Structural and mechanical characterization of biofilm-associated bacterial polymer in the emulsification of petroleum hydrocarbon

Abstract

The marine bacterium Pseudomonas furukawaii PPS-19 isolated from the oil-polluted site of Paradip port, Odisha, India, was found to form a strong biofilm in 2% (v/v) crude oil. Confocal Laser Scanning Microscopy (CLSM) revealed biofilm components along with multi-layered dense biofilm of rod-shaped cells with 64.7 µm thickness. Scanning electron micrographs showed similar biofilm architecture covered with a gluey matrix of extracellular polymeric substances (EPS) in the presence of 2% (v/v) crude oil. The architecture of purified EPS was also studied through FESEM that exposed its porous and three-dimensional flakes-like structure. The structural characterization by FTIR revealed that EPS was composed of primary alkane, amines, halide, hydroxyl groups, uronic acid, and saccharides. The XRD profile exhibited an amorphous phase of the EPS with a crystallinity index of 0.336. The EPS showed three-step thermal decomposition and thermal stability up to 600 °C, as confirmed by TGA and DSC thermogram. EPS produced by marine bacterium P. furukawaii PPS-19 could act as bioemulsifier and showed the highest emulsifying activity of 66.23% on petrol. The emulsifying ability of the EPS was superior to the commercial polymer xanthan. The emulsion also showed high stability with time and temperature exposure. The marine bacterium P. furukawaii PPS-19 and the EPS complex showed 89.52% degradation of crude oil within 5 days. These properties demonstrated the potential of biofilm-forming marine bacterium as bioemulsifier for its application in the bioremediation of oil-polluted sites.

Supplementary Information

The online version contains supplementary material available at 10.1007/s13205-021-02795-8.

Introduction

Marine environments have been polluted by a variety of organic pollutants due to catastrophic events and anthropogenic activities. Petroleum hydrocarbons have been the principal cause of marine contamination due to their use as an energy source in various industries (Varjani et al. 2015; Varjani and Upasani 2016). There are many severe effects on the environment of these pollutants as they are highly toxic to the marine life and surrounding ecosystem (Iwabuchi et al. 2002). Every year, about 1.3 million tons of petroleum hydrocarbons are released into the ocean from industries or accidental oil spills. Therefore, the U.S Environmental Protection Agency classified petroleum as priority environmental pollutants (Todd et al. 1999; Costa et al. 2012). However, the microbial community residing in the polluted sites form biofilm as a strategy for growth and survival in adverse conditions and produce extracellular polymeric substances (EPS). EPS are three-dimensional, gel-like, highly hydrated matrix, self-produced by biofilm microorganisms in which they are embedded and more or less immobilized. Therefore, the biofilm is metaphorically called a “city of microbes” (Watnick and Kolter 2000), and the EPS represents the “house of biofilm cells” (Flemming et al. 2007).

EPS are biopolymers of a complex mixture of polysaccharides, proteins, nucleic acids, lipids, uronic acids, humic substances etc. (Pal and Paul 2008; Jayathilake et al. 2017). EPS are the major structural and functional component of microbial biofilm. EPS provide structural stability to the biofilms and maintain their metabolic activity (Mangwani et al. 2016). The central role of EPS is to facilitate the attachment of microbial cells to biotic and abiotic surfaces. Marine environments are dynamic where marine bacteria are adapted to survive in extreme environmental conditions like temperature, pH shift, salinity, pressure fluctuation, osmotic stress, nutrient availability, heavy metal accumulation (Bhaskar and Bhosle 2005). The survival strategy in such a harsh environment includes the biosynthesis of EPS. The functional groups in the biopolymer facilitate natural ligand binding interaction with other groups with opposite charges. This way, the EPS matrix defends against harsh environmental conditions (Chowdhury et al. 2011). EPS influences the physicochemical environment around the bacterial cell that enhances the survival of marine bacteria (Sun et al. 2015; Shukla et al. 2017). Changes in environmental conditions lead microorganisms to the production of novel biomolecules. Bacteria inhabiting extreme environments adopt unique metabolic pathways and defensive mechanisms to persist adverse conditions (Nicolaus et al. 2010; Poli et al. 2010). The composition and structure of EPS generally depend on the environment and ecological niche.

The high molecular weight EPS having amphiphilic nature acts as an emulsifier (Sheng et al. 2010). Bioemulsifier forms and stabilize oil-in-water or water-in-oil emulsion by increasing their kinetic stability. Bioemulsifier gets accumulated at the interface between two immiscible phases by reducing the surface tension. Thus, it increases the solubility and emulsification of immiscible phases leading to enhanced degradation of petroleum hydrocarbon (Uzoigwe et al. 2015; Karlapudi et al. 2018). This makes hydrophobic substrates easily accessible to the indigenous degrading microorganism. The accumulation of emulsion at the oil–water interface in the form of a closely packed layer provides mechanical stability by preventing both coalescence and flocculation by a steric barrier (Peele et al. 2016). To circumvent the limitations of hydrocarbon availability, some bacteria produce emulsifiers to increase the availability of these compounds for their energy and carbon source, resulting in biodegradation of hydrocarbons. The role of EPS as an emulsifier could be valuable in detoxifying petrochemical oil-polluted areas (Banat et al. 2000; Batista et al. 2006; Zheng et al. 2011).

