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. 2021 Jan 3;11(1):29. doi: 10.1007/s13205-020-02592-9

Isolation of a soil bacterium for remediation of polyurethane and low-density polyethylene: a promising tool towards sustainable cleanup of the environment

Rusha Roy 1, Goutam Mukherjee 1, Anirban Das Gupta 1, Prosun Tribedi 1,2, Alok Kumar Sil 1,✉
PMCID: PMC7779388  PMID: 33442527

Abstract

A soil bacterium, designated strain AKS31, was isolated on the plastic polyurethane (PUR) and based on the molecular and biochemical analysis was tentatively assigned to the genus Pseudomonas. Preliminary studies suggested that strain AKS31 had the capability of biodegrading polyurethane (PUR) and low-density polyethylene (LDPE). This observation was confirmed by the analysis of the biodegradation products. The hydrolyzed products of PUR analyzed sequentially by High-Performance Liquid Chromatography (HPLC) and Matrix Assisted Laser Desorption Ionization Time of Flight Mass Spectrometry (MALDI-TOF MS) showed the presence of diethylene glycol suggesting the presence of an esterase. A gene that could be involved in producing an esterase-like activity (PURase gene) was identified after the amplification and sequencing of a PCR product. Fourier Transformed Infrared (FTIR) spectrophotometric analysis of AKS31-treated LDPE film revealed the incorporation of hydroxyl groups suggesting the involvement of a hydroxylase in the degradation of LDPE. It is established that plastics form microplastics and microbeads in soils which negatively impact the health of living organisms and there have been concentrated research efforts to remediate this problem. Microcosm studies revealed that when strain AKS31 was bioaugmented with soil both the polymers were degraded during which time the heterotrophic plate counts, soil respiration and soil organic carbon content increased but this was not the case with the control nonbioaugmented microcosm. The results demonstrate that the strain AKS31 may have the potential in biodegradation of PUR and LPDE present as plastic microbeads and thereby improving soil health. Further studies in this direction are warranted.

Supplementary Information

The online version contains supplementary material available at 10.1007/s13205-020-02592-9.

Keywords: Biodegradation, Plastic, PUR, LDPE, Pseudomonas

Introduction

Plastic materials include a plethora of synthetic or semi-synthetic materials that have a wide range of industrial as well as domestic applications owing to their stability, malleability, high tensile strength, low cost, and ease of manufacturing (Mukherjee et al. 2011). Plastics are polymeric substances, oftentimes petroleum by-products, that contain linear or branched chains of hydrocarbon molecules. Other elements, such as oxygen, nitrogen, chlorine, etc., may be present as part of the functional groups within the polymer structure (Das and Kumar 2014). The indiscriminate use and irresponsible disposal of plastics have led to widespread pollution that threatens ecosystems in marine to terrestrial environments (Mukherjee et al. 2011; Bhatia et al. 2014; Muenmee et al. 2015). Moreover, larger pieces of plastic litter serve as sources of microplastics (< 5 mM), including microbeads (10–500 μM in size), as they undergo physical degradation by natural processes in the open environment. As microplastics can persist in the environment for a long period of time, studies show an alarming rate of accumulation of these micropollutants in terrestrial, freshwater, and marine ecosystems (Andrady 2011; Wu et al. 2017). Owing to their minute size and widespread distribution, these microparticles pose a serious threat to these diverse ecosystems, more so, because their high surface to volume ratio allows faster leaching of pollutants.

Two plastic polymers that are of particular concern for their widespread use, as well as resistance to biodegradation, are polyurethane (PUR) and low-density polyethylene (LDPE). PUR is synthesized by the condensation of isocyanates and poly-alcohols and contains an ester linkage. PUR is hydrophilic and compared to many other plastic polymers, is relatively susceptible to environmental degradation. It is largely used as a synthetic foam that has extensive application in diverse fields (Schmidt et al. 2017). On the other hand, LDPE is used in the manufacturing of plastic bags, bottles, and disposable containers and accounts for 60% of all kinds of synthetic polymers used (Ambika et al. 2015; Byuntae et al. 1991; Zahra et al 2010). LDPE is hydrophobic and does not have any functional group, making the polymer acutely resistant to environmental degradation (Tribedi et al. 2012; Bhatia et al. 2014). Therefore, this plastic polymer can persist in the environment for extended periods. The accumulation of this polymer in the environment is extremely high, about 25 million tons per year (Soni et al. 2009; Zahra et al. 2010).

