HOST MICROBE INTERACTIONS

Fungal Propagules and DNA in Feces of TwoDetritus-Feeding Amphipods

Kandikere Ramaiah Sridhar & Margaret Beaton &

Felix Bärlocher

Received: 28 April 2010 /Accepted: 16 July 2010 /Published online: 10 August 2010# Springer Science+Business Media, LLC 2010

Abstract Aquatic shredders (leaf-eating invertebrates)preferentially ingest and digest leaves colonized by aquatichyphomycetes (fungi). This activity destroys leaf-associatedfungal biomass and detritial resources in streams. Fungalcounter-adaptations may include the ability to survive passagethrough the invertebrate's digestive tract. When fecal pelletsof Gammarus tigrinus and Hyalella azteca were incubatedwith sterile leaves, spores of nine (G. tigrinus) and seven(H. azteca) aquatic hyphomycete species were subsequentlyreleased from the leaves, indicating the presence of viablefungal structures in the feces. Extraction, amplification, andsequencing of DNA from feces revealed numerous fungalphylotypes, two of which could be assigned unequivocallyto an aquatic hyphomycete. The estimated contributions ofmajor fungal groups varied depending on whether 18S orITS sequences were amplified and cloned. We conclude thata variable proportion of fungal DNA in the feces ofdetritivores may originate from aquatic hyphomycetes.Amplified DNA may be associated with metabolicallyactive, dormant, or dead fungal cells.

Introduction

Aquatic hyphomycetes are a polyphyletic group of fungi thatdominate decomposition of autumn-shed leaves in streams [6,

23]. Their growth “conditions” leaves and makes them morepalatable to invertebrate shredders [4, 6, 17, 43]. Shredderstypically prefer nitrogen-rich leaf species (e.g., alder) overlignin-rich leaves (e.g., oak or beech). These preferences canbe overcome by inoculating less-preferred leaves with anappropriate fungal species [9]. Invertebrates also differentiateamong patches on the same leaf colonized by different fungi[1]. No consistent correlations have been found betweenpalatability and leaf parameters such as toughness, proteincontent, or fungal biomass [24, 44]. However, the order ofpreference can be reproduced by extracts of fungal myceliaapplied to fungus-free leaf disks [40].

It is difficult to evaluate shredder preferences for certainfungus–leaf combinations in streams. Examination of gutcontents reveals primarily vascular plant remains, fine organicor mineral particles, and animal remains (e.g., [3]). Fungalhyphae or spores are rare and cannot usually be identified.Immunological assays may result in more detailed informa-tion (e.g., [34]), especially when based on monoclonalantibodies [25]. Bermingham et al. [14, 15] developedseveral monoclonal antibodies specific to aquatic hyphomy-cetes. However, this approach works best for tracing the fateof a few chosen species or for documenting predation onspecific life stages [41, 46]. It is less suitable when trying toestablish the full range of prey items. For this purpose,characterization of DNA fragments in guts, feces, orregurgitated bird pellets are preferred [29]. This typicallyinvolves extraction of DNA, amplification of selected geneswith group-specific primers, and analysis of PCR productsthrough fingerprinting techniques (e.g., denaturing gradientgel electrophoresis, terminal restriction fragment lengthpolymorphism) or cloning and sequencing. In principle, thiscan provide unambiguous evidence that tissue of one specieshas passed through the gut of another species. However,none of the DNA-based techniques can distinguish between

K. R. SridharDepartment of Biosciences, Mangalore University,Mangalagangotri,Mangalore 574 199 Karnataka, India

M. Beaton : F. Bärlocher (*)Department of Biology, Mt. Allison University,Sackville, NB E4L1G7, Canadae-mail: fbaerlocher@mta.ca

Microb Ecol (2011) 61:31–40DOI 10.1007/s00248-010-9732-4

predation and secondary predation [46], nor do they provideany information on the condition of the organism providingthe DNA. It may have been dead before it was consumed bya scavenger or it may have been killed by a predatorimmediately before ingestion. Alternatively, some preytissues or cells may still be viable after passing through thedigestive tract of the consumer, for example, in the form ofinvertebrate eggs, plant seeds, or fungal spores and sclerotia(e.g., [50]). To document the potential impact of leaf-eatinginvertebrates on aquatic hyphomycete communities, there-fore, requires knowing which fungal species are consumedand to what extent their reproductive structures survivepassage through the gut ([2]). To approach these objectives,we fed Hyalella azteca Saussure and Gammarus tigrinusSexton with leaves precolonized by a pure culture of one ofthree fungi. We analyzed their fecal pellets for fungalreproductive potential and for the presence of species-specific DNA sequences. The same analyses were performedon fecal pellets of field-collected individuals. In addition, thepellets of these invertebrates were analyzed by extractingand amplifying fungal DNA (SSU rDNA, ITS), followed bycloning and sequencing.

