- Original article
- Open Access
Diversity and biochemical features of culturable fungi from the coastal waters of Southern China
AMB Express volume 4, Article number: 60 (2014)
Fungi play a major role in various biogeochemical cycles of terrestrial and marine ecosystems. However, fungi in marine environments remain to be one of the most under-studied microbial groups. This study investigates the diversity of planktonic fungi from the coastal habitat off Pearl River Delta (China) using culture-dependent approach. A total of 22 fungi and 9 yeast isolates were recovered from 30 seawater and 2 sediment samples. Microscopic and ITS rRNA gene sequence analyses revealed that most of the fungi belonged to the phylum Ascomycota and Basidiomycota with a very small percentage (3%) of the subphylum Mucoromycotina of the Phylum Zygomycota. Most of these fungal isolates exhibited considerable production of extracellular enzymes, cellulase, lipase and laccase. Fungal isolates of two genera Mucor and Aspergillus sp. demonstrated pelletization capability over a wide range of pH, suggesting them as potential agents towards algae harvesting and wastewater treatment.
Coastal marine habitats have been characterized as the most variable, highly diverse and rich in primary production (Jickells ; Danovaro and Pusceddu ). The primary production in coastal habitats sometimes reaches very high levels resulting into availability of a great fraction of organic matter for consumers as detritus even after consumption of herbivores (Newell ). Different types of microbes in coastal waters degrade a large proportion of this detritus actively (Manini et al. ; Pusceddu et al. ). Among these microbes, heterotrophic bacteria and archaea have been described for their degradation abilities towards such detritus to a greater extent (Moran and Miller ; Mou et al. ). In spite of being a significant component of coastal waters, the diversity and ecology of heterotrophic eukaryotes however has not been received much attention (Giovannoni and Stingl ; Hallam et al. ; Fenchel ; Strom ).
Among eukaryotes, fungi have been reported to exhibit as individual filaments or aggregates in coastal waters (Gutiérrez et al. ). However, in comparison with terrestrial environments, fungi in the world’s oceans remain largely unknown (Gao et al. ). Despite of a few reports on diversity of fungi from the oceans, the diversity and ecology of their planktonic forms (mycoplankton) have barely been explored (Richards et al. ). Mycoplankton include free-living filamentous fungi, yeasts, fungal-like protists, and those associated with planktonic particles or phytoplankton (Wang and Johnson ; Gao et al. ).
Fungi are a key component of the biosphere, fulfilling a wide range of biogeochemical and ecological functions in natural environments (Christensen ; Pang and Mitchell ). They are best known as decomposers of organic matter and play major role in nutrient regeneration in the detrital ecosystems. The filamentous mycelia may greatly enhance the efficient mineralization of particulate organic matter (Tisdall and Oades ; Damare and Raghukumar ) and thus benefit the growth of planktonic microbial communities (Kiørboe and Jackson ; Gutiérrez et al. ). The biomass of planktonic fungi has been reported to be comparable with prokaryotes including both Bacteria and Archaea (Gutiérrez et al. ).
Fungi occupy distinct ecological niches from that of bacterioplankton in detritus ecosystems with the ability to utilize large lignocellulose-predominated substrates with high C: N ratio (Newell ; Raghukumar ). They possess the ability to penetrate relatively persistent particulate detritus much more efficiently than bacterioplankton (Raghukumar ). The endophytic fungi have been demonstrated to reside within marine plants intra or intercellularly, and produce a variety of bioactive and chemically active metabolites (Kaul et al. ). The bioactive metabolites produced by endophytic fungi originate from different biosynthetic pathways and belong to different groups of terpenoids, steroids, quinones, phenols and coumarins (Kaul et al. ). Therefore, the endophytes represent a potential chemical reservoir for anticancer, antioxidant, antiviral and insecticidal compounds for pharmaceutical and agrochemical industries. Two new benzopyranones, diaportheone A and B, were obtained via bioassay-guided isolation of the secondary metabolites from the endophytic fungus Diaporthe sp. P133 isolated from Pandanus amaryllifolius leaves (Bungihan et al. ). These benzopyranones have been successfully used as antimicrobial compounds towards several microorganisms. One of the fungal isolate belonging to Aureobasidium pullulans has been exhibited as a reservoir of biotechnologically active products in previous reports (Chi et al. ).
