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Environmental fungal diversity in the upwelling ecosystem off central Chile and potential contribution to enzymatic hydrolysis of macromolecules in coastal ecotones
Fungal Ecology 29 (2017) 90e95
Contents lists available at ScienceDirect
Fungal Ecology journal homepage: www.elsevier.com/locate/funeco
Environmental fungal diversity in the upwelling ecosystem off central Chile and potential contribution to enzymatic hydrolysis of macromolecules in coastal ecotones rrez a, b, *, Diana V. Garce s c, d, Silvio Pantoja a, b, Rodrigo R. Gonza lez a, b, Marcelo H. Gutie a , d, e ~ ones Renato A. Quin n, Concepcio n, Chile Department of Oceanography, University of Concepcio n, Concepcio n, Chile COPAS Sur-Austral, University of Concepcio Graduate Program in Oceanography, Department of Oceanography, University of Concepcion, Concepcion, Chile d n, Concepcio n, Chile Interdisciplinary Center for Aquaculture Research (INCAR), University of Concepcio e n Marina de Excelencia (PIMEX), Faculty of Natural and Oceanographic Sciences, University of Concepcio n, Concepcio n, Chile Programa de Investigacio a
b c
a r t i c l e i n f o
a b s t r a c t
Article history: Received 15 November 2016 Received in revised form 6 March 2017 Accepted 6 July 2017
Even though occurrence of fungi in several marine environments has been documented, their inclusion within the marine microbial loop is not fully recognized. A major constraint is whether fungi in coastal waters are truly marine or represent transient microorganisms transported from terrestrial environments. We approached this issue by analyzing ambient fungal composition and hydrolytic activity of culturable fungi along a nearshore-offshore gradient in the upwelling ecosystem off central Chile, a region of high marine productivity strongly influenced by river discharges. We detected different communities of fungi in nearshore and offshore waters, with near estuary strains hydrolyzing proteins and carbohydrates faster than those from offshore sites. We conclude that coastal waters off central Chile comprise distinct fungal communities representative of offshore and nearshore environments, and provide new evidence for fungi processing organic matter in coastal ecotones, opening a fresh perspective for disappearance of organics carried by rivers in the coastal ocean. © 2017 Elsevier Ltd and British Mycological Society. All rights reserved.
Corresponding Editor: Kevin D. Hyde Keywords: Fungal composition Coastal upwelling Nearshore-offshore gradient Enzymatic hydrolysis
1. Introduction Major roles in biogeochemical cycling of elements in marine ecosystems are attributed to bacteria (e.g. Fuhrman and Azam, 1982; Azam et al., 1983) and archaea (e.g. Wuchter et al., 2006; Levipan et al., 2007, Offre et al., 2013). Even though the occurrence of fungi has been reported for several marine environments, such as deep-sea sediments (Damare and Raghukumar, 2008), water and sediment in oxygen minimum zones (Jebaraj and rrez et al., 2011) and Raghukumar 2009; Jebaraj et al., 2010; Gutie rrez coastal oligotrophic (Gao et al., 2010) and upwelling (Gutie et al., 2010, 2016) waters, impact of fungi in ocean biogeochemistry is less acknowledged than in terrestrial environments (Fell and Newell, 1998; Clipson et al., 2006; Richards et al., 2012).
n, * Corresponding author. Department of Oceanography, University of Concepcio n. Chile. Concepcio rrez). E-mail address: [email protected] (M.H. Gutie http://dx.doi.org/10.1016/j.funeco.2017.07.002 1754-5048/© 2017 Elsevier Ltd and British Mycological Society. All rights reserved.
Although evidence of the role of marine filamentous fungi has been rrez et al., 2011), evidence is documented (Jebaraj et al., 2010; Gutie needed to assess their distribution in relation to marine physical, chemical and biological spatial gradients, which would help to determine whether fungal communities contribute significantly to processing carbon in the coastal ocean. We studied water column fungal diversity and activity of culturable fungi in a coastal area in the Humboldt Current System off n in central Chile, one of the most productive ecosystems Concepcio in the world with primary production rates >15 g C m2 d1 associated with upwelling during austral spring and summer (Montero et al., 2007). In this environment, we previously detected high fungal biomass during the productive season in the coastal upwelling ecosystem off central Chile and attributed to fungi a role rrez in breakdown of biologically derived organic polymers (Gutie et al., 2011). Since we are interested in elucidating whether fungi could be active in the coastal ocean and whether fungal communities contribute significantly to processing carbon in the coastal ocean in addition to other microbes, here we investigated: (a) how
M.H. Gutierrez et al. / Fungal Ecology 29 (2017) 90e95
operational taxonomic units (OTUs) of water column fungal rRNA genes distribute in relation to marine properties, and (b) the potential role of fungi in decomposition of organic matter by testing extracellular hydrolysis of biopolymers by culturable fungal strains. Extracellular hydrolysis of macromolecules is the initial reaction to degrade organic matter in the ocean (Hoppe et al., 2002; Arnosti, 2003; Pantoja et al., 2011) and as such it would point to a role in marine carbon biogeochemistry.
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various depths between 0 and 110 m below the surface, and surface sediment samples were obtained with a Van Veen Grab (sampling area 0.1 m2). One-liter water samples were filtered through 0.45 mm cellulose ester sterile filters to collect suspended particulate matter, and filters were immediately stored in liquid nitrogen for DNA analysis. Unfiltered seawater and sediment samples were stored in sterile containers at 4 C for isolation of fungi.
2.2. DNA extraction and PCR amplification of fungi 2. Materials and methods 2.1. Study site and sampling The study was conducted in the coastal zone of the southern Humboldt Current System off central Chile at ca. 36 S (Fig. 1). The n is characterized by a relatively continental shelf off Concepcio straight coastline and is limited by submarine canyons of the Itata River (average runoff of 300 m3 s1) to the north and Biobío River ~ ones and Montes, (average runoff of 900 m3 s1) to the south (Quin 2001). Continental shelf break is located ~60 km from the coastline at 150 m depth. Seawater and surface marine sediments were sampled during August 2007 (austral winter) onboard R/V Kay Kay II (Department n, Chile), at sampling sites of Oceanography, University of Concepcio A, B and C (Fig. 1), located 30, 15 and 2 km from the Itata River mouth. The water column was sampled with Niskin bottles at
Environmental DNA from filters was extracted using PowerSoil DNA Kit (MO BIO Laboratories, Inc.). Template DNA (1e3 mL) was subjected to standard PCR using general fungal primers NS1 and ITS4 (White et al., 1990). PCR parameters: initial denaturation of 2 min at 95 C, 30 amplification cycles of 94 C of 1 min denaturation, annealing/extension at 55 C for 4 min, and final extension of 5 min at 72 C. One mL of PCR product was used for nested PCR with NS1 and GC-Fung primers, which amplify a segment of small sub unit (SSU) 18s rDNA suitable for separation by denaturing gradient gel electrophoresis (DGGE) (May et al., 2001). Nested PCR parameters were initial denaturation for 2 min at 94 C, 35 amplification denaturation cycles at 94 C, annealing for 1 min at 50.3 C, extension for 1 min at 72 C, and extension for 5 min at 72 C. PCR reaction mixtures (50 mL) contained 200 mM each DNTP, 3.5 mM MgCl2, 0.4 mM of each primer, and one unit Taq DNA polymerase (GoTaq® Flexi DNA Polymerase, Promega).