Materials and Methods

Collection Site

Decaying leaf litter and invertebrates were collected in theSackville Waterfowl Park (Sackville, New Brunswick, Can-ada; N latitude 45° 54′, W longitude 64° 80′; [26]). It consistsof 19 ha of shallow (0.3–1 m depth) freshwater wetlands and3 ha of adjacent uplands. The park has a shoreline length of2.4 km and drains a 207-ha watershed. The water level iscontrolled by a 10-m weir that incorporates a 2-m stop-logsection to enable manipulation of water levels. The water isgenerally neutral to slightly alkaline and has been characterizedas eutrophic, with phosphorus being a limiting factor forprimary production [26]. Lesser duckweed (Lemna minor),pondweed (Potamogeton spp.), and algae are the dominantfloating or submerged vegetation. Wild rice (Zizaniaaquatica), bur-reed (Sparganium eurycarpum), and cattail(Typha spp.) are the dominant emergents. Purple loosestrife(Lythrum salicaria) has been present since 1963. The mainriparian trees species are alder (Alnus rugosa (Du Roi)Spreng.), birch (Betula papyrifera Marsh.), willow (Salixnigra Marsh.), maple (Acer saccharum Marsh.), andAmerican mountain ash (Sorbus americana Marsh.).

Naturally decaying leaf litter of riparian trees wascollected from the Waterfowl Park in May and July, 2009.The leaves were rinsed and portions corresponding toapprox. 200 mg dry mass were placed in separate 250 mlErlenmeyer flasks filled with 100 ml of sterile distilled

water. The flasks were placed on a shaker (130 rpm) andincubated at 16°C for 48 h. The suspension was thensucked through an 8-μm membrane filter. The leaf materialwas dried to constant weight at 50°C. The spores retainedon the filter were stained with cotton blue in lactophenol(50 mg l−1). The entire filter was scanned at ×200magnification, and fungal spores were identified andcounted. On both sampling dates, ten filters were scanned.

Preparation of Invertebrate Food

Autumn-shed leaves of sugar maple (A. saccharum L.) andelm (Ulmus americana L.) were collected from a tree on thecampus of Mt. Allison University, Sackville, NB, Canada.They were cut into 18-mm disks, which were leached for48 h under running tap water, air dried, and stored untilneeded. Before being used as food, leaf disks wereautoclaved and subsequently used as sterile food. Addi-tional maple leaf disks were used to prepare fungal-colonized substrates. Twelve maple leaf disks were placedin 250 ml Erlenmeyer flasks, autoclaved, and supplied with150 ml of autoclaved stream water (Boss Brook, Fenwick,Nova Scotia; [36]). Each flask was inoculated with a 5-mmagar plug overgrown with a pure culture of Articulosporatetracladia Ingold, Clavariopsis aquatica de Wild. orHeliscus lugdunensis Sacc. & Thérry, all isolated fromsingle spores from foam collected in Boss Brook. The diskswere incubated for 14 days at 15°C on a shaker (130 rpm)before being fed to invertebrates.

Fungi in Fecal Pellets

Pellets as Inocula for Sterile Leaves Specimens ofG. tigrinus Sexton and H. azteca Saussure were collectedfrom the Waterfowl Park in Sackville (New Brunswick,Canada) in the last week of September and first week ofOctober 2008, and in July 2009. They were kept in aeratedwell water at 16°C and were provided with autoclavedmaple leaves (four animals in 150 ml water with six leafdisks per 250 ml Erlenmeyer flask). After 1, 7, 14, and28 days of feeding, visible fecal pellets were collectedaseptically with Pasteur pipettes at least every 2 h. Nofecal pellets that had been sitting in the flask for >2 h wereused. Pellets were added to three sterile leaf disks in100 ml of sterile stream water in 250 ml Erlenmeyerflasks. For each invertebrate/date combination, four flaskswere prepared and placed on a shaker for 14 days(130 rpm). Leaf disks were then harvested and aerated in100 ml of sterile distilled water for 48 h. The suspensionwas filtered through 8 μm membrane filters, which werestained with cotton blue in lactophenol. The filterswere scanned for spores of aquatic hyphomycetes, whichwere counted and identified.

32 K. R. Sridhar et al.

To further document viable structures of aquatic hypho-mycetes that had passed through invertebrate digestivetracts, specimens of G. tigrinus and H. azteca were kept insterilized stream water for 6 h without food. Twoconspecifics were then placed in 150 ml of sterilized streamwater in a 250 ml Erlenmeyer and provided with six mapleleaf disks inoculated with one of three fungal species(A. tetracladia, C. aquatic, or H. lugdunensis). Fecalpellets, produced 2 days after feeding had started, werecollected aseptically and analyzed for viable structures ofaquatic hyphomycetes (Table 3, Expt. 1) and for DNAspecific to the fungal strain present on the food (see below).Fecal pellets released in one flask over 2 days were addedto three disks of sterile leaf disks in 100 ml of sterile streamwater in 250 ml Erlenmeyer flasks. These flasks were puton a shaker for 14 days (130 rpm). Leaf disks were thenharvested, aerated in 100 ml of sterile distilled water for48 h, and dried to constant weight. The suspension wasfiltered through 8 μm membrane filters, which were stainedwith cotton blue in lactophenol. The filters were scannedfor spores of aquatic hyphomycetes, which were countedand identified. Results are expressed as conidia per day permilligram dry leaf mass. Four replicates were evaluated.

