Marine Pollution Bulletin xxx (2015) xxx–xxx Contents lists available at ScienceDirect Marine Pollution Bulletin journal homepage: www.elsevier.com/locate/marpolbul Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India Rajendran Sangeetha Devi, Velu Rajesh Kannan ⇑, Duraisamy Nivas, Kanthaiah Kannan, Sekar Chandru, Arokiaswamy Robert Antony Rhizosphere Biology Laboratory, Department of Microbiology, School of Life Sciences, Bharathidasan University, Tiruchirappalli 620 024, Tamil Nadu, India a r t i c l e i n f o Article history: Received 10 February 2015 Revised 11 May 2015 Accepted 19 May 2015 Available online xxxx Keywords: HDPE Aspergillus tubingensis VRKPT1 Aspergillus flavus VRKPT2 Fourier transformed infra-red spectroscopy Scanning electron microscopy a b s t r a c t High density polyethylene (HDPE) is the most commonly found non-degradable solid waste among the polyethylene. In this present study, HDPE degrading various fungal strains were isolated from the polyethylene waste dumped marine coastal area and screened under in vitro condition. Based on weight loss and FT-IR Spectrophotometric analysis, two fungal strains designated as VRKPT1 and VRKPT2 were found to be efficient in HDPE degradation. Through the sequence analysis of ITS region homology, the isolated fungi were identified as Aspergillus tubingensis VRKPT1 and Aspergillus flavus VRKPT2. The biofilm formation observed under epifluorescent microscope had shown the viability of fungal strains even after one month of incubation. The biodegradation of HDPE film nature was further investigated through SEM analysis. HDPE poses severe environmental threats and hence the ability of fungal isolates was proved to utilize virgin polyethylene as the carbon source without any pre-treatment and pro-oxidant additives. Ó 2015 Elsevier Ltd. All rights reserved. 1. Introduction Plastics are man-made synthetic organic polymers that have found wide variety of applications in every aspect of life and industries. The versatility of these materials has lead to a great increase in their use over the past three decades and they have rapidly moved into all types of utility. These widespread applications of plastics are not only due to their favorable mechanical and thermal properties but mainly due to the stability and durability (Hansen, 1990). Among the various types of plastic polymers, the most popular and convenient plastic polymers include low-density polyethylene (LDPE), high-density polyethylene (HDPE), polypropylene (PP), polyvinyl chloride (PVC), polystyrene (PS), nylon, polyethylene terephthalate (PET), polyurethanes, etc. Among the synthetic plastics waste produced, polythene shares about 64% and it is considered as the most commonly found solid waste that has been recognized as a major threat to marine life (Lee et al., 1991). Annually 25 million tons of synthetic plastics are being accumulated in the coastal and terrestrial environment (Derraik, 2002). The increasing quantities of plastic waste and their effective and safe disposal has become a matter of public concern. However, this extensive usage has not been accompanied with development of safe deposition or degradation protocols. Owing ⇑ Corresponding author. E-mail address: uvrajesh@gmail.com (V. Rajesh Kannan). to their inert nature and degradation resistant property, their accumulation in environment has become enormous. Accumulation of plastic products in the environment adversely affects wildlife, habitat of lands and oceans. It also poses variety of problems like choking drainage systems and importantly health hazards to humans and animals. Chlorinated plastic releases harmful chemicals in the soil and it leads to soil leaching and contamination of ground water ecosystem. In the marine environment alone, out of total marine waste, plastic shares about 60–80% (Derraik, 2002). As per literature, every year hundred thousand tons of plastics have been dumped in the marine environment causing fatal effects (Rutkowska et al., 2002). Due to plastic pollution in the marine environment, minimum 267 species are being affected including all mammals, sea turtles (86%) and seabirds (44%) (Coe and Rogers, 1997). The usage of plastic materials is increasing day-by-day. The threat associated with all ecosystems makes their degradation and deterioration necessary. Plastics are resistant against microbial attack, since during their short time of presence in nature evolution could not design new enzyme structures capable of degrading synthetic polymers. Biodegradation of polymers primarily focuses on increasing the surface hydrophobicity, thereby enhancing microbial attachment. Hence, most of the researchers recommend pretreatment (Arkatkar et al., 2010; Balasubramanian et al., 2014) for efficient microbial adherence. Such treatments result in the formation of carbonyl, carboxyl and ester functional groups that decreases the hydrophobicity. http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 0025-326X/Ó 2015 Elsevier Ltd. All rights reserved. Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 2 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx There is a growing interest in synthetic polymer biodegradation using effective microorganisms. Developments of microbial communities attached to the synthetic wastes have been found to be powerful degrading agents in nature. Studies on fungal mediated degradation of polyethylene has previously been reported using Mucor rouxii and Aspergillus flavus (El-Shafei et al., 1998), Phanerochaete chrysosporium (Liyoshi et al., 1998), Penicillium simplicissimum YK (Yamada-Onodera et al., 2001) and Aspergillus niger ITCC no. 6052 (Mathur et al., 2011). This study aims to isolate and identify the polyethylene degrading efficient fungal strains without any pre-treatment, pro-oxidant additives and also attempts to understand the degradation ability of two fungal strains, Aspergillus tubingensis VRKPT1 and A. flavus VRKPT2. 2. Materials and methods 2.1. Sample collection Partially degraded polyethylene along with soil samples, adhering and adjacent to it, was collected from the plastic waste dumped marine environmental site in the coastal area of Gulf of Mannar, Tuticorin, Tamil Nadu, India. 2.2. Polyethylene Commercially available HDPE (40 lm in thickness and 0.95 g/cm3 in density) materials were purchased from the local market, Tiruchirappalli, Tamil Nadu, India. The composition of commercially available HDPE varies from pure polyethylene by the addition of additives like antioxidants and colorant. Prior to the experiments, HDPE films were cut into small strips, sterilized with 70% ethanol and dried in sterile condition. The amplified products were analyzed through electrophoresis on 1.2% agarose gel (Sigma) and stained with 0.5 lg ml1 ethidium bromide. The PCR products obtained were purified and sequenced using Sanger’s dideoxynucleotide chain termination method and were carried out at Eurofins genomics India Pvt. Ltd. (Bangalore). The gene sequences obtained was analyzed by using BLAST program (http://www.ncbi.nlm.nih.gov/blast). The phylogenetic and evolutionary analyses were conducted using MEGA 5 software (Tamura et al., 2011). 2.5. Screening of HDPE degrading fungi The isolated fungal strains were screened for HDPE degradation efficiency according to the method followed by Gilan et al., 2004. Each individual isolate was grown in SM amended with HDPE as a sole carbon source. Preweighed disinfected untreated films were aseptically added to flasks containing 50 ml of SM. The isolated fungal strains were inoculated and incubated at 30 °C for one month. SM containing HDPE without the microorganisms were maintained as positive controls. In some experiments, mineral oil (light white oil, d = 0.84 g l1; Difco,) was added to the medium to test the effect of colonization of fungal strains over the HDPE surface. After 30 days of incubation, the HDPE samples were removed subsequently, washed, dried at 60 °C and weighed. Furthermore, the degradation was confirmed with the aid of Fourier Transform Infrared Spectroscopy (FT-IR). 2.6. Methods of analysis The physical, chemical and mechanical changes in the polyethylene that occurred during biodegradation were monitored as described below. 2.7. Determination of the dry weight of the residual HDPE 2.3. Isolation of HDPE degrading fungal species Isolation was performed according to the method proposed by Sivan et al. (2006). From the collected samples, 10 g of soil was inoculated in 100 ml synthetic media (SM) containing (per liter of distilled water): NH4NO3 1.0, MgSO47H2O – 0.2, K2HPO4 – 1.0, CaCl22H2O – 0.1, KCl – 0.15, yeast extract – 0.1 (Difco) (g/l) and micronutrients for 1.0 mg l1 of each of the following: FeSO46H2O, ZnSO47H2O and MnSO4. In addition, the collected partially degraded polyethylene samples were added as substrate to the medium. The samples were incubated for 12 weeks at 30 °C with shaking (150 rpm min1). After incubation, the mixtures of fungi were isolated and purified by spread plate technique. The purified cultures were maintained in potato dextrose agar (PDA) slants. The cultures were maintained at ambient temperature and frequently revived to sustain its viability. 