BIOTROPICA 38(1): 42–51 2006 10.1111/j.1744-7429.2006.00110.x Successional Patterns of Microfungi in Fallen Leaves of Ficus pleurocarpa (Moraceae) in an Australian Tropical Rain Forest1 Barbara Paulus2,3 , Paul Gadek School of Tropical Biology, James Cook University, Cairns, QLD 4870, Australia and Kevin Hyde Centre for Research in Fungal Diversity, Department of Ecology & Biodiversity, The University of Hong Kong, Pokfulam Road, Hong Kong, SAR, People’s Republic of China ABSTRACT Successional patterns of microfungi on decaying leaves of Ficus pleurocarpa were assessed as part of a study to enumerate microfungi in tropical rain forest leaf litter. Leaves degraded into fragments over a period of 3 mo. Two methods were applied, a direct observational method and a particle filtration protocol. Using a direct method, 104 species were observed, while 53 sporulating taxa and 100 sterile morphotaxa were isolated by particle filtration. Overall patterns of succession were confirmed by both methods, but the relative abundance of species detected differed between the two methods. Nonmetric Multidimensional Scaling identified at least four successional stages and suggested that microfungal communities increased in similarity with advancing decay. Data collected by the direct method indicated a slow but steady decline of diversity with advancing decay, whereas an increase in diversity was detected by particle filtration. Synecological succession studies provide a useful tool to identify patterns and generate hypotheses. Understanding the underlying causes of successional patterns will require further autecological studies. Key words: Australia; decomposition; direct method; fungal ecology; particle filtration; tropical microfungi; tropical rain forest. THE TERM “SUCCESSION” DENOTES THE CHANGES OR TURNOVER OF (Walker & del Moral 2003). Within this broad definition, a rich variety of types and trajectories of succession over a range of temporal and spatial scales has been described (Walker & del Moral 2003). Plant successions, particularly in complex systems such as forests, may take place over decades or centuries and their study requires painstaking monitoring of mortality and recruitment in plots or the study of chronosequences (Rees et al. 2001). Along with successional changes among plants, concurrent changes among associated fungal species have been reported (Suzuki 2002). This fungal succession at the level of ecosystems (macroscale) differs from the replacement of fungal species within particular substrata with advancing decay (microscale; Suzuki 2002). Successional changes at the level of substrata may occur rapidly, and thus can provide a model system for which results can be obtained over short periods of time. Due to the importance of fungi in nutrient immobilization and mineralization (Lodge 1992), fungal successions on plant substrata have been well documented and a discussion of definitions and hypotheses has been provided in a number of reviews (Park 1968; Rayner & Todd 1979; Dix & Webster 1985; Frankland 1992, 1998; Neville & Webster 1995; Fryar 2002). In addition, a recent volume of Fungal Diversity (vol. 10, 2002) was devoted solely to this topic. Numerous early studies were undertaken in temperate regions of SPECIES OVER TIME 1 Received 25 August 2003; revision accepted 19 April 2005. Corresponding author; e-mail: paulusb@landcareresearch.co.nz 3 Present address: Landcare Research, Private Bag 92-170, Auckland, New Zealand. 2 42 the Northern Hemisphere (e.g., Webster 1956, 1957; Kendrick & Burgess 1962; Frankland 1966; Hudson 1968; Kuter 1986) and to lesser extent in tropical and subtropical regions (Hudson 1962c, Meredith 1962, Sandhu & Sidhu 1980, Cornejo et al. 1994). Recent years have shown a renewed interest in studying fungal succession, particularly in subtropics and tropics (e.g., Promputtha et al. 2002, Sivichai et al. 2002, Somrithipol et al. 2002, Yanna et al. 2002, Zhou & Hyde 2002) and in Japan (Osono 2002, Suzuki et al. 2002, Tokumasu & Aoiki 2002). Many of these studies have taken a synecological approach, recording species assemblages at different stages of decay. Fewer studies have been concerned with unraveling the underlying causes of fungal succession (Frankland 1966, 1998; Rosenbrock et al. 1995; Fryar 2002; Osono 2002). The term “fungal succession” has been applied to both the replacement of fungal mycelium and the sequence of fungal sporulation (Fryar 2002). As our aim was to detect successional patterns of both fruiting bodies and mycelia, we applied two methods to study fungal succession in decaying leaves of Ficus pleurocarpa F. Muell. The direct method, in which the substratum is examined in the field or laboratory for fruiting bodies (Booth 1971), provides information only about the sequence of fungal sporulation (Fryar 2002) and may underestimate the fungal diversity in a substratum (Bills & Polishook 1994, Paulus et al. 2003). In contrast, indirect isolation approaches bypass observations of fungal reproductive structures by detecting the presence of a fungus in its vegetative state (Booth 1971). The study was carried out over a period of 3 mo during which the leaves decomposed to a fragmented state.