In a second experiment, feeding ofG. tigrinus and H. azteca(animals collected in July 15, 2009) with fungal-colonizedmaple leaves was continued for 14 days (Table 3, Expt. 2),with the food and water being replaced every third day. Fecalpellets were collected on day 15 and evaluated as describedabove.

Fungal DNA from Pellets of Amphipods Feeding on LeavesInoculated with a Pure Fungal Culture Pellets, produced2 days after feeding on leaves inoculated with a pure culture,were collected aseptically and analyzed for the presence ofDNA specific to the fungal strains present on the food.

Fungal DNA from Pellets of Freshly Collected AmphipodsSpecimens of both G. tigrinus and H. azteca were collectedand fed autoclaved leaves of U. americana. Fecal pelletswere collected at least every 2 h during the first 2 days andstored at −80°C (pellets left in the flasks for >2 h werediscarded). They were used for extraction and amplificationof fungal DNA, followed by cloning and sequencing.

Molecular Analyses

All DNA amplifications were performed in an iCycler(BioRad) with GoTaq™Green Master Mix (Promega,Madison, WI). Each analysis included a negative controlwith sterile buffer instead of DNA containing extract.

To evaluate fecal DNA in invertebrates feeding onautoclaved leaves inoculated with one of the pure cultures,

DNA was extracted with a Forensic DNA Purification Kit(ChargeSwitch®, Invitrogen; [13]). The manufacturer'sinstructions were followed, except that 150 μl of lysisbuffer (L13) and 2 l of proteinase K were used in the firststep. After binding the DNA and washing the beads, theDNA was eluted in 20 μl buffer, and 2 μl were transferredto a 0.2 ml Thermo Strip Tube with ABgene Flat Cap Strip(Thermo Scientific, USA). A nested PCR procedure wasused; the first with 1.0 μl of NS5 (forward) and LR16(reverse) primer, 10.0 μl of DEPC water (Invitrogen), and14.0 μl of Promega GoTaq® Hot Start Master Mix DNAPolymerase were added. The following cycling parameterswere used: initial denaturation, 2 min at 94°C; 38 cycles ofdenaturation, 45 s at 94°C, annealing 45 s at 55°C, extension1.5 min at 73°C; final extension 10 min at 73°C. After the firstPCR, 1.0μl of the product was mixed with 8.0μl DEPCwater(Invitrogen), 1.0μl of forward primer ITS1 and reverse primerITS4, and 14.0 μl of Promega GoTaq® Hot Start Master MixDNA Polymerase. The cycling parameters were the same asfor the first PCR. The final products were sent to GenomeQuebec Innovation Centre at McGill University for sequencing(http://www.genomequebecplatforms.com/mcgill/home/index.aspx).

To characterize the fungal diversity in pellets of freshlycollected invertebrates, a cloning–sequencing approach wasused [12]. Individual animals were placed in cups containingautoclaved well water and supplied with an excess ofautoclaved leaves as a food source. Over the next 2 days,feces were collected from each individual and transferred tosterile 1.5 ml microcentrifuge tubes and stored at −20°C.

Genomic DNA was extracted from the fecal materialusing the Soil DNA extraction kit (MoBio Laboratories,Solana Beach, CA) following the manufacturers' protocols.In separate amplifications, partial SSU rRNA and ITSsequences were targeted. For SSU sequences, PCR wasperformed with the fungal-specific primers AU2 and AU4[12, 48] using the following protocol: (1) initial denaturation,94°C, 2 min; (2) denaturation, 94°C, 30 s; (3) annealing,51°C, 30 s; (4) extension, 70°C, 90 s; (5) repeat (2)–(4) for35 cycles; (5) final extension, 70°C, 5 min; (6) pause at 4°Cuntil retrieved. For ITS sequences, the primers ITS4 andITS1F were used with the following protocol: (1) initialdenaturation, 94°C, 2 min; (2) denaturation, 94°C, 30 s; (3)annealing, 51°C, 30 s; (4) extension, 73°C, 90 s; (5) repeat(2)–(4) for 35 cycles; (5) final extension, 73°C, 5 min; (6)pause at 4°C until retrieved.