Plastics are mostly dumped into landfills or are incinerated. While incineration generates copious toxic fumes, landfill sites, which are suitable for the safe disposal of plastic polymers, are increasingly becoming limited (Al-Salem et al. 2009; Crowley et al. 2003; Jumaah 2017). Therefore, there is a pressing need to adopt biological approaches to solve the environmental challenges related to plastic pollution. In this context, several studies have shown that microorganisms are capable of efficient degradation of different plastic polymers (Tribedi and Sil 2013; Bhatia et al. 2014; Caruso 2015; Muhonja et al. 2018). To this end multiple microorganisms (mostly bacteria and fungi) have been isolated that can degrade PUR (Oceguera- Cervantes et al. 2007; Mukherjee et al. 2011; Cregut et al. 2013; Gamerith et al. 2016; Osman et al. 2018; Magnin et al. 2019). Fungal species like Aspergillus, Penicillium, and Trichoderma can effectively degrade PUR (Álvarez-Barragán et al. 2016; Osman et al. 2018; Magnin et al. 2019). Similarly, several bacteria, including Corynebacterium, Bacillus, Pseudomonas, and Enterobacter, were also shown to degrade PUR efficiently (Kay et al. 1991; Oceguera-Cervantes et al. 2007; Mukherjee et al. 2011; Cregut et al. 2013; Shah et al. 2013; Kumar Gupta and Devi 2019). Besides PUR, microorganisms like Pseudomonas, Staphylococcus, Bacillus, Arthrobacter, Aspergillus, Penicillium, etc. have shown immense potential in degrading polyethylene samples (Chatterjee et al. 2010; Yang et al. 2014). Interestingly, several bacteria including Pseudomonas, Micrococcus, Staphylococcus, Streptococcus, and fungi that include Aspergillus glaucus, Aspergillus niger, and Trichoderma have been found to degrade both biodegradable (PUR) and non-biodegradable (LDPE) polymers efficiently (Howard et al. 1999; Nakajima-Kambe et al. 1999; Roy et al. 2008; Shah et al. 2008; Tribedi et al. 2012; Bhatia et al. 2014; Gajendiran et al. 2016; Wei and Zimmermann 2017).

Typically microorganisms attach themselves to the polymer surface, and subsequently, mineralize the plastic polymer with the help of enzymes (Silva et al. 2010; Das and Kumar 2014; Chaisu 2015). Alternatively, microorganisms that do not attach to the polymer surface, produce extracellular enzyme(s) capable of degrading the target polymer (Shah et al. 2013). To this end, the activity of both cell-associated and extracellular esterases has been shown to be involved in the microbial degradation of PUR (Nomura et al. 1998; Akutsu et al. 1998; Shah et al. 2013). Towards LDPE degradation, it was documented that a hydroxylase enzyme that belongs to the AlkB family initiates the oxidation process. (Yoon et al. 2012. In a separate study, a copper binding bacterial enzyme, laccase, was shown to be capable of degrading LDPE (Santo et al. 2013). Treatment of polyethylene by this enzyme reduced the molecular weight of the polymer with a concomitant increase in keto-carbonyl index.

Despite continuous research in this field and some notable success, biodegradation of PUR and LDPE derived contaminants from the environment is still an ongoing challenge. It has been noticed that for most of the isolated organisms either the efficiency of polymer degradation is not high or their effect on soil health remains unexplored. The test microorganism being used for biodegradation within a given natural environment, such as a landfill, must interact with the existing natural indigenous microorganisms of the given soil and these interactions may affect its activity as well as the natural microbial population of the given site. Therefore, the impact of introducing a given microbial isolate on soil health must be taken into consideration while evaluating the effectiveness of an isolate towards the biodegradation of plastics. In this context, the present study reports the isolation of a natural isolate of Pseudomonas origin for the biodegradation of PUR and LDPE. This isolate degrades these polymers efficiently both in laboratory conditions as well as in soil microcosm. The results also indicate that it does not damage soil health.

Materials and methods

Preparation of polymer films

In the current study, two polymer films namely PUR and LDPE were used. For the preparation of PUR film, 2 g of solid polyurethane (Sigma Aldrich, USA) was dissolved in 100 mL of tetrahydrofuran and the solution was then kept in desiccators for 2 days at room temperature. However, LDPE films used in the present study were collected from the local markets of Kolkata, India.

Growth medium

The selection of desired microorganisms having the ability to degrade the polymers was carried out by preparing a minimal medium in which all the growth factors were available excluding the carbon source. The minimal medium was prepared in 100 mL in which 0.2 g KH2PO4; 0.7 g K2HPO4; 0.1 g (NH4)2SO4; 0.01 g MgSO4∙7H2O; 0.0001 g ZnSO4∙7H2O; 0.00001 g CuSO4∙7H2O; 0.001 g FeSO4∙7H2O; 0.0002 g MnSO4∙H2O was added. The pH was set at 7.2 before the medium was sterilized by autoclaving. Thereafter, 0.3% PUR (Impranil DLN™) was added to the medium at permissible temperature. For solid medium, agar (20 g/L) was added to the minimal medium before autoclaving. In some experiments, PUR film was used instead of liquid PUR (Impranil DLN™) as the sole carbon source. The preparation of the PUR film was described above. For the LDPE degradation study, the same minimal medium was used to which LDPE film was added as the sole carbon source.

Soil sample collection and isolation of bacteria

Soil samples were collected in sterile containers from 5 cm below the land surface of Kolkata municipal solid waste dumping ground (DHAPA) and kept refrigerated until further processed. One gram of the collected soil was then mixed thoroughly with 10 mL of sterile saline (0.85%) and allowed to stand for 30 min at room temperature. Following the sedimentation of the soil to the bottom, the supernatant was collected and serially diluted in sterile 0.85% saline. An aliquot (100 µL) from the appropriate dilution was spread on agar plates containing PUR (Impranil DLN™) as the main carbon source and incubated further at 30 °C for 3–5 days.