C 2005 The Author(s) C 2005 by The Association for Tropical Biology and Conservation Journal compilation
Succession of Microfungi on Ficus pleurocarpa METHODS STUDY SITE AND SUBSTRATUM.—The study site was near Topaz, Queensland, Australia (17◦ 24′ 00′′ S, 145◦ 43′ 30′′ E) at an altitude of approximately 700 m. Characteristically, a high percentage (79%) of the mean annual rainfall (3810 mm) occurs during the months of December to May (Bureau of Meteorology 2000), but the rainfall during the study period was well below average. The rain forest of the site has been classified as complex mesophyll rain forest (Type 1b; Tracey 1982) and is a highly diverse mosaic of both early and late successional tree species. The tropical rain forests of Australia accommodate 32 species of Ficus, which belongs to the pantropical family Moraceae (Hyland & Whiffin 1993). Decaying leaves of one representative species, F. pleurocarpa, were selected as substratum for this study. This strangler fig is endemic to an area between Cape Tribulation and Tully, North East Queensland, occurring in well-developed rain forest at an altitudinal range from near sea level to 1000 m (Hyland & Whiffin 1993). Typically, its leaves are large (16–28 × 7–11 cm) and usually glabrous. They have a thick, leathery lamina with a prominent midrib and a long, robust petiole. Midribs and petioles of attached and freshly fallen leaves produce a white, milky exudate (latex). Freshly fallen leaves are usually yellow/green or yellow and have a fresh abscission scar, while green leaves are encountered only infrequently on the forest floor. RESEARCH DESIGN.—As few individual F. pleurocarpa trees were present in the study area, it was impractical to randomize the selection; hence, five trees of F. pleurocarpa were selected on the basis of adequate leaffall. A total of 120 green/yellow or yellow leaves with relatively fresh abscission scars were collected on 6 January 2002 (wet season). Twenty-four leaves were placed under each of five trees and secured to a tent peg with nylon string. Half of these leaves were placed with the abaxial and half with the adaxial surface in contact with the soil. As a pilot study had shown that the nylon string was frequently chewed, presumably by rats, we laid plastic mesh (15 × 15 mm grid) over groups of leaves and secured them with tent pegs. Ten leaves (two per tree) were removed on days 0, 7, 14, 30, 46, 62, 78, and 94, and transported in plastic bags to the laboratory within 4 h. Collections were abandoned after approximately 3 mo when 80 leaves had been examined, as leaves had disintegrated to an extent that fragments could not readily be identified as belonging to the study leaves. Ten green leaves were also collected across the first six visits (N = 10). DIRECT ISOLATIONS.—All leaves were incubated in separate humid chambers containing tissue paper moistened with sterile distilled water. To ensure that a large proportion of fungi fruiting on the leaves was captured, leaves were initially observed after 2 d (Hudson 1968) and again after a longer period of incubation (Hering 1965), i.e., between 7 and 17 d. As our aim was to maximize detection of fungi on leaves, we used a 15 × 15-mm sampling grid. We assumed that some fungal taxa would superficially resemble each other when viewed with a stereomicroscope. Therefore, a slide was prepared for one representative fruiting structure of each morphologically dis- 43 tinct fungal entity in each 15 × 15 mm quadrat. To manage the large number of observations, only preliminary notes were taken while examining leaves. Slides were rendered semipermanent by the addition of 90 percent lactic acid and identifications were completed once all leaves had been examined. Herbarium specimens were prepared for each taxon by removing a section of leaf with a scalpel blade and drying it at 37◦ C for 3 d. Single spore isolations were undertaken for selected taxa (Booth 1971). In view of the relative paucity of taxonomic treatise available for microfungi of the tropics in general and of North Queensland in particular, species identifications were deliberately conservative. Specimens that did not agree fully with a published description were designated as “sp.” Nevertheless, it cannot be excluded that identifications based on descriptions of temperate fungi may harbor some cryptic species. Fungal specimens of taxa, which were recognized as new to science, were deposited at the Queensland Plant Pathology Herbarium (BRIP), Department of Primary Industries. INDIRECT ISOLATIONS.