ATOPO TA Cloning®Kit For Sequencing (Invitrogen) wasused to clone the amplified inserts. Selective Luria–Bertani(LB) plates were prepared with 70 μg ml−1 ampicillin (SigmaAldrich) and spread with 70 μl of Ultra Pure X–GAL(20 mg ml−1 in DMF, Invitrogen). Escherichia coli cells werethawed on ice and transformed according to the One Shot®Chemical Transformation Protocol. Volumes of 15, 30, and

Fungal Propagules and DNA in Feces of Detritus-Feeding Amphipods 33

50 μl were spread onto LB plates and incubated overnight at37°C. Sterilized wooden toothpicks were used to transferclones from a single white colony onto a labeled grid plate ofLB media containing ampicillin and to a 0.2-μl tube for PCRamplification of the inserted fragment. PCR reactions wereperformed as described above. The final products were sent toGenome Quebec Innovation Centre at McGill University forsequencing. Where forward and reverse sequences wereavailable, consensus sequences were created with Se-Alv2.0. from fasta formats uploaded as text files. Discrepancieswere resolved by verifying the chromatogram for bothforward and reverse sequences. Consensus sequences weresubmitted to GenBank. Taxonomic relationships wereestablished by using the BLAST search option for “short,exact nucleotide matches”. “Closest hits” were subdi-vided into four major fungal categories: uncultured,Ascomycetes, Basidiomycetes, and Chytridiomycetes.

Results

Aquatic Hyphomycetes on Naturally Decaying Leaves

In total, ten species of aquatic hyphomycetes wererecovered from naturally decaying leaves in the WaterfowlPark (Table 1). Five species were found on both dates.Spore numbers released per day per milligram dry leafmaterial reached a maximum of 88±14 (average of tensamples, ±SD) in the July sample.

Fecal Pellets as Inocula for Sterile Leaves

Amphipods Feeding on Sterile Leaves The fecal pelletsof freshly collected G. tigrinus and H. azteca after 1 day offeeding on sterile leaves contained viable structures ofaquatic hyphomycetes that were able to colonize sterile

leaves and produce active conidiophores on these leaves(Table 2). After feeding on sterile maple leaves for 7 days,both amphipods still produced pellets that resulted in sporeproduction by five (G. tigrinus) and four (H. azteca)aquatic hyphomycetes. Inoculation of sterile leaves withpellets from amphipods that had been on a sterile diet for 14or 28 days no longer resulted in spore production. Two ofthe species present on fecal pellets were not found on leaves(collected in May and July), and two species identified fromleaves were not recovered from the feces (animals collected inSeptember/October and July; Tables 1 and 2).

Amphipods Feeding on Leaves Inoculated with a PureFungal Culture Three aquatic hyphomycete species growingaxenically on maple leaves retained the ability to colonizesterile substrates and sporulate after having passed through thedigestive tracts ofG. tigrinus or H. azteca (Table 3, Expt. 1).Without exception, they were the most prolific sporeproducers on the target leaves, but up to seven additionalspecies (on G. tigrinus, feeding on H. lugdunensis inoculatedleaves) were present as well, suggesting that up to 2 days offeeding on leaves with a monoculture had not completelydisplaced aquatic hyphomycete species acquired on earlierfood. In a second experiment, where feeding with fungal-colonized leaves was continued for 14 days, only the speciesprovided with the food was found (Table 3, Expt. 2).

Molecular Analyses

Amphipods Feeding on Leaves Inoculated with a PureFungal Culture When amphipods were fed with fungal-colonized leaves for up to 2 days following field collection,their fecal pellets generally contained enough amplifiablefungal DNA to produce clear bands on the gel. However,attempts to sequencing failed except for one test withH. azteca feeding on C. aquatica. In a second series ofexperiments, where animals were allowed to feed for14 days on leaves inoculated with a pure culture,unequivocal sequences were found in 4 out of 18 attempts(3 attempts for shredder/fungus combination; once eachwith G. tigrinus or H. aquatica on C. aquatica; twice withH. azteca on Articulospora aquatica). This suggests thatthe amplicons contained mixtures of fungal sequences,some presumably present in the gut before feeding onleaves colonized by a pure culture began.

Freshly Collected Amphipods Table 4 lists the 161 DNAsequences recovered from fecal pellets of freshly inverte-brates collected in September/October 2008 and feeding onautoclaved leaves for up to 2 days. Only two could beassigned to an aquatic hyphomycete species, A. tetracladia(H. azteca, ITS region).

Table 1 Aquatic hyphomycetes identified from spores released byleaves collected in the Waterfowl Park (2009)

5 May 1 July

Anguillospora filiformis Greath. + +

A. longissima Ingold +

Anguillospora sp. + +

Alatospora acuminata Ingold +

Articulospora tetracladia Ingold + +

Clavariopsis aquatica Ingold + +

Flagellospora curvula Ingold + +

Heliscus lugdunensis Sacc. & Thérry +

Lemonniera terrestris Tubaki +

Varicosporium elodeae Kegel +

34 K. R. Sridhar et al.

The taxonomic distribution of closest blast hits for thefour sources of clones (SSU and ITS regions for G. tigrinusand H. azteca) is summarized in Table 5. Sequences withidentical closest hits were placed in a common set (Table 5,column 1); however, individual sequences within a setdiffering by >1 bp were submitted separately to GenBank.

