991) Microbial ecology of leaves. Springer,
Berlin
Angst Š, Harantová L, Baldrian P, Angst G, Cajthaml T, Straková P,
Blahut J, Veselá H, Frouz J (2019) Tree species identity alters
decomposition of understory litter and associated microbial com-
munities: a case study. Biol Fertil Soils 55:525–538
Arslan H, Gurcan G, Kirmizi S (2010) Nitrogen mineralization in the
soil of indigenous oak and pine plantation forests in a Mediter-
ranean environment. Eur J Soil Biol 46:11–17
Bani A, Pioli S, Ventura M, Panzacchi P, Borruso L, Tognetti R,
Tonon G, Brusetti L (2018) The role of microbial community
in the decomposition of leaf litter and deadwood. Appl Soil
Ecol 126:75–84
Bargali SS, Pandey CB, Sharma DK (2006) Weight loss and nitrogen
release pattern in leaf and wood litter of Gliricidia sepium
(Jacq.) Walp. Bull Natl Inst Ecol 17:25–29
Bargali SS, Shukla K, Singh L, Ghosh L, Lakhera ML (2015) Leaf-
litter decomposition and nutrient dynamics in four tree species
of dry deciduous forest. Trop Ecol 56:191–200
Berg B (2014) Foliar litter decomposition: a conceptual model with
focus on pine (Pinus) litter—a genus with global distribution.
ISRN Forestry 2014:22
Berg B, McClaugherty C (2003) Plant litter. Decomposition, humus
formation, carbon sequestration. Springer, Berlin
Berg B, Laskowski R (2006) Litter decomposition: a guide to carbon
and nutrient turnover. Academic Press, Amsterdam
Blackwell M (2011) The fungi: 1, 2, 3 … 5.1 million species? Am
J Bot 98:426–438
Boddy L, Hiscox J (2017) Fungal ecology: principles and mecha-
nisms of colonization and competition by saprotrophic fungi.
The Fungal Kingdom 45:293–308
Bohara M, Acharya K, Perveen S, Manevski K, Hu C, Yadav RKP,
Shrestha K, Li X (2020) Insitu litter decomposition and nutri-
ent release from forest trees along an elevation gradient in
Central Himalaya. CATENA 194:104698
Bothwell LD, Selmants PC, Giardina CP, Litton CM (2014) Leaf
litter decomposition rates increase with rising mean annual
temperature in Hawaiian tropical montane wet forests. Peer
J 2:e685
Cannon PF (1997) Strategies for rapid assessment of fungal diversity.
Biodivers Conserv 6:669–680
Chethana KW, Jayawardena RS, Chen YJ, Konta S, Tibpromma S,
Phukhamsakda C, Abeywickrama PD, Samarakoon MC, Sen-
wanna C, Mapook A, Tang X (2021) Appressorial interactions
with host and their evolution. Fungal Divers 110:75–107
Chomel M, Guittonny-Larchevêque M, DesRochers A, Baldy V (2016)
Effect of mixing herbaceous litter with tree litters on decomposi-
tion and N release in boreal plantations. Plant Soil 398:229–241
Chomnunti P, Hongsanan S, Aguirre-Hudson B, Tian Q, Peršoh D,
Dhami MK, Alias AS, Xu J, Liu X, Stadler M, Hyde KD (2014)
The sooty moulds. Fungal Divers 66:1–36
Chou CH, Tang HY (2016) Conservation of biodiversity in Taiwan.