Considering the crucial role of planktonic fungi in versatile oceanic biogeochemical cycles, their diversity needs to be addressed from different ecosystems. The coastal habitats of Pearl River Delta, China have been highly productive ecosystems of China, being the major source for fishing industries. However, recently there has been an increased pollution level detected in these ecosystems which may further enhance detritus levels available for degradation. Therefore, the mycoplankton diversity studies from these still unexplored habitats may provide greater insight on potential fungal isolates playing significant role in ecological cycles of Pearl River Delta. This study is the first report on diversity of planktonic fungi based on culture-dependent approach from the coastal habitats of Pearl River Delta.. Potential fungal isolates were further investigated for the production of different extracellular enzymes such as laccase, lipase and cellulase in order to understand their active role in ecological cycles of coastal ecosystems.
Materials and methods
Sample collection and isolation of fungi
Seawater and sediment samples were collected from coastal marine habitats of Pearl River Delta region of China during March, 2012 (Table 1). These samples were carried in sterile, screw capped plastic bottles and bags immediately back to the laboratory for isolation. The isolation of fungi from the seawater samples was done within one hour of collection using the membrane filtration technique. Briefly, 15 ml triplicate water samples were filtered through sterile 0.45 μm cellulose ester membranes (Millipore, USA). These membranes were then placed on solid media plates, Malt Extract Agar (MEA), Sabouraud Dextrose Agar (SDA), Potato Dextrose Agar (PDA), Czapek Dox Agar (CDA) and Corn Meal Agar (CMA), supplemented with antibiotics (0.075% streptomycin and 0.05% ampicillin) to suppress bacterial growth. Sediment samples (0.1 g) were suspended in 10 ml sterile seawater and 100 μl of the resulting suspension was plated directly on the above media plates containing antibiotics. The plates were incubated at room temperature (28°C) and examined daily for the growth of fungi. Fungal colonies that developed were subcultured onto fresh MEA plate for pure, single colony isolation and identification. The identification of filamentous fungi was done by macroscopic and microscopic morphology (Additional file 1: Figure S1). Three promising strains (Rhodosporidium sp. PKU Y5, Rhodotorula sp. PKU Y7 and Cladosporium sp. PKU F16) have been deposited in China General Microbiological Culture Collection Center (CGMCC No. 2.5198, CGMCC No. 2.5199 and CGMCC No.3.17121).
Isolation and sequencing of ITS rRNA gene from the fungal isolates
All the isolated fungi were grown in MEB for 4–5 days for DNA isolation. Yeasts were grown in YPD (yeast extract peptone and dextrose) medium and shaken at 170 rpm for 3–4 days. Mycelia and cells were harvested, lyophilized and crushed in a mortar and pestle to fine powder. Isolation of DNA was carried out using the Ezup Soil DNA extraction kit (Sangon Biotech, China), following the manufacturer’s guideline. The small subunit ITS rRNA gene was amplified by polymerase chain reaction (PCR) in the DNA T100™ Thermal cycler (Bio-Rad, USA) using the ITS rRNA gene specific primers ITS1 (5’-TCCGTAGGTGAACCTGCGG-3’) and ITS4 (5’-TCCTCCGCTTATTGATATGC-3’) (White et al. ). One microliter of DNA (~25 ng) was added to 50 μl reaction volume containing 25 μl of Taq PCR mix (Generay, China) , 23 μl dd water and 5 pmoles of each primer. The PCR program was run for initial denaturation step at 95°C for 3 min, followed by 35 cycles of 1 min at 94°C, 0.5 min at 50°C and 1 min at 72°C, and a final extension at 72°C for 5 min. The PCR products were purified using Gel DNA extraction kit (NewTopBio, China). Amplified products were transformed into Escherichia coli DH5α cells (Invitrogen, Carlsbad, CA, USA), following the manufacturer’s instructions. Transformants were grown overnight at 37°C in Luria-Bertani broth containing 100 mg of ampicillin. The presence of insert was confirmed by PCR with M13 forward and reverse primers. One ml of the broth containing the clone was added to 25 ml of PCR reaction mixture. PCR protocol included an initial hot start incubation (5 min at 94°C) followed by 34 cycles of denaturation at 94°C for 30 s, annealing at 55°C for 30 s, and extension at 72°C for 1 min followed by a final extension at 72°C for 5 min. Clones containing positive insert were further processed for plasmid isolation and purification using Millipore plasmid preparation kit (Millipore, USA). Clones containing positive insert were sent to BGI (Shenzhen, China) for sequencing analysis using M13 primers.