The EPS production is essential for biofilm formation, stability, architecture, and metabolism, which help biofilm survival in a petroleum hydrocarbon polluted environment. Thus, the characterization of EPS from the relevant bacteria present in the marine environment is essential for gaining a better understanding of the structure–function interactions in biofilm. The involvement of marine bacteria in mobilized or immobilized conditions to remove petroleum hydrocarbon pollution has been extensively studied. However, the characterization of EPS and its utilization in the degradation of petroleum hydrocarbon from biofilm-forming marine bacteria from the petroleum hydrocarbon polluted environment has not received much attention. Consequently, there is a lack of knowledge about the formation of stable oil droplets in the marine environment through bacterial biofilm.

The present work is distinct as we report the biofilm formation of the marine bacterium P. furukawaii PPS-19 at the oil–water interface and the role of biofilm-associated EPS in the stabilization of the interface to prevent coalescence of oil droplets. Thus the physicochemical characterization of biofilm-associated EPS was studied. Functional properties of EPS, including its emulsion forming and stabilizing capacity and its degradability of petroleum hydrocarbon, are also reported. The present study aims to provide an environmentally sustainable and natural material for the emulsification of petroleum hydrocarbon. EPS produced by marine bacterium P. furukawaii PPS-19 could be a rational and natural method in controlling oil contamination in the marine environment.

Materials and Methods

Materials

Crude oil (ASTM D5307, Supelco) and ethanol were purchased from Merck Millipore, India. All the dyes used, such as Concanavalin A (Con A) Alexa fluor 633 conjugate, FITC, Calcofluor white, SYTO 9, and the Nile red procured from Invitrogen by Thermo Fisher Scientific, USA. Toluene and Hexane were purchased from Sigma-Aldrich, USA. Culture media were procured from Hi-media, India. Petrol and diesel were purchased from an oil petrol station.

Identification and culture condition of the bacterial strain

The bacterial strain was isolated from the sediment of oil-polluted site of Paradip port, Odisha, India. The bacterial strain was identified by 16S rRNA partial gene sequencing. The contig sequence was matched with those available in the GenBank database using BLASTn (https://blast.ncbi.nlm.nih.gov/Blast.cgi?PAGE_TYPE=BlastSearch) as well as with EzBioCloud database (https://www.ezbiocloud.net). The related FASTA format sequences were downloaded and aligned using Clustal W (1.6) DNA weight matrix in MEGA 6. The aligned sequences were used to construct the phylogenetic tree using the neighbour-joining statistical method. Tamura-Nei model was used to compute the evolutionary distances clustered together in 1000 bootstrap replicates. A partial sequence of 16S rRNA gene has been submitted to the NCBI GenBank database. The bacterial strain has been deposited to Microbial Culture Collection (MCC), National Centre for Microbial Resources (NCMR), Pune, India. The working culture was maintained on Luria Bertani agar (Hi-media, India) by incubation at 37 °C for 24 h and stored at 4 °C.

Biofilm assay

Biofilm assay was performed by microtiter plate method following O'Toole (2011) with slight modifications (Mangwani et al. 2014). 20 µl of the overnight grown culture (O.D_600nm = 0.4) was inoculated into 180 µl of Bushnell Hass broth medium (BHB) (Hi-media, India), supplemented with 2% crude oil. 1% Glucose supplemented in BHB medium was used as the positive control. The microtiter plate was incubated for 48 h in a moist chamber at 37 °C in static condition. After 48 h, the used media and the planktonic cells were carefully taken off from the wells without disturbing the biofilm layer formed on the microtiter plate wells. After that, the plate was washed with 1X PBS (pH 7.4) and air-dried. The attached biofilm was stained with 200 µl of 0.2% crystal violet for 10 min, followed by washing with 1X PBS. The plate was again air dried, and then 200 µl of 30% acetic acid was added to each well of the microtiter plate followed by incubation for 30 min. Optical density was measured using a 96-well Plate Reader (Victor X3, Perkin Elmer, USA) at 562 nm. The assay was conducted in triplicate, and the mean ± standard deviations was calculated. Cut-off OD (ODcut) was the three standard deviations above the mean OD of the negative control. The isolate was further characterized as weak, moderate, and strong biofilm formers following Stepanović et al. (2007).