Botanica Orientalis. J Plant Sci 10:1–5
Cooke RC, Rayner ADM (1984) Ecology of saprotrophic fungi. Long-
man, London and New York
Cotrufo MF, Briones MJI, Ineson P (1998) Elevated CO2 affects field
decomposition rate and palatability of tree leaf litter: importance
of changes in substrate quality. Soil Biol Biochem 30:1565–1571
De Silva NI, Phillips AJ, Liu JK, Lumyong S, Hyde KD (2019) Phy-
logeny and morphology of Lasiodiplodia species associated with
Magnolia forest plants. Sci Rep 9:1–11
De Silva N, Maharachchikumbura SSN, Thambugala KM, Bhat DJ,
Karunarathna SC, Tennakoon DS, Phookamsak R, Jayawardena
RS, Lumyong S, Hyde KD (2021) Morpho-molecular taxonomic
studies reveal a high number of endophytic fungi from Magnolia
candolli and M. garrettii in China and Thailand. Mycosphere
12:163–237
Dix NJ, Webster J (1995) Fungal ecology. Chapman & Hall, London
Doilom M, Dissanayake AJ, Wanasinghe DN, Boonmee S, Liu JK,
Bhat DJ, Taylor JE, Bahkali AH, McKenzie EH, Hyde KD
(2017) Microfungi on Tectona grandis (teak) in Northern Thai-
land. Fungal Divers 82:107–182
Dossa GG, Yang YQ, Hu W, Paudel E, Schaefer D, Yang YP, Cao KF,
Xu JC, Bushley KE, Harrison RD (2021) Fungal succession in
decomposing woody debris across a tropical forest disturbance
gradient. Soil Biol Biochem 155:108142
Duong LM, McKenzie EHC, Lumyong S, Hyde KD (2008) Fungal
succession on senescent leaves of Castanopsis diversifolia in Doi
Suthep-Pui National Park, Thailand. Fungal Divers 30:23–36
Dutta RK, Agrawal M (2001) Litterfall, litter decomposition and
nutrient release in five exotic plant species planted on coal mine
spoils. Pedobiologia 45:298–312
Esperschütz J, Zimmermann C, Dümig A, Welzl G, Buegger F, Elmer
M, Munch JC, Schloter M (2013) Dynamics of microbial com-
munities during decomposition of litter from pioneering plants in
initial soil ecosystems. Biogeosciences 10:5115–5124
Frankland JC (1998) Fungal succession—unraveling the unpredictable.
Mycol Res 102:1–15
Fröhlich J, Hyde KD (1999) Biodiversity of palm fungi in the tropics:
are global fungal diversity estimates realistic? Biodivers Conserv
8:977–1004
Fryar SC (2002) Fungal succession or sequence of fruit bodies. Fungal
Divers 10:5–10
Gangadevi V, Muthumary J (2008) Taxol, an anticancer drug produced
by an endophytic fungus Bartalinia robillardoides Tassi, isolated
from a medicinal plant, Aegle marmelos Correa ex Roxb. World
J Microbiol Biotechnol 24:717–724
Garrett SD (1963) Soil fungi and soil fertility. Pergamon Press Ltd,
London
Ge J, Xie Z, Xu W, Zhao C (2017) Controls over leaf litter decomposi-
tion in a mixed evergreen and deciduous broad-leaved forest,
Central China. Plant Soil 412:345–355
Gessner O (2005) Proximate lignin and cellulose. In: Graca MAS, Bär-
locher F, Gessner MO (eds) Methods to Study Litter Decomposi-
tion. Springer Verlag, Dordrecht, A Practical Guide, pp 115–120
101Fungal Diversity (2022) 115:73–103
1 3
Gil-Martínez M, López-García Á, Domínguez MT, Kjøller R, Nav-
arro-Fernández CM, Rosendahl S, Marañón T (2021) Soil fungal
diversity and functionality are driven by plant species used in
phytoremediation. Soil Biol Biochem 153:108102
Glassman SI, Weihe C, Li J, Albright MB, Looby CI, Martiny AC,
Treseder KK, Allison SD, Martiny JB (2018) Decomposition
responses to climate depend on microbial community composi-
tion. Proc Natl Acad Sci 115:11994–11999
Gomes RR, Glienke C, Videira SIR, Lombard L, Groenewald JZ, Crous
PW (2013) Diaporthe: a genus of endophytic, saprobic and plant
pathogenic fungi. Persoonia 31:1–41
Goya JF, Frangi JL, Pérez C, Dalla F (2008) Decomposition and nutri-