Forward and reverse sequences were edited and assembled using Chromas Pro version 1.34 (Technelysium Pty Ltd, Tewantia, Queensland, Australia). The final sequences were compared to the nucleotide sequences of reference organisms available in the GenBank database using Blastn (Altschul et al. ). The ITS1-5.8S-ITS4 gene sequences obtained for the organisms were aligned with their closest match using the program, ClustalW (Thompson et al. ). Gaps and ambiguously aligned sequences were removed manually from further analyses. Phylogenetic analyses were carried out using distance setting (Maximum parsimony) in MEGA 4 software (Tamura et al. ) with 1,000 bootstrap replicates. The resulting ITS1-5.8S-ITS4 gene sequences were submitted to GenBank under the accession number of KC113282-KC113312.
Qualitative assay for extracellular enzymes
The enzymatic activity of all the fungal isolates was analyzed in this study. The strains were screened initially using qualitative plate assay for three different enzymes (laccase, cellulase and lipase) by streaking them on the media plates, supplemented with specific substrates. Laccase activity was detected using MEA plates amended with ABTS (2, 2’-azino-bis-3-ethylbenzothiazoline-6-sulfonic acid) (Srinivasan et al. ). The fungal isolates were grown on these media plates at room temperature for 4 days. Green color produced around the fungal colonies on media plates indicated laccase activity.
Cellulase (CMCase) activity was detected using carboxymethylcellulose (CMC)-MEA plates (Carder et al. ). The CMC-MEA plates comprised 0.5% carboxymethylcellulose-sodium salt (CMC-Na), 1.0% glucose, 0.15% peptone, 0.01% yeast extract, 100% seawater, and 2.0% agar. After growing the fungal isolates for 3 days at room temperature, these plates were stained with 0.1% Congo red solution for 20 min at room temperature. The resulting plates were washed twice with 1.0 M NaCl, and were kept overnight at 4°C. Clear zones around the colonies indicated the CMCase activities (Nagano and Fraser ; Carder et al. ). Lipase activity was detected using MEA plates supplemented with 0.01% phenol red, 1% olive oil and 10 mM CaCl2. The pH was adjusted to 7.3-7.4 with 1.0 M NaOH (Singh et al. ). After incubation for 3 days, a change in color from pink to yellow indicated the lipase activity.
Quantitative analysis of enzymatic activities
On the basis of qualitative screening, four fungal strains (PKU F16, PKU F18, PKU Y5 and PKU Y8) were selected for quantitative studies. Fungal inoculums were prepared by growing isolates in MEB medium and yeast in YPD medium, respectively, for four days at 28°C. These inoculums were further subcultured to fresh MEB medium containing individual enzyme specific substrates and incubated on shaker at 30°C, 150 rpm for 4 days. All these experiments were performed in triplicates. Individual enzymes were quantified in the supernatants of the isolates.
The laccase activity was assayed using glycine-HCl (pH3.0) buffer and ABTS as substrate (Niku-Paavola et al. 1988). Five hundred μl of crude culture filtrate was incubated with equal volumes of buffer containing ABTS and laccase activity was measured at 405 nm. The enzyme units were expressed as μM of substrate transformed per minute per liter of culture filtrate i.e. as enzyme units per liter of culture filtrate (UL−1). In the absence of the enzyme activity, no increase in the rate of absorbance was observed.
Cellulase activity was assayed following the method described by Raghukumar et al. (). A reaction mixture containing 200 μl culture filtrate and 200 μl of 0.5% CMC in 0.05 M sodium phosphate buffer, pH 7 was incubated at 37°C for 30 min. To terminate the reaction, 1 ml of DNS (dinitrosalicylic acid) reagent was added to above reaction mixture and then was boiled for 5 min. CMCase activity was measured at 575 nm. One unit of CMCase activity was defined as the amount of enzyme liberating 1 μM of reducing sugar per minute under the above assay conditions.
For lipase assay, 6 mg para-nitrophenyl palmitate (pNPP), dissolved in 2 ml isopropanol and 18 ml 50 mM sodium phosphate buffer (pH 8) was used as the substrate. One ml each of culture filtrate and sodium phosphate buffer was added to 1 ml of the pNPP substrate solution. After incubating at 37°C for 30 min, the released product, pNP (para-nitrophenol) was measured at 410 nm. One unit of lipase activity was defined as the amount of enzyme liberating 1 μM of pNP per minute under these assay conditions.