—Leaves collected on days 0, 46, and 94 were also analyzed in replicate using an adapted particle filtration protocol (Paulus et al. 2003). Leaves were cleansed, surface treated with sodium hypochlorite, homogenized in a blender, and washed through a series of sieves. Leaf particles between 105 and 210 µm in size were plated onto triplicate plates of four isolation media (see below). Plates were incubated at room temperature (ca 23◦ C) and checked daily over a period of 3 wk. Leaf particles from which hyphae emerged were transferred to a Petri dish of potato dextrose agar (Merck). Isolates were then sorted into groups based on morphological characters such as hyphal and soluble pigments, shape of margin, colony surface textures, and growth rates (Bills & Polishook 1994, Lacap et al. 2003). For the purpose of this discussion, each group of isolates defined either by a taxonomic identification or by a colony morphology is termed a “morphotaxon.” These groups were reconfirmed after 4 wk. To induce sporulation, representative strains of each morphotaxon were subcultured onto Potato Carrot Agar (Johnston & Booth 1983) containing a 0.02 m2 piece of banana leaf and incubated with a 12-h photo period under black light (Booth 1971). Most sporulating cultures were identified to genus, and isolates of different species within the same genus were designated species 1, species 2, etc. (Bills & Polishook 1994). To supplement information on the species composition in green leaves, cultures were also isolated from a small number of naturally fallen green leaves. It was not our intention to compare the observed diversity, as the applied methodology is not comparable, and microfungal diversity and the presence of slower growing cultures may be better reflected using smaller leaf pieces. Green leaves were surface sterilized (96% ethanol for 30 sec, 33% sodium hypochlorite for 5 min, four washes with sterile distilled water; Petrini & Dreyfuss 1981). Ten squares (5 × 5 mm) were cut from each of ten leaves with a sterile scalpel blade and placed on replicate plates of cornmeal agar (CMA; Merck) containing streptomycin 50 mg/l. Petri dishes were checked daily and emerging hyphae were subcultured onto malt yeast extract (MYA). Cultures were treated as described above. 44 Paulus, Gadek, and Hyde MEDIA.—As the isolation media may potentially influence the diversity observed, four different media types were utilized. These were based on media described by Bills and Polishook (1994) and differed in nutrient content and the method of restricting colony growth. All media used for the isolation of fungal strains were made up to 1 liter of distilled water. Bandoni’s medium contained 4 g L-sorbose, 0.5 g yeast extract, and 20 g agar (Bandoni 1981). MYA agar contained 10 g malt extract, 2 g yeast extract, 20 g agar, and 25 mg rose bengal (adapted from Dreyfuss 1986). CMA (Merck) was made up to manufacturer’s instructions and rose bengal (25 mg/l) was added. An adapted medium based on dichloran rose bengal chloramphenicol agar (DRBC; Gams et al. 1998) contained 5 g yeast extract, 10 g dextrose, 1 g potassium phosphate, 0.5 g magnesium sulfate, and 25 mg rose bengal. After cooling to approximately 37◦ C, the following antibiotics were added to the respective media: Bandoni’s and MYA contained and chlortetracycline (50 mg/l), CMA streptomycin (50 mg/l) and DRBC streptomycin (50 mg/l) and chloramphenicol (10 mg/l). STATISTICAL ANALYSES AND DEFINITIONS.—Fungal species sporulating on incubated leaves were recorded as either present or absent for each leaf, as it was not possible to determine whether fruiting structures occurring in different quadrats on the same leaf were produced by the same mycelium. The number of leaves on which a particular fungal species was found was then designated the “occurrence of a fungus” and was used to calculate the “percent occurrence” of a species during each time period using the following formula (Yanna et al. 2002): Percent occurrence of taxon A = Occurrence of taxon A × 100 . Occurrence of all taxa The percent abundance was assessed for morphotaxa isolated by particle filtration. “Abundance” in this context refers to the relative frequencies of isolates within