Extraction of EPS

EPS was extracted by ethanol precipitation method following Chowdhury et al. (2011) with modifications. Mature biofilm was developed on glass beads (2 mm diameter) for 48 h at 37 °C in static condition. After 48 h, the consumed media was discarded and washed with 1X PBS. The glass beads were vortexed to disrupt the biofilm. The biofilm sample was collected and harvested at 7000 rpm for 10 min at 4 °C. EPS was precipitated from the supernatant by mixing with the triple volume of prechilled absolute ethanol. The mixture was then incubated overnight at 4 °C. The precipitated EPS obtained was separated by centrifugation at 7000 rpm at 4 °C for 20 min and purified through 14 KDa molecular weight cut-off dialysis membrane (cellulose) against deionized water at 4 °C for 24 h. The dialyzed EPS solution was then lyophilized and used for further characterization studies.

Structural characterization of biofilm-associated EPS

Biofilm was characterized by Scanning Electron Microscopy (SEM) and Confocal Laser Scanning Microscopy (CLSM). For CLSM, overnight grown culture was inoculated in LB medium in a falcon with a glass coverslip of dimension 5 × 2 cm and incubated at 37 °C under the static condition for 48 h. The glass coverslips were washed twice with 1X PBS. Biofilm and EPS were stained with 5 µmol l⁻¹ SYTO 9 and 100 µg ml⁻¹ Concanavalin A (Con A) Alexa Fluor 633 conjugate, respectively, for 2 min. Calcofluor white (5 mM), FITC (15 mM), and Nile red (0.5 µg ml⁻¹) were used to stain biofilm-associated EPS components like β-polysaccharides, proteins, and lipids, respectively. The coverslip was then washed with 1X PBS to remove the excess stain. It was then viewed under Leica TCS SP8 confocal system (Leica Microsystems, Wetzlar, Germany) with water immersed 63× objective lens. Confocal micrograms were analyzed by Image J 1.46 software to calculate biofilm thickness (Nancharaiah et al. 2005).

For SEM, overnight grown culture was inoculated in the BHB medium supplemented with 2% crude oil in a six-well polystyrene plate containing a cut glass slide (1 cm × 1 cm). BHB medium supplemented with 1% Glucose was used as the positive control. The plate was incubated at 37 °C under static conditions for 48 h. The glass slides were taken out slowly, washed twice with 1X PBS, and then fixed with 2.5% (v/v) glutaraldehyde. It was then incubated at 4 °C for 4–6 h. After that, it was dehydrated sequentially with an ethanol gradient of 20%, 40%, 60%, 80%, and 100% (Asahi et al. 2015) and gold coated. The image was captured with ×20,000 and ×80,000 magnifications using SEM (FEI Quanta 250, Thermo, USA).

Surface morphology and microstructure of the extracted EPS were also observed using field emission scanning electron microscopy (FESEM) (Nova NanoSEM 450/FEI, USA). The lyophilized EPS was mounted on a glass slide (1 mm X 1 mm) with carbon tape and placed on the metal stab for gold coating. Micrographs were recorded at ×2000 and ×10,000 magnification. EPS elemental composition was also determined by an Energy Dispersive X-ray analyzer (EDAX) equipped with FESEM, which revealed different elements such as weight and atomic percentage.

Chemical composition and functional group analysis of biofilm-associated EPS

Total carbohydrate content in the EPS solution was determined quantitatively by the Phenol–sulphuric acid method (Dubois et al. 1956). The absorbance was taken at 490 nm wavelength using UV–Vis Spectrophotometer (Lambda 35 Perkin Elmer, USA). The protein content of the aqueous EPS was determined following the Bradford method (Bradford 1976). The absorbance was taken at 595 nm wavelength using UV–Vis Spectrophotometer (Lambda 35 Perkin Elmer, USA).

The attenuated total reflection-Fourier transform infrared (ATR FT-IR) (Alpha ATR FT-IR, Bruker, Germany) spectrum was obtained to detect the major functional group in the EPS. Spectrum was recorded in the transmission mode from 4000 to 400 cm⁻¹ with a resolution of 4 cm⁻¹, and 32 scans were averaged. X-ray diffraction (XRD) study was performed on an X-ray diffractometer (X-Pert, PANalytical, PW 3040/00, Netherlands) using Ni-filtered CuKα (λ = 1.54056 Å). The powdered form of lyophilized EPS was examined by maintaining the 2θ range between 10° and 90°, and other running conditions were 0.005° scan speed, 0.1 s time/scan, and 0.2 mm slit. The intensities of diffracted peaks were continuously recorded and plotted as a 2θ value. The interplanar d-spacing at different θ values was calculated using Bragg's law (d = λ/2 Sinθ). The crystallinity index (CI_XRD) was calculated from the ratio of areas under the crystalline peaks to the sum of total diffracted area using equation (Ricou et al. 2005) CI_XRD = ∑ A_{Crystalline peaks}/(∑ A_{Crystalline peaks} + ∑ A_{amorphous peaks}).