Biodegradation assay

To examine the biodegradation of PUR and LDPE by the isolated organism, PUR and LDPE films were cut into small pieces, weighed, and subsequently made sterile by washing with 70% alcohol. The sterile PUR (~ 0.125 g) and LDPE (~ 0.125 g) films were separately added to conical flasks containing 50 ml of sterile minimal medium. An aliquot (100 µl) of the overnight saturated culture of the isolated organism was then independently added to each growth medium containing either PUR or LDPE. Also, a control set was prepared in which the minimal medium was supplemented with either PUR or LDPE but the inoculum was not added. All growth media were incubated at 30 °C for different times as per the need of the experiment. After the incubation, the polymer films were recovered, washed with 2% SDS, and dried. The weight loss of each strip of polymer (either PUR or LDPE) was measured. To monitor the change in mechanical properties of the recovered strips of polymers, the tensile strength of the polymers was measured using a tensometer. Moreover, the recovered polymer films were observed under an atomic force microscope (AFM) to see the change in surface topography. To do that, dried films were examined under AFM (Veeco instruments 3100, Dresden, Germany) and images were taken. Experiments were done at least three times.

Microcosm preparation and analysis of in-situ degradation of polymers by the isolated organism AKS 31

To assess the PUR and LDPE degradation ability of the isolated organism in soil, degradation of the polymers was carried out in soil microcosms. For this study, two separate microcosms were prepared with the soil (mentioned above): (i) Naturally attenuated, wherein natural soil was used for the degradation, (ii) Bioaugmented, wherein natural soil was mixed with isolated polymer degrading organism and used for degradation study. For each soil microcosm preparation, 20 g of -unsterile soil was taken in a sterile glass beaker. Three replicates were used for each type of microcosm. For bioaugmented microcosm (BIOAUG), the soil was inoculated with 2 g of the isolated bacteria. PUR and LDPE films (0.125 g) were separately added into each microcosm, covered with sterile aluminium foil, and incubated at 30 °C for different periods. A control microcosm was also set up, wherein sterile soil was used for the degradation without exogenous addition of isolated microorganism. Each microcosm was kept moistened by adding sterile water (Milli Q) at regular time intervals. After the incubation, soil samples and the polymer films were recovered from each microcosm and analyzed for both biotic activities and PUR/LDPE degradation. Experiments were done at least three times.

Microbial count analysis

One gram soil sample collected from different microcosms was separately added to 9 mL of 0.85% sterile NaCl in sterile test tubes. Thereafter, a series of dilutions from 10–2 to 10–6 were prepared in sterile 0.85% NaCl. An aliquot (0.1 mL) of the appropriate dilution from each microcosm was spread aseptically onto agar plates containing PUR as the sole carbon source for enumeration of PUR-degrading organisms and Luria agar (LA) plates for enumeration of heterotrophic organisms. The PUR plates were incubated at 30 °C for 5 days and LA plates were incubated at 30 °C for 2 days. Experiments were done at least three times.

Estimation of soil organic carbon

Soil organic carbon was measured by the modified Walkley–Black method as described previously (Alef and Nannipieri 1995). Briefly, ~ 1 g of soil was taken, mixed with 10 mL 5% (w/v) potassium dichromate, and shaken gently. Subsequently, 20 mL of concentrated sulphuric acid was added and the mixture was allowed to cool. Thereafter, the mixture was treated with (0.4%) barium chloride and allowed to stand for 8 h. Subsequently, an aliquot of clear supernatant was collected, and OD was measured at 600 nm. The experiment was done at least three times.

Soil respiration

Basal soil respiration for the bioaugmented and non-bioaugmented soils was measured by following standard protocols as described previously (Alef and Nannipieri 1995). Briefly, 10 g of soil sample was taken in a 500 mL conical hanging tube apparatus and was sealed with an airtight glass cork. Ten milliliters of 0.05 N NaOH was taken in a hanging glass tube and suspended from the glass cork. For control experiments, two sets of blanks were taken which were devoid of any soil sample. This apparatus was incubated at 25 °C for a period of 24 h. Following incubation, 5 mL of barium chloride was added to the NaOH solution and the mixture was titrated with 0.05 M HCl. The CO2 liberated was calculated according to the following formula:

CO2producedinmg/h/kgofdrysoil=(Vo-V)×1.1/D.

Where Vo = Mean volume of HCl required for titration of the blank sample; V = Volume of HCl required for titration of soil samples; 1.1 is the conversion factor [1 mL of 0.05 NaOH is equivalent to 1.1 mg of CO2] (Alef and Nannipieri 1995).

Soil respiration was also measured after the soil was subjected to heat as well as desiccation stress. For heat stress, the soil was exposed to a temperature of 70 °C for half an hour. For desiccation, soil samples were desiccated for 24 h for dehydration and subsequently, the soil respiration was measured for the treated soils. Experiments were done at least three times.

Ribotyping

Genomic DNA was extracted from the isolated polymer degrading organism by following the procedure described by Ausubel et al. (1994). Partial amplification of the 16S rRNA gene of the isolated organism AKS31 was performed by polymerase chain reaction (PCR) using genomic DNA as the template and 16S rRNA gene-specific universal primers; 27F and 1492R. The amplified PCR product was run through an agarose gel, purified, and subsequently sequenced. The obtained sequences of the nucleotides were then subjected to BLAST analysis to identify the organism. After that, the generated nucleotide sequences were submitted to GenBank (http://www.ncbi.nlm.nih.gov/genbank). To build the phylogenetic tree, the partial 16S rRNA gene sequence of the isolated organism (AKS31) and 16S rRNA gene sequences of other neighboring microorganisms having similarities of more than 98% were aligned through the MUSCLE tool of MEGA 7 (Kumar et al. 2016). Then, the phylogenetic tree was constructed from the aligned sequences of the 16S rRNA gene using the maximum parsimony algorithm of MEGA 7.