2.4. Identification of fungal strains The fungal isolates were identified based on morphological characteristics and nucleotide sequence analysis of internal transcribed space (ITS) region. After 5 days of growth at 28 ± 2 °C, approximately 100 mg of the mycelial biomass was harvested. The total genomic DNA was extracted using the CTAB method (Moller et al., 1992). The ITS region was amplified by PCR using the following universal primers: ITS1 (50 -TCCGTAGGTGAAC CTGCGG-30 ) and ITS4 (50 -TCCTCCGCTTATTGATATGC-30 ) (White et al., 1990). The conditions used for PCR amplification were as follows: initial denaturation stage of 95 °C for 5 min followed by 35 cycles of 95 °C for 30 s, 55 °C for 1 min, 72 °C for 1 min and a final stage of 72 °C for 6 min. The microbial biofilm colonizing from the HDPE surface was washed off with 2% (v/v) of aqueous sodium dodecyl sulfate (SDS) solution for 4 h at 50 °C and further washed with warm distilled water to facilitate accurate measurement of the residual HDPE weight (Sivan et al., 2006). The washed HDPE samples were collected on filter paper, rinsed with distilled water and then dried overnight at 60 °C before they were finally weighed. The weight loss was calculated using the following formula: Percentage of weight loss = [(Final weight  Initial weight)/Original weight]  100. 2.8. FT-IR analysis Changes in the polyethylene structure and the formation or disappearance of functional groups in the polyethylene during the process of degradation can be monitored by FT-IR (Perkin Elmer, Spectrum RX, USA). Spectra in the frequency range of 4000–400 cm1 were used at a resolution of 2 cm1. The relative absorbance intensities of the ester carbonyl bond at 1740 cm1, keto-carbonyl bond at 1715 cm1, terminal double bond (vinyl) at 1650 cm1 and internal double bond at 908 cm1 to that of the methylene bond at 1465 cm1 were evaluated using the following formula (Albertsson et al., 1987): Keto-carbonyl bond index (KCBI) = I1715/ I1465; Ester-carbonyl bond index (ECBI) = I1740/I1465; Vinyl bond index (VBI) = I1650 / I1465 and Internal double bond index (IDBI) = I908/I1465. The crystallinity (%) of the HDPE was measured based on the method suggested by Zerbi et al. (1989) and calculated by the following formula: Percentage of crystallinity ¼ 100  ½f1  ðIa=1:233IbÞ=1 þ ðIa=IbÞg  100; Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx 3 Carlsbad, CA, USA), according to the manufacturer’s instructions. After 30 days, HDPE containing biofilms were removed from the medium and washed in sterile water. The films were stained and viewed under epifluorescent microscope equipped with a fluorescein isothiocyanate filter (Chavant et al., 2002). Live cells (green in color) can be differentiated from dead cells, which emit red color. 2.11. Spectrometric determination of fungal biomass The growth of the fungal strains in the liquid media due to the presence of HDPE film was determined according to the method adopted by Banerjee et al. (1993). During degradation study, the liquid cultures were withdrawn at weekly intervals during one month of incubation. The turbidity of the liquid medium was measured by detecting its absorbance in UV–Visible spectrophotometer (Elico SL-159) at 450 nm against a blank of uninoculated sterile medium. 2.12. Estimation of protein content of the biofilms The total protein content of the biofilms was determined after alkaline hydrolysis treatment. The HDPE films were sampled from the flasks containing the fungal cultures, washed gently with water to remove medium debris and boiled for 30 min in 4 ml of NaOH (0.5 M). The extracts were centrifuged to precipitate cell-debris fragments. The supernatants were collected and the pellets were subjected to the same procedure. The collected supernatants were combined and the protein concentration was determined spectrophotometrically by Lowry’s method (Lowry et al., 1951). 2.13. SEM analysis Fig. 1. Gel electrophoresis of PCR products amplified from DNA extracted from fungal strains. The PCR products were amplified by the primers ITS1 and ITS4. 1: Molecular weight marker (Lambda DNA/Hind III and uX 174 DNA/Hae III digest); 2: A. tubingensis VRKPT1; 3: A. flavus VRKPT2. where Ia and Ib are absorbance values from the bands at 1474 and 1464 cm1 or at 730 and 720 cm1, respectively. 2.9. Evaluation of fungal cell surface hydrophobicity Determination of fungal cell-surface hydrophobicity was evaluated by the microbial adhesion to hydrocarbons (MATH) assay, which is based on the affinity of fungal cells toward organic hydrocarbon. The more hydrophobic of a fungal cell has greater affinity toward hydrocarbon, resulting in transfer of cells from aqueous to organic phase. Cell surface hydrophobicity was performed as described by Smith et al. (1998). Briefly, blastospores, aerial and submerged conidia were suspended in 10 mM potassium phosphate buffer (pH 7.0). Hexadecane (300 ll) were added to the set of test tubes containing cell suspension and the test tubes were shaked for 10 min and allowed to stand for 2 min. The absorbance of the aqueous suspensions before (A0) and after (At) were measured at 470 nm. Cell-free buffer served as a blank. The percentage of cells adhering to organic phase was calculated using the formula: Percentage of hydrophobicity = (A0  At)/A0  100. Hydrophobicity was expressed as the percentage of cells removed from the aqueous phase. 