isolation periods and not to the relative number of “individuals” (Magurran 1988). Shannon’s diversity index (H ′ ) was used to express the species diversity of microfungal assemblages at each time point (Magurran 1988). We used regression analysis to detect the best model for the relationship between Shannon’s diversity indices and the number of days after baiting as well as days after baiting squared (SPSS 2001). The relative similarity of microfungal assemblages from leaves at different stages of decay identified by the direct method was visualized in two-dimensional space by Nonmetric Multidimensional Scaling (NMDS). This method was chosen because our data did not conform to the assumptions of normality, linearity, and similar variance of data and independence of variables, which need to be met for most other ordination methods (Shepard 1962, Kruskal 1964, Clarke & Warwick 1994). Calculations were based on a matrix of Bray–Curtis distances (Bray & Curtis 1957, SPSS 2001). For the purpose of this article, the term “assemblage” instead of “community” was applied to indicate that groups of fungi on a resource unit are not “a crisply defined entity” (Frankland 1992). The term “dominant species” refers to species with percent abundances over 5 percent and is not intended to express area coverage of a mycelium on individual leaves. RESULTS A total of 629 “occurrences,” i.e., individual observations of a fungal taxon on a leaf, were recorded in 105 taxa over 94 d using a direct observational method (Appendix 1). If known anamorph– teleomorph connections were taken into account, 104 species were recorded. Other anamorphs were suspected to be asexual states of observed teleomorphs (sexual states), such as Guignardia sp. and a Leptodothierella state and Chaetosphaeria sp. and Dictyochaeta sp., but these connections have not been confirmed. Indirect isolations yielded 562 cultures of which 265 (47%) sporulated. Sporulating cultures comprised 53 taxa, while the remaining 297 sterile cultures comprised 100 morphotaxa. No attempt was made to determine the equivalency of sterile cultures from different isolation periods. The percent abundance of all species at different time points are provided in Appendix 1 for direct isolation data and in Appendix 2 and Figure 1 for indirect isolation data. Dominant taxa, i.e., those with an abundance of greater than 5 percent, in direct observations are shown in Figures 2–4. Early species included Colletotrichum sp. 1 and sp. 2, a Gaeumannomyces-like ascomycete, Meliola sp., Pestalotiopsis sp., Phomopsis sp. 1, Zygosporium echinosporum and Z. mansonii, an unidentified ascomycete (F450), and the anamorph of a discomycete (F472) new to science. Some of the early species sporulated up to day 14 and a small number up to days 30 and 46 (Fig. 2). Although the overall species numbers were high between days 7 and 30 (Table 1), the group of dominant taxa was comparatively smaller (Fig. 3). These included Volutella ramkurii, Selenodriella fertilis, Phomopsis sp. 2, Ophiognomonia elasticae, Lanceispora amphibia, and discomycete F472. Some species commenced sporulation as early as day 7 but did not become dominant until later in decay, while others only formed fruiting bodies from day 62 onward (Fig. 4). These included Dactylaria ficusicola, FIGURE 1. Percent abundance of sporulating microfungi isolated from leaves of Ficus pleurocarpa using a particle filtration protocol. Microfungal species were ordered by their abundance in naturally abscised green (GL) and yellow leaves (day 0) and in leaves, which had been on the ground for 46 and 94 d. Succession of Microfungi on Ficus pleurocarpa 45 FIGURE 2. Percentage abundance and distribution of dominant species on green leaves and freshly fallen leaves of Ficus pleurocarpa. Chaetospermum camelliae, Gliocladiopsis tenuis, Helicosporium griseum, Helicosporium sp., Speiropsis pedatospora, an ascomycete in the Hysteriaceae, and a Xenogliocladiopsis-like species. An Asterina sp. was present for much of the study period with fruiting structures deteriorating in the latter stages. Species numbers, occurrence, and Shannon’s diversity index (H ′ ) are presented in Table 1. The best model for Shannon’s diversity indices and number of days after baiting was an inverse linear relationship (R2 = 0.655, P = 0.017; Fig. 5) for data collected by the direct method. In contrast, the correlation for the three data points obtained by particle filtration was not significant (R2 = 0.615, P = 0.426). A two-dimensional NMDS representation of direct observation data is provided in Figure 6. Although time between collections was shorter for days 0, 7, 14, and 