Open Access

Diversity and biochemical features of culturable fungi from the coastal waters of Southern China

AMB Express20144:60

DOI: 10.1186/s13568-014-0060-9

Received: 17 May 2014

Accepted: 9 July 2014

Published: 30 August 2014

Abstract

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.

Keywords

Marine-derived fungi Diversity Hydrolytic enzymes Pelleterization

Introduction

Coastal marine habitats have been characterized as the most variable, highly diverse and rich in primary production (Jickells [1998]; Danovaro and Pusceddu [2007]). 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 [1982]). Different types of microbes in coastal waters degrade a large proportion of this detritus actively (Manini et al. [2003]; Pusceddu et al. [2003]). Among these microbes, heterotrophic bacteria and archaea have been described for their degradation abilities towards such detritus to a greater extent (Moran and Miller [2007]; Mou et al. [2008]). 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 [2005]; Hallam et al. [2006]; Fenchel [2008]; Strom [2008]).

Among eukaryotes, fungi have been reported to exhibit as individual filaments or aggregates in coastal waters (Gutiérrez et al. [2010]). However, in comparison with terrestrial environments, fungi in the world’s oceans remain largely unknown (Gao et al. [2010]). 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. [2012]). Mycoplankton include free-living filamentous fungi, yeasts, fungal-like protists, and those associated with planktonic particles or phytoplankton (Wang and Johnson [2009]; Gao et al. [2010]).

Fungi are a key component of the biosphere, fulfilling a wide range of biogeochemical and ecological functions in natural environments (Christensen [1989]; Pang and Mitchell [2005]). 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 [1982]; Damare and Raghukumar [2008]) and thus benefit the growth of planktonic microbial communities (Kiørboe and Jackson [2001]; Gutiérrez et al. [2010]). The biomass of planktonic fungi has been reported to be comparable with prokaryotes including both Bacteria and Archaea (Gutiérrez et al. [2011]).

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 [1994]; Raghukumar [2004]). They possess the ability to penetrate relatively persistent particulate detritus much more efficiently than bacterioplankton (Raghukumar [2004]). 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. [2013]). 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. [2013]). 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. [2011]). 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. [2009]).

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).
Table 1

Details regarding colony forming units (CFUs) of fungal colonies on the media plates

Sampling date

Location

Lat(°N)

Long (°E)

Habitat

Depth (m)

Temperature (°C)

Salinity (ppt)

Media

No. of colonies

CFU/L

2012-03-05

Shenzhen Bay

22°31'19.776"

113°57'4.284"

Seawater

0

16.25

31.02

MEA

3

120

22°31'19.776"

113°57'4.284"

Seawater

0

16.25

31.02

SDA

2

80

22°31'19.776"

113°57'4.284"

Sediment

0

16.25

31.02

MEA

12

480

22°31'19.776"

113°57'4.284"

Sediment

0

16.25

31.02

SDA

55

2200

Daya Bay

22°35'34.224"

114°30'28.800"

Seawater

0

16.25

31.02

MEA

4

160

22°35'34.224"

114°30'28.800"

Seawater

0

16.25

31.02

SDA

5

200

22°35'34.224"

114°30'28.800"

Sediment

0

16.25

31.02

MEA

6

240

22°35'34.224"

114°30'28.800"

Sediment

0

16.25

31.02

SDA

8

320

2012-03-21

Mirs Bay

22°29'44.02"

114°27'34.38"

Seawater

0

17.37

31.37

MEA

1

40

22°29'44.02"

114°27'34.38"

Seawater

0

17.37

31.37

SDA

3

120

22°29'44.02"

114°27'34.38"

Seawater

5

17.37

31.37

MEA

2

80

22°29'44.02"

114°27'34.38"

Seawater

5

17.37

31.37

SDA

4

160

22°29'44.02"

114°27'34.38"

Seawater

10

17.37

31.37

MEA

3

120

22°29'44.02"

114°27'34.38"

Seawater

10

17.37

31.37

SDA

5

200

Mirs Bay

22°31'33.14"

114°27'51.59"

Seawater

0

17.74

31.37

MEA

3

120

22°31'33.14"

114°27'51.59"

Seawater

0

17.74

31.37

SDA

2

80

22°31'33.14"

114°27'51.59"

Seawater

5

17.74

31.37

MEA

4

160

22°31'33.14"

114°27'51.59"