2.10. Viability test Viability of the fungal biofilm was determined by the LIVE/DEADÒ FungalLight™ Yeast viability kit (Molecular Probes, After 30 days of incubation, the HDPE film was taken and washed by the previously defined method (Sivan et al., 2006). The samples were fixed in 2% glutaraldehyde for 2 h, washed twice in 50% ethanol, incubated overnight in 70% ethanol and finally washed again in 100% ethanol. The processed films were cut into 1  1 cm size and subjected to SEM for the examination of surface erosion in HDPE film. The samples were coated with gold on the surfaces of HDPE film before observing the micrographs. After fixation, the samples were taken into deep vacuum and visualized under scanning electron microscope (Vega 3, Tescan). 2.14. Statistical analysis All the experiments were performed in triplicates. Statistical analysis was performed using GraphPad Prism software by one-way analysis of ANOVA. The data obtained were expressed as mean ± standard deviation (SD). Differences were considered to be significant at p < 0.05. 3. Results and discussion High-Density Polyethylene degrading fungal strains were isolated from the partially degraded polyethylene wastes along with adhered soil in marine ecosystem samples by enrichment technique. The enrichment technique gave rise to a mixture of fungus capable of growing in liquid synthetic medium (SM) containing HDPE as a sole carbon source. A number of fungal populations were obtained from the fungal mixtures, among these, two isolates designated as VRKPT1 and VRKPT2 exhibited the fastest growth and showed increased biofilm formation over the HDPE surface. The fungal isolates were screened for HDPE degradation efficiency in SM supplemented with HDPE. During degradation study, the Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 4 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx Fig. 2. Molecular phylogenetic analysis of fungal strains (a) VRKPT1 and (b) VRKPT2 showed the relationship between the ITS rRNA gene sequences retrieved from GenBank. The evolutionary relationship was inferred by using the Maximum Likelihood method based on the Kimura 2-parameter model. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The analysis involved 11 nucleotide sequences. All positions containing gaps and missing data were eliminated. Evolutionary analyses were conducted in MEGA 5. Fig. 3. Weight loss of HDPE film after one month of incubation with A. tubingensis VRKPT1 and A. flavus VRKPT2 in SM (50 ml) containing pre-weighed HDPE as a sole carbon source in the medium. Data represent means of three replicates ± SD. fungal isolates were colonized over the HDPE surface within a few days. The fungal isolates were identified based on the sequence analysis of ITS region homology. Amplification of fungal genomic DNA by primers ITS1 and ITS4 yielded 603 bps fragments (Fig. 1). The isolates VRKPT1 and VRKPT2 showed 100% homology with A. tubingensis and A. flavus respectively. The evolutionary history was inferred by using the Maximum Likelihood method based on the Kimura 2-parameter model (Kimura, 1980). A phylogenetic Fig. 4. Percentage weight loss of HDPE exposed to A. tubingensis VRKPT1 and A. flavus VRKPT2 with the presence and absence of mineral oil. The data represent the means of three replicates, and the error bars indicate the percent error. analysis confirmed their similarity to the respective species (Fig. 2). The sequence of fungal isolates A. tubingensis VRKPT1 (Accession No. KM873026) and A. flavus VRKPT2 (Accession No. KM873027) were deposited in GenBank Sequence repository. After 30 days of incubation, the HDPE degradation was monitored by weight loss. Our isolates showed better HDPE biodegradation ability than the previously reported work. The weight loss observed by the fungal isolates VRKPT1 and VRKPT2 was Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx Fig. 5. Fourier transform infrared analysis of HDPE incubated with A. tubingensis VRKPT1 and A. flavus VRKPT2 for one month. (KCBI – Keto Carbonyl Bond Index; ECBI – Ester Carbonyl Bond Index; VBI – Vinyl Bond Index; IDBI – Internal Double Bond Index.) Data represent means of three replicates ± SD. 6.02 ± 0.2% and 8.51 ± 0.1% (Fig. 3). El-Shafei et al. (1998) investigated the ability of fungi and Streptomyces strains to attack disposed polyethylene bags, and isolated eight different Streptomyces strains and two fungi M. rouxii NRRL 1835 and A. flavus from sewage sludge. Seneviratne et