Biochemical tests

Several biochemical tests, as mentioned in the respective Figure, were performed by following the protocol described in the ninth edition of Bergey’s manual to characterize the isolated polymer degrading organism (Holtz 1993). Each experiment was repeated at least three times.

Identification of the PUR breakdown product

The microbial breakdown products of polyurethane were analyzed as follows. The cells were grown at 30 °C for 5 days and 10 days in a sterile minimal medium containing PUR as the sole carbon source. Thereafter, the cells were harvested and 20 µL of the cell-free supernatant was applied to the C18 column (Waters) for the separation of the components through high-performance liquid chromatography (HPLC) (Waters). Before injection, the column was equilibrated with 0.01 M KH2PO4 buffer (pH 2.5). After the binding, elution was carried out with the 0.02 M KH2PO4 buffer containing CH3CN (1:1). The flow rate was adjusted at 1 mL/min at ambient temperature. Two different concentrations of diethylene glycol (10 mg/ml and 100 mg/ml) were used as standard. The eluted compound that corresponds to standard diethylene glycol was subjected to matrix assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry (MS) (Bruker Daltonics, Bremen, Germany) for the determination of mass. Experiments were done at least three times.

Thin layer chromatography (TLC)

TLC aluminium sheet coated with silica gel 60 F254 (Merck) was used for this assay. A mutant of AKS31 was isolated after UV mutagenesis as described previously (Mukherjee et al. 2011). Thereafter, wild type and the mutant AKS31 cells were grown in the minimal medium containing PUR as the sole carbon source for10 days and harvested. Next, 10 µL of cell-free conditioned medium from wild type and mutant cultures were spotted on the TLC plate. Along with this uninoculated medium was also spotted. A single lane was also used for pure PUR as a positive control. Thereafter, the samples were run in a solvent system of ethyl acetate: n-hexane: methanol in the ratio 3:5:1. Color development was carried out using 5% ethanolic sulphuric acid. The experiment was done at least three times.

p-nitro phenyl acetate (pNPA) assay

Wild type and mutant cells were grown in minimal medium containing PUR as the sole carbon at 30 °C to mid-log phase and harvested. Cell lysates were made in lysis buffer containing 50 mM Tris–HCl (pH 8), 10% glycerol, 0.1% Triton X 100, 2 mM MgCl2, 1 mM PMSF and100 µg/mL lysozyme. Esterase activity was measured by incubating cell lysates containing 50 µg of total protein with 50 mM of pNPA (Hi-Media) at 37 °C for 1 h. The protein concentration was measured using the BCA protein assay kit (Thermo Fischer, USA). After the incubation, reactions were stopped by the addition of 0.1 N NaOH. The absorbance of the esterase breakdown product, pNP, was measured spectrophotometrically at 405 nm. The experiment was done at least three times.

Partial amplification of polyurethanase (PURase) gene by polymerase chain reaction (PCR)

Three known sequences of the PURase gene namely pueA, pueB, and pul A were retrieved from the GenBank database and subsequently aligned using multiple sequence alignment tool. Primers were then designed from the conserved sequences of the mentioned genes (Table 1). PCR amplification was carried out using these primers and AKS31 genomic DNA as template to examine the presence of the PURase gene. After the PCR, the amplified product (if any) was run through 1.5% agarose gel. Experiments were done at least three times. Subsequently, PCR product was gel purified, sequenced and identity with genes and proteins was searched by blastn and blastx (NCBI) analysis.

Table 1.

Primer sequences

Name of the primer Sequence (5′ to 3′)
27F AGA GTT TGA TCM TGG CTC AG
1492 R GGT TAC CTT GTT ACG ACT T
PURase F TCG CCG ACC CAG AGC GCC
PURase R TCG CTG CCG ATG ATG AAG GTG
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Fourier transformed infrared (FTIR) spectrophotometry of LDPE

For FTIR analysis, cells were grown in sterile minimal medium supplemented with LDPE films at 37 °C for 30 days. After the incubation, LDPE films were recovered and subsequently analyzed by FTIR spectroscopy (Jasco FT/IR-6300, Tsukubai, Japan). Experiments were done at least three times.

Measurement of cell surface hydrophobicity

Bacterial cell surface hydrophobicity was measured by following the protocol of bacterial adhesion to hydrocarbon (BATH) assay as described by (Tribedi et al. 2012). For the assay, cells were separately grown in sterile minimal medium supplemented with either PUR or LDPE as the sole source of carbon for 5 days and 30 days, respectively. After the incubation, cells were harvested, washed with sterile double distilled water, and re-suspended in phosphate urea magnesium (PUM) buffer [K2HPO4 (17 g/L), KH2PO4 (726 g/L), urea (18 g/L) and MgSO4, 7H2O (0.2 g/L)] such that OD400 becomes 1.0–1.2. One milliliter of this cell suspension was mixed with 0.2 mL of n-hexadecane. Shaking of the solution was carried out for 10 min following which it was made to stand for 15 min for completion of phase separation. The aqueous part was collected and the absorbance at 400 nm was measured. Cell-free PUM buffer was served as the blank. The formula used for determining microbial cell surface hydrophobicity is as follows:

Cell surface hydrophobicity (in %) = 100 X {(initial OD- final OD)/initial OD}. Experiments were done at least three times.

Statistical analysis

One-way analysis of variance (ANOVA) for the experimental results was done to evaluate statistically significant differences among samples. Mean values were compared at different levels of significance using the software Minitab 16. All experiments were performed in triplicate.