Seawater

5

17.74

31.37

SDA

13

520

22°31'33.14"

114°27'51.59"

Seawater

10

17.74

31.37

MEA

3

120

22°31'33.14"

114°27'51.59"

Seawater

10

17.74

31.37

SDA

10

400

Mirs Bay

22°31'32.02"

114°28'26"

Seawater

0

17.69

31.38

MEA

2

80

22°31'32.02"

114°28'26"

Seawater

0

17.69

31.38

SDA

8

320

22°31'32.02"

114°28'26"

Seawater

5

17.69

31.38

MEA

5

200

22°31'32.02"

114°28'26"

Seawater

5

17.69

31.38

SDA

7

280

22°31'32.02"

114°28'26"

Seawater

10

17.69

31.38

MEA

5

200

22°31'32.02"

114°28'26"

Seawater

10

17.69

31.38

SDA

17

680

Daya Bay

22°34'19.99"

114°31'30"

Seawater

0

19.24

31.00

MEA

2

80

22°34'19.99"

114°31'30"

Seawater

0

19.24

31.00

SDA

3

120

22°34'19.99"

114°31'30"

Seawater

5

19.24

31.00

MEA

2

80

22°34'19.99"

114°31'30"

Seawater

5

19.24

31.00

SDA

7

280

22°34'19.99"

114°31'30"

Seawater

10

19.24

31.00

MEA

3

120

22°34'19.99"

114°31'30"

Seawater

10

19.24

31.00

SDA

12

480

Daya Bay

22°34'34.37"

114°30'30.46"

Seawater

0

19.69

30.86

MEA

3

120

22°34'34.37"

114°30'30.46"

Seawater

0

19.69

30.86

SDA

9

360

22°34'34.37"

114°30'30.46"

Seawater

10

19.69

30.86

MEA

10

400

22°34'34.37"

114°30'30.46"

Seawater

10

19.69

30.86

SDA

8

320

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. [1990]). 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.

Phylogenetic analyses

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. [1990]). The ITS1-5.8S-ITS4 gene sequences obtained for the organisms were aligned with their closest match using the program, ClustalW (Thompson et al. [1994]). 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. [2007]) 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. [1995]). 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. [1986]). 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 [2011]; Carder et al. [1986]). 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. [2006]). 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. ([1994]). 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).

Results

Culturable diversity

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).
Table 2

Phylogenetic affiliations of culturable fungi based on of ITS rDNA gene sequences

Isolate ID

GenBank accession No.

Closest identified relative

Phylum of closest relative

Source of isolation of the closest relative

% Identity

Species

(GenBank accession No.)

PKU F1

KC113282

Mucor sp. FJ09

(HQ019160)

Zygomyceta

Water hyacinth leaf

99

PKU F2

KC113283

Hypocrea koningii strain JZ-25

(HQ637343)

Ascomycota

Soil from Sichuan

99

PKU F3

KC113284

Arthrinium phaeospermum isolate T57

(FJ462766 )

Ascomycota

Not known

99

PKU F4

KC113285

Diaporthe sp. H4236

(GU595056 )

Ascomycota

Mangrove in China

98

PKU F5

KC113286

Phoma sp. ZH2.1

(FJ450059 )

Ascomycota

Argyrosomus argentatus

99

PKU F6

KC113287

Trichoderma piluliferum strain wxm37

(HM037939 )

Ascomycota

River water

99

PKU F7

KC113288

Trichoderma asperellum isolate T29

(JN108927)

Ascomycota

Rhizospheric soil

99

PKU F8

KC113289

Aspergillus nomius strain NRRL 26885

(JF824686)

Ascomycota

Philanthus triangulum

99

PKU F9

KC113290

Aspergillus flavipes isolate BCT2-3

(JQ082507 )

Ascomycota

Shark gills

100

PKU F10

KC113291

Dothideomycetes sp. OY307

(FJ571450 )

Ascomycota

Not known

99

PKU F11

KC113292

Arthrinium phaeospermum isolate DFFSCS004

(JX156350)

Ascomycota

Deep-sea sediments

99

PKU F12

KC113293

Hypocrea lixii isolate SZMC 20858

(JX173851)

Ascomycota

Hungarian vegetables

100

PKU F13

KC113294

Trichoderma hamatum strain LXM1

(GQ220703 )