al. (2006) reported the degradation ability of HDPE by Penicillium frequentans and Bacillus mycoides through biofilm formation. A. flavus, isolated from sanitary landfills was also found to be capable of degrading polyethylene (Mendez et al., 2007). Yamada-Onodera et al. (2001) suggested the pretreatment of polyethylene using a fungus P. simplicissimum YK, makes degradation easier and faster. Most of the previously reported work in Aspergillus spp. like Aspergillus terreus (Balasubramanian et al., 2014), A. niger (Volke et al., 2001), Aspergillus cremeus, Aspergillus ornatus, Aspergillus glaucus, Aspergillus candidus, Aspergillus nidulans, A. flavus and Aspergillus oryzae (Konduri et al., 2010) suggested the pretreatment of HDPE. Hence, the potential HDPE degraders, A. tubingensis VRKPT1 and A. flavus VRKPT2 has shown up highest degradation rate without any pre-treatment and pro-oxidant additive (Fig. 3). The biofilm formation can be influenced by the cellular surface hydrophobicity (Gilan et al., 2004; Balasubramanian et al., 2010). Mineral oil has promoted the hydrophobic interaction between the fungal strains and HDPE surface. It has been previously reported that the agents such as mineral oil and Tween 80 act as a modulators of hydrophobic interaction between the polymer and microorganisms, which increases the rate of biofilm formation and biodegradation of polyethylene (Gilan et al., 2004; Tribedi and Sil, 2013). Mor and Sivan (2008) evaluated the three non-ionic surfactants (Tween 20, Tween 60 and Tween 80) act as potential Fig. 6. Percentage hydrophobicity of fungal strains (A. tubingensis VRKPT1 and A. flavus VRKPT2) toward the hydrocarbon (hexadecane). The data represent the means of three replicates, and the error bars indicate the percent error. 5 enhancers of biofilm formation. Tribedi and Sil (2013) reported that the degradation of LDPE by Pseudomonas sp. AKS2 was found to increase with the addition of mineral oil. Yamada-Onodera et al. (2001) reported that the nonionic surfactant Triton X-100 improved the growth of P. simplicissimum in a medium containing polyethylene without being utilized by the fungus. HDPE degradation was also carried out in the presence and absence of mineral oil. The present study revealed that the increased attachment of fungal strains to the polymer surface when mineral oil was added to the medium. In the absence of mineral oil, the rate of biodegradation was found to be 6.02 ± 0.2% and 8.51 ± 0.1% by A. tubingensis VRKPT1 and A. flavus VRKPT2 but in the presence of mineral oil, the biofilm formation and the rate of biodegradation of HDPE film was increased to 6.88 ± 0.1% and 9.34 ± 0.2% (Fig. 4). Consistent with the above results, the mineral oil not only helps in fungal attachment to the HDPE surface but also accelerates the formation of a biofilm and contributes higher degradation of HDPE. The biodegradation of polyethylene was originally initiated by abiotic process. Oxidation of polymer chain occurred due to the dissolved oxygen or that which is present in the ambient leading to the formation of carbonyl groups. Later, the carbonyl groups form the carboxylic groups, undergo b-oxidation and finally enters the citric acid cycle resulting in the formation of CO2 and H2O (Albertsson et al., 1987). During the degradation process, changes in functional groups and/or side chain modification occur due to the action of microbes over HDPE surface. Monitoring the formation or disappearance of acids (1740 cm1), ketones (1715 cm1) and double bonds (1640 and 915 cm1) during HDPE biodegradation process was examined using FT-IR analysis. Previous findings reported that there was a continuous increase in the amount of carbonyl groups with exposure in an abiotic environment (Gilan et al., 2004). The 1700–1800 cm1 region in FT-IR spectra indicates the presence of oxidized groups. Initially, carbonyl index has increased due to the oxidation of dissolved oxygen. Prolonged exposure to fungal strains A. tubingensis VRKPT1 and A. flavus VRKPT2, leads to decrease in carbonyl index probably due to biodegradation through the formation of ester or Norrish-type mechanism (Fig. 5). Previous reports are also proved that the amount of carbonyl groups decreased with prolonged exposure to a biotic environment (Dolezel, 1967). Albertsson et al. (1987) have reported that biotic environment supports the formation of terminal double bonds (IR 905–915 cm1). In this study, the same was observed for terminal double bonds when treated with fungal strains. The fraction of internal double bonds (ACH@CHA) was higher than that of terminal/vinyl double bond (ACH@CH2) (Fig. 5). The formation of keto, ester, vinyl and double bond index was observed in both A. tubingensis VRKPT1 and A. flavus VRKPT2. However, in control