Thermal gravimetric analysis (TGA) and Differential scanning calorimetry (DSC)

The pyrolysis pattern of the lyophilized EPS was studied using a thermal analyzer (NETZSCH, STA409C, Germany). The experiment was carried out in a dynamic argon atmosphere at a heating rate of 10 °C min⁻¹ by gradually raising the temperature in a range of 29 °C to 800 °C. TGA and DSC thermograms were plotted as the weight change (percentage) and heat flow as a temperature function, respectively (Mishra et al. 2011). The enthalpy transition and crystallinity index of the EPS (CI_DSC) were calculated following Singh et al. (2011) using equation CI_DSC = ΔH_Net/H_Total where ΔH_Net is the difference between the heat of crystallization and melting.

Emulsifying activity of EPS

The potential of EPS to emulsify petroleum oils (petrol and diesel) and hydrocarbons (hexane, toluene) was carried out following the method described by Cooper and Goldenberg (1987) and modified by Freitas et al. (2011). Briefly, an aqueous solution (2% w/v) of EPS was added to each hydrophobic compound in a ratio of 3:2 (v/v). A screw cap glass test tube of dimension 20 mm × 180 mm was used to perform the assay. The mixture was vortexed at maximum speed for 2 min and left to settle at room temperature for 24 h. Tween 80 and Xanthan gum were taken as the positive control and water as the negative control. The emulsion indexes (E) of petroleum oils and hydrocarbons were expressed as the percentage of the total height occupied by the emulsion after 24, 168, and 360 h. E = (h_E/h_t) × 100, where h_E is the height of the emulsion and h_t is the total height of the mixture.

The effect of different concentrations of EPS and temperature on the stability of emulsion was determined on the basis of the E indexes of the most promising emulsion. The standard assay for emulsion stability was based on the method described by Castellane et al. (2015) and Kielak et al. (2017). The new emulsions with different EPS concentrations (0, 0.5, 1, 1.5, and 2 g l⁻¹) were prepared, and the emulsion indexes (E) were calculated after 24 h. Additionally, to evaluate the thermal stability of the emulsion, emulsions were heated to 85 °C for 50 min and then cooled to room temperature. The emulsification capacity of the EPS before and after heat treatment was measured and compared. All the tests were conducted in triplicate.

Microcosm study

Microcosm study was performed in four sets of experiments in an artificial seawater medium (ASW) following Gong et al. (2015) with slight modification. The first set included 5 ml ASW medium amended with 2% (v/v) crude oil; the second set included 5 ml ASW medium amended with 2% (v/v) crude oil, and 2 g l⁻¹ EPS; the third set included 5 ml ASW medium amended with 2% (v/v) crude oil and P. furukawaii PPS-19, and the fourth set included 5 ml ASW medium amended with 2% (v/v) crude oil, 2 g l⁻¹ EPS, and P. furukawaii PPS-19. For the degradation of petroleum hydrocarbon, P. furukawaii PPS-19 was inoculated in sets three and four following Mahto and Das (2020). Briefly, P. furukawaii PPS-19 was grown overnight (O.D₆₀₀ 0.4) and then diluted to 1:100 in LB medium in a conical flask. It was further incubated at 37 °C in static for 48 h. After that, planktonic cells were decanted carefully, and biofilm was washed with 1X PBS and inoculated in sets three and four. For sets two and four, UV sterilized extracted EPS was added. Set one was considered as control. All sets of conical flasks were incubated at 30 °C in shaking (140 rpm). Residual crude oil was extracted using an equal volume of n-hexane at every 24-h time interval for 5 days. All sets of conical flasks were vortexed vigorously for 2 min. It was then centrifuged at 7000 rpm for 10 min for separation of the aqueous and organic phase. The upper organic phase was taken out carefully and analyzed using high-performance liquid chromatography (HPLC) (Agilent 1260 Infinity Series, USA) equipped with a photodiode array detector at 254 nm. Total viable cells inoculated at the set three and four were also counted.

For calculation, a standard of crude oil in n-hexane concentrations from 1%, 2%, 3%, 4%, and 5% were prepared and analyzed under the same conditions as samples. The ratios of the peak areas to corresponding concentrations were linear with very high correlations (r² ≥ 0.998). Quantification of residual crude oil in all sets was performed by integrating peak areas, and the concentration of residual crude oil was determined with the standard curve. The degradation efficiency of each set was calculated using the following equation (Bao et al. 2013): Degradation % = (Co–C)/Co × 100 where Co and C represent the concentration of crude oil at the control and concentration of crude oil in the degraded sample, respectively. All the experiments were conducted in triplicate.