Ascomycota

Saline-alkali soil

100

PKU F14

KC113295

Cladosporium sp. JS1043

(AM176680)

Ascomycota

Deep sea

100

PKU F15

KC113296

Curvularia sp. B34

(HQ696021)

Ascomycota

Moso Bamboo Seeds

99

PKU F16

KC113297

Cladosporium sphaerospermum isolate IJL07

( EU823317 )

Ascomycota

Soybean

99

PKU F17

KC113298

Arthrinium sp. LH11

(HQ832842 )

Ascomycota

Tea plants

99

PKU F18

KC113299

Ascomycota sp. AR-2010 isolate TR063

(HQ608095 )

Ascomycota

Soil

92

PKU F19

KC113300

Pleosporales sp. LH241

(HQ832825)

Ascomycota

Tea plants

100

PKU F20

KC113301

Cladosporium cladosporioides isolate 2728

(EU272532)

Ascomycota

Plant

99

PKU F21

KC113302

Pyrenochaeta sp. CF-2008

(EU885415)

Ascomycota

Corneal ulcer

99

PKU F22

KC113303

Aspergillus terreus strain 1B61

(KF572455)

Ascomycota

Geothermal soil

99

PKU Y1

KC113304

Rhodotorula mucilaginosa strain KDLYC24-1

(HQ909092 )

Basidiomycota

Not known

99

PKU Y2

KC113305

Rhodotorula glutinis var. salinaria strain ZH7-(7)

(FJ487944)

Basidiomycota

Mangrove in China

99

PKU Y3

KC113306

Aureobasidium pullulans strain KDLYC4-9

(HQ909088 )

Ascomycota

Not known

99

PKU Y4

KC113307

Rhodosporidium sphaerocarpum isolate 2223

(HQ670691)

Basidiomycota

Shrimp Culture

99

PKU Y5

KC113308

Rhodosporidium diobovatum strain IWBT-Y840

(JQ993385)

Basidiomycota

grapevines

99

PKU Y6

KC113309

Trichosporon brassicae

(NR_073251)

Basidiomycota

Not known

99

PKU Y7

KC113310

Rhodotorula mucilaginosa strain WC53-2

(EF190221)

Basidiomycota

Not known

99

PKU Y8

KC113311

Candida parapsilosis IFM52618

(AB109284)

Ascomycota

Braziland Japan

99

PKU Y9

KC113312

Rhodotorula mucilaginosa strain R-4685

(KF726105)

Basidiomycota

Aortic valve

100

Figure 1

Phylum affiliation of culturable fungi with the existing sequences of NCBI database.

Figure 2

NJ phylogenetic tree based on ITS rRNA genes from 22 fungi and 9 yeast derived from coastal ecosystems of Pearl River Delta. Numbers at nodes indicate bootstrap values of neighborjoining analysis for 1,000 replicates (values below 50% not shown).

Enzymatic activities

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).
Table 3

Qualitative assay of enzyme activities on media plates amended with specific substrates

Isolate ID

Laccase (4thday)

Lipase (3rdday)

Cellulase (3rdday)

Source of isolation

Depth (m)