ent release from leaf litter in Eucalyptus grandis plantations on
three different soils in Entre Ríos, Argentina. Bosque 29:217–226
Hammer O, Harper DA, Ryan PD (2001) Palaeontological statistics
software package for education and data analysis. Palaeontol
Electronn 4:2–9
Hawksworth DL (1991) The fungal dimension of biodiversity: mag-
nitude, significance, and conservation. Mycol Res 95:641–655
Hawksworth DL (2001) The magnitude of fungal diversity: the 1.5
million species estimate revisited. Mycol Res 105:1422–1432
Hawksworth DL, Lücking R (2017) Fungal diversity revisited: 2.2 to
3.8 million species. Microbiol Spectr 5:1–17
Heberle H, Meirelles GV, da Silva FR, Telles GP, Minghim R (2015)
InteractiVenn: a web-based tool for the analysis of sets through
Venn diagrams. BMC Bioinform 16:1–7
Hou PCL, Zou X, Huang CY, Chien HJ (2005) Plant litter decomposi-
tion influenced by soil animals and disturbance in a subtropical
rainforest of Taiwan. Pedobiologia 49:539–547
Huang Y, Ma Y, Zhao K, Niklaus PA, Schmid B, He JS (2017) Positive
effects of tree species diversity on litterfall quantity and quality
along a secondary successional chronosequence in a subtropical
forest. J Plant Ecol 10:28–35
Hudson HJ (1968) The ecology of fungi on plant remains above the
soil. New Phytol 67:837–874
Hyde KD (2001) Where are the missing fungi? Does Hong Kong have
any answers? Mycol Res 105:1514–1518
Hyde KD, Soytong K (2008) The fungal endophyte dilemma. Fungal
Divers 33:163–173
Hyde KD, Bussaban B, Paulus B, Crous PW, Lee S, Mckenzie EH,
Photita W, Lumyong S (2007) Diversity of saprobic microfungi.
Biodivers Conserv 16:7–35
Hyde KD, Jeewon R, Chen YJ, Bhunjun CS, Calabon MS, Jiang HB,
Lin CG, Norphanphoun C, Sysouphanthong P, Pem D, Tib-
promma S (2020) The numbers of fungi: is the descriptive curve
flattening? Fungal Divers 103:219–271
Index Fungorum (2021) http:// www. index fungo rum. org/ Names/
Names. asp. Accessed on March 2021
Jayasiri CS, Hyde KD, Ariyawansa HA, Bhat J, Buyck B, Cai L, Dai
YC, Abd-Elsalam KA, Ertz D, Hidayat I, Jeewon R, Jones EBG,
Bahkali AH, Karunarathna SC, Liu JK, Luangsa-ard JJ, Lumb-
sch HT, Maharachchikumbura SSN, McKenzie EHC, Moncalvo
JM, GhobadNejhad M, Nilsson H, Pang KL, Pereira OL, Phil-
lips AJL, Raspé O, Rollins AW, Romero AI, Etayo J, Selçuk F,
Stephenson SL, Suetrong S, Taylor JE, Tsui CKM, Vizzini A,
AbdelWahab MA, Wen TC, Boonmee S, Dai DQ, Daranagama
DA, Dissanayake AJ, Ekanayaka AH, Fryar SC, Hongsanan S,
Jayawardena RS, Li WJ, Perera RH, Phookamsak R, De Silva NI,
Thambugala KM, Tian Q, Wijayawardene NN, Zhao RL, Zhao
Q, Kang JC, Promputtha I (2015) The Faces of fungi database:
fungal names linked with morphology, phylogeny and human
impacts. Fungal Divers 74:3–18
Johnston SR, Boddy L, Weightman AJ (2016) Bacteria in decompos-
ing wood and their interactions with wood-decay fungi. FEMS
Microbiol Ecol 92:179
Jones EBG, Hyde KD (2002) Succession: where do we go from here?
Fungal Divers 10:1–4
Kara O, Bolat I, Cakıroglu K, Senturk M (2014) Litter decomposi-
tion and microbial biomass in temperate forests in Northwestern
Turkey. J Soil Sci Plant Nut 14:31–41
Kim C, Lim JH, Shin JH (2003) Nutrient dynamics in litterfall and
decomposing leaf litter at the Kwangneung deciduous broad-
leaved natural forest. Korean J Agric for Meteorol 5:87–93
Kirby R (2005) Actinomycetes and lignin degradation. Adv Appl
Microbiol 58:125–168
Kjøller AH, Struwe S (2002) Fungal communities, succession,
enzymes, and decomposition. In: Burns RG, Dick RP (eds)
Enzymes in the environment: activity, ecology and applications.