Statistical analysis

All the data reported in this manuscript were performed in triplicate, and the results are presented as mean ± SD. To evaluate the significant change in biofilm growth, an unpaired t test was performed. Emulsification activity result was subjected to One-way analysis of variance (ANOVA), and differences among mean were tested using Tukey’s test. The two-way ANOVA was used to analyze the effects of different concentrations of EPS and temperature on emulsion stability. P < 0.05 was considered a statistically significant value. GraphPad (Prism version 5.01) software was used for all statistical analysis.

Results

Phylogenetic analysis and the growth conditions of the bacterial strain

Pseudomonas furukawaii strain PPS-19 was isolated from the sediment sample of petroleum hydrocarbon contaminated site of Paradip port, Odisha coast, India using crude oil amended basal medium. The partial sequence (1420 bp) of 16S rRNA gene of the marine bacterium P. furukawaii PPS-19 was submitted to the NCBI GenBank (Accession number MT588911). The EzBioCloud database analysis revealed a 99.65% similarity with P. furukawaii KF707. The phylogenetic tree analysis designated P. furukawaii PPS-19 and P. furukawaii KF707 under the single node and indicated a close relationship of P. furukawaii PPS-19 with P. furukawaii KF707 (Fig. S1). The strain has been submitted to the Microbial Culture Collection (MCC), National Centre for Microbial Resources (NCMR), Pune, India (Accession number MCC 4630).

Quantitative assessment of biofilm-forming potential in presence and absence of petroleum hydrocarbon

Biofilm was visible as a purple ring of the attached cells formed on the wall of the 96-well microtiter plates at the oil–water and air–water-interface both in the presence and absence of crude oil, respectively. P. furukawaii PPS-19 showed strong biofilm potential with an absorbance of 0.46 in the presence of 2% (v/v) crude oil, and the control showed an absorbance value of 1.2 (P = 0.0014) (Fig. S2). The ODcut value was 0.07. The robust biofilm formation at the interface showed the potential of EPS in oil–water stabilization. Statistical analysis indicated a significant difference in the effect of carbon source on the biofilm formation ability.

Characterization of biofilm and EPS structure

CLSM images were obtained to study the biofilm community and the distribution pattern of biofilm and matrix composition using ConA, Calcofluor white, SYTO 9, FITC, and Nile red dyes (Fig. 1). SYTO 9 and ConA were used to stain cells and α-polysaccharides (Fig. 1(i) and (ii)). Figure 1(iii) represents 3-D of Z- stack (upper) and 3-D cross-section of Z-stack (lower). CLSM in conjugation with image analysis software Image J was used for the in-depth analysis of biofilm that revealed biofilm thickness of 64.7 µm. Calcofluor white, FITC, and Nile red stained images revealed the presence of β-polysaccharides, proteins, and lipids, respectively (Fig. 1(iv), (v) and (vi)). CLSM micrographs led to a better understanding of the assemblage of biofilm components into a three-dimensional structure. The biofilm thickness showed the dense biofilm-forming ability of P. furukawaii PPS-19.

Fig. 1.

Confocal Laser Scanning Microscopy (CLSM) images of mature biofilm (grown for 48 h at 37 °C) of P. furukawaii PPS-19 showing cells and different components of the EPS matrix, (i) cells stained with SYTO 9; (ii) α- polysaccharides stained with ConA Alexa Fluor 633 conjugate; (iii) 3-D image; (iv) β- polysaccharide stained with Calcofluor white; (v) proteins stained with FITC; and (vi) lipids stained with nile red

Under the scanning electron microscope, biofilm morphology was studied. Short rod-shaped cells were observed both in the presence and absence of crude oil (Fig. 2a). Under both conditions, cells were aggregated together and covered with a layer of EPS matrix. The control biofilm (Fig. 2a(i)) appeared as distinct compared to the biofilm grown in the presence of 2% (v/v) crude oil (Fig. 2a(ii)). The biofilm grown under oil appeared bulkier with firmness. EPS provided the architecture and stability to the biofilm. The formation of mature biofilm in crude oil showed adhesion of biofilm cells, and EPS contributes to the stabilization of the oil–water interface. The scanning electron micrographs of purified EPS at ×2000 and ×10,000 magnification revealed the three-dimensional structure with irregular flakes. The EPS possess porous nature with little compactness (Fig. 2a(iii)). The quantitative analysis of the elemental composition of EPS by the EDX revealed the weight and atomic percentage of nine elements carbon, nitrogen, phosphorus, sodium, sulphur, calcium, iron, potassium, and magnesium (Fig. 2b).

Fig. 2.