Fungal pelletization experiment
Three of the fungal isolates (Mucor sp., PKU F1), (Aspergillus sp., PKU F8) and (Cladosporium sp., PKU F14) were investigated for pellet formation in this study. The spore suspension from these species was obtained by rinsing the mycelia on media plate with distilled water containing 10% Tween 20. The number of spores in the suspension was counted using an optical microscope (Olympus BX53 Manual fluorescence microscope). The spore suspensions were added to the 250-mL Erlenmeyer flasks containing 50 ml of nutrient media (Malt Extract Broth). The pellet formation experiments were performed by placing these Erlenmeyer flasks on a horizontal shaker (100 rpm) at room temperature for 4 days. The pellet formation was analyzed at different spore concentrations and pH ranges (2–8).
Fungi were isolated from all the seawater depths in the present study (Table 1). However, maximum numbers of fungal colonies were recovered from surface sediments (0 m depth) and seawater samples at 10 m depth. Fungi were also recovered from sediment samples using particle plating technique (Table 1). SDA was found to be better media than MEA for the isolation of fungi (Table 1). However, there was no statistical significance observed between different depths, media and the number of fungal colonies. A total of 22 fungi and 9 yeasts were isolated in this study. The fungal isolates mostly belonged to Ascomycota, Basidiomycota and Zygomycota based on ITS rRNA gene analysis (Table 2, Figure 1). Fungal isolates showing similarity with the phylum Ascomycota were dominating among above three, accounting for 74%. Members of Basidiomycota and Zygomycota made up for 23% and 3%, respectively (Figure 1). Of the Ascomycetes, isolates belonged to 13 genera i.e. Aspergillus, Hypocraea, Arthrinium, Diaporthe, Phoma, Trichoderma, Dothideomycetes, Cladosporium, Curvularia, Pleosporales, Pyrenochaeta, Aureobasidium and Candida. Isolates of Basidiomycota were affiliated with Rhodotorula, Rhodosporidium and Trichosporon sp. Only Mucor sp. of Zygomycota was identifed in this study (Table 2, Figure 2). All the fungal ITS rRNA gene sequences showed 100 or 99% identity with the existing sequences of NCBI database except PKU F18, showing 92% identity with Ascomycota sp. AR-2010 (Table 2).
Qualitative analyses of different enzymes demonstrated positive result for most of the fungal isolates (Table 3, Figure 3). The majority of fungal isolates (~84%) exhibited cellulase and lipase activities. In comparison, only a few of the isolates (~38%) showed positive laccase activity (Table 3, Figure 3). The quantitative analyses for production of these three enzymes were done for four of these fungal isolates (Table 4). Cladosporium sp. (PKU F16), Rhodosporidium sp. (PKU Y5) and Candida sp. (PKU Y8) showed considerable production of lipase enzyme, reaching up to 21.94 U ml−1 for Candida sp. In contrast, cellulase production was not very high for any of these isolates, being maximum of 1.3 U ml−1 demonstrated by Candida sp. PKU F16 (Cladosporium sp.) and PKU F18 (Ascomycota sp.) were the best producers of laccase enzyme showing activities of 14.6 U ml−1 and 10.5 U ml−1 respectively (Table 4). All the isolates showed maximum production of these extracellular enzymes on 6th day of their growth (Table 4).
Fungal pellet formation
Among three fungal isolates i.e. Aspergillus sp. (PKU F8), Mucor (PKU F1) and Cladosporium sp. (PKU F14), PKU F1 was found to be the best agent for pellet formation (Table 5, Additional file 1: Figure S2). The pH range of 6 and 8 were most suitable for all of these four isolates for pellet formation. Pellet sizes were comparative bigger with higher concentration of spore inoculums for Mucor sp. (Table 5). However, Aspergillus sp. did not show much variation with inoculum spore concentrations (Table 5). Maximum size of pellet of 7 mm was demonstrated by Aspergillus sp. (PKU F8) at pH 8.
Diversity of fungi has been reported from various environments such as freshwater (Gulis et al. ), marine environments such as coastal waters (Gao et al. ), deep-sea sediments (Damare et al. ; Singh et al. ), hypersaline waters (Buchalo et al. ), methane hydrates (Lai et al. ), oxygen deficient ecosystems (Cathrine and Raghukumar ), mangroves and salt marshes (Hyde et al. ; Raghukumar ), and hydrothermal vents (Le Calvez et al. ). This study is the first report on culturable diversity of fungi from coastal ecosystems of Pearl River Delta, China.