there was no change in keto, ester, vinyl and double bond index. Synthetic polymers are water insoluble, which is due to their crystallinity. Generally, a water uptake was higher in polyethylene when overseeded with growing molds. When the polyethylene exposed to a biotic environment, the increase in water uptake was accompanied by an increase in degree of crystallinity (Dolezel, 1967). The percentage of crystallinity was increased at about 37.52 ± 0.3% and 29.47 ± 0.2% after incubated with the fungal strains A. tubingensis VRKPT1 and A. flavus VRKPT2. Experimentally this is observed as an initial increase in percentage crystallinity due to the consumption of amorphous portions (Santo et al., 2012). Later, the depleted smaller crystals have been consumed by microorganisms resulting in the proportion of larger crystals (Albertsson et al., 1995). The formation of functional groups and side chain modifications confirms the HDPE biodegradation. Thus, the FT-IR data strongly suggests the biotic factors which play a major role in the biodegradation of HDPE film. Hydrophobicity is an important property of the surface in biodegradation studies. The ability of microorganisms to utilize Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 6 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx Fig. 7. Fluorescent photomicrographs showing the viability of fungal colonization and biofilm formation on the surface of the HDPE film even after 30 days of incubation (a) control, (b) A. tubingensis VRKPT1, (c) A. tubingensis VRKPT1 + mineral oil, (d) A. flavus VRKPT2 and (e) A. flavus VRKPT2 + mineral oil. any substrate depends on its growth and adherence to that substrate. In general, it is accepted that more hydrophilic surfaces are more easily colonized by microorganisms. Several reports have demonstrated that the correlation between cell surface hydrophobicity and carbon starvation. Sanin et al. (2003) have reported that microbial cell surface become more hydrophobic and adhesive in Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx Fig. 8. Spectrometric determination of fungal biomass monitored at weekly intervals during HDPE degradation with the fungal strains VRKPT1 and VRKPT2. carbon-starved cultures than with non-starved cultures. To determine the fungal interaction with HDPE film, MATH assay has been carried out to test the hydrophobicity of the fungal strains. Strains with hydrophobicity more than 40% were considered hydrophobic (Del Re et al., 2000). The MATH assay showed the greater affinity toward the solvent hexadecane. The strain A. flavus VRKPT2 showed 55.49% affinity toward hydrocarbon whereas A. tubingensis VRKPT1 showed 48.47% (Fig. 6). HDPE is non-polar in nature; an organism with higher cell surface hydrophobicity is likely to utilize the HDPE more efficiently because cell surface hydrophobicity favors closer association between organism and HDPE. Our result demonstrates that an increase in cell surface hydrophobicity of fungal strains resulted in an enhanced microbial adherence to HDPE surface, lead to the increase in HDPE degradation by A. tubingensis VRKPT1 and A. flavus VRKPT2. Since, the biodegradation is a slow process; it requires the biofilm to be active over a long period of time. After a month of incubation, the HDPE film was subjected for the fungal biofilm viability test using LIVE/DEADÒ FungalLight™ Yeast viability kit. The fungal mycelia and spores attached over the surface of the HDPE indicated the viability of both the fungal strains even after one month of incubation (Fig. 7). It is clearly evident that increase in biofilm formation over the HDPE surface confirms the degradation ability of the fungal strains (A. tubingensis VRKPT1 and A. flavus VRKPT2) on utilization of HDPE as a sole carbon source. The turbidity of the SM increased with time on being inoculated with the fungal strains was measured at 450 nm (Fig. 8). The fungal isolates that survived in the SM supplemented with HDPE as its 7 only sole carbon source should possess some mechanism degrading the HDPE to access its carbon content. The increase in turbidity and the displayed colloidal suspensions were observed within one week of incubation. But in control set up, the SM supplemented with HDPE did not show any turbidity. The slow increase of the turbidity could be due to the formation of biofilms and the residues occurred during the biodegradation process (Chatterjee et al., 2010). When compared to A. tubingensis VRKPT1, the turbidity was observed higher in A. flavus VRKPT2. The increase in turbidity in the following weeks indicates that the enhanced growth and biofilm formation by the fungal strains. Protein assays were proved as an efficient tool for determining the state of polyethylene colonization and biofilm