SEM micrograph of mature biofilm of P. furukawaii PPS-19 a rod-shaped cells embedded in mucilaginous EPS (inset) (i) grown in BHB medium supplemented with 1% Glucose, and (ii) grown in BHB medium supplemented with 2% (v/v) crude oil, (iii) FESEM micrograph analysis showing microstructure and exterior architecture of the purified EPS, and b EDX spectrum of the purified EPS showing elemental composition (inset). Wt% represents percent by weight, and the At% represents percent by atomic weight

Chemical composition and functional groups of EPS

The bacterium produced 1 g l⁻¹ EPS, and after purification and lyophilization, this EPS was used for chemical analysis. The carbohydrate and protein content per mg of EPS was estimated to be 837 µg mg⁻¹ and 382 µg mg⁻¹, respectively. The ATR FT-IR spectrum showed several strong frequency bands associated with proteins and polysaccharides components of EPS extracted from the bacterial strain P. furukawaii PPS-19. The position of the characteristic functional groups of the EPS is represented in Fig. 3a. The spectrum showed a broad stretching vibration of the hydroxyl group (O–H) at 3461 cm⁻¹. The medium broad stretching peak at 2924 cm⁻¹ was observed corresponding to the C–H vibration of CH₂. A sharp peak at 2360 cm⁻¹ represents the stretching of the amine group (N–H). The C–N and C=O stretching peak at 1643 cm⁻¹, and N–H stretching peak at 1544 cm⁻¹ represent amide I or carbonyl stretching of acetamido groups and amide II, respectively. The short stretch of the absorption peak at 1405 cm⁻¹ was attributed to the symmetric stretching of carboxyl groups (–COO). Another intense peak at 1232 cm⁻¹ represents the stretching of amides (C–N). A long stretch of the peak at 1079 cm⁻¹ was designated to the O-acetyl ester linkage bond of uronic acid. Stretch of weak absorption peaks observed in the region from 800–926 cm⁻¹ corresponds to the C–C and C–O–C vibration bands of polysaccharides. The absorption peak at 682 cm⁻¹ was assigned to the alkyl-halides (C–Br). This result evidenced the functional nature of EPS. The hydrophobic groups present in the EPS impart emulsification activity.

Fig. 3.

Characterization of EPS of P. furukawaii PPS-19 a FTIR spectrum of the purified EPS scanned in the range 500–4000 cm⁻¹, and b XRD profile of the purified, powdered EPS examined with 2θ in the range 10°–90°. The diffractogram represents the sharp diffracted peaks of crystalline portion and broad peaks amorphous part

The X-ray diffraction pattern of EPS is shown in Fig. 3b. The diffractogram exhibited both the sharp narrow peaks and broad peaks that correspond to the crystalline and amorphous parts of the EPS. X-ray diffraction analysis of EPS exhibited characteristic diffraction peaks at 22.12, 24.45, 26.94, 30.28, 36.06, 38.36, 48.08, 53.38, 55.53, 61.45, 62.97, 68.25 and 76.44 with interplanar spacing (d-spacing) of 4.66, 4.22, 3.83, 3.42, 2.88, 2.81, 2.72, 2.19, 1.99, 1.91, 1.75, 1.71, 1.59 and 1.44. The crystallinity index (CI_XRD) of the EPS was found to be 0.336, which revealed the semi-crystallinity with the dominant amorphous phase.

Thermal stability of the EPS

The thermogravimetric analysis of EPS showed three significant weight loss steps (Fig. 4a). In the first step, about 13.15% weight loss was recorded from 35 to 106 °C. The second step degradation from 198 to 332 °C corresponds to another significant weight loss of 46.83%. The third step degradation showed 27.95% weight loss from 492 to 600 °C. The thermostability of EPS up to 600 °C enables additional characteristics for further application. The differential scanning calorimetric thermogram (Fig. 4b) depicted the exothermic transition of the EPS with crystallization temperature (T_c) of 69.18 °C (onset temperature 30.18 °C) accompanied with 343.15 mJ latent energy. After that melting transition started, and the DSC graph showed two melting peaks. The melting temperatures were (T_m1) 424.76 °C (onset temperature 382.18) and (T_m2) 556.90 °C (onset temperature 478.18 °C), with 211.73 mJ and 2030.44 mJ latent energy, respectively. The crystallinity index of EPS (CI_DSC = 0.30) from DSC was in coherence with the XRD. Some variation was due to uncertainties in placing baseline for area integration.

Fig. 4.