Results

Isolation of a soil bacterium that can degrade PUR and LDPE films

PUR degrading microbes were isolated by selecting the soil microbes on plates containing the polymer as the sole carbon source (as described in materials and methods). The colony exhibiting the most robust growth on this selective medium was designated AKS31. AKS31 cells were spotted on a plate containing PUR as the sole carbon source and allowed to grow for 5 days at 30 °C. A clear zone surrounding the microbial growth was observed on the plate (Fig. 1a). In contrast, E. coli failed to grow on the same plate. Survival of AKS31 on plates having PUR as the sole C-source indicated that the isolated organism was capable of degrading this polymer. For further support, the organism was grown in a minimal medium containing PUR films as the sole carbon source. At different time points, films were verified for tensile strength. Tensile strength indicates the resistance of a material to break under tension. Generally, if a material undergoes degradation, it becomes weak and exhibits reduced tensile strength compared to its original counterpart. The result showed that relative to the control, there was a significant reduction of the tensile strength of PUR films that were incubated with AKS31 and this reduction correlated positively with the time of incubation (Fig. 1b). Thus, this result again indicated the degradation of PUR by the isolated organism. To further confirm the ability of AKS31 to degrade PUR, the surface topology of the treated-and -untreated polymer films were examined by atomic force microscopy (AFM). Consistent with our expectation, while the AFM image revealed a smooth surface for the untreated control film, peaks and troughs were present on the films grown in the presence of the isolated organism (Fig. 1c). This change in the surface topology clearly indicated the degradation of the polymer. Taken together, these results documented the degradation of PUR by the isolated organism.

Fig. 1.

Fig. 1

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AKS31 degrades PUR and LDPE. a AKS31 produced a clear zone surrounding its growth on PUR plate. An equal number of cells (~ 104) from saturated cultures of the soil isolate (AKS31) and E. coli was added to wells punched in plate containing PUR as the sole C-source. The plate was incubated at 30 °C for 5 days. The experiment was repeated at least three times. b The tensile strength of PUR films was reduced after treatment with AKS31. AKS31 was allowed to grow in minimal medium containing PUR films as the sole carbon source. Films were removed from the culture at indicated time points, washed and subjected to tensile strength measurement. The result represents the average of three independent experiments. The error bar indicates the standard deviation (± SD). **p < 0.001. c AKS31 altered surface topography of PUR film. AKS31 was inoculated in minimal medium containing PUR films as the sole carbon

source and incubated. Films were removed from the culture at the indicated time points, washed and their surface topology was examined by AFM. Image is representative of three independent experiments. d AKS31 could grow in a minimal medium containing LDPE as the sole carbon source. An aliquot of the saturated culture of AKS31 was added to different tubes containing the minimal medium with or without LDPE films as indicated, and turbidity was monitored after 15 days of incubation. This experiment was repeated at least three times. e The tensile strength of LDPE films was reduced after treatment with AKS31. AKS31 was allowed to grow in minimal medium containing LDPE films as the sole carbon source. Films were removed at the indicated time points, washed, and subjected to tensile strength measurement. The result represents the average of three independent experiments. The error bar indicates standard deviation (± SD).*p < 0.05, **p < 0.001. f AKS31 altered the surface topography of LDPE film. AKS31 was inoculated in minimal medium containing LDPE films as the sole carbon source and incubated. Films were removed from the culture at indicated time points, washed and their surface topology was examined by AFM. Image is representative of three independent experiments. All the above experiments entailed incubation at 30 °C

AKS31 also exhibited significant growth in minimal medium containing LDPE films, but was unable to grow in medium lacking the film, which indicated that this organism was also capable of using LDPE as a carbon source (Fig. 1d). Assessment of the tensile strength of LDPE films showed a significant reduction in the tensile strength for the films that were treated with the organism compared to the untreated films and the level of reduction was positively correlated with the treatment period (Fig. 1e). AFM was also used for further confirmation of the LDPE-degradation ability of the isolated organism and similar to PUR results, AFM images showed marked alteration in the surface topology of the treated films, compared with the one that was not treated with the isolated organism (Fig. 1f). Collectively, the results indicate that AKS31 is able to degrade both PUR and LDPE.

To quantify the degradation of polymers by the isolated organism, the weight loss of the polymer films during its growth in minimal medium containing either PUR or LDPE films as the sole carbon source was measured at different time points as indicated in Fig. 2. Approximately 30% and 70% reduction of the initial weight of the PUR film was observed after 5 and 10 days of incubation, respectively (Fig. 2a, left panel). The maximum reduction of the weight of the films took place between 5 and 10 days. Similarly, there was also a marked degradation of the LDPE film with time. After 5, 15, 30, and 45 days of incubation, a loss of ~ 4, ~ 12%, ~ 18%, and ~ 20% of the initial weight was observed, respectively (Fig. 2a, right panel). Thus, these results showed that with time, AKS31 caused significant weight reduction of both the PUR and LDPE films.

Fig. 2.