formation (Gilan et al., 2004). The growth kinetics of the fungal isolates on the HDPE film was monitored by quantifying the total protein content extracted from the surface of the HDPE. The continuous and slow increase in extractable protein suggests a regular growth of fungal isolates over the HDPE surface. During first week of incubation, there was a slight increase in protein content, reflecting an increase in the biofilm biomass. During the second week of incubation period, the protein content remains constant. There was a steep increase in protein content in both the fungal strains during third week of incubation. But during the fourth week of degradation period, there was a rapid increase in protein content, which indicates the strong affinity of fungal isolates to the HDPE (Fig. 9). This is due to the rate of biofilm formation over the surface of the HDPE film. The extractable protein content was found to be higher in A. flavus VRKPT2 than that of A. tubingensis VRKPT1. The changes in surface topography of the HDPE films were examined by SEM analysis. Microbial adhesion over the HDPE surface during biofilm formation led to the corrosion on the surface of HDPE. In a study by Bonhomme et al. (2003), SEM evidence confirmed that fungi has build up on the surface of the HDPE; and after removal of the fungi, the surface became physically pitted and eroded. Similar to that, the SEM images showed an alteration in the surface topology of HDPE films treated by fungal strains. The number of cracks and grooves were observed on the surface of the fungal treated films. The fungal treated films became rough, whereas the untreated film retained a smooth surface even after 30 days of incubation under the same condition (Fig. 10). Such patterns have previously been observed for biodegradable polymers (Otake et al., 1995). Based on the results of weight loss, viability of fungal biofilm, FT-IR spectral analysis and surface topography analysis, both the fungal strains are considered to be potent HDPE degraders but when compared to both fungal strains A. flavus VRKPT2 showed higher degradation efficiency than A. tubingensis VRKPT1. 4. Conclusion Fig. 9. The protein content of the surface attached biofilm of the fungal strains (A. tubingensis VRKPT1 and A. flavus VRKPT2) in SM supplemented with HDPE as a sole source of carbon. The data represent the means of three replicates, and the error bars indicate the percent error. HDPE have become an integral part in our daily life as a key material of basic needs. Indiscriminate littering of unskilled recycling and non-biodegradability of HDPE raises several environmental issues. In general, biodegradation of HDPE by microorganisms and enzymes seems to be the most effective process. The current study demonstrates that the degradation of HDPE film by two marine fungi namely, A. tubingensis VRKPT1 and A. flavus VRKPT2 without any pre-treatment and pro-oxidant additives. Between these two strains, the colonization, biofilm formation and biodegradation of HDPE film by A. flavus VRKPT2 was higher than A. tubingensis VRKPT1. The smooth surface of the HDPE film has turned to rough with cracking; this indicates the presence of enzyme activities. It is evident that both the fungal strains release the extracellular enzymes to degrade the HDPE film, but the detailed characterization of these enzymes is Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 8 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx Fig. 10. SEM images showing surface modification of HDPE after biodegradation (a) untreated film (control) (b) film treated with A. tubingensis VRKPT1; (b) film treated with A. tubingensis VRKPT1 + mineral oil; (c) film treated with A. flavus VRKPT2; (d) film treated with A. flavus VRKPT2 + mineral oil. still needed to be carried out. The experimental results demonstrated the degradation ability of the fungal strains under in vitro condition and also provide a feasible solution to the environmental threat posed by the HDPE polymer. Hence, further study will be focused in the field of genomics and proteomics, which could speed up the rate of degradation. Please cite this article in press as: Sangeetha Devi, R., et al. Biodegradation of HDPE by Aspergillus spp. from marine ecosystem of Gulf of Mannar, India. Mar. Pollut. Bull. (2015), http://dx.doi.org/10.1016/j.marpolbul.2015.05.050 R. Sangeetha Devi et al. / Marine Pollution Bulletin xxx (2015) xxx–xxx Acknowledgements The first author (R. Sangeetha Devi) is thankful to Department of Science and Technology, New Delhi, India for their financial support in the form of DST-INSPIRE fellowship to carry out this research work. We would like to express our sincere gratitude to Dr. M. 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