Thermal stability of EPS of P. furukawaii PPS-19, a thermogram of the EPS showing thermal stability, and b DSC profile of EPS showing the thermal transition

Emulsifying activity of EPS

Emulsification indices measured for emulsions of EPS formed with petroleum oils and hydrocarbons are shown in Table 1. The emulsifying ability of EPS produced by marine bacterium P. furukawaii PPS-19 with the synthetic surfactant Tween 80 and a commercial and widely used EPS, xanthan, was compared. EPS showed a higher E₂₄ value for petrol (66.23%) (P < 0.0001). The emulsification ability of EPS was more as compared to xanthan. However, Tween 80 was more efficient than bacterial EPS and xanthan, with 100% emulsification efficiency. The lowest emulsification activity was observed for hexane (50%), xanthan (27%), and Tween 80 (90%) (P < 0.0001). All emulsions were stable at room temperature during 15 days of incubation with no sign of droplet coalescence, flocculation, or sedimentation. The effect of EPS concentration (0, 0.5, 1, 1.5, and 2 g l⁻¹) on emulsion stability was studied for petrol. Heat treatment at 85 °C for 50 min showed no reduction of the emulsion forming capacity of EPS (Fig. 5). EPS exhibited the best emulsifying activity at 2 g l⁻¹ concentration. Emulsification indices were slightly lower for the remaining tested concentrations. No significant difference between EPS concentrations was found (P > 0.05).

Table 1.

Emulsification activity of 2% (w/v) EPS solution, Tween 80 and xanthan with various oils and hydrocarbons in 3:2 ratio (v/v)

Oil/hydrocarbons	Tween 80	Xanthan			EPS
	E24	E24	E168	E360	E24	E168	E360
Petrol	100 ± 0.01	64.25 ± 4.25	44 ± 4.96	38 ± 9.05	66.23 ± 0.39	62 ± 0.05	56.45 ± 7.75
Diesel	91 ± 0.05	60 ± 4.8	35 ± 5.0	28.21 ± 2.03	59.53 ± 1.9	61.02 ± 4.61	52 ± 1.12
Toluene	90 ± 1.71	24 ± 3.97	19 ± 2.63	9 ± 2.9	58 ± 2	46 ± 4	40.32 ± 0.53
Hexane	91 ± 2	27 ± 2	17 ± 1.01	5.04 ± 1.42	50 ± 2.1	50.01 ± 4.04	34 ± 1

Results are presented as percentages of total height occupied by emulsion. The experiment was performed in triplicate ± SD (P < 0.0001). E₂₄ emulsification index after 24 h; E₁₆₈ emulsification index after 168 h; E₃₆₀ emulsification index after 360 h

Fig. 5.

Effect of different concentrations of EPS on emulsification index (%). The emulsification index was calculated after 24 h (E₂₄) of incubation of different concentrations of EPS with petrol at room temperature and after an additional 50 min at 85 °C. Error bars represent standard error for three replicates

Biodegradability of petroleum hydrocarbon by bacteria-EPS complex

The degradation rate of crude oil by P. furukawaii PPS-19 in the presence and absence of EPS was carried out by preparing microcosm using ASW medium supplemented with 2% (v/v) crude oil (Fig. 6). The total viable cells in biofilm culture of the set two and three were 2.35 ± 0.2 × 10⁷ cells ml⁻¹. The degradation rate of crude oil in control (set one) was found to be 9.7% that might be due to physico-chemical factors. In P. furukawaii PPS-19 (set three), the degradation rate increased significantly by 67.83%; however, in the presence of bacteria-EPS complex (set four), the degradation rate was increased still further to 89.52% after 5 days. Pure EPS (set two) showed no contribution to the degradation of crude oil.

Fig. 6.

Biodegradation process of crude oil by the biofilm-forming marine bacterium P. furukawaii PPS-19 and the bacterium-EPS complex in an artificial seawater microcosm. The bacterial load inoculated in ASW medium amended with 2% (v/v) crude oil included 2.35 × 10⁷ cells ml⁻¹ with 2 g l⁻¹ EPS

Discussion

Microbial biofilm is considered as a dynamic structural community crucial to confer protection against the hostile marine environment. Biofilm formation in a stressful environment with increased petroleum hydrocarbon is a natural strategy to maintain a favourable niche. In general, biofilm is formed at the air–liquid interface of the static liquids. In a biofilm, microbial cells occupy 10% while 90% is occupied by EPS matrix (Flemming and Wingender 2010; Kavita et al. 2013). The marine ecosystem is called the “mother of origin of life,” encasing vast bacterial diversity. Unlike clinical isolates, marine biofilm offers several advantages, including driving biogeochemical cycles and supplying materials and energy to higher trophic levels (de Carvalho 2018). Due to several unique features of indigenous bacteria in the marine habitat, biofilm-associated EPS of marine bacteria are regarded as the vital component.