Fig. 1. Location of sampling sites (red circles) in the upwelling ecosystem off central Chile adjacent to the Itata River (inset shows turbid river plumes as seen in a 250 m resolution MODIS true color image from August 21, 2008). The image was provided by M Sobarzo and G Saldías.
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2.3. Denaturing gradient gel electrophoresis of DNA fragments of fungi DGGE profiles of fungal community were developed on 8% acrylamide/bisacrylamide (37.5:1) gel with 20e60% denaturing gradient (1 mm thick, 1X TAE buffer, 20 cm 20 cm). Following polymerization, gel was placed in the buffer chamber of a DCode universal mutation detection unit (Bio-Rad) in 1X TAE buffer at 60 C. Nested PCR products (40 mL) were loaded onto gels with 10 mL 2X gel-loading dye and electrophoresis proceeded at 100 V and 60 C for 10 h. Gel was stained in a 0.5 mg L1 1X TAE/ethidium bromide solution and photographed on a UV transilluminator. DGGE band patterns were identified by analysis of digital image of gels (Quantity One, Bio-Rad). Similarity analysis of DGGE profiles was carried out using band presence and intensity, and clusters were produced from UPGMA (Unweighted Pair-Group Method with Arithmetic mean) statistical analysis. Richness (S) of operational taxonomic units (OTUs) was estimated as number of bands in DGGE gel. Statistical differences in OTUs composition among sampling sites were tested by PERMANOVA analysis. 2.4. Isolation of fungal strains and extracellular enzymatic hydrolysis Fungi were isolated and cultured with culture media used for marine fungi (Kohlmeyer and Kohlmeyer, 1979) and yeasts (Yarrow, 1998). Seawater aliquots (100e150 mL) filtered through sterile cellulose ester filters (0.22 mm) and subsamples of surface sediments (ca. 100 mg) were placed on solid culture media. Successive streaking and spreading on agar plates were carried out to select and isolate pure fungal colonies. Fifteen isolated strains from sites A, B, and C were assayed for extracellular activity of aminopeptidase and glucosidase (Hoppe, 1983); two strains from surface water (5 m) of each site, and 9 strains from surface sediments (2 from site A, 3 from site B, and 4 from site C). Five mL duplicate aliquots of cell suspensions prepared in filtered seawater (coastal seawater filtered through 0.2 mm) were incubated in darkness at 10 mM final concentration of fluorogenic substrates L-leucine-4-methylcoumarinyl-7-amide (MCA-Leu) for aminopeptidase activity and 4-methylumbelliferone b -D-glucoside (MUF-Glu) for glucosidase activity. Fluorescence was measured on subsamples removed at time zero and every hour for 6e7 h. Abiotic hydrolysis and adsorption were evaluated by incubating fluorogenic substrates in boiled (10 min) cells suspensions. Fluorescence was measured in a Turner fluorometer (TBS-380 Mini) using wavelength 365 nm for excitation and 455 nm for emission (Meyer-Reil, 1987). Calibration curves were constructed by measuring fluorescence of 7-amino-4methylcoumarin (MCA) and 4-methylumbelliferone (MUF) in sterile seawater at concentrations ranging between 0.03 and 0.5 mM. Rates of hydrolysis by fungal strains, calculated according to Pantoja and Lee (1994), were normalized to fungal carbon estimated from dry weight of fungal tissue using a conversion factor 1 g dry weight fungal biomass ¼ 0.35 g fungal carbon (FC) (Newell and Statzeif-Taliman, 1982). The Mann-Whitney non-parametric U test was applied to test the statistical differences between sampling sites for OTUs richness and extracellular enzymatic activity. All statistical analyses were carried out using the PAST software (Hammer et al., 2001).
A ¼ 8.2 ± 1.3, site B ¼ 12.4 ± 1.5, site C ¼ 12.3 ± 1.5) in waters of sites B and C, located at 15 and 2 km from the river mouth, than at offshore site A located 30 km from the river mouth (Fig. 2). Similarity analysis showed three distinctive water column fungal communities (ca. 50% similarity): (i) a community grouping surface coastal water of riverine influence (Site C; Fig. 3, Group 1), (ii) a community including marine waters below the halocline (>30 m depth in Sites A and B; Fig. 3, Group 2), and (iii) a surface water (<30 m depth) community from coastal marine sites (Fig. 3, Group 3). Segregation of the structure of the fungal community with respect to depth and distance from the coast has been reported for the Hawaiian coast (Gao et al., 2010) and along a saline gradient in estuarine waters (Burgaud et al., 2013). The warmest temperature was 12.6 ± 0.3 C between 5 and 15 m depth at the three marine sites, with cooler waters in the top 1 m, probably due to mixing with riverine waters (Fig. 4). Site C consistently showed the influence of the Itata River with lower salinity as low as 22.7 psu near the estuary, that contrasts with values of 34.5 ± 0.1 observed below 40 m depth (Fig. 4) and with salinities >34 measured in austral summer in the area (Pantoja et al., 2011). OTU composition in the water column provided evidence of distinctive communities in (a) waters near the Itata River mouth, and (b) in surface and (c) in subsurface waters of offshore sites (Fig. 3), suggesting the occurrence of fungal communities representative of offshore and nearshore environments. Both salinity values (Fig. 4) and a distinctive river plume (Fig. 1) are evidence of the Itata River favourably influencing the transport of riverine fungi to the coastal ocean. Average fungal richness decreased with distance from the coast (Fig. 2), suggesting that freshwater can be a source of fungi in these coastal waters, increasing the richness of the fungal community in the area under direct influence of river discharge. A previous study showed both a decrease in fungal diversity with salinity along the Delaware River and coastal ocean and a high relative abundance (37%) of halotolerant strains of terrestrial origin in the estuarine area (Burgaud et al., 2013). In addition, our hypothesis is also consistent with
3. Results and discussion 3.1. Compositional patterns of fungal community OTUs richness was 34% (4.2 OTUs) higher (Richness: site
Fig. 2. Vertical profile of OTUs richness of fungi derived from DGGE profiles in sites A, B and C. Significant statistical differences were observed between sites A and B (MannWhitney U ¼ 0, p-value ¼ 0.01) and A and C (Mann-Whitney U ¼ 7, p-value ¼ 0.03).
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Fig. 3. Dendrograms derived from similarity analysis of DGGE profiles along the nearshore-offshore gradient in the water column of the coastal area adjacent to the Itata River mouth. Composition of fungal communities was significantly different between clusters identified from OTUs similarity analysis (Pseudo-F ¼ 6.92, p-value ¼ 0.0001).