We tested for the independent distribution of four majorfungal categories: uncultured, Ascomycetes (to which weadded the two aquatic hyphomycete clones), Basidiomycetes,and Chytridiomycetes among the four sequence categories(two invertebrates×two targeted sequences). Since severalcounts were still low (e.g., no close relatives of Chytridiomy-cetes were found among ITS sequences), a χ2 test was notappropriate, and we used a randomization test with the sum

of squared deviations from expected values as test statistics[7]. Distribution of the taxonomic groups among sourcesdiffered significantly (p<0.0001). We also tested for inde-pendence between the combined data for G. tigrinus vs. H.azteca (p=0.053) and the combined data for SSU vs. ITSsequences (p<0.0001).

Discussion

The main habitat and substrate of aquatic hyphomyceteswere discovered by Ingold [27]. Most limnologistsremained unaware of their existence until the importance

Table 3 Sporulation rates of aquatic hyphomycetes from maple leaves (per day per milligram dry leaf mass) exposed to fecal pellets from G.tigrinus or H. azteca fed on leaves with Clavariopsis aquatica (CLAQ), Articulospora tetracladia (ARTE), or Heliscus lugdunensis (HELU)

G. tigrinus inoculated with H. azteca inoculated with

ARTE CLAQ HELU ARTE CLAQ HELU

Expt. 1 Anguillospora filiformis 0.04±0.99 0.02±0.04 0.04±0.09 0.04±0.09 0.02±0.04

A. longissima 0.08±0.2 0.04±0.09 0.04±0.08

ARTE 6.7±4.7 0.3±0.4 0.2±0.2 19.5±22.2 0.08±0.1 0.06±0.05

CLAQ 1.1±0.9 0.3±0.1

Heliscus tentaculus Umplett 0.05±0.1 0.06±0.1

Flagellospora curvula Ingold 0.04±0.05

HELU 4.6±2.5 3.6±2.6

Lemonniera aquatica 0.08±0.2 0.2±0.5

L. centrosphaera 0.06±0.1 0.4±0.9

Expt. 2 ARTE 8.5±2.7 12.6±9.8

CLAQ 5.3±4.9 1.3±2.5

HELU 7.2±5.1 6.8±5.5

Averages ± SD, n=10

Expt. 1: collection in October 2008, pellets collected after 2 days of feeding

Expt. 2: collection in July 2009, pellets collected after 14 days of feeding

G. tigrinus H. azteca

1 7 1 7

Anguillospora filiformis Greath. + + + +

A. longissima (Sacc. & Syd.) Ingold + + +

Anguillospora sp. + +

Articulospora tetracladia Ingold + + + +

Clavariopsis aquatica Ingold +

Clavatospora longibrachiata (Ingold) Marvanová & Nilsson +

Flagellospora curvula Ingold + + +

Heliscus lugdunensis Sacc. & Thérry +

Tricladium angulatum Ingold + +

Varicosporium elodeae Kegel + + + +

Total species number 9 5 8 4

Table 2 Aquatic hyphomycetesidentified from spores releasedfrom leaves inoculated withfeces from H. azteca or G.tigrinus

Feces were collected within1 day from freshly collected(October 2008), unfed animalsafter 1 day (1), or from animalsthat had been feeding on auto-claved maple leaves for 7, 14, or28 days. No spores were foundafter 14 and 28 days

Fungal Propagules and DNA in Feces of Detritus-Feeding Amphipods 35

Table 4 Sets of SSU and ITS sequences recovered from G. tigrinus and H. azteca fecal pellets associated with closest BLAST hits

Set # Clones Accession (clone sequence) Closest hit Accession (closest hit) Taxonomic group