Marcel Dekker, New York, pp 267–284
Koide K, Osono T, Takeda H (2005) Fungal succession and decom-
position of Camellia japonica leaf litter. Ecol Res 20:599–609
Krishna MP, Mohan M (2017) Litter decomposition in forest ecosys-
tems: a review. Energy Ecol Environ 2:236–249
Kubartová A, Ranger J, Berthelin J, Beguiristain T (2009) Diversity
and decomposing ability of saprophytic fungi from temperate
forest litter. Microb Ecol 58:98–107
Lin Y, He X, Ma T, Han G, Xiang C (2015) Priority colonization of
Cinnamomum camphora litter by endophytes affects decomposi-
tion rate, fungal community and microbial activities under field
conditions. Pedobiologia 58:177–185
Lodge DJ, Cantrell S (1995) Fungal communities in wet tropical for-
ests: variation in time and space. Can J Bot 73:1391–1398
Lynd LR, Weimer PJ, Van Zyl WH, Pretorius IS (2002) Microbial cel-
lulose utilization: fundamentals and biotechnology. Microbiol
Mol Biol Rev 66:506–577
Ma Y, Huang S, Gan Z, Xiong Y, Cai R, Liu Y, Wu L, Ge G (2020) The
succession of bacterial and fungal communities during decompo-
sition of two hygrophytes in a freshwater lake wetland. Ecosphere
11:e03242
Mapook A, Hyde KD, McKenzie EH, Jones EG, Bhat DJ, Jeewon
R, Stadler M, Samarakoon MC, Malaithong M, Tanunchai B,
Buscot F (2020) Taxonomic and phylogenetic contributions to
fungi associated with the invasive weed Chromolaena odorata
(Siam weed). Fungal Divers 101:1–175
May RM (2000) The dimensions of life on earth. In: Raven PH, Wil-
liams T (eds) Nature and human society: the quest for a sustain-
able world. National Academy Press, Washington, pp 30–45
McNaughton SJ, Wolf LL (1973) General Ecology. Holt, Rinehard and
Winston Inc., New York
Mutshekwa T, Cuthbert RN, Wasserman RJ, Murungweni FM, Dalu
T (2020) Nutrient release dynamics associated with native and
invasive leaf litter decomposition: a mesocosm experiment.
Water 12:2350
O’Brien HE, Parrent JL, Jackson JA, Moncalvo JM, Vilgalys R (2005)
Fungal community analysis by large-scale sequencing of environ-
mental samples. Appl Environ Microbiol 71:5544–5550
Oksanen J, Blanchet FG, Kindt R, Legendre P, Minchin PR, O’hara RB,
Simpson GL, Solymos P, Stevens MHH, Wagner H, Oksanen MJ
(2013) Package ‘vegan.’ Community Ecology Package, R Pack-
age Version 2:1–295
Osono T (2002) Phyllosphere fungi on leaf litter of Fagus crenata:
occurrence, colonization, and succession. Can J Bot 80:460–469
Osono T (2006) Role of phyllosphere fungi of forest trees in the devel-
opment of decomposer fungal communities and decomposition
processes of leaf litter. Can J Microbiol 52:701–716
Osono T (2017) Leaf litter decomposition of 12 tree species in a sub-
tropical forest in Japan. Ecol Res 32:413–422
Osono T (2020) Functional diversity of ligninolytic fungi associated
with leaf litter decomposition. Ecol Res 35:30–43
Osono T, Bhatta BK, Takeda H (2004) Phyllosphere fungi on living and
decomposing leaves of giant dogwood. Mycoscience 45:35–41
102 Fungal Diversity (2022) 115:73–103
1 3
Osono T, Ishii Y, Takeda H, Seramethakun T, Khamyong S, To-Anun
C, Hirose D, Tokumasu S, Kakishima M (2009) Fungal suc-
cession and lignin decomposition on Shorea obtusa leaves in
a tropical seasonal forest in northern Thailand. Fungal Divers
36:101–119
Osono T, Matsuoka S, Hirose D (2020) Diversity and Geographic
Distribution of Ligninolytic Fungi Associated with Castanopsis
sieboldii Leaf Litter in Japan. Front Microbiol 11:2911
Pascoal C, Cássio F, Marcotegui A, Sanz B, Gomes P (2005) Role of
fungi, bacteria, and invertebrates in leaf litter breakdown in a
polluted river. J North Am Benthol Soc 24:784–797
Paulus B, Gadek P, Hyde KD (2003) Estimation of microfungal