Despite of being isolated from coastal environment, the diversity of fungi was observed to be comparatively less, resulting into a total of 22 filamentous fungi and 9 yeast isolates. The isolated fungi belonged mostly to Ascomycota, Basidiomycota and at a very less percentage of Zygomycota (Table 2). Previous studies have also reported abundance of above fungal phylotypes in coastal seawater ecosystems (Gao et al. ). The results obtained in the present study are also in concordance with earlier reports where surface water of coastal ecosystems has been reported to contain higher mycoplankton diversity compared with open-ocean ecosystems (Gao et al. ). However, use of only culture-dependent approach in this study for estimation of diversity may have limited the isolation of other genera of fungi which are present as uncultivable forms in the oceanic habitats. Previous studies have reported a greater percentage of fungal diversity when assayed using combined approach of culture-dependent and culture-independent methods (Singh et al. ). Therefore, a detailed study on fungal diversity from Pearl River Delta using both the approaches may provide a greater insight on hidden, still unexplored mycoplankton communities in future. The diversity of fungi has also been reported to vary with the nutrient contents of any particular habitat, which can be available as detritus for the fungal population (Newell ).
Isolation of fungi was attempted using five different media (MEA, SDA, CMA, PDA and CMA). However, fungal isolates were recovered only on two media i.e., MEA and SDA (Table 3). In contrast, the fungal isolates were obtained with all the above five media during isolation from deep-sea sediments by Singh et al. (). Most of the isolates affiliated with the existing sequences of NCBI data base at the percentage identity of 100 or 99. However, the 92% identity of the fungal sequence PKU F18 with Ascomycota sp., suggests its probability of being novel. On the contrary, the insufficient database for ITS sequences may also be one of the reasons for such low similarity values (Zachow et al. ; Anderson et al. ).
The production of extracellular enzymes i.e., laccase. cellulase and lipase was demonstrated by most of the fungal isolates in the present study, suggesting their active role in various ecological cycles of coastal ecosystems off China. Laccase in one of the important lignin degrading enzymes, demonstrated to decolorize a range of dyes and toxic industrial effluent in earlier reports (Nyanhongo et al. ). The fungal isolates PKU F16 and PKU F18, showing considerable quantitative production of laccase in the present study render them possible candidates for industrial application (Table 4). Cellulose is the most common substrate present in the seawater column in the form of plant biomass. It is found in nature exclusively in plant cell walls, although it is produced by some animals also e.g., tunicates and few bacteria (Lynd et al. ). Fungi are well known agents of decomposition of organic matter composed of cellulosic substrate in particular (Lynd et al. ). Therefore, cellulase production by the fungal isolates in the present study suggest their active involvement in the mineralization and leaf litter degradation in the coastal sea water habitats. Lipases are ubiquitous enzymes of considerable industrial and physiological significance. They catalyze the hydrolysis of triacylglycerols to glycerol and free fatty acids. Lipases are widely used in the processing of fats and oils, detergents and degreasing formulations, food processing, the synthesis of fine chemicals and pharmaceuticals, paper manufacture, and production of cosmetics, and pharmaceuticals (Rubin and Dennis [1997a], [b]; Kazlauskas and Bornscheuer ). Lipase production up to 21.9 U ml−1 by Candida sp. in the present study reveals the potential of this fungus towards lipolytic degradation abilities. Additionally, the optimization of media and nutrient condition for enhanced lipase production from above isolate may be applied in future to maximize their industrial application and commercialization.
Fungi have been characterized widely for their efficient role in pellet formation towards harvesting of algae and wastewater treatment (Zhou et al. ). The microalgae cells can be processed into a broad spectrum of biofuels by the transesterification process. These biofuels include biodiesel, green diesel and gasoline, being produced by transformation of algal biomass using various technologies (Chisti ). However, many challenges have restricted the development of algal biofuel technology to commercial practicality that could allow for sustainable production and utilization (Brennan and Owende ). Among these, one is the harvesting process of algae, which can be improved by application of agents, causing aggregation of algal cells. The pellet formation capabilities exhibited by three of the fungal isolates in the present study (Table 5, Additional file 1: Figure S2) opens new avenues for their efficient utilization towards algae harvesting and wastewater treatment. In addition, the pellet formation capabilities of fungi have been shown to be affected widely by various factors such as pH, inoculum concentration and trace metals (Zhou et al. ). A co-cultivation method using fungi of the present study with algal strains under different pH, trace metals and inoculum concentration for efficient optimization of the harvesting process is suggested for future studies.
In conclusion, the fungal diversity obtained from this study was low. The fungal isolates belonged to three major phyla i.e., Ascomycota, Basidiomycota and Zygomycota, with Ascomycota being the dominant forms. The different qualitative as well as quantitative levels of extracellular enzymes produced by these isolates suggests them as significant component of the ecological cycles of coastal ecosystems off Pearl River Delta. Finally, the production of enzymes and pellet formation abilities of a few fungal isolates indicate their possible utilization in biotechnological industries.
Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ: Basic local alignment search tool. J Mol Biol 1990, 215: 403–410. 10.1016/S0022-2836(05)80360-2
Anderson IC, Campbell CD, Prosser JI: Potential bias of fungal 18S rDNA and internal transcribed spacer polymerase chain reaction primers for estimating fungal biodiversity in soil. Environ Microbiol 2003, 5: 36–47. 10.1046/j.1462-2920.2003.00383.x
Brennan L, Owende P: Biofuels from microalgae-a review of technologies for production, processing, and extractions of biofuels and co-products. Renew Sustain Energy Rev 2010, 14: 557–577. 10.1016/j.rser.2009.10.009
Buchalo AS, Nevo E, Wasser SP, Volz PA: Newly discovered halophilic fungi in the Dead Sea (Israel). In Journey to diverse microbial worlds. Edited by: Seckbach J. Kluwer, Dordrecht, The Netherlands; 2000:239–252. 10.1007/978-94-011-4269-4_17
Bungihan ME, Tan MA, Kitajima M, Kogure N, Franzblau SG, Cruz TEE, Takayama H, Nonato MG: Bioactive metabolites of Diaporthe sp. P133, an endophytic fungus isolated from Pandanus amaryllifolius . J Nat Med 2011, 65: 606–609. 10.1007/s11418-011-0518-x
Carder KL, Steward RG, Paul JH, Vargo GA: Relationships between chlorophyll and ocean color constituents as they affect remote-sensing reflectance models. Limnol Oceanogr 1986, 31: 403–413. 10.4319/lo.1986.31.2.0403
Cathrine SJ, Raghukumar C: Anaerobic denitrification in fungi from the coastal marine sediments off Goa, India. Mycol Res 2009, 11: 100–109. 10.1016/j.mycres.2008.08.009
Chi Z, Wang F, Chi Z, Yue L, Liu G, Zhang T: Bioproducts from Aureobasidium pullulans , a biotechnologically important yeast. Appl Microbiol Biotechnol 2009, 82: 793–804. 10.1007/s00253-009-1882-2
Chisti Y: Biodiesel from microalgae. Biotechnol Adv 2007, 25: 294–306. 10.1016/j.biotechadv.2007.02.001
Christensen M: A view of fungal ecology. Mycol 1989, 81: 1–19. 10.2307/3759446
Damare V, Raghukumar S: Abundance of thraustochytrids and bacteria in the equatorial Indian Ocean, in relation to transparent exopolymeric particles (TEPs). FEMS Microbiol Ecol 2008, 65: 40–49. 10.1111/j.1574-6941.2008.00500.x
Damare S, Raghukumar C, Raghukumar S: Fungi in deep-sea sediments of the Central Indian Basin. Deep-Sea Res Part I 2006, 53: 14–27. 10.1016/j.dsr.2005.09.005
Danovaro R, Pusceddu A: Biodiversity and ecosystem functioning in coastal lagoons: does microbial diversity play any role? Estuar Coast Shelf Sci 2007, 75: 4–12. 10.1016/j.ecss.2007.02.030
Fenchel T: The microbial loop-25 years later. J Exp Mar Biol Ecol 2008, 366: 99–103. 10.1016/j.jembe.2008.07.013
Gao Z, Johnson ZI, Wang G: Molecular characterization of the spatial diversity and novel lineages of mycoplankton in Hawaiian coastal waters. ISME J 2010, 4: 111–120. 10.1038/ismej.2009.87
Giovannoni SJ, Stingl U: Molecular diversity and ecology of microbial plankton. Nature 2005, 437: 343–348. 10.1038/nature04158
Gulis V, Kuehn K, Suberkropp K: The role of fungi in carbon and nitrogen cycles in freshwater ecosystems. In Fungi in biogeochemical cycles. Edited by: Gadd GM. Cambridge University Press, New York; 2006:405–435.
Gutiérrez MH, Pantoja S, Quiňones RA, González RR: First record of filamentous fungi in the coastal upwelling ecosystem off central Chile. Gayana 2010, 74: 66–73.