GtSSU1 3 6963–6964 Bullera taiwanensis AB072234.1 BAS

GtSSU2 1 6965 Cryptococcus gastricus DQ645513 BAS

GtSSU3 1 6966 Cryptococcus skinneri AB032646.1 BAS

GtSSU4 3 6967–6969 Endochytrium sp. AY635844.1 CHY

GtSSU5 3 6970–6971 Filobasidium globisporum AB075546 BAS

GtSSU6 2 6972 Kriegeria eriophori DQ419918.1 BAS

GtSSU7 1 6973 Mrakia frigid DQ831017.1 BAS

GtSSU8 1 6974 Nowakowskiella sp. AY635835.1 CHY

GtSSU9 1 6975 Olpidium brassicae DQ322624.1 CHY

GtSSU10 1 6976 Powellomyces sp. AF164245.2 CHY

GtSSU11 2 6977–6978 Rhodotorula hordea AY657013.1 BAS

GtSSU12 2 6979–6980 Rhodotorula marina AB126645.1 BAS

GtSSU13 1 6981 Rhodotorula pinicola AB126652.1 BAS

GtSSU14 1 6982 Sporobolomyces falcatus AB021670.1 BAS

GtSSU15 1 6983 Sporobolomyces fushanensis AB176530.1 BAS

GtSSU16 2 6984 Taphrina pruni AJ495828.1 ASC

GtSSU17 1 6985 Tremellales sp. EU517058.1 BAS

GtSSU18 1 6986 Trimorphomyces papilionaceus AB085808.1 BAS

GtSSU19 2 NA Uncultured alveolate EU91606.1 OEU

GtSSU20 1 6987 Uncultured alveolate EU910604.1 OEU

GtSSU21 2 6988 Uncultured Chytridiomycete EU162638.1 CHY

GtSSU22 7 6989–6995 Uncultured eukaryote AY916571.1 OEU

GtSSU23 11 6996–7004 Uncultured fungus DQ244004.1 UFU

GtSSU24 1 7005 Uncultured fungus AM114806.1 UFU

GtSSU25 8 7006–7011 Uncultured eukaryote AJ130850.1 OEU

GtITS1 3 7012–7013 Mycosphaerella berberidis EU167603.1 ASC

GtITS2 5 7014–7018 Mycosphaerella graminicola EU019297.1 ASC

GtITS3 1 7019 Ramularia pratensis EU019284.1 ASC

GtITS4 1 7020 Septoria passerinii AF181700.1 ASC

GtITS5 3 7021 Sporobolomyces xanthus AF444547.1 BAS

GtITS6 2 7022 Uncultured Thelephoraceae AY945290.1 BAS

GtITS7 2 7023 Uncultured fungus EU516682.1 UFU

GtITS8 1 7024 Uncultured Helotiaceae FJ554107.1 ASC

GtITS9 3 7025 Uncultured Helotiales FJ553766.1 ASC

GtITS10 1 7026 Uncultured fungus EU480318.1 ASC

GtITS11 5 7027–7028 Uncultured Venturia FJ553066.1 ASC

GtITS12 1 NA Venturia hystrioides EU035459.1 ASC

HaSSU1 2 7029–7030 Cladochytrium replicatum AY546683.1 CHY

HaSSU2 1 7031 Cryptococcus aquaticus AB032621.1 BAS

HaSSU3 1 7032 Crytococcus sp. EF363152.1 BAS

HaSSU4 7 7033–7038 Endochytrium sp. AY635844.1 CHY

HaSSU5 1 7039 Kriegeria eriophori DQ419918.1 BAS

HaSSU6 9 7040–7044 Rhizophydium sp. AY635821.1 CHY

HaSSU7 1 7045 Rhodotorula marina AB126645.1 BAS

HaSSU8 1 NA Rhodotorula sp. FJ153119.1 BAS

HaSSU9 1 NA Trichosporon pullulans AB001766.2 BAS

HaSSU10 3 7046–7047 Uncultured alveolate EU910604.1 OEU

HaSSU11 4 7048–7049 Uncultured eukaryote AY916571.1 OEU

HaSSU12 2 7050 Uncultured fungus DQ244004.1 UFU

36 K. R. Sridhar et al.

of allochthonous plant litter for the stream food web wasrecognized, and Kaushik and Hynes [28] demonstrated thecrucial involvement of fungi in making deciduous leavesacceptable to invertebrates [5]. Even today, researchremains primarily focused on the fungal role in theprocessing of plant detritus [8]. Complementary aspects,such as how the fungi respond to invertebrate activity, havereceived less attention. Hynes was probably the first toobserve star-shaped spores of aquatic hyphomycetes in theguts of stone fly larvae [5]. Provided they survive passagethrough the digestive system, this zoochory might contributeto the fungi's ability to persist in a habitat characterized byunidirectional displacement, even though they lack activemovement. This possibility was first addressed byBärlocher [2]. In fecal pellets of Gammarus fossarum,conidia of eight species of aquatic hyphomycetes were

identified. On average, their ability to germinate was36.6% and varied between 0 (Flagellospora curvula) and74.5 % (H. lugdunensis). When fecal pellets were aerated,additional conidia belonging to seven species werereleased, 76.5% of which were able to germinate.