diversity in tropical rainforest leaf litter using particle filtra-
tion: the effects of leaf storage and surface treatment. Mycol
Res 107:748–756
Paulus B, Gadek P, Hyde K (2006) Successional patterns of microfungi
in fallen leaves of Ficus pleurocarpa (Moraceae) in an Australian
tropical rain forest. Biol Conserv 38:42–51
Perez-Harguindeguy N, Diaz S, Cornelissen JHC, Venramini F, Cabido
M, Castellanos A (2000) Chemistry and toughness predict leaf
litter decomposition rates over a wide spectrum of functional
types and taxa in central Argentina. Plant Soil 218:21–30
Peršoh D, Stolle N, Brachmann A, Begerow D, Rambold G (2018)
Fungal guilds are evenly distributed along a vertical spruce for-
est soil profile while individual fungi show pronounced niche
partitioning. Mycol Prog 17:925–939
Photita W, Lumyong S, Lumyong P, Ho WH, McKenzie EHC, Hyde
KD (2001) Fungi on Musa acuminata in Hong Kong. Fungal
Divers 6:99–106
Phukhamsakda C, McKenzie EH, Phillips AJ, Jones EG, Bhat DJ,
Stadler M, Bhunjun CS, Wanasinghe DN, Thongbai B, Campo-
resi E, Ertz D (2020) Microfungi associated with Clematis
(Ranunculaceae) with an integrated approach to delimiting spe-
cies boundaries. Fungal Divers 102:1–203
Pointing SB, Pelling AL, Smith GJD, Hyde KD, Reddy CA (2005)
Screening of basidiomycetes and xylariaceous fungi for lignin
peroxidase and laccase gene-specific sequences. Mycol Res
109:115–124
Põlme S, Abarenkov K, Nilsson RH, Lindahl BD, Clemmensen KE,
Kauserud H, Nguyen N, Kjøller R, Bates ST, Baldrian P, Frøslev
TG (2020) Fungal Traits: a user-friendly traits database of fungi
and fungus-like stramenopiles. Fungal Divers 105:1–16
Promputtha I, Lumyong S, Lumyong P, McKenzie EC, Hyde KD
(2002) Fungal succession on senescent leaves of Manglietia
garrettii in Doi Suthep-Pui National Park, northern Thailand.
Fungal Divers 10:89–100
Promputtha I, Lumyong S, Dhanasekaran V, McKenzie EHC, Hyde
KD, Jeewon R (2007) A phylogenetic evaluation of whether
endophytes become saprotrophs at host senescence. Microb Ecol
53:579–590
Promputtha I, Hyde KD, McKenzie EH, Peberdy JF, Lumyong S (2010)
Can leaf degrading enzymes provide evidence that endophytic
fungi becoming saprobes? Fungal Divers 41:89–99
Promputtha I, McKenzie EH, Tennakoon DS, Lumyong S, Hyde KD
(2017) Succession and natural occurrence of saprobic fungi on
leaves of Magnolia liliifera in a tropical forest. Cryptogamie
Mycol 38:213–225
Purahong W, Hyde KD (2011) Effects of fungal endophytes on grass
and non-grass litter decomposition rates. Fungal Divers 47:1–7
Purahong W, Kapturska D, Pecyna MJ, Schulz E, Schloter M, Buscot
F, Hofrichter M, Krüger D (2014) Influence of different forest
system management practices on leaf litter decomposition rates,
nutrient dynamics and the activity of ligninolytic enzymes: a
case study from Central European forests. PLoS ONE 9:e93700
Purahong W, Durka W, Fischer M, Dommert S, Schöps R, Buscot F,
Wubet T (2016) Tree species, tree genotypes and tree genotypic
diversity levels affect microbe-mediated soil ecosystem functions
in a subtropical forest. Sci Rep 6:36672
Qu H, Pan C, Zhao X, Lian J, Wang S, Wang X, Ma X, Liu L (2019)
Initial lignin content as an indicator for predicting leaf litter
decomposition and the mixed effects of two perennial grami-
neous plants in a desert steppe: a 5-year long-term study. Land
Degrad Dev 30:1645–1654
Rahman MM, Tsukamoto J, Rahman MM, Yoneyama A, Mostafa KM
(2013) Lignin and its effects on litter decomposition in forest
ecosystems. Chem Ecol 29:540–553
Reddy MS, Murali TS, Suryanarayanan TS, Rajulu MG, Thiruna-
vukkarasu N (2016) Pestalotiopsis species occur as generalist
endophytes in trees of Western Ghats forests of southern India.