Fig. 4. Vertical profiles of temperature and salinity during austral winter 2007 at sites A, B, C.
findings of freshwater phylotypes of bacterioplankton inhabiting et al., 2000). However, coastal seawater (Crumb et al., 1999; Rappe others have found dominance of distinctive phylotypes of prokaryotes in freshwater and marine ecosystems (Bouvier and del Giorgio, 2002; Cottrell and Kirchman, 2004; Campbell and Kirchman, 2013). For the same study area in central Chile, freeliving prokaryote communities showed differences in
composition between fresh and coastal waters and a singular pattern of composition suggesting development of native prokaryote assemblages in estuarine zones (Levipan et al., 2012). For fungi, coastal transitions in community composition have been evidenced in tidal marshes along a salinity gradient in Rhode Island (Mohamed and Martiny, 2011), and in estuarine sediments of Delaware Bay (Burgaud et al., 2013).
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Fig. 5. Extracellular hydrolytic activity on protein- and carbohydrate-like substrates by fungal strains isolated from the water column and surface sediments. Hydrolysis rates were normalized by fungal carbon (FC). Significant statistical differences were observed between proteolytic activity of strains isolated from sites C and B (Mann-Whitney U ¼ 3, p-value ¼ 0.036).
In offshore waters (sites A, B in Fig. 1), fungal communities were distinct between the top 30 m depth and deeper waters (Fig. 3). Since this change coincided with the depth of the base of thermocline (ca. 30 m), we suggest that vertical segregation is associated with thermal stratification of the water column (Fig. 4). This vertical zonation correlated with physicochemical properties is characteristic of most microplankton in the ocean (e. g. Lalli and Parsons, 1996), suggesting a marine source for fungal communities in offshore waters. Consistently, prokaryote bacteria in offshore waters of the same study area showed vertical segregation in phylotypes (Levipan et al., 2012). 3.2. Extracellular enzymatic hydrolysis of fungal strains One hundred and six mycelial and 90 yeast colonies were isolated from seawater, with most of the colonies cultured from marine sites A and B, averaging 11 and 23 strains per depth versus 9 in site C. From surface sediments, 42 fungal and 10 yeast colonies were isolated, with the highest number of strains from site C. No yeasts grew in isolation media from sediments of sites A and B, located at 15 and 30 km from the river mouth. In addition, a higher average number of colonies per depth were cultured from waters of the top 30 m (20 ± 19) than from deeper waters (12 ± 6) in offshore sites, which is consistent with higher fungal biomass found in surface than deeper waters of the coastal upwelling ecosystem off rrez et al., 2010, 2011). Along the nearshorecentral Chile (Gutie offshore gradient, most culturable fungi from the water column were recovered from offshore sites A (11 ± 1) and B (23 ± 4) than site C (9 ± 0.6), whereas in surface sediments most fungal strains were isolated near the river mouth (site C ¼ 40 vs 4 and 8 in sites A and B respectively). All assayed strains hydrolyzed fluorogenic proteinaceous (MCALeu) and carbohydrate-like (MUF- Glu) substrates. Rates of hydrolysis of protein substrate were five times higher than those of carbohydrate substrates and hydrolysis rates of strains from site C near the estuary were generally higher (Fig. 5), suggesting that during winter culturable fungi inhabiting seawater and sediments of the coastal zone adjacent to the river mouth have the ability to rapidly process organic matter. This result is consistent with the traditional view of fungi as key decomposers in detrital trophic webs of coastal
ecotones (Fell and Newell, 1998; Hyde et al., 1998; Raghukumar 2005) and provides a new perspective for the disappearance of organics exported from soils by rivers in the coastal zone that, despite their apparent refractory nature, are poorly represented in both water column and sedimentary marine carbon pools (Hedges, 1992). Acknowledgments n Marina This study was funded by the Programa de Investigacio de Excelencia Pimex-Nueva Aldea (PIMEX; Universidad de Con n & Celulosa Arauco y Constitucio n S.A.) and COPAS Surcepcio ~ ones received also partial Austral CONICYT PIA PFB31. R. Quin funding from the Interdisciplinary Center for Aquaculture Research ~ ez, J Soto, and (INCAR; FONDAP 1511002). We thank R Castro, L Nun E Tejos for their help during field and laboratory work. We also thank the valuable guidance of Professor J Fell from University of rrez during a research visit in his Miami who hosted MH Gutie laboratory for training in marine mycology (Capacity Building Grant E-13852 FA-UDEC-WHOI). M Sobarzo and G Saldías kindly provided MODIS image of Fig. 1. We are very grateful to the HanseWissenschaftskolleg (HWK), Delmenhorst, Germany, for Fellowship awarded to S Pantoja. References Arnosti, C., 2003. Microbial extracellular enzymes and their role in DOM cycling. In: Findley, S., Sinsabaugh, R.S. (Eds.), Aquatic Ecosystems: Interactivity of Dissolved Organic Matter. iAcademic Press, California, pp. 315e342. Azam, F., Fenchel, T., Field, J.G., Gray, J.S., Meyerreil, L.A., Thingstad, F., 1983. The ecological role of water-column microbes in the sea. Mar. Ecol. Prog. Ser. 10, 257e263. Bouvier, T.C., del Giorgio, P.A., 2002. Compositional changes in free-living bacterial communities along a salinity gradient in two temperate stuaries. Limnol. Oceanogr. 47, 453e470. Burgaud, G., Woehlke, S., Redou, V., Orsi, W., Beaudoin, D., Barbier, G., Biddle, J.F., Edgcomb, V.P., 2013. Deciphering presence and activity of fungal communities in marine sediments using a model estuarine system. Aquat. Microb. Ecol. 70, 45e62. Campbell, B.J., Kirchman, D.L., 2013. Bacterial diversity, community structure and potential growth rates along an estuarine salinity gradient. ISME J. 7, 210e220. Clipson, N., Otte, M., Landy, E., 2006. Biogeochemical roles of fungi in the marine and estuarine habitats. In: Gadd, G.M. (Ed.), Fungi in Biogeochemical Cycles. Cambridge university press, pp. 436e461.