Gutiérrez MH, Pantoja S, Tejos E, Quiňones RA: Role of fungi in processing marine organic matter in the upwelling ecosystem off Chile. Mar Biol 2011, 158: 205–219. 10.1007/s00227-010-1552-z
Hallam SJ, Mincer TJ, Schleper C, Preston CM, Roberts K, Richardson PM, DeLong EF: Pathways of carbon assimilation and ammonia oxidation suggested by environmental genomic analyses of marine Crenarchaeota. PLoS Biol 2006, 4: 520–536. 10.1371/journal.pbio.0040095
Hyde KD, Jones EBG, Leano E, Pointing SB, Poonyth AD, Vrijmoed LLP: Role of fungi in marine ecosystems. Biodivers Conserv 1998, 7: 1147–1161. 10.1023/A:1008823515157
Jickells TD: Nutrient biogeochemistry of the coastal zone. Science 1998, 281: 217–222. 10.1126/science.281.5374.217
Kaul S, Gupta S, Ahmed M, Dhar MK: Endophytic fungi from medicinal plants: a treasure hunt for bioactive metabolites. Phytochem Rev 2013, 11: 487–505. 10.1007/s11101-012-9260-6
Kazlauskas RJ, Bornscheuer UT: Biotransformations with lipases. In Biotechnology. Edited by: Rehm HJ, Pihler G, Stadler A, Kelly PJW. VCH, New York; 1998:37–192.
Kiørboe T, Jackson GA: Marine snow, organic solute plumes, and optimal sensory behaviour of bacteria. Limnol Oceanogr 2001, 46: 1309–1318. 10.4319/lo.2001.46.6.1309
Lai X, Cao L, Tan H, Fang S, Huang Y, Zhou S: Fungal communities from methane hydrate-bearing deep-sea marine sediments in South China Sea. ISME J 2007, 1: 756–762. 10.1038/ismej.2007.51
Le Calvez T, Burgaud G, Mahe S, Barbier G, Vandenkoornhuyse P: Fungal diversity in deep-sea hydrothermal ecosystems. Appl Environ Microbiol 2009, 75: 6415–6421. 10.1128/AEM.00653-09
Lynd LR, Weimer PJ, Van Zyl WH, Pretorius IS: Microbial cellulose utilization: fundamentals and biotechnology. Microbiol Mol Biol Rev 2002, 66: 506–577. 10.1128/MMBR.66.3.506-577.2002
Manini E, Fiordelmondo C, Gambi C, Pusceddu A, Danovaro R: Benthic microbial loop functioning in coastal lagoons: a comparative approach. Oceanolog Acta 2003, 26: 27–38. 10.1016/S0399-1784(02)01227-6
Moran MA, Miller WL: Resourceful heterotrophs make the most of light in the coastal ocean. Na Rev Microbiol 2007, 5: 792–800. 10.1038/nrmicro1746
Mou XZ, Sun SL, Edwards RA, Hodson RE, Moran MA: Bacterial carbon processing by generalist species in the coastal ocean. Nature 2008, 451: 708–711. 10.1038/nature06513
Nagano T, Fraser P: No-nonsense functions for long noncoding RNAs. Cell 2011, 145: 178–181. 10.1016/j.cell.2011.03.014
Newell RC: The energetics of detritus utilisation in coastal lagoons and nearshore waters. In Proceedings of International Symposium on Coastal Lagoos; 8–14 September 1981; Oceanologica Acta (Special issue). Edited by: Laserre P, Postma H. SCOR/IABO/UNESCO, Bordeaux; 1982:347–355.
Newell SY: Ecomethodology for organo-osmotrophs, prokaryotic unicellular versus eukaryotic mycelial. Microb Ecol 1994, 28: 151–157. 10.1007/BF00166803
Niku-Paavola ML, Karhunen E, Salola P, Raunio V: Lignolytic enzymes of the white-rot fungus Phlebia radiata. Biochem J 1988, 254: 877–884.
Nyanhongo GS, Gomes J, Gübitz GM, Zvauya R, Read J, Steiner W: Decolorization of textile dyes by laccases from a newly isolated strain of Trametes modesta. Wat Res 2002, 36: 1449–1456. 10.1016/S0043-1354(01)00365-7
Pang KL, Mitchell JI: Molecular approaches for assessing fungal diversity in marine substrata. Bot Mar 2005, 48: 332–347. 10.1515/BOT.2005.046
Pusceddu A, Dell’Anno A, Danovaro R, Manini E, Sara G, Fabiano M: Enzymatically hydrolysable protein and carbohydrate sedimentary pools as indicators of the trophic state of detritus sink systems: a case study in a Mediterranean coastal lagoon. Estuar 2003, 26: 641–650. 10.1007/BF02711976
Raghukumar S: The role of fungi in marine detrital processes. In Marine Microbiology. Edited by: Ramaiah N. Facets & Opportunities. National Institute of Oceanography, Goa, India; 2004:91–101.