30, these collections did not cluster closely while those of days 46–94 leaves did. A low scatter in a Shepard’s plot (not shown) and a very low stress value FIGURE 3. Percentage abundance and distribution of dominant species on Ficus pleurocarpa leaf baits, which had been on the ground for 7–30 d. FIGURE 4. Percentage abundance and distribution of dominant species on Ficus pleurocarpa leaf baits, which had been on the ground for 46–94 d. of 0.02 indicate that the “map” is a good representation of the data (Clarke & Warwick 1994). DISCUSSION A precise definition of fungal succession was provided by Rayner and Todd (1979) who termed it “the sequential occupation of the same site by thalli (normally mycelia) either of different fungi or of different associations of fungi.” Many studies of microfungal succession in substrata, including this study, fall short of demonstrating “sequential occupation of the same site” as most methods are destructive (Fryar 2002) and, unlike in large pieces of wood (Rayner & Todd 1979), individual mycelia are rarely clearly delimited in substrata such as leaves. The challenge of directly demonstrating sequential occupation of species is also known from the study of plant succession where sequential occupation is often inferred through the study of chronosequences (Walker & del Moral 2003). In contrast to chronosequences, microfungal systems have the advantage that they are amenable to experimental manipulation. Therefore, the potential disadvantage of space-for-time substitution in chronosequences, i.e., that older sites have different histories from younger ones, is circumvented. Changes in microfungal species associations with advancing decay were observed with both direct and indirect methods in decaying leaves of F. pleurocarpa (Figs. 1–4). The species composition of the particle filtration cohorts in our study was distinct for each isolation (Appendix 2; Fig. 1), suggesting that the observed changes in microfungal assemblages represent mycelial replacement and are true successive waves of invasion (Neville & Webster 1995). While most dominant sporulating species isolated by the indirect methods 46 Paulus, Gadek, and Hyde TABLE 1. Number of species, total occurrence, and Shannon’s diversity indices for direct isolations and number of morphotaxa, number of cultures, and Shannon’s diversity index for indirect isolations. Sampling time GLa Day 0 Day 7 Day 14 Day 30 Day 46 Day 62 Day 78 Day 94 21 66 2.74 33 111 3.12 39 109 3.43 28 79 3.00 23 59 2.83 31 65 3.16 20 60 2.64 23 56 2.84 13 27 2.36 Direct No. of species Total occurrenceb H′ Indirect No. of morphotaxa No. of cultures H′ a GL 38 109 3.11 23 146 2.37 54 169 3.34 52 138 3.20 = green leaves; data not directly comparable due to longer period of collection and different indirect isolation protocol. b Occurrence is the number of leaves on which a fungal species was observed. were also detected by the direct method in this study, the relative abundances obtained by the two methods differed markedly in some instances (Appendices 1 and 2). This discrepancy may represent sampling error (Ludwig & Reynolds 1988, Henderson 2003), but the labor-intensive nature of isolating fungi by an indirect method prohibited more intensive sampling. Identification of sterile morphotaxa using molecular methods would have allowed a more comprehensive comparison of species composition between different collections using the same method of isolation and between the same collections using different methods but this was considered beyond the scope of this study. An analysis of microfungal assemblages by two-dimensional NMDS suggested at least two successional stages (Fig. 6). The similarity of microfungal assemblages increased with advancing decay with the greatest distance between microfungal assemblages of day 0 leaves to day 30 leaves. A similar trend was inferred from the study of bamboo baits (Zhou & Hyde 2002) and of palm frond baits of Phoenix hanceana (Yanna et al. 2002). The underlying mechanism for this trend is not known, but it can be hypothesized that distinct, early microfungal assemblages may include species adapted to breaking down inhibitory compounds such as latex and alkaloids contained in plant tissues (Swenson et al. 1989, Isaac 1992, Neville & Webster 1995). Rapid degradation of these compounds may lead to equally rapid changes in fungal associations during early decay. It is likely that processes similar to those reported for plant successions may shape the course of succession after biologically active compounds have been broken down (Frankland 1992, Neville & Webster 1995). These may include a variety of