Fig. 2

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Degradation kinetics of PUR-and LDPE films by AKS31. a Degradation profile of polymers under the in vitro laboratory conditions. AKS31 was grown in minimal medium containing either PUR (left panel) or LDPE (right panel) as the sole carbon source. The films were removed from the culture at different time points as indicated, washed and the dry weight of the films was measured. The results represent an average of at least three independent experiments. Error bars indicate standard deviation (± SD). *p < 0.05, **p < 0.001, ***p < 0.0001. b Degradation profile of PUR and LDPE-films in various soil microcosms. Either PUR (left panel) or LDPE (right panel) films were added to three different microcosms- sterile, naturally attenuated, and bioaugmented and incubated at 30 °C for 30 days and 45 days, respectively. Following incubation, PUR and LDPE films were recovered, washed, and weighed. The results shown are the average of three independent experiments. Error bars indicate standard deviation (± SD). *p < 0.05, ***p < 0.0001

The ability of AKS31 to degrade the polymers in the soil was also examined. Towards this, two sets of soil microcosms; (i) attenuated containing naturally occurring microbes, and (ii) bioaugmented, containing naturally occurring microbes and exogenous AKS31, were set up using soil collected from the local solid waste dumping ground. Sterile soil was used as a negative control. PUR and LDPE films were inserted in these microcosms and their weight loss was examined after 30 and 45 days of incubation, respectively. The result showed that in bioaugmented microcosm, there was a loss of ~ 45% and ~ 14% of the initial weight of PUR and LDPE films, respectively (Fig. 2b, left panel). In contrast, films inserted in naturally attenuated microcosm exhibited a reduction of ~ 12% and ~ 2% of their weight, respectively (Fig. 2b, right panel). Based on the increased degradation of the polymers by the bioaugmented microcosm compared to the naturally attenuated one, we conclude that the isolated organism retains its ability to degrade both PUR and LDPE in the soil environment.

Isolated organism is a strain of Pseudomonas

To identify AKS31, its 16S rRNA gene was sequenced and the sequence was submitted to GenBank(accession number: KY849590). The 16S rRNA gene of AKS 31 has significant identity with the previously-reported 16S rRNA gene sequences of different members of Pseudomonas genera (maximum identity of 99% with Pseudomonas sp. CYEB-6, having accession number FJ422398.1). A phylogenetic tree, constructed on the basis of sequence similarity, showed that AKS31 is strongly associated with Pseudomonas sp. (Fig. 3a). To gain further confidence, several biochemical tests were also conducted for AKS31. Consistent with the ribotyping data, results of the biochemical tests also indicated that the isolated organism belongs to the genus Pseudomonas (Fig. 3b). Hence, the isolated organism was named as Pseudomonas sp. AKS31.

Fig. 3.

Fig. 3

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Characterization of AKS31. a Phylogenetic tree of AKS31. The unrooted tree was constructed based on the 16S rDNA sequences. Numbers at branches indicate the corresponding bootstrap value. Mesorhizobium loti LMG 17826t2 (AJ315352) was used as an outgroup. b Biochemical characterization of AKS31. The indicated biochemical tests were performed for the characterization of AKS31. Experiments were done at least three times

Exogenous addition of AKS31 to soil maintains soil health

For the verification of soil health after the addition of AKS31, heterotrophic organism counts along with the number of PUR degraders present in different microcosms were determined. The result showed the presence of a greater number of heterotrophs and PUR degraders in bioaugmented soil than naturally attenuated soil (Fig. 4a). Although the number of both heterotrophs and PUR degraders increased in the bioaugmented soil, the heterotrophic count is ~ 1500 fold more than the count of PUR degraders in this microcosm. This result indicates that the exogenous addition of AKS31 positively influences the growth of other heterotrophs resulting in higher heterotrophic counts.

Fig. 4.

Fig. 4

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Exogenous addition of AKS31 does not affect soil health. a Heterotrophic count. One gram of each soil sample, from sterile, naturally attenuated (NA) and bioaugmented microcosms, was serially diluted in 0.85% NaCl and then spread onto either LA plates, or plates containing PUR as the sole carbon source, to determine the heterotrophic counts and PUR degrader count, respectively. PUR plates were incubated at 30 °C for 5 days and LA plates were incubated at 30 °C for 2 days. Colonies that appeared on each plate were counted. The results represent the average of three independent experiments. The error bar indicates the standard deviation (± SD). *p < 0.05, ***p < 0.0001. b Soil organic carbon content. Soil samples from sterile, naturally attenuated (NA) and bioaugmented microcosms were taken and analyzed for their organic carbon content. The results represent the average of three independent experiments. The error bar indicates the standard deviation (± SD). **p < 0.001. c Soil respiration profile. Soil samples from sterile, naturally attenuated (NA) and bioaugmented microcosms were taken and subjected to various stress conditions as indicated in the figure, followed by measurement of soil respiration by the hanging tube method. The results represent the average of three independent experiments. The error bar indicates the standard deviation (± SD). **p < 0.005, ***p < 0.0001

The experiment described above investigated only the culturable population. To assess the status of unculturable organisms as well, we measured the soil organic carbon content of bioaugmented and naturally attenuated microcosms. It was observed that after an incubation of 30 days, the bioaugmented microcosm showed significantly higher levels of soil organic carbon content than the naturally attenuated microcosm (Fig. 4b). We also examined the metabolic activities of soil microorganisms in different microcosms by performing soil respiration assay. Consistent with the previous results, an increase in respiration was observed for the bioaugmented sample, compared to the attenuated microcosm (Fig. 4c, upper panel). Soil respiration was also measured under heat and desiccation stress and the results showed that under both these conditions soil respiration was higher in the bioaugmented sample than in the naturally attenuated one (Fig. 4c, middle and lower panels). The above results document that the exogenous addition of AKS31 helps in increasing biomass by positively influencing metabolic activity and proliferation of the organisms present in the soil. Thus, the exogenous addition of AKS31 to the soil is not deleterious for soil health.