M.H. Gutierrez et al. / Fungal Ecology 29 (2017) 90e95 Cottrell, M.T., Kirchman, D.L., 2004. Single-cell analysis of bacteria growth, cell size, and community structure in the Delaware stuary. Aquat. Microb. Ecol. 34, 139e149. Crumb, B.C., Armbrust, E.V., Baros, J.A., 1999. Phylogenetic analysis of particuleattached and free-living bacterial communities in the Columbia River, its stuary, and the adjacent coastal ocean. Appl. Environ. Microb. 65, 3192e3204. Damare, S., Raghukumar, C., 2008. Fungi and macroaggregation in deep-sea sediments. Microb. Ecol. 56, 168e177. Fell, J.W., Newell, S.Y., 1998. Biochemical and molecular methods for the study of marine fungi. In: Cooksey, K.E. (Ed.), Biochemical and Molecular Methods for the Study of Marine Fungi. Chapman and Hall, London, pp. 259e283. Fuhrman, J.A., Azam, F., 1982. Thymidine incorporation as a measure of heterotrophic bacterioplankton production in marine surface waters: evaluation and field results. Mar. Biol. 66, 109e120. Gao, Z., Johnson, Z.I., Wang, G., 2010. Molecular characterization of the spatial diversity and novel lineages of mycoplankton in Hawaiian coastal waters. ISME J. 4, 111e120. rrez, M.H., Pantoja, S., Quin ~ ones, R.A., Gonza lez, R.R., 2010. First record of Gutie filamentous fungi in the coastal upwelling ecosystem off central Chile. Gayana 74, 66e73. rrez, M.H., Pantoja, S., Tejos, E., Quin ~ ones, R.A., 2011. The role of fungi in Gutie processing marine organic matter in the upwelling ecosystem off Chile. Mar. Biol. 158, 205e219. rrez, M.H., Jara, A.M., Pantoja, S., 2016. Fungal parasites infect marine diatoms Gutie in the upwelling ecosystem of the Humboldt current system off central Chile. Environ. Microbiol. 18, 1646e1653. Hammer, Ø., Harper, D.A.T., Ryan, P.D., 2001. PAST: paleontological statistics software package for education and data analysis. Palaeontol. Electron. 4, 9 pp. Hedges, J.I., 1992. Global geochemical cycles: progress and problems. Mar. Chem. 39, 67e93. Hoppe, H.G., 1983. Significance of exoenzymatic activities in the ecology of brackish water: measurements by means of methylumbelliferyl-substrates. Mar. Ecol. Prog. Ser. 11, 299e308. Hoppe, H.G., Arnosti, C., Herndel, G.F., 2002. Ecological significance of bacterial enzymes in marine environment. In: Burns, R.C., Dick, R.P. (Eds.), Microbial Enzymes in the Environment Activity, Ecology, and Applications. Marcel Dekker, Inc., New York, pp. 73e107. ~ o, E., Pointing, S., Poonyth, A.D., Vrijmoed, L., 1998. Role Hyde, K., Jones, E.B.G., Lean of fungi in marine ecosystem. Biodivers. Conserv. 7, 1147e1161. Jebaraj, C.S., Raghukumar, C., 2009. Anaerobic denitrification in fungi from the coastal marine sediments off Goa, India. Mycol. Res. 113, 100e109. Jebaraj, C.S., Raghukumar, C., Behnke, A., Stoeck, T., 2010. Fungal diversity in oxygen-depleted regions of the Arabian Sea revealed by targeted environmental sequencing combined with cultivation. FEMS Microbiol. Ecol. 71, 399e412. Kohlmeyer, J., Kohlmeyer, E., 1979. Marine mycology, the Higher Fungi. Academic press, New York. Lalli, C.M., Parsons, T.R., 1996. Biological Oceanography: an Introduction. Butterworth/Heinemann, Oxford, 320 pp. n, W.O., Saldías, G.S., 2012. Figerprinting analysis of the Levipan, H.A., Alarco
95
prokaryote community along a marine-freshwater transect in central-southern Chile. Ann. Microbiol. 62, 1121e1140. ~ ones, R.A., Urrutia, H., 2007. A time series of prokaryote secLevipan, H.A., Quin ondary production in the oxygen minimum zone of the Humboldt Current System, off central Chile. Prog. Oceanogr. 75, 531e549. May, L.A., Smiley, B., Schmidt, M.G., 2001. Comparative denaturing gradient gel electrophoresis analysis of fungal communities associated with whole plant corn silage. Can. J. Microbiol. 47, 829e841. Meyer-Reil, L.-A., 1987. Seasonal and spatial distribution of extracellular enzymatic activities and microbial incorporation of dissolved organic substrates in marine sediments. Appl. Environ. Microb. 53, 1748e1755. Mohamed, D.J., Martiny, J.B., 2011. Patterns of fungal diversity and composition along a salinity gradient. ISME J. 5, 379e388. rraga, L., Montero, P., Daneri, G., Cuevas, L.A., Gonz alez, H.E., Jacob, B., Liza Menschel, E., 2007. Productivity cycles in the coastal upwelling area off Con n: the importance of diatoms and bacterioplankton in the organic carbon cepcio flux. Prog. Oceanogr. 75, 518e530. Newell, S.Y., Statzeif-Taliman, A., 1982. Factors for conversion of fungal biovolume values to biomass, carbon and nitrogen: variation with mycelial ages, growth conditions, and strains of fungi from a salt marsh. Oikos 39, 261e268. Offre, P., Spang, A., Schleper, C., 2013. Archaea in biogeochemical cycles. Annu. Rev. Microbiol. 67, 437e457. Pantoja, S., Lee, C., 1994. Cell-surface oxidation of amino acids in seawater. Limnol. Oceanogr. 39, 1718e1726. rrez, M.H., Ampuero, P., Tejos, E., 2011. Degradation capability of Pantoja, S., Gutie the coastal environment adjacent to the Itata River in central Chile (36.5 S). Biogeosciences 8, 2063e2074. ~ ones, R.A., Montes, R., 2001. Relationship between freshwater input to the Quin coastal zone and the historical landings of the benthic/demersal fish Eleginops maclovinus in central-south Chile. Fish. Oceanogr. 10, 311e328. Raghukumar, S., 2005. The role of fungi in marine detrital processes. In: Ramaiah, N. (Ed.), Marine Microbiology: Facets & Opportunities. National Institute of Oceanography, Goa, pp. 91e101. , M.S., Vergin, K., Giovannoni, S.J., 2000. Phylogenetic comparisons of a coastal Rappe bacterioplankton community with its counterparts in open ocean and freshwater systems. FEMS Microbiol. Ecol. 33, 219e232. Richards, T.A., Jones, M.D.M., Leonard, G., Bass, D., 2012. Marine fungi: their ecology and molecular diversity. Annu. Rev. Mar. Sci. 4, 495e522. White, T.J., Bruns, T.D., Lee, S.B., Taylor, J.W., 1990. Analysis of phylogenetic relationships by amplification and direct sequencing of ribosomal DNA genes. In: Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols: a Guide to Methods and Applications. Academic Press, San Diego, pp. 315e322. Wuchter, C., Abbas, B., Coolen, M.J.L., Herfort, L., Van Bleijswijk, J., Timmers, P., Strous, M., Teira, E., Herndl, G.J., Middleburg, J.J., Schouten, S., Sinninghe Damste, J.S., 2006. Archaeal nitrification in the ocean. Proc. Nat. Acad. Sci. U. S. A. 103, 12317e12322. Yarrow, D., 1998. Methods for the isolation, maintenance and identification of yeasts. In: Kurtzman, C.P., Fell, J.W. (Eds.), The Yeasts, a Taxonomic Study. Elsevier, Amsterdam, pp. 77e100.