Fungal Ecol 24:70–75
Romaní AM, Fischer H, Mille-Lindblom C, Tranvik LJ (2006) Inter-
actions of bacteria and fungi on decomposing litter: differential
extracellular enzyme activities. Ecology 87:2559–2569
Rossman AY (1994) A strategy for an all-taxa inventory of fungal
diversity. In: Peng CI, Chou CH (eds) Biodiversity and terrestrial
ecosystems. Academia Sinica Monograph Series No. 14. Taipei,
pp 169–194
Saitta A, Anslan S, Bahram M, Brocca L, Tedersoo L (2018) Tree spe-
cies identity and diversity drive fungal richness and community
composition along an elevational gradient in a Mediterranean
ecosystem. Mycorrhiza 28:39–47
Semwal RL, Maikhuri RK, Rao KS, Sen KK, Saxena KG (2003) Leaf
litter decomposition and nutrient release patterns of six multipur-
pose tree species of central Himalaya, India. Biomass Bioenergy
24:3–11
Seta T, Demissew S, Woldu Z, Lemenih M (2016) Carbon and nutrient
release patterns during leaf litter decomposition in Boter-Becho
Forest, Southwestern Ethiopia. J Ecosystem Ecography 6:1–9
Shannon CE, Weaver W (1949) A mathematical model of communica-
tion. University of Illinois Press, Urbana, IL, p 11
Shearer CA (1995) Fungal competition. Can J Bot 73:1259–1264
Shirouzu T, Hirose D, Fukasawa Y, Tokumasu S (2009) Fungal suc-
cession associated with the decay of leaves of an evergreen oak,
Quercus myrsinaefolia. Fungal Divers 34:87–109
Subedi R, Bhatta B (2010) Decomposition of forest leaf litters for bet-
ter nitrogen release. In: Proceedings of National Conference on
Forest-People Interaction, Pokhara, Nepal
Tang AM, Hyde JR (2005) Succession of microfungal communi-
ties on decaying leaves of Castanopsis fissa. Can J Microbiol
51:967–974
Tennakoon DS, Jeewon R, Gentekaki E, Kuo CH, Hyde KD (2019a)
Multi-gene phylogeny and morphotaxonomy of Phaeosphaeria
ampeli sp. nov. from Ficus ampelas and a new record of P. musae
from Roystonea regia. Phytotaxa 406:111–128
Tennakoon DS, Thambugala KM, Jeewon R, Hongsanan S, Kuo CH,
Hyde KD (2019b) Additions to Chaetothyriaceae (Chaetothyri-
ales): Longihyalospora gen. nov. and Ceramothyrium longi-
volcaniforme, a new host record from decaying leaves of Ficus
ampelas. MycoKeys 61:91–109
Tennakoon DS, Kuo CH, Hyde KD (2020) Multi-locus phylogeny
reveals Phaeodothis mori sp. nov. (Didymosphaeriaceae, Ple-
osporales) from dead leaves of Morus australis. Phytotaxa
428:241–254
Tennakoon DS, Gentekaki E, Jeewon R, Kuo CH, Promputtha I, Hyde
KD (2021a) Life in leaf litter: Fungal community succession
during decomposition. Mycosphere 12:406–429
Tennakoon DS, Kuo CH, Maharachchikumbura SS, Thambugala KM,
Gentekaki E, Phillips AJ, Bhat DJ, Wanasinghe DN, de Silva NI,
Promputtha I, Hyde KD (2021b) Taxonomic and phylogenetic
contributions to Celtis formosana, Ficus ampelas, F. septica,
Macaranga tanarius and Morus australis leaf litter inhabiting
microfungi. Fungal Divers 108:1–215
103Fungal Diversity (2022) 115:73–103
1 3
Thambugala KM, Daranagama DA, Phillips AJ, Bulgakov TS, Bhat
DJ, Camporesi E, Bahkali AH, Eungwanichayapant PD, Liu
ZY, Hyde KD (2017) Microfungi on Tamarix. Fungal Divers
82:239–306
Tian G, Brussaard L, Kang BT, Swift MJ (1997) Soil fauna-mediated
decomposition of plant residues under constrained environmental
and residue quality conditions. In: Cadisch G, Giller KE (eds)
Driven by Nature: Plant Litter Quality and Decomposition. CAB
International, London
Tsui KM, Hyde KD, Hodgkiss IJ (2000) Biodiversity of fungi on
submerged wood in Hong Kong streams. Aquat Microb Ecol
21:289–298
Veen GF, Snoek BL, Bakx-Schotman T, Wardle DA, van der Putten
WH (2019) Relationships between fungal community composi-
tion in decomposing leaf litter and home-field advantage effects.