In the current study, we introduced an intermediate step totest for the presence of aquatic hyphomycetes: fecal pelletswere incubated with sterile leaf disks, allowing resistant fungalstructures to colonize a new substrate, presumably building uptheir biomass and then reproduce. A potential pitfall of thisapproach is contamination of fecal pellets before they wereanalyzed. We tried to minimize this by never collecting fecesthat had been sitting in the flasks for >2 h. Microscopicexamination of the pellets never revealed any attached conidia,and 14 attempts to extract and amplify fungal DNA from 20μlwater samples without pellets from experimental flasks werenot successful (unpublished data). We are therefore confidentthat the majority of our positive results were due to fungalstructures present in the fecal pellets.

In total, we recovered ten species of aquatic hyphomycetesfrom invertebrate feces (Table 2), which is comparable to theearlier study by Bärlocher [2]. Four species had never beenbefore recovered from fecal pellets (Anguillospora filiformis,an unknown Anguillospora species, Lemonniera terrestris,and Varicosporium elodeae). However, the habitat of thecurrent study is not optimal for aquatic hyphomycetes. Thewater is essentially lentic, without noticeable current. Thiswas reflected by the low number of spores released fromrandomly collected leaves; the maximum was 88 conidia permilligram leaf matter per day. In near-by streams, this canroutinely reach 2,000–4,000 (e.g., [36]), and other studieshave reported values >8,000 [22].

Table 4 (continued)

Set # Clones Accession (clone sequence) Closest hit Accession (closest hit) Taxonomic group

HaSSU13 1 7051 Uncultured zygomycete AJ506030.1 ZYG

HaSSU14 1 7052 Uncultured Tremellaceae EF023436.1 BAS

HaSSU15 2 7053 Uncultured eukaryote AJ130850.1 OEU

HaITS1 2 7054–7055 Articulospora tetracladia EU998923.1 AQH

HaITS2 2 7056 Mycosphaerella berberidis EU167603.1 ASC

HaITS3 10 7057–7060 Mycosphaerella graminicola EU019297.1 ASC

HaITS4 9 7061–7062 Uncultured Thelephoraceae AY945290.1 BAS

HaITS5 1 NA Uncultured fungus EF619900.1 UFU

HaITS6 2 7063 Uncultured Helotiaceae FJ554107.1 ASC

HaITS7 8 7064 Uncultured fungus EU480318.1 UFU

# Clones: clones within set giving either uni- or bidirectional sequence. Last four digits of accession numbers from HM486963–HM487064 aregiven for clones within the set where both forward and reverse sequences were recovered

Key: GtSSU1=G. tigrinus SSU rDNA, set 1; GtITS1=G. tigrinus ITS, set 1; HaSSU1=H. azteca SSU rDNA, set 1; HaITS1=H. azteca ITS, set 1.NA: only unidirectional sequence was obtained

Taxonomic group: AQH aquatic hyphomycetes; ASC Ascomycetes; BAS Basidiomycetes; CHY Chytridiomycetes; EUK other Eukarya; UFUuncultured fungus; ZYG Zygomycetes

Table 5 Taxonomic distribution of closest hits to cloned SSU rDNAand ITS sequences from fecal pellets of G. tigrinus and H. azteca

G. tigrinus H. azteca

Taxonomic group SSU ITS SSU ITS

Aquatic hyphomycetes 2

Uncultured fungi 12 5 2 9

Ascomycetes 2 20 14

Basidiomycetes 20 5 7 9

Chytridiomycota 8 18

Zygomycota 1

Other Eukarya 18 9

Total 60 30 37 34

Fungal Propagules and DNA in Feces of Detritus-Feeding Amphipods 37

Surprisingly, inocula for several aquatic hyphomycetespecies were still present in feces of invertebrates that hadbeen feeding on sterile leaves for up to 7 days. Thedigestive tracts of amphipods are essentially straight tubes,and gut passage times are relatively short, ranging between0.50–2 h and 12 h in Gammarus pulex [50]. In G.pseudolimnaeus, food retention was typically between0.75 and 2 h in the presence of food [11]; when food wasabsent, however, guts retained some food particles forseveral weeks. In our experiments, both amphipods wereobserved feeding and continued to release fecal pelletsthroughout the experiments, particularly when providedwith leaves conditioned by one of the three pure cultures.Surprisingly, even after 2 days on that diet, fecal pelletscontained inocula for several other aquatic hyphomycetes(Table 3, Expt. 1). This was no longer the case after 14 days(Table 3, Expt. 2). In both experiments, however, we wererarely able to amplify unambiguous sequences that could beconnected to the one species present on the food. It is likelythat the fecal pellets still contained other sources of fungalDNA which may or may not have been associated withaquatic hyphomycetes. Regardless, these observationssuggest that dispersal of aquatic hyphomycetes may befacilitated by their ability to remain viable during theirpassage through an amphipod gut. Comparable relation-ships exist between hypogeous ectomycorrhizal fungi,whose fruit bodies (truffles sensu lato, including ascomaor basidioma) are frequently consumed by small mammals.Spores within the truffle survive passage through thedigestive tract [16]. This mechanism may facilitate recolo-nization of new habitats by plants and their mycorrhizalpartners. Possible examples include the invasion by trees ofthe forefront of a receding glacier [19], of abandonedbeaver ponds [47], and areas altered by forest fires [20].