Contents lists available at ScienceDirect
Fungal Ecology journal homepage: www.elsevier.com/locate/funeco
Environmental fungal diversity in the upwelling ecosystem off central Chile and potential contribution to enzymatic hydrolysis of macromolecules in coastal ecotones rrez a, b, *, Diana V. Garce s c, d, Silvio Pantoja a, b, Rodrigo R. Gonza lez a, b, Marcelo H. Gutie a , d, e ~ ones Renato A. Quin n, Concepcio n, Chile Department of Oceanography, University of Concepcio n, Concepcio n, Chile COPAS Sur-Austral, University of Concepcio Graduate Program in Oceanography, Department of Oceanography, University of Concepcion, Concepcion, Chile d n, Concepcio n, Chile Interdisciplinary Center for Aquaculture Research (INCAR), University of Concepcio e n Marina de Excelencia (PIMEX), Faculty of Natural and Oceanographic Sciences, University of Concepcio n, Concepcio n, Chile Programa de Investigacio a
b c
a r t i c l e i n f o
a b s t r a c t
Article history: Received 15 November 2016 Received in revised form 6 March 2017 Accepted 6 July 2017
Even though occurrence of fungi in several marine environments has been documented, their inclusion within the marine microbial loop is not fully recognized. A major constraint is whether fungi in coastal waters are truly marine or represent transient microorganisms transported from terrestrial environments. We approached this issue by analyzing ambient fungal composition and hydrolytic activity of culturable fungi along a nearshore-offshore gradient in the upwelling ecosystem off central Chile, a region of high marine productivity strongly influenced by river discharges. We detected different communities of fungi in nearshore and offshore waters, with near estuary strains hydrolyzing proteins and carbohydrates faster than those from offshore sites. We conclude that coastal waters off central Chile comprise distinct fungal communities representative of offshore and nearshore environments, and provide new evidence for fungi processing organic matter in coastal ecotones, opening a fresh perspective for disappearance of organics carried by rivers in the coastal ocean. © 2017 Elsevier Ltd and British Mycological Society. All rights reserved.
Corresponding Editor: Kevin D. Hyde Keywords: Fungal composition Coastal upwelling Nearshore-offshore gradient Enzymatic hydrolysis
1. Introduction Major roles in biogeochemical cycling of elements in marine ecosystems are attributed to bacteria (e.g. Fuhrman and Azam, 1982; Azam et al., 1983) and archaea (e.g. Wuchter et al., 2006; Levipan et al., 2007, Offre et al., 2013). Even though the occurrence of fungi has been reported for several marine environments, such as deep-sea sediments (Damare and Raghukumar, 2008), water and sediment in oxygen minimum zones (Jebaraj and rrez et al., 2011) and Raghukumar 2009; Jebaraj et al., 2010; Gutie rrez coastal oligotrophic (Gao et al., 2010) and upwelling (Gutie et al., 2010, 2016) waters, impact of fungi in ocean biogeochemistry is less acknowledged than in terrestrial environments (Fell and Newell, 1998; Clipson et al., 2006; Richards et al., 2012).
n, * Corresponding author. Department of Oceanography, University of Concepcio n. Chile. Concepcio rrez). E-mail address: [email protected] (M.H. Gutie http://dx.doi.org/10.1016/j.funeco.2017.07.002 1754-5048/© 2017 Elsevier Ltd and British Mycological Society. All rights reserved.
Although evidence of the role of marine filamentous fungi has been rrez et al., 2011), evidence is documented (Jebaraj et al., 2010; Gutie needed to assess their distribution in relation to marine physical, chemical and biological spatial gradients, which would help to determine whether fungal communities contribute significantly to processing carbon in the coastal ocean. We studied water column fungal diversity and activity of culturable fungi in a coastal area in the Humboldt Current System off n in central Chile, one of the most productive ecosystems Concepcio in the world with primary production rates >15 g C m2 d1 associated with upwelling during austral spring and summer (Montero et al., 2007). In this environment, we previously detected high fungal biomass during the productive season in the coastal upwelling ecosystem off central Chile and attributed to fungi a role rrez in breakdown of biologically derived organic polymers (Gutie et al., 2011). Since we are interested in elucidating whether fungi could be active in the coastal ocean and whether fungal communities contribute significantly to processing carbon in the coastal ocean in addition to other microbes, here we investigated: (a) how
M.H. Gutierrez et al. / Fungal Ecology 29 (2017) 90e95
operational taxonomic units (OTUs) of water column fungal rRNA genes distribute in relation to marine properties, and (b) the potential role of fungi in decomposition of organic matter by testing extracellular hydrolysis of biopolymers by culturable fungal strains. Extracellular hydrolysis of macromolecules is the initial reaction to degrade organic matter in the ocean (Hoppe et al., 2002; Arnosti, 2003; Pantoja et al., 2011) and as such it would point to a role in marine carbon biogeochemistry.
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various depths between 0 and 110 m below the surface, and surface sediment samples were obtained with a Van Veen Grab (sampling area 0.1 m2). One-liter water samples were filtered through 0.45 mm cellulose ester sterile filters to collect suspended particulate matter, and filters were immediately stored in liquid nitrogen for DNA analysis. Unfiltered seawater and sediment samples were stored in sterile containers at 4 C for isolation of fungi.
2.2. DNA extraction and PCR amplification of fungi 2. Materials and methods 2.1. Study site and sampling The study was conducted in the coastal zone of the southern Humboldt Current System off central Chile at ca. 36 S (Fig. 1). The n is characterized by a relatively continental shelf off Concepcio straight coastline and is limited by submarine canyons of the Itata River (average runoff of 300 m3 s1) to the north and Biobío River ~ ones and Montes, (average runoff of 900 m3 s1) to the south (Quin 2001). Continental shelf break is located ~60 km from the coastline at 150 m depth. Seawater and surface marine sediments were sampled during August 2007 (austral winter) onboard R/V Kay Kay II (Department n, Chile), at sampling sites of Oceanography, University of Concepcio A, B and C (Fig. 1), located 30, 15 and 2 km from the Itata River mouth. The water column was sampled with Niskin bottles at
Environmental DNA from filters was extracted using PowerSoil DNA Kit (MO BIO Laboratories, Inc.). Template DNA (1e3 mL) was subjected to standard PCR using general fungal primers NS1 and ITS4 (White et al., 1990). PCR parameters: initial denaturation of 2 min at 95 C, 30 amplification cycles of 94 C of 1 min denaturation, annealing/extension at 55 C for 4 min, and final extension of 5 min at 72 C. One mL of PCR product was used for nested PCR with NS1 and GC-Fung primers, which amplify a segment of small sub unit (SSU) 18s rDNA suitable for separation by denaturing gradient gel electrophoresis (DGGE) (May et al., 2001). Nested PCR parameters were initial denaturation for 2 min at 94 C, 35 amplification denaturation cycles at 94 C, annealing for 1 min at 50.3 C, extension for 1 min at 72 C, and extension for 5 min at 72 C. PCR reaction mixtures (50 mL) contained 200 mM each DNTP, 3.5 mM MgCl2, 0.4 mM of each primer, and one unit Taq DNA polymerase (GoTaq® Flexi DNA Polymerase, Promega).