Funct Ecol 33:1524–1535
Voříšková J, Baldrian P (2013) Fungal community on decomposing leaf
litter undergoes rapid successional changes. ISME J 7:477–486
Voříšková J, Dobiášová P, Šnajdr J, Vaněk D, Cajthaml T, Šantrůčková
H, Baldrian P (2011) Chemical composition of litter affects the
growth and enzyme production by the saprotrophic basidiomy-
cete Hypholoma fasciculare. Fungal Ecol 4:417–426
Wang HK, Hyde KD, Soytong K, Lin FC (2008) Fungal diversity on
fallen leaves of Ficus in northern Thailand. J Zhejiang Univ Sci
B 9:835–841
Wikee S, Lombard L, Crous PW, Nakashima C, Motohashi K, Chuke-
atirote E, Alias SA, McKenzie EH, Hyde KD (2013) Phyllosticta
capitalensis, a widespread endophyte of plants. Fungal Divers
60:91–105
Wu SH, Hsieh CF, Rejmánek M (2004) Catalogue of the naturalized
flora of Taiwan. Taiwania-Taipei 49:16–31
Yanna Ho WH, Hyde KD (2002) Fungal succession on fronds of Phoe-
nix hanceana in Hong Kong. Fungal Divers 10:185–211
Zhang J, Li J, Fan Y, Mo Q, Li Y, Li Y, Li Z, Wang F (2020) Effect of
nitrogen and phosphorus addition on litter decomposition and
nutrients release in a tropical forest. Plant Soil 454:139–153
Zhao B, Xing P, Wu QL (2020) Interactions between bacteria and
fungi in macrophyte leaf litter decomposition. Environ Micro-
biol 23:1130–1144
Zhou D, Hyde KD (2001) Host-specificity, host-exclusivity, and host-
recurrence in saprobic fungi. Mycol Res 105:1449–1457
Zhou G, Zhang J, Qiu X, Wei F, Xu X (2018) Decomposing litter and
associated microbial activity responses to nitrogen deposition in
two subtropical forests containing nitrogen-fixing or non-nitro-
gen-fixing tree species. Sci Rep 8:1–11
Zhuang L, Liu Q, Liang Z, You C, Tan B, Zhang L, Yin R, Yang K, Bol
R, Xu Z (2020) Nitrogen additions retard nutrient release from
two contrasting foliar litters in a subtropical forest, southwest
China. Forests 11:377
Authors and Aliations
DanushkaS.Tennakoon1,2,3,4· Chang‑HsinKuo5· WitoonPurahong6· EleniGentekaki1,2· ChayakornPumas4·
ItthayakornPromputtha4,8· KevinD.Hyde1,2,3,4,7,8
1 Center ofExcellence inFungal Research, Mae Fah Luang
University, ChiangRai57100, Thailand
2 School ofScience, Mae Fah Luang University,
ChiangRai57100, Thailand
3 Research Center ofMicrobial Diversity andSustainable
Utilization, Chiang Mai University, ChiangMai50200,
Thailand
4 Department ofBiology, Faculty ofScience, Chiang Mai
University, ChiangMai50200, Thailand
5 Department ofPlant Medicine, National Chiayi University,
300 Syuefu Road, ChiayiCity60004, Taiwan
6 Department ofSoil Ecology, UFZ-Helmholtz Centre
forEnvironmental Research, Theodor-Lieser-Str. 4,
06120Halle(Saale), Germany
7 Innovative Institute forPlant Health, Zhongkai University
ofAgriculture andEngineering, Haizhu District,
Guangzhou510225, People’sRepublicofChina
8 Environmental Science Research Center, Faculty ofScience,
Chiang Mai University, ChiangMai50200, Thailand