The incomplete digestion of fungal structures may alsoindirectly benefit leaf-shredding invertebrates by ensuring asufficient supply of fungal inocula to condition newlyimmersed leaves despite the unidirectional downstreamdisplacement by the current. The effect may be comparableto obligate fungal farming in ants, termites, and beetles[33], or, to the marine snail Littoraria irrorata feeding onlive Spartina alterniflora to prepare substrate for fungalgrowth [42]. In the case of stream amphipods, one might betempted to revive the concept of “prudent predators”.However, this behavior is generally not considered to beevolutionarily stable, and the same outcome can bepredicted more parsimoniously by natural selection actingindependently on prey and predator (e.g., [30], but see[39]). In addition, fungal conditioning, while beneficial, isnot an absolute requirement for most stream invertebrates[4, 24].

A major surprise of our study was the discordancebetween molecular and microscopic data. Five of the ten

aquatic hyphomycete species recovered from feces arerepresented in GenBank by both 18S and ITS sequencesand two more species by 18S sequences. But of the 161cloned sequences, only two could unequivocally beconnected to an aquatic hyphomycete (A. tetracladia;Table 4). Conceivably, some or most of the sequencesclosest to uncultured fungi or Ascomycetes (Table 5)belonged to aquatic hyphomycete taxa. The rest clearly donot. Forty-one were related to Basidiomycetes and 26 toChytridiomycetes (another 27 were assigned to otherEukarya). Members of both groups are rarely encounteredwhen leaf decomposition is complemented by the charac-terization of released spores, which favors the detection ofaquatic hyphomycetes), but they have repeatedly beenreported when molecular methods were applied (e.g., [8,12, 35, 36]). We therefore believe that these fungi are wellrepresented on fecal pellets, even though the scarcity ofaquatic hyphomycete sequences may at least partly be dueto biases in DNA extraction, amplification, and cloning togaps in the database and possibly to selective feeding bythe invertebrates. The fact that percentage distributionsamong the four major taxa varied between SSU and ITSsequences also suggest that our approach did not accurate-ly reflect their actual frequencies of occurrence in thefeces. Clearly, more extensive studies, including calibra-tion of known fungal mixtures with a variety of primers,are needed.

The molecular methods used do not allow us todistinguish between metabolically active or inactive struc-tures or even between intact cells and cell fragments (sporeproduction proves the presence of viable propagules).Furthermore, they provide no information on whether theywere present on the ingested food or are part of expelled gutsymbionts. Chytridiomycetes and other zoosporic fungi arecommonly recovered from cellulosic baits exposed instreams and may well make a substantial contribution toleaf decomposition [21, 31, 32, 37, 38].

Basidiomycetous yeasts were also well representedamong our fungal sequences. Very little is knownconcerning their ecology [49]. They occur in marine andfreshwater habitats, in the soil, on rhizospheres, and onplant surfaces such as tree bark, leaves, flowers, and fruits.Sporobolomyces and Rhodotorula spp. are major compo-nents of the phylloplane yeast populations, together withthe black yeast Aureobasidium pullulans, which wasfrequently isolated from senescent maple leaves [10], andcan survive passage through the gut of G. pulex [50].

Alternatively, fungal sequences found in fecal pelletsmay originate from gut symbionts. Suh et al. [45]isolated over 650 yeasts from a variety of beetles, atleast 200 of which were undescribed taxa. It is possiblethat they are also present in the guts of amphipods and otheraquatic invertebrates. Zoosporic fungi (rumen fungi;

38 K. R. Sridhar et al.

Neocallimastigales) inhabit the foreguts of ruminants. Sincethey are obligate anaerobes, it seems unlikely that they wouldpersist in amphipod guts with their relatively quick processingof food. Nevertheless, a more directed study might beworthwhile.

Traditional methods of food web analysis and trophicecology have relied on morphological identification oforganismal remains in guts or feces. These techniques haveoften given biased results due to inaccurate identifications andvariations in digestion rates [18]. Some of these limitationscan be mitigated by various immunological approachesusing polyclonal and monoclonal antibodies to detect proteinepitopes, and recently, developed polymerase chain reaction(PCR) based methods for detecting prey DNA [29, 51, 52].Symondson [46] concluded that PCR-based techniques arehighly effective and versatile and will likely displace allother approaches. Without calibration, however, we cannotassess if their relative frequencies truly reflect the contribu-tions of various food sources. In addition, they provide noinformation on whether recovered sequences were associatedwith metabolically active, viable cells.

Acknowledgements The National Engineering and Research Councilsupported this research through a discovery grant to FB.We are grateful toGeorge C. Carroll, University of Oregon, for advice concerning primers.

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