environmental factors, such as microclimate, and population processes, such as competition and chance (Rees et al. 2001, Walker & del Moral 2003). Further work is required to elucidate the relationship between enzymatic capabilities of early successional species and leaf chemistry and to determine the effect of environment and population processes on microfungal assemblages. Changes in functional groups, which have been described for a number of fungal successions, include the replacement of parasitic and endophytic fungi by primary saprotrophs followed by a rather FIGURE 5. Shannon’s diversity indices for microfungal assemblages on fallen FIGURE 6. Nonmetric Multidimensional Scaling ordination of Bray–Curtis leaves of Ficus pleurocarpa for direct and indirect isolation methods. The regression line refers to direct data points only. distances between microfungal assemblages on fallen leaves of Ficus pleurocarpa collected at different stages of decay. Succession of Microfungi on Ficus pleurocarpa diverse group of secondary saprotrophs (Hudson 1968, Frankland 1998). Both endophytic and parasitic fungi have the advantage of colonizing the leaf ahead of other species (Hudson 1968, Neville & Webster 1995) and some may be able to switch to a saprobic mode of nutrition once the leaf senesces (Cooke & Whipps 1993). Primary saprotrophs include species with fast germination, rapid growth, and high inoculum potential (Neville & Webster 1995). Similar traits have also been reported for early successional plant species, which rapidly colonize disturbed areas under resource-rich conditions (Rees et al. 2001). During later stages of decay, fungal assemblages may be dominated by secondary saprotrophs (Hudson 1968, Frankland 1998) and may undergo successive waves of invasion (Neville & Webster 1995). Secondary saprotrophs are analogous to late successional plant species that are able to outcompete early successional plant species when resources are rare (Rees et al. 2001). These previously described patterns could also be discerned from leaves of F. pleurocarpa. Fungi observed on freshly fallen leaves included opportunistic or obligate parasitic or endophytic genera such as Colletotrichum (anamorphic Glomerella), Cylindrocladium, Meliola, Pestalotiopsis, Phomopsis (Shivas & Hyde 1997), xylariaceous species (Rodrigues & Petrini 1997), and “discomycete F472” (Appendices 1 and 2). Two species of Zygosporium and Beltrania rhombica (Appendices 1 and 2) appear to be primary colonizers with rapid growth and high inoculum potential. Decaying leaves of F. pleurocarpa lacked species commonly reported from the Northern Hemisphere such as Alternaria, Aureobasidium, Cladosporium, and Epicoccum (Frankland 1998). Secondary saprotrophs were a diverse assemblage, which reached dominance either at earlier or at later stages of decay (Appendices 1 and 2; Figs. 3 and 4). Observations undertaken subsequently indicated that species composition in leaf baits studied during this succession study was similar to that of naturally occurring leaves. In contrast, the species composition differed significantly in naturally occurring leaves of other tree species in the same rain forest (B. Paulus, pers. obs.). Two models describe the changes in diversity of microfungi during succession (Dix & Webster 1985, Neville & Webster 1995). According to the first model, “pioneer communities” have a low species diversity while “mature communities” are species rich. Eventually, species diversity may decline toward an “impoverished community” (Dix & Webster 1985). In contrast, Neville and Webster (1995) proposed that species diversity is richest and number of “individuals” greatest during earliest stages of colonization and following a period of stability, diversity and total numbers begin to decline. In our study, we observed both patterns depending on the methodology applied. Direct isolation data showed a slow but steady decline in species numbers, occurrence, and Shannon’s diversity index with advancing decay (Fig. 5) as predicted by Neville and Webster (1995). In contrast, diversity measures for the particle filtration cohort appeared to increase with advancing decay (Figs. 6 and 7), thus mirroring the prediction of Dix and Webster (1985). Contradictory results have been also observed in previous studies. For example, Kuter (1986) noted a marked increase in species diversity with advancing leaf decay using an indirect isolation protocol, while Hogg and Hudson (1966) observed an increase in species abundance with a direct method and a decrease with an indirect 47 isolation protocol. A satisfactory explanation for these observed differences is still