AKS31-mediated PUR and LDPE breakdown involves esterase and hydroxylase activities, respectively

Since PUR contains hydrolyzable ester bonds, the activity of esterase is likely to be involved in its breakdown with the accompanying production of diethylene glycol (DEG). To verify the involvement of esterase activity, HPLC, and MALDI-TOF MS was performed sequentially to detect the presence of DEG in the conditioned media. The components of the conditioned media, obtained after growth at 30 °C for 5 days and 10 days, were separated by HPLC using the C18 column. Figure 5a i, ii, and iii shows the HPLC profiles for the standard DEG, conditioned media, and the uninoculated medium, respectively. The result showed the presence of peaks (marked with *) corresponding to DEG in the conditioned media (Fig. 5a i and ii). These peaks were not present in the uninoculated medium (Fig. 5a iii) and thus the generation of these peaks can be attributed to microbial metabolic activity. The subsequent MALDI-TOF MS analysis of the peak marked with ‘a’ detected sodiated DEG at m/z 129 (MW. of DEG is 106). Besides, MALDI-TOF MS analysis also detected dimeric sodium DEG adducts at 277.09 (Fig. 5b). Thus, this result demonstrated that peak ‘a’ indeed contains DEG. The absence of this peak in the uninoculated medium indicates that diethylene glycol is a PUR-breakdown product. Since DEG can only be obtained by hydrolytic cleavage of the ester bond present in PUR, AKS31-mediated PUR degradation can be attributed to its esterase activity.

Fig. 5.

Fig. 5

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Esterase activity is involved in PUR degradation. a High Performance Liquid Chromatography (HPLC) profile. AKS31 cells were grown separately at 30 °C for 5 days and 10 days in minimal media containing PUR as the sole carbon source. Cells were harvested and 20 µL of the cell-free supernatant was applied to the C18 column for the separation of the components through HPLC (i). HPLC chromatogram of pure diethylene glycol. Black and blue colours represent the spectrum for 10 mg/mL and 100 mg/mL of DEG, respectively. Green colour represents the profile for the buffer. (ii). HPLC chromatogram of conditioned media. Black colour represents 10 days old conditioned medium and blue represents 5 days old conditioned medium. (iii). HPLC chromatogram of the uninoculated medium. Peaks corresponding to diethylene glycol are marked with *. The experiment was done three times. The result shown is representative of these three. b Matrix assisted laser desorption ionization time−of−flight (MALDI−TOF) mass spectrum profile. HPLC fraction corresponds to peak ‘a’ obtained from 10 days old conditioned medium was subjected to MALDI−TOF MS. The peaks of the spectrum are marked with their m/z ratio. c PCR amplified fragment of the PURase gene. PCR was carried out using genomic DNA from AKS31 as template and PURase specific degenerate primers. PCR product was run on 1.5% agarose gel and visualized under UV after staining with ethidium bromide. L represents the DNA ladder

To investigate the presence of PURase gene in AKS31 genome, we have aligned the published sequences of the PURase gene (Supplementary Figure S1) and designed primers from the conserved region to amplify the part of the gene using AKS31 genomic DNA as template. As per the aligned sequences the expected size of the PCR products will be ~ 350 bp. Consistent with this, the result showed amplification of ~ 350 bp nucleotides (Fig. 5c). Subsequent sequencing of this PCR product and blastn (NCBI) analysis of the obtained sequence exhibited a high level of identity (> 90%) with many of the published lipase genes that encode PURase. The nucleotide sequence of the PCR product and alignment with one of such hits have been shown in Supplementary Table S1. Direct blastx (NCBI) analysis of this sequence also showed several hits to PURase enzyme with high level of identity (> 95%). The bottom panel of Supplementary Table 1 shows one such alignment. Taken together these results indicate that an esterase activity is involved in this AKS31-mediated PUR degradation.

Existing literature documents that incorporation of hydroxyl (-OH) group facilitates the breakdown of LDPE. To verify the incorporation of -OH group, FTIR analysis of the treated and untreated films was carried out. The result showed an appearance of a sharp trough (marked with *) at ~ 1025 cm−1 with a concomitant appearance of a broad trough in between 3200 to 3600 cm−1 in the treated film only (Fig. 6). This result indicated the incorporation of –OH group into the polymer suggesting the involvement of hydroxylase enzyme(s) for AKS31 mediated LDPE degradation.

Fig. 6.

Fig. 6

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Fourier transformed infrared (FTIR) analysis profile of LDPE films. AKS31 cells were grown in sterile minimal medium supplemented with LDPE films at 30 °C for 30 days. After the incubation, LDPE films were recovered and subsequently subjected to FTIR analysis. LDPE film not treated with AKS31 was used as control. Upper and lower panels represent FTIR profiles for AKS31−untreated and −treated films, respectively. Images are representative of three independent experiments

Isolated organism exhibited altered surface hydrophobicity for different polymers

Attachment of a microbe to a polymer film increases the efficiency of polymer degradation by its enzyme(s). During this study, it has been noticed that the isolated organism can attach to both PUR and LDPE films (data not shown). In a previous report, we have shown that bacterial attachment on the polymer surface is directly associated with cell surface hydrophobicity [Tribedi et al. 2012]. Since PUR and LDPE are hydrophilic and hydrophobic in nature, respectively, attachment of the organism to these two different polymers is possible only if the isolated organism has the ability to modulate its cell surface hydrophobicity accordingly. To investigate this, the cell surface hydrophobicity of the organism was examined in the presence of either PUR or LDPE films by BATH assay as described in the methods and materials. Interestingly, we observed that the cell surface hydrophobicity changed with the hydrophobicity of the polymers (Fig. 7). The organism exhibited high surface hydrophobicity when it was exposed to hydrophobic polymer LDPE (Fig. 7). In contrast, hydrophobicity was markedly decreased when it was exposed to PUR film, which is hydrophilic in nature (Fig. 7). Thus, the isolated organism shows a unique feature, wherein it can change its surface hydrophobicity on encountering different polymeric surfaces.