PKU F1

-

+

+

Seawater

0

PKU F2

+

-

+

Seawater

0

PKU F3

+

++

++

Seawater

0

PKU F4

++

+

+

Seawater

0

PKU F5

-

++

-

Sediment

0

PKU F6

-

-

+

Seawater

0

PKU F7

++

+

++

Sediment

0

PKU F8

-

+

++

Sediment

0

PKU F9

-

++

++

Sediment

0

PKU F10

+

-

-

Sediment

0

PKU F11

+

++

+++

Seawater

10

PKU F12

-

++

++

Seawater

10

PKU F13

-

++

+++

Seawater

0

PKU F14

++

+++

++

Seawater

0

PKU F15

+

-

-

Seawater

10

PKU F16

+++

+++

++

Seawater

5

PKU F17

-

++

+

Seawater

10

PKU F18

+++

+

+++

Seawater

0

PKU F19

-

+

-

Seawater

0

PKU F20

++

++

++

Seawater

0

PKU F21

+

++

-

Seawater

0

PKU F22

-

+++

++

Seawater

0

PKU Y1

-

++

+

Seawater

0

PKU Y2

-

+

++

Seawater

0

PKU Y3

-

++

+

Seawater

10

PKU Y4

-

+

++

Seawater

0

PKU Y5

-

++

++

Seawater

10

PKU Y6

-

++

++

Seawater

5

PKU Y7

-

-

++

Seawater

0

PKU Y8

-

+++

+++

Seawater

0

PKU Y9

-

++

++

Seawater

10

Figure 3

Qualitative assay of enzyme activities of fungi on substrate specific media plates. A. Plate assay for cellulose activities (PKU Y7 Rhodotorula mucilaginosa sp. (left) and PKU Y9 Rhodotorula mucilaginosa sp. showing good cellulose activity (rght)); B. Plate assay for laccase activities (PKU F11 Arthrinium phaeospermum sp. showing no laccase activity (left); PKU F16 Cladosporium sphaerospermum showing considerable laccase activities (right)); C. Plate assay for lipase activities on plate (PKU Y7 Rhodotorula mucilaginosa sp. showing little lipase activity (left); PKU Y9 Rhodotorula mucilaginosa sp. showing good lipase activity (right)).

Table 4

Quantitative assay of enzyme activities for four fungal isolates

Isolate ID

Fungal genera

Growth

date

Lipase (Umin−1 ml−1)

Cellulase (Umin−1 ml−1)

Laccase (Umin−1 ml−1)

PKU F16

Cladosporium sp.

6th day

14.384 ± 0.994

0.843 ± 0.084

10.495 ± 1.300

9th day

12.285 ± 0.772

0.443 ± 0.032

5.105 ± 0.943

PKU F18

Ascomycota sp.

6th day

6.605 ± 0.959

1.133 ± 0.096

14.609 ± 1.039

9th day

4.630 ± 0.812

1.001 ± 0.049

7.581 ± 1.642

PKU Y5

Rhodosporidium sp.

6th day

17.174 ± 0.742

0.333 ± 0.040

-

9th day

12.186 ± 1.705

0.371 ± 0.028

-

PKU Y8

Candida sp.

6th day

21.940 ± 1.582

1.311 ± 0.013

-

9th day

18.767 ± 1.554

1.132 ± 0.029

-

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.
Table 5

Details regarding pellet formation by the fungal isolates

Isolate ID

Species

No. of spores (per L)

Pellet color

Different pH

PH = 2

PH = 4

PH = 6

PH = 8

Pellet size(mm)

Pellet size(mm)

Pellet size(mm)

Pellet size(mm)

PKU F1

Muco r sp. FJ09

4.00*106

yellow

-

cotton- shaped

5

4

1.60*107

yellow

-

cotton- shaped

2 pellets (20*10)/(5*10)

1 pellet (20*10)

PKU F8

Aspergillus nomius strain

1.59*107

white

<0.5

4-6

3

3-7

6.34*107

white

<0.5

1-2

3

4

PKU F14

Cladosporium sp.

3.85*108

black

<0.1

0.5

0.5-1

1-2

1.54*109

black

-

-

-

1

Discussion

Diversity of fungi has been reported from various environments such as freshwater (Gulis et al. [2006]), marine environments such as coastal waters (Gao et al. [2010]), deep-sea sediments (Damare et al. [2006]; Singh et al. [2010]), hypersaline waters (Buchalo et al. [2000]), methane hydrates (Lai et al. [2007]), oxygen deficient ecosystems (Cathrine and Raghukumar [2009]), mangroves and salt marshes (Hyde et al. [1998]; Raghukumar [2004]), and hydrothermal vents (Le Calvez et al. [2009]). 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. [2010]). 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. [2010]). 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. [2012]). 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 [1982]).

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. ([2012]). 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. [2009]; Anderson et al. [2003]).

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. [2002]). 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. [2002]). Fungi are well known agents of decomposition of organic matter composed of cellulosic substrate in particular (Lynd et al. [2002]). 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 [1998]). 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. [2012]). 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 [2007]). 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 [2010]). 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. [2000]). 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.

Additional file

Declarations

Acknowledgements

This work was partially funded by National Science Foundation of China grant 31170109 (GYW) and Shenzhen Development and Reform Commission grant 835 (GYW).

Authors’ Affiliations

(1)
School of Environment and Energy, Peking University Shenzhen Graduate School
(2)
Department of Biological Science, National University of Singapore
(3)
Tianjin University Center for Marine Environmental Ecology, School of Environmental Science and Engineering, Tianjin University
(4)
Department of Microbiology, University of Hawaii at Manoa

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