Fig. 1. Location of sampling sites (red circles) in the upwelling ecosystem off central Chile adjacent to the Itata River (inset shows turbid river plumes as seen in a 250 m resolution MODIS true color image from August 21, 2008). The image was provided by M Sobarzo and G Saldías.
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2.3. Denaturing gradient gel electrophoresis of DNA fragments of fungi DGGE profiles of fungal community were developed on 8% acrylamide/bisacrylamide (37.5:1) gel with 20e60% denaturing gradient (1 mm thick, 1X TAE buffer, 20 cm 20 cm). Following polymerization, gel was placed in the buffer chamber of a DCode universal mutation detection unit (Bio-Rad) in 1X TAE buffer at 60 C. Nested PCR products (40 mL) were loaded onto gels with 10 mL 2X gel-loading dye and electrophoresis proceeded at 100 V and 60 C for 10 h. Gel was stained in a 0.5 mg L1 1X TAE/ethidium bromide solution and photographed on a UV transilluminator. DGGE band patterns were identified by analysis of digital image of gels (Quantity One, Bio-Rad). Similarity analysis of DGGE profiles was carried out using band presence and intensity, and clusters were produced from UPGMA (Unweighted Pair-Group Method with Arithmetic mean) statistical analysis. Richness (S) of operational taxonomic units (OTUs) was estimated as number of bands in DGGE gel. Statistical differences in OTUs composition among sampling sites were tested by PERMANOVA analysis. 2.4. Isolation of fungal strains and extracellular enzymatic hydrolysis Fungi were isolated and cultured with culture media used for marine fungi (Kohlmeyer and Kohlmeyer, 1979) and yeasts (Yarrow, 1998). Seawater aliquots (100e150 mL) filtered through sterile cellulose ester filters (0.22 mm) and subsamples of surface sediments (ca. 100 mg) were placed on solid culture media. Successive streaking and spreading on agar plates were carried out to select and isolate pure fungal colonies. Fifteen isolated strains from sites A, B, and C were assayed for extracellular activity of aminopeptidase and glucosidase (Hoppe, 1983); two strains from surface water (5 m) of each site, and 9 strains from surface sediments (2 from site A, 3 from site B, and 4 from site C). Five mL duplicate aliquots of cell suspensions prepared in filtered seawater (coastal seawater filtered through 0.2 mm) were incubated in darkness at 10 mM final concentration of fluorogenic substrates L-leucine-4-methylcoumarinyl-7-amide (MCA-Leu) for aminopeptidase activity and 4-methylumbelliferone b -D-glucoside (MUF-Glu) for glucosidase activity. Fluorescence was measured on subsamples removed at time zero and every hour for 6e7 h. Abiotic hydrolysis and adsorption were evaluated by incubating fluorogenic substrates in boiled (10 min) cells suspensions. Fluorescence was measured in a Turner fluorometer (TBS-380 Mini) using wavelength 365 nm for excitation and 455 nm for emission (Meyer-Reil, 1987). Calibration curves were constructed by measuring fluorescence of 7-amino-4methylcoumarin (MCA) and 4-methylumbelliferone (MUF) in sterile seawater at concentrations ranging between 0.03 and 0.5 mM. Rates of hydrolysis by fungal strains, calculated according to Pantoja and Lee (1994), were normalized to fungal carbon estimated from dry weight of fungal tissue using a conversion factor 1 g dry weight fungal biomass ¼ 0.35 g fungal carbon (FC) (Newell and Statzeif-Taliman, 1982). The Mann-Whitney non-parametric U test was applied to test the statistical differences between sampling sites for OTUs richness and extracellular enzymatic activity. All statistical analyses were carried out using the PAST software (Hammer et al., 2001).
A ¼ 8.2 ± 1.3, site B ¼ 12.4 ± 1.5, site C ¼ 12.3 ± 1.5) in waters of sites B and C, located at 15 and 2 km from the river mouth, than at offshore site A located 30 km from the river mouth (Fig. 2). Similarity analysis showed three distinctive water column fungal communities (ca. 50% similarity): (i) a community grouping surface coastal water of riverine influence (Site C; Fig. 3, Group 1), (ii) a community including marine waters below the halocline (>30 m depth in Sites A and B; Fig. 3, Group 2), and (iii) a surface water (<30 m depth) community from coastal marine sites (Fig. 3, Group 3). Segregation of the structure of the fungal community with respect to depth and distance from the coast has been reported for the Hawaiian coast (Gao et al., 2010) and along a saline gradient in estuarine waters (Burgaud et al., 2013). The warmest temperature was 12.6 ± 0.3 C between 5 and 15 m depth at the three marine sites, with cooler waters in the top 1 m, probably due to mixing with riverine waters (Fig. 4). Site C consistently showed the influence of the Itata River with lower salinity as low as 22.7 psu near the estuary, that contrasts with values of 34.5 ± 0.1 observed below 40 m depth (Fig. 4) and with salinities >34 measured in austral summer in the area (Pantoja et al., 2011). OTU composition in the water column provided evidence of distinctive communities in (a) waters near the Itata River mouth, and (b) in surface and (c) in subsurface waters of offshore sites (Fig. 3), suggesting the occurrence of fungal communities representative of offshore and nearshore environments. Both salinity values (Fig. 4) and a distinctive river plume (Fig. 1) are evidence of the Itata River favourably influencing the transport of riverine fungi to the coastal ocean. Average fungal richness decreased with distance from the coast (Fig. 2), suggesting that freshwater can be a source of fungi in these coastal waters, increasing the richness of the fungal community in the area under direct influence of river discharge. A previous study showed both a decrease in fungal diversity with salinity along the Delaware River and coastal ocean and a high relative abundance (37%) of halotolerant strains of terrestrial origin in the estuarine area (Burgaud et al., 2013). In addition, our hypothesis is also consistent with
3. Results and discussion 3.1. Compositional patterns of fungal community OTUs richness was 34% (4.2 OTUs) higher (Richness: site
Fig. 2. Vertical profile of OTUs richness of fungi derived from DGGE profiles in sites A, B and C. Significant statistical differences were observed between sites A and B (MannWhitney U ¼ 0, p-value ¼ 0.01) and A and C (Mann-Whitney U ¼ 7, p-value ¼ 0.03).
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Fig. 3. Dendrograms derived from similarity analysis of DGGE profiles along the nearshore-offshore gradient in the water column of the coastal area adjacent to the Itata River mouth. Composition of fungal communities was significantly different between clusters identified from OTUs similarity analysis (Pseudo-F ¼ 6.92, p-value ¼ 0.0001).