Fig. 7.

Fig. 7

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AKS31 displays differential surface hydrophobicity in presence of different polymers. AKS31 cells were grown separately in sterile minimal medium supplemented with either PUR or LDPE as the sole

source of carbon for 5 days and 30 days, respectively. Next, cells were harvested, washed with sterile water, and subsequently BATH assay was performed to determine the cell surface hydrophobicity. The results represent the average of three independent experiments. The error bar indicates the standard deviation (± SD). *p < 0.01***p < 0.0001

Discussion

To address a critical environmental problem arising from the burden of plastic, the present study isolated a bacterium from the soil that belongs to Pseudomonas sp. that is capable of degrading two widely-used polymers, PUR and LDPE. The experimental results showed that the isolated bacterium not only degrades the polymers under laboratory conditions but also exhibits substantial activity in the soil environment without negatively affecting soil health. In fact, important parameters like soil respiration and soil organic carbon were found to be more for the bioaugmented microcosm, compared to the corresponding naturally attenuated one. Additionally, a higher level of soil respiration was also evident under heat and desiccation stress. These results indicated the presence of higher microbial diversity in the bioaugmented microcosm compared to its attenuated counterpart. This was further supported by the fact that the heterotrophic microbial count in the bioaugmented soil is markedly higher than the naturally attenuated microcosm. The increased growth can be attributed to the fact that degradation of PUR by AKS31 generates some products, such as adipic acid and DEG, which then serve as good nutrients for other organisms to grow; the result is a microbial population with higher metabolic diversity.

To understand the mechanism of PUR degradation by AKS31, PUR-hydrolyzed products in the growth medium were looked for and detected the generation of DEG. Since DEG will be produced from PUR by the hydrolysis of ester bonds within the polymer, it indicates that AKS31 hydrolyses PUR in the growth medium by producing esterase(s). This conclusion is supported by the identification of a mutant of AKS31 that has reduced esterase activity and less PUR degradation ability compared to the wild type (Supplementary Figure S2). Thus, there is a positive correlation between esterase activity and PUR degradation. The presence of a PURase gene in AKS31 lends further support to this notion. Thus, all these results indicate that esterase activity plays an important role in the AKS31 mediated PUR degradation process.

Towards gaining an insight into LDPE degradation, FTIR spectroscopy was done for the AKS31-treated and -untreated LDPE films as this is a useful tool to determine the presence of different functional groups, including unsaturation. Since polyethylene biodegradation entails the oxidation of the polyethylene chain, it leads to the incorporation of OH/carbonyl group (Yoon et al. 2012). Consistent with this, the FTIR data of AKS31-treated LDPE showed incorporation of -OH within the polymer backbone. Since this incorporation arises due to AKS31 activity, this result indicates that AKS31 produces enzyme(s) capable of introducing –OH group into LDPE that leads to the degradation.

Finally, it can be noted that AKS31 exhibited an interesting and unique property of altering its surface hydrophobicity in response to the change in its environment. This property enables it to degrade both PUR and LDPE efficiently, even though their surface property, with respect to hydrophobicity, is vastly different. Since attachment of the microbe to a given polymer substrate is an important contributor to subsequent efficient biodegradation, this ability of AKS31 to change its surface hydrophobicity is a useful feature that is likely to greatly enhance its potential as an effective tool for biodegradation of various polymers.

In conclusion, the present study reports the isolation of a naturally occurring soil microorganism named Pseudomonas sp. AKS31 capable of degrading two-widely used polymers, LDPE and PUR to a significant level in both the laboratory and soil microcosm. This microorganism positively affected soil health and the environment when it was exogenously added to the soil. Thus, this organism could potentially be a good candidate for the biodegradation of PUR-and LDPE-derived contaminants from the environment.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOC 1330 KB) (1.3MB, doc)

Acknowledgements

Authors would like to thank Professor Srimonti Sarkar, Bose Institute, Kolkata for the critical reading and constructive criticism during the manuscript preparation. This work is supported by a grant from the Department of Science and Technology, Government of West Bengal, India (701/(Sanc.)/ST/P/S & T/2G-3/2010, Dated 03.12.2015). RR (UGC-NET Fellow) and GM (CSIR-NET Fellow) extend sincere thanks to the UGC and CSIR, Govt. of India, respectively, for providing financial assistance for carrying out the research work. Thanks to DBT-IPLS programme at the University of Calcutta for various instrumental support.

Compliance with ethical standards

Conflict of interest

The authors declare that they do not have any conflict of interest.

Contributor Information

Rusha Roy, Email: rusha.r19@gmail.com.

Goutam Mukherjee, Email: rkmv.raja426@gmail.com.

Anirban Das Gupta, Email: joy.dasgupta1987@gmail.com.

Prosun Tribedi, Email: tribedi.prosun@gmail.com.

Alok Kumar Sil, Email: alokksil7@gmail.com.

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