lacking. From our observations, we hypothesize that with advancing decay leaves may accumulate greater numbers of propagules as earlier fungi may form dormant resting structures (Griffin 1972, Neville & Webster 1995). In contrast, only metabolically active mycelia-producing fruiting structures would be observed by the direct method. In addition, the indirect method could detect mycelia of fungal species that are not adapted to the substratum and thus would have limited vegetative growth without sporulation (D.J. Lodge, pers. comm.). Investigating the ecology of microfungi, particularly those on leaf litter of tropical rain forest trees, can be a daunting task due to the great number of species and the limitations of currently available methods to rapidly and reliably detect fungal diversity. The most effective approach may be to maximize the information obtained from tightly defined samples of greatest similarity using standardized methods (Colwell & Coddington 1994, Cannon 1999). The study of fungal succession meets this objective and has the potential to elucidate patterns and generate hypotheses. Improving our understanding of the processes that underlie fungal succession and other aspects of fungal ecology, especially in the tropics, will require further testing utilizing autecological approaches (e.g., Frankland 1998, Jones & Hyde 2002). ACKNOWLEDGMENTS The following institutions are gratefully acknowledged for providing funding for this project: the Centre for Research of Fungal Diversity at the Department of Ecology & Biodiversity, University of Hong Kong, the Cooperative Research Centre for Rainforest Ecology and Management and the School of Tropical Biology, James Cook University. BP would also like to thank C. Pearce, I. Steer, and U. Hiller, and J. Hiller for their company and help on collection trips, and B. Bussaban and H. Leung for assistance in obtaining literature. L. 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Fungal succession on bamboo in Hong Kong. Fungal Divers. 10: 213–227. Succession of Microfungi on Ficus pleurocarpa 49 APPENDIX 1. Percent occurrence of fungi observed on decaying leaves of Ficus pleurocarpa using a direct observational method in North Queensland, Australia (ten subsamples per sampling time). Higher taxonomic ranks are indicated in brackets next to the species names. “A” indicates Ascomycota and “B” Basidiomycota; anamorphic fungi are denoted by “C” for coelomycetes and “H” for hyphomycetes. Fungal code Species name F579 Acanthophyses-like structures (B) F473 Acremonium spp. (H) F556 Anthostomella reniformis (A) F569 Asterina sp. (A) F548 Basidiomycete A (B) F573 Basidiomycete B (B) F530 Beltrania concurvispora (H) F509 Beltrania rhombica (H) F518 Beltraniella portoricensis (H) F456 Brooksia tropicalis (A) Green leaves Day 0 Day 7 Day 14 6.2 5.5 2.8 5.1 4.6 4.6 F515 Chaetospermum camelliae (C) F495 Chaetosphaeria sp. (A) F552 Circinotrichum maculiforme (H) F589 Cladosporium sp. 1 (H) F590 Cladosporium sp. 2 (H) F455 Coelomycete A (anamorph F472) (C) F531 Coelomycete B (C) F465 Coelomycete C (C) 1.5 0.9 9.2 9.2 1.8 9.2 2.8 F520 Colletotrichum sp. 3 (C) F588 Curvularia sp. (H) Cylindrocladiella elegans (H) F489 Cylindrocladium colhounii var. colhounii (H) 3.1 F524 Cylindrocladium floridanum (H) 3.1 Cylindrocladium ilicola (H) 0.9 0.9 F505 Cylindrosympodium cryptocaryae (H) F549 Cylindrosympodium variabile (H) F484 Dactylaria belliana (H) F535 Dactylaria ficusicola (H) F587 Dactylaria sp. (H) F508 Gliocladiopsis tenuis (H) F481 Gliocladium ?solani (H) F511 Gliocladium sp. 2 (H) F544 Gliocladium sp. 3 (H) 10.7 14.8 1.5 0.9 5.1 6.2 13.3 0.9 1.3 3.1 1.7 6.4 11.4 1.5 0.9 1.8 0.9 Cylindrocladium retaudii (H) Falcocladium sp. (H) 1.7 3.4 3.7 F493 Gaeumannomyces-like sp. (A) 1.3 2.5 1.7 F559 F543 7.4 0.9 F532 F468 1.8 1.7 Colletotrichum sp. 1 (C) Dermateaceae (gen. nov.) (A) 7.1 3.7 7.7 Colletotrichum sp. 2 (C) Discostroma ficusicola (A) 5.0 3.7 F462 F528 4.6 3.4 F463 F472 5.1 0.9 0.9 Dictyochaeta sp. 2 (H) 5.1 0.9 3.1 Chaetopsina fulva (H) F533 12.5 0.9 F517 Dictyochaeta sp. 1 (H) 11.7 Day 94 1.7 0.9 F507 9.2 1.5 Cephalosporiopsis sp. 1 (H) Dactylella sp. (H) Day 78 1.7 Cephalosporiopsis sp. 2 (H) Dendrosporium lobatum (H) Day 62 1.3 F482 F572 Day 46 3.6 F491 F586 Day 30 0.9 3.7 1.7 1.8 0.9 3.8 5.1 12.3 16.7 8.9 7.4 3.7 1.5 3.7 0.9 2.5 3.7 1.3 4.6 7.3 12.7 1.5 16.9 1.8 1.5 3.1 3.4 7.3 2.8 6.3 4.6 2.8 1.7 1.3 2.8 1.3 1.5 1.8 1.7 4.6 6.7 1.5 1.7 5.4 3.4 1.3 F480 Gliomastix ?macrocerealis (H) 0.9 F557 Gliomastix elasticae (H) 0.9 F558 Guignardia sp. (A) 1.5 F503 Hansfordia pulvinata (H) 1.5 F584 Helicosporium griseum (H) 4.6 1.7 1.8 22.2 50 Paulus, Gadek, and