Fig. 4. Vertical profiles of temperature and salinity during austral winter 2007 at sites A, B, C.
findings of freshwater phylotypes of bacterioplankton inhabiting et al., 2000). However, coastal seawater (Crumb et al., 1999; Rappe others have found dominance of distinctive phylotypes of prokaryotes in freshwater and marine ecosystems (Bouvier and del Giorgio, 2002; Cottrell and Kirchman, 2004; Campbell and Kirchman, 2013). For the same study area in central Chile, freeliving prokaryote communities showed differences in
composition between fresh and coastal waters and a singular pattern of composition suggesting development of native prokaryote assemblages in estuarine zones (Levipan et al., 2012). For fungi, coastal transitions in community composition have been evidenced in tidal marshes along a salinity gradient in Rhode Island (Mohamed and Martiny, 2011), and in estuarine sediments of Delaware Bay (Burgaud et al., 2013).
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Fig. 5. Extracellular hydrolytic activity on protein- and carbohydrate-like substrates by fungal strains isolated from the water column and surface sediments. Hydrolysis rates were normalized by fungal carbon (FC). Significant statistical differences were observed between proteolytic activity of strains isolated from sites C and B (Mann-Whitney U ¼ 3, p-value ¼ 0.036).
In offshore waters (sites A, B in Fig. 1), fungal communities were distinct between the top 30 m depth and deeper waters (Fig. 3). Since this change coincided with the depth of the base of thermocline (ca. 30 m), we suggest that vertical segregation is associated with thermal stratification of the water column (Fig. 4). This vertical zonation correlated with physicochemical properties is characteristic of most microplankton in the ocean (e. g. Lalli and Parsons, 1996), suggesting a marine source for fungal communities in offshore waters. Consistently, prokaryote bacteria in offshore waters of the same study area showed vertical segregation in phylotypes (Levipan et al., 2012). 3.2. Extracellular enzymatic hydrolysis of fungal strains One hundred and six mycelial and 90 yeast colonies were isolated from seawater, with most of the colonies cultured from marine sites A and B, averaging 11 and 23 strains per depth versus 9 in site C. From surface sediments, 42 fungal and 10 yeast colonies were isolated, with the highest number of strains from site C. No yeasts grew in isolation media from sediments of sites A and B, located at 15 and 30 km from the river mouth. In addition, a higher average number of colonies per depth were cultured from waters of the top 30 m (20 ± 19) than from deeper waters (12 ± 6) in offshore sites, which is consistent with higher fungal biomass found in surface than deeper waters of the coastal upwelling ecosystem off rrez et al., 2010, 2011). Along the nearshorecentral Chile (Gutie offshore gradient, most culturable fungi from the water column were recovered from offshore sites A (11 ± 1) and B (23 ± 4) than site C (9 ± 0.6), whereas in surface sediments most fungal strains were isolated near the river mouth (site C ¼ 40 vs 4 and 8 in sites A and B respectively). All assayed strains hydrolyzed fluorogenic proteinaceous (MCALeu) and carbohydrate-like (MUF- Glu) substrates. Rates of hydrolysis of protein substrate were five times higher than those of carbohydrate substrates and hydrolysis rates of strains from site C near the estuary were generally higher (Fig. 5), suggesting that during winter culturable fungi inhabiting seawater and sediments of the coastal zone adjacent to the river mouth have the ability to rapidly process organic matter. This result is consistent with the traditional view of fungi as key decomposers in detrital trophic webs of coastal
ecotones (Fell and Newell, 1998; Hyde et al., 1998; Raghukumar 2005) and provides a new perspective for the disappearance of organics exported from soils by rivers in the coastal zone that, despite their apparent refractory nature, are poorly represented in both water column and sedimentary marine carbon pools (Hedges, 1992). Acknowledgments n Marina This study was funded by the Programa de Investigacio de Excelencia Pimex-Nueva Aldea (PIMEX; Universidad de Con n & Celulosa Arauco y Constitucio n S.A.) and COPAS Surcepcio ~ ones received also partial Austral CONICYT PIA PFB31. R. Quin funding from the Interdisciplinary Center for Aquaculture Research ~ ez, J Soto, and (INCAR; FONDAP 1511002). We thank R Castro, L Nun E Tejos for their help during field and laboratory work. We also thank the valuable guidance of Professor J Fell from University of rrez during a research visit in his Miami who hosted MH Gutie laboratory for training in marine mycology (Capacity Building Grant E-13852 FA-UDEC-WHOI). M Sobarzo and G Saldías kindly provided MODIS image of Fig. 1. We are very grateful to the HanseWissenschaftskolleg (HWK), Delmenhorst, Germany, for Fellowship awarded to S Pantoja. References Arnosti, C., 2003. Microbial extracellular enzymes and their role in DOM cycling. In: Findley, S., Sinsabaugh, R.S. (Eds.), Aquatic Ecosystems: Interactivity of Dissolved Organic Matter. iAcademic Press, California, pp. 315e342. Azam, F., Fenchel, T., Field, J.G., Gray, J.S., Meyerreil, L.A., Thingstad, F., 1983. The ecological role of water-column microbes in the sea. Mar. Ecol. Prog. Ser. 10, 257e263. Bouvier, T.C., del Giorgio, P.A., 2002. Compositional changes in free-living bacterial communities along a salinity gradient in two temperate stuaries. Limnol. Oceanogr. 47, 453e470. Burgaud, G., Woehlke, S., Redou, V., Orsi, W., Beaudoin, D., Barbier, G., Biddle, J.F., Edgcomb, V.P., 2013. Deciphering presence and activity of fungal communities in marine sediments using a model estuarine system. Aquat. Microb. Ecol. 70, 45e62. Campbell, B.J., Kirchman, D.L., 2013. Bacterial diversity, community structure and potential growth rates along an estuarine salinity gradient. ISME J. 7, 210e220. Clipson, N., Otte, M., Landy, E., 2006. Biogeochemical roles of fungi in the marine and estuarine habitats. In: Gadd, G.M. (Ed.), Fungi in Biogeochemical Cycles. Cambridge university press, pp. 436e461.