Raghukumar S, Sharma S, Raghukumar C, Sathe-Pathak V, Chandramohan D: Thraustochytrid and fungal component of marine detritus. IV. Laboratory studies on decomposition of leaves of the mangrove Rhizophora apiculata Blume. J Exp Mar Biol Ecol 1994, 183(1):113–131. 10.1016/0022-0981(94)90160-0
Richards TA, Jones MD, Leonard G, Bass D: Marine fungi: their ecology and molecular diversity. Ann Rev Mar Sci 2012, 4: 495–522. 10.1146/annurev-marine-120710-100802
Rubin B, Dennis EA: Lipases: Part A. Biotechnology Methods in enzymology. Academic Press, New York; 1997a.
Rubin B, Dennis EA: Lipases: Part B. Enzyme characterization and utilizationMethods in enzymology. Academic Press, New York; 1997b.
Singh R, Gupta N, Goswami VK, Gupta R: A simple activity staining protocol for lipases and esterases. Appl Microbiol Biot 2006, 70: 679–682. 10.1007/s00253-005-0138-z
Singh P, Raghukumar C, Verma P, Shouche Y: Phylogenetic diversity of culturable fungi from the deep-sea sediments of the Central Indian Basin and their growth characteristics. Fungal Divers 2010, 40: 89–102. 10.1007/s13225-009-0009-5
Singh P, Raghukumar C, Verma P, Shouche Y: Fungal diversity in deep-sea sediments revealed by culture-dependent and culture-independent approaches. Fungal Ecol 2012, 5: 543–553. 10.1016/j.funeco.2012.01.001
Srinivasan C, Dsouza TM, Boominathan K, Reddy CA: Demonstration of laccase in the white rot basidiomycete phanerochaete chrysosporium BKM-F1767. Appl Environ Microbiol 1995, 61: 4274–4277.
Strom SL: Microbial ecology of ocean biogeochemistry: A community perspective. Science 2008, 320: 1043–1045. 10.1126/science.1153527
Tamura K, Dudley J, Nei M, Kumar S: MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol 2007, 24: 1596–1599. 10.1093/molbev/msm092
Thompson JD, Higgins DG, Gibson TJ: CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 1994, 22: 4673–4680. 10.1093/nar/22.22.4673
Tisdall JM, Oades JM: Organic matter and water-stable aggregates in grassland soils. J Soil Sci 1982, 33: 141–163. 10.1111/j.1365-2389.1982.tb01755.x
Wang GY, Johnson ZI: Impact of parasitic fungi on the diversity and functional ecocology of marine phytoplankton. In Marine Phytoplankton. Edited by: Kersey TW, Munger SP. Nova Science Publishers, Hauppauge, NY; 2009:211–228.
White TJ, Bruns T, Lee S: Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. PCR Protocols: a Guide to Methods and Applications 1990, 18: 315–322. 10.1016/B978-0-12-372180-8.50042-1
Zachow C, Berg C, Muller H, Meincke R, Komon-Zelazowska M, Druzhinina IS, Kubicek CP, Berg G: Fungal diversity in the rhizosphere of endemic plant species of Tenerife (Canary Islands): relationship to vegetation zones and environmental factors. ISME J 2009, 3: 79–92. 10.1038/ismej.2008.87
Zhou Y, Du J, Tsao GT: Mycelial Pellet Formation by Rhizopus oryzae ATCC 20344. Appl Biochem Biotechnol 2000, 84: 779–789. 10.1385/ABAB:84-86:1-9:779
Zhou WG, Cheng YL, Li Y, Wan YQ, Liu YH, Lin XY, Ruan R: Novel fungal pelletization assisted technology for algae harvesting and wastewater treatment. Appl Biochem Biotechnol 2012, 167: 214–228. 10.1007/s12010-012-9667-y
This work was partially funded by National Science Foundation of China grant 31170109 (GYW) and Shenzhen Development and Reform Commission grant 835 (GYW).
The authors declare that they have no competing interests.
GW & PS conceived and designed the experiments; LL & YL performed the experiments; PS & LL analyzed the data; GW & PS contributed reagents/materials/analysis tools; PS, LL, GW & SQP wrote the paper. All authors read and approved the final manuscript.
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Li, L., Singh, P., Liu, Y. et al. Diversity and biochemical features of culturable fungi from the coastal waters of Southern China. AMB Expr 4, 60 (2014). https://doi.org/10.1186/s13568-014-0060-9
- Marine-derived fungi
- Hydrolytic enzymes