Hyde APPENDIX 1. Continued. Fungal code Species name F574 Helicosporium sp. (H) F570 Hysteriaceous ascomycete (A) F563 Idriella cagnizarii (H) F475 Idriella lunata (H) F521 Idriella ramosa (H) F540 Idriella sp. (H) F467 Lanceispora amphibia (A) F525 Leptodothierella state of Guignardia sp. (C) F550 Leptothyrium sp. (C) F464 Meliola sp. (A) F561 Microdochium phragmites (H) F582 Microdochium sp. (H) F541 Myrothecium ?lachastrae (H) F458 Nigrospora panici (H) F486 Ochroconis humicola (H) F513 Ophiognomonia elasticae (A) F564 Parasympodiella sp. (H) F496 Penicillium sp. 1 (H) F453 Penicillium sp. 2 (H) F457 Pestalotiopsis ?breviseta (C) F502 Phoma sp. 1 (C) F527 Phoma sp. 2 (C) F461 Phomopsis sp. 1 (C) Green leaves Day 0 Day 7 Day 14 4.6 3.7 0.9 0.9 3.4 7.7 3.1 1.8 1.7 1.5 0.9 3.7 5.1 1.7 3.1 1.5 1.8 15.4 9.2 3.1 0.9 1.3 1.8 1.5 1.5 6.4 2.8 2.5 F478 Phomopsis sp. 4 (C) 5.5 1.8 1.8 1.5 F512 Selenodriella fertilis (H) 5.5 F504 Selenosporella cristata (H) 0.9 1.7 5.1 3.1 F560 Sphaeridium sp. (H) Sporodesmium-like gen. nov. (H) 1.5 F466 Sterile brown mycelium with chlamydospores 1.5 F506 Subulispora procurvata (H) F566 Thozetella falcata (H) 3.1 F567 Trichoderma sp. (H) 1.5 1.5 0.9 F485 Trichoderma viride (H) 0.9 Trichothecium sp. (H) 0.9 F578 Unidentified ascomycete (A) F450 Unidentified ascomycete (A) F547 Unidentified hyphomycete (H) F538 Unidentified hyphomycete (H) Verticillium sp. (H) Volutella ramkurii (H) F546 Wiesneriomyces javanicus (H) F519 Wiesneriomyces sp. (H) F534 Xenogliocladiopsis-like gen. nov. (H) F471 Zygosporium echinosporium (H) F460 Zygosporium mansonii (H) 5.4 1.8 F483 F474 1.7 1.8 F449 F510 1.7 1.8 3.8 Speiropsis pedatospora (H) 7.4 12.5 1.5 5.5 Sphaeridium pilosum (H) 7.1 6.7 6.3 0.9 F542 3.3 1.7 3.7 0.9 F580 Day 94 1.8 Phomopsis sp. 2 (C) Polyscytalum sp. (H) Day 78 11.9 Phomopsis sp. 3 (C) Pseudospiropes pinarensis (H) Day 62 1.5 1.8 F454 F501 Day 46 4.6 F497 F451 Day 30 1.3 1.8 1.7 1.8 12.3 1.8 3.4 1.3 4.6 1.3 1.5 6.4 1.7 2.8 5.1 3.1 1.8 1.5 8.3 3.7 1.3 7.3 0.9 1.3 0.9 1.8 10.1 0.9 0.9 1.3 1.5 1.8 1.7 10.2 3.1 3.3 1.8 5.1 10.8 11.7 3.6 11.1 8.5 1.5 5.0 1.8 7.4 Associated microorganisms F487 Actinomycete 1 F492 Actinomycete 2 4.6 Succession of Microfungi on Ficus pleurocarpa APPENDIX 2. Percent abundance of fungi found on fallen leaves of Ficus pleurocarpa during succession process in North Queensland, Australia [replicate particle filtrations for each sampling period with exception of green leaves (GL)]. GLa Species Acremonium sp. 1b Aureobasidium pullulans Day 0 Day 46 0.7 0.7 Beltrania rhombicab Beltrania concurvisporab Beltraniella portoricensisb 18.5 0.7 3.6 5.9 0.6 sp.b 0.7 0.6 1.2 7.3 Cylindrocarpon sp. Cylindrocladiella elegansb Cylindrosympodium cryptocaryaeb Continued. GLa Species Phomopsis sp.b Unidentified ascomycete Ab Unidentified ascomycete Bb Unidentified hyphomycete A Unidentified hyphomycete B Unidentified hyphomycete C 0.7 0.7 0.6 3.0 Day 0 0.6 0.9 0.9 2.1 1.4 0.7 0.7 Verticillium sp.b Xenogliocladiopsis-like gen. nov.b Xylariaceous sp. 1 9.2 Xylariaceous sp. 2 Xylariaceous sp. 3 4.6 3.7 Xylariaceous sp. 4 Xylariaceous sp. 5 Sterile mycelium 1 3.7 0.9 5.5 Sterile mycelium 2 Sterile mycelium 3 Sterile mycelium 4 4.6 3.7 2.8 1.8 0.9 2.4 Dactylaria sp. Dendrosporium lobatumb Dictyochaeta sp.b 0.6 1.8 Sterile mycelium 5 and 6 Sterile mycelium 7–23 Sterile mycelium 24 0.6 3.0 Sterile mycelium 25 Sterile mycelium 26 Sterile mycelium 27 6.8 3.4 2.7 Sterile mycelium 28 Sterile mycelium 29–32 Sterile mycelium 33 2.1 0.7 Geniculosporium sp. Geotrichum sp. Gliocladiopsis tenuisb 0.9 Gliocladium sp. 2b Glomerella sp. and anamorphb 4.6 Guignardia sp. 1b Guignardia sp. 2 ?Hansfordia sp. 1.8 0.9 3.7 0.7 0.6 1.2 Idriella lunatab Idriella sp.b Lanceispora amphibiab 17.8 0.6 1.2 Lemonniera terrestris Microdochium sp.b Ochroconis humicolab 0.6 0.6 0.7 3.0 0.7 Ochroconis sp. 2 Pestalosphaeria sp. Pestalotiopsis ?brevisetab Phoma sp. 1b Phoma sp. 2b 19.6 0.7 0.7 2.1 0.9 20.2 5.5 0.6 26.0 4.1 2.2 1.4 0.7 1.8 1.4 Day 94 0.7 Cylindrosympodium variabileb Dactylaria bellianab Dactylaria ficusicolab 1.4 17.4 Day 46 13.7 Unidentified hyphomycete D Unidentified hyphomycete E Unidentified hyphomycete F 0.6 Cephalosporiopis sp. 1b Cephalosporiopis sp. 2b Cladosporium Coelomycete Bb Cylindrocladium retaudiib Day 94 APPENDIX 2. 51 8.2 11.8 Sterile mycelium 34 Sterile mycelium 35 and 36 4.1 3.6 Sterile mycelium 37–40 Sterile mycelium 41 Sterile mycelium 42–63 2.4 1.8 0.6 Sterile mycelium 64 Sterile mycelium 65–67 Sterile mycelium 69–74 8.7 2.9 1.4 Sterile mycelium 75–100 0.7 a Green leaf isolation differ in collection times and isolation method. also by direct isolation. b Identified