M.H. Gutierrez et al. / Fungal Ecology 29 (2017) 90e95 Cottrell, M.T., Kirchman, D.L., 2004. Single-cell analysis of bacteria growth, cell size, and community structure in the Delaware stuary. Aquat. Microb. Ecol. 34, 139e149. Crumb, B.C., Armbrust, E.V., Baros, J.A., 1999. Phylogenetic analysis of particuleattached and free-living bacterial communities in the Columbia River, its stuary, and the adjacent coastal ocean. Appl. Environ. Microb. 65, 3192e3204. Damare, S., Raghukumar, C., 2008. Fungi and macroaggregation in deep-sea sediments. Microb. Ecol. 56, 168e177. Fell, J.W., Newell, S.Y., 1998. Biochemical and molecular methods for the study of marine fungi. In: Cooksey, K.E. (Ed.), Biochemical and Molecular Methods for the Study of Marine Fungi. Chapman and Hall, London, pp. 259e283. Fuhrman, J.A., Azam, F., 1982. Thymidine incorporation as a measure of heterotrophic bacterioplankton production in marine surface waters: evaluation and field results. Mar. Biol. 66, 109e120. Gao, Z., Johnson, Z.I., Wang, G., 2010. Molecular characterization of the spatial diversity and novel lineages of mycoplankton in Hawaiian coastal waters. ISME J. 4, 111e120. rrez, M.H., Pantoja, S., Quin ~ ones, R.A., Gonza lez, R.R., 2010. First record of Gutie filamentous fungi in the coastal upwelling ecosystem off central Chile. Gayana 74, 66e73. rrez, M.H., Pantoja, S., Tejos, E., Quin ~ ones, R.A., 2011. The role of fungi in Gutie processing marine organic matter in the upwelling ecosystem off Chile. Mar. Biol. 158, 205e219. rrez, M.H., Jara, A.M., Pantoja, S., 2016. Fungal parasites infect marine diatoms Gutie in the upwelling ecosystem of the Humboldt current system off central Chile. Environ. Microbiol. 18, 1646e1653. Hammer, Ø., Harper, D.A.T., Ryan, P.D., 2001. PAST: paleontological statistics software package for education and data analysis. Palaeontol. Electron. 4, 9 pp. Hedges, J.I., 1992. Global geochemical cycles: progress and problems. Mar. Chem. 39, 67e93. Hoppe, H.G., 1983. Significance of exoenzymatic activities in the ecology of brackish water: measurements by means of methylumbelliferyl-substrates. Mar. Ecol. Prog. Ser. 11, 299e308. Hoppe, H.G., Arnosti, C., Herndel, G.F., 2002. Ecological significance of bacterial enzymes in marine environment. In: Burns, R.C., Dick, R.P. (Eds.), Microbial Enzymes in the Environment Activity, Ecology, and Applications. Marcel Dekker, Inc., New York, pp. 73e107. ~ o, E., Pointing, S., Poonyth, A.D., Vrijmoed, L., 1998. Role Hyde, K., Jones, E.B.G., Lean of fungi in marine ecosystem. Biodivers. Conserv. 7, 1147e1161. Jebaraj, C.S., Raghukumar, C., 2009. Anaerobic denitrification in fungi from the coastal marine sediments off Goa, India. Mycol. Res. 113, 100e109. Jebaraj, C.S., Raghukumar, C., Behnke, A., Stoeck, T., 2010. Fungal diversity in oxygen-depleted regions of the Arabian Sea revealed by targeted environmental sequencing combined with cultivation. FEMS Microbiol. Ecol. 71, 399e412. Kohlmeyer, J., Kohlmeyer, E., 1979. Marine mycology, the Higher Fungi. Academic press, New York. Lalli, C.M., Parsons, T.R., 1996. Biological Oceanography: an Introduction. Butterworth/Heinemann, Oxford, 320 pp. n, W.O., Saldías, G.S., 2012. Figerprinting analysis of the Levipan, H.A., Alarco
95
prokaryote community along a marine-freshwater transect in central-southern Chile. Ann. Microbiol. 62, 1121e1140. ~ ones, R.A., Urrutia, H., 2007. A time series of prokaryote secLevipan, H.A., Quin ondary production in the oxygen minimum zone of the Humboldt Current System, off central Chile. Prog. Oceanogr. 75, 531e549. May, L.A., Smiley, B., Schmidt, M.G., 2001. Comparative denaturing gradient gel electrophoresis analysis of fungal communities associated with whole plant corn silage. Can. J. Microbiol. 47, 829e841. Meyer-Reil, L.-A., 1987. Seasonal and spatial distribution of extracellular enzymatic activities and microbial incorporation of dissolved organic substrates in marine sediments. Appl. Environ. Microb. 53, 1748e1755. Mohamed, D.J., Martiny, J.B., 2011. Patterns of fungal diversity and composition along a salinity gradient. ISME J. 5, 379e388. rraga, L., Montero, P., Daneri, G., Cuevas, L.A., Gonz alez, H.E., Jacob, B., Liza Menschel, E., 2007. Productivity cycles in the coastal upwelling area off Con n: the importance of diatoms and bacterioplankton in the organic carbon cepcio flux. Prog. Oceanogr. 75, 518e530. Newell, S.Y., Statzeif-Taliman, A., 1982. Factors for conversion of fungal biovolume values to biomass, carbon and nitrogen: variation with mycelial ages, growth conditions, and strains of fungi from a salt marsh. Oikos 39, 261e268. Offre, P., Spang, A., Schleper, C., 2013. Archaea in biogeochemical cycles. Annu. Rev. Microbiol. 67, 437e457. Pantoja, S., Lee, C., 1994. Cell-surface oxidation of amino acids in seawater. Limnol. Oceanogr. 39, 1718e1726. rrez, M.H., Ampuero, P., Tejos, E., 2011. Degradation capability of Pantoja, S., Gutie the coastal environment adjacent to the Itata River in central Chile (36.5 S). Biogeosciences 8, 2063e2074. ~ ones, R.A., Montes, R., 2001. Relationship between freshwater input to the Quin coastal zone and the historical landings of the benthic/demersal fish Eleginops maclovinus in central-south Chile. Fish. Oceanogr. 10, 311e328. Raghukumar, S., 2005. The role of fungi in marine detrital processes. In: Ramaiah, N. (Ed.), Marine Microbiology: Facets & Opportunities. National Institute of Oceanography, Goa, pp. 91e101. , M.S., Vergin, K., Giovannoni, S.J., 2000. Phylogenetic comparisons of a coastal Rappe bacterioplankton community with its counterparts in open ocean and freshwater systems. FEMS Microbiol. Ecol. 33, 219e232. Richards, T.A., Jones, M.D.M., Leonard, G., Bass, D., 2012. Marine fungi: their ecology and molecular diversity. Annu. Rev. Mar. Sci. 4, 495e522. White, T.J., Bruns, T.D., Lee, S.B., Taylor, J.W., 1990. Analysis of phylogenetic relationships by amplification and direct sequencing of ribosomal DNA genes. In: Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols: a Guide to Methods and Applications. Academic Press, San Diego, pp. 315e322. Wuchter, C., Abbas, B., Coolen, M.J.L., Herfort, L., Van Bleijswijk, J., Timmers, P., Strous, M., Teira, E., Herndl, G.J., Middleburg, J.J., Schouten, S., Sinninghe Damste, J.S., 2006. Archaeal nitrification in the ocean. Proc. Nat. Acad. Sci. U. S. A. 103, 12317e12322. Yarrow, D., 1998. Methods for the isolation, maintenance and identification of yeasts. In: Kurtzman, C.P., Fell, J.W. (Eds.), The Yeasts, a Taxonomic Study. Elsevier, Amsterdam, pp. 77e100.