In the present study, EPS of the marine bacterium P. furukawaii PPS-19 was characterized. Considering molecular characterization, P. furukawaii PPS-19 was classified under Gammaproteobacteria. The biofilm formation by P. furukawaii PPS-19 was quantified spectroscopically after 48 h of growth, and denser biofilm was observed at 48 h. However, Al-kafaween et al. (2019) found that Pseudomonas aeruginosa ATCC 10145 and Streptococcus pyogenes ATCC 19615 showed strong biofilm 72 h of growth. It is well known that switching from planktonic to the biofilm mode of growth is an intrinsic process in response to environmental changes. Adhesion to a solid surface is the first step in the formation of biofilm. So, biofilm formation depends on the ability of bacteria to adhere to the surface of the microtiter plate by creating hydrophobic interaction (Merritt et al. 2005; Achinas et al. 2019). The marine bacterium was able to form a strong biofilm in 2% (v/v) crude oil at the interface because of the strong attachment of the cells and their self-locking with the produced EPS. EPS alters the cell surface hydrophobicity of the bacteria within the biofilm to regulate their partitioning in the oil–water interface. Bacteria within biofilm utilize hydrocarbons as a source of energy. Marine bacteria were predominantly found to form biofilm and degrade various polycyclic aromatic hydrocarbons (Mangwani et al. 2019). The ability of marine bacteria to form biofilm on multiple substrates is advantageous in survival, adaptation, metabolism, and propagation. One of the major limitations of bioremediation is the bioavailability of organic compounds that could be overcome by biofilm. Hence, marine biofilm could efficiently degrade petroleum hydrocarbon and is an environmentally friendly method for enhanced oil recovery.

Biofilm morphotype and matrix structural elements were observed in the present study after staining with the dyes such as ConA, Calcofluor white, SYTO 9, FITC, and Nile red. Major biofilm matrix components include polysaccharides, proteins, lipids, and nucleic acids, which are involved in complex interaction with each other (Schlafer and Meyer 2017). The SYTO 9 specifically stains live cells as it intercalates to double-stranded DNA present in the cell and eDNA of the matrix (Stiefel et al. 2015). The eDNA helps in the initial stage of biofilm development, strengthens biofilm, and acts as a gene pool during horizontal gene transfer (Wang et al. 2015). Furthermore, staining with ConA, Calcofluor white, FITC, and the Nile red indirectly distinguish the matrix components. ConA is a lectin binding dye that binds specifically to α-d-mannosyl and α-d-glucosyl groups while Calcofluor white binds with β-1,3 and β-1,4 glucans. In general, the biofilm matrix constitutes about 70% of polysaccharides providing a framework to biofilm (Chen et al. 2007). FITC is commonly used to stain protein. The isothiocyanate reactive group of FITC binds with amine and sulfhydryl groups of the protein. Extracellular proteins primarily help in bacterial adhesion to the substratum (Costerton et al. 1995), and lipids play a crucial role in maintaining biofilm integrity (Andrews et al. 2010).

Nile red is a lipophilic stain with distinguished fluorochromes properties at two spectral emissions. The 535 nm emission setting is responsible for the emission of golden yellow colour staining neutral lipid (intracellular). The 590 nm emission setting is for red colour staining polar or hydrophobic region (membrane lipid). The Nile red with emission set at 590 nm was used to study matrix and membrane lipid molecules (Diaz et al. 2008). The biofilm thickness of 64.7 µm demonstrated the dense biofilm forming potential of the marine bacterium P. furukawaii PPS-19. A previous study showed the biofilm thickness of P. aeruginosa and Klebsiella pneumoniae was 29 µm and 100 µm, respectively (Murga et al. 1995). Mean thickness represents the spatial dimension of biofilm and is the most common variable used in biofilm (Heydorn et al. 2000).

Biofilm formation depends on the ability of bacteria to adhere to the solid surface through hydrophobic interaction and form an organized microbial community. SEM and FESEM tools used in the study of biofilm and its polymer help to predict its common physical properties. SEM study revealed that EPS coats biofilm cells and immobilizes them to form a three-dimensional rigid biofilm network. EPS provided the mechanical stability to biofilm both in the presence and absence of crude oil. Kumari and Das (2019) observed a similar study on stress tolerance to metal (Pb) by P. aeruginosa N6P6. The microstructure analysis of EPS using FESEM displayed a porous, compact flake-like structure. The porous structure is essential for water holding capacity. Such porous structure with compactness was observed in EPS obtained from Azotobacter sp. (Gauri et al. 2009). Lactobacillus fermentum CFR2195 also showed a similar flake-like structure with more compactness (Yadav et al. 2011). Besides, EDX microanalysis revealed nine elemental constituents of EPS with carbon as the primary component with 74.72% and nitrogen with 13.43%. Other elements (phosphorus, sodium, sulphur, calcium, iron, potassium, and magnesium) were also present in trace amounts. The presence of cations in the EPS indicated their binding potential to the negatively charged groups of the EPS. The chemical binding helps in greater accessibility of essential elements required for their growth and development. In the marine environment, the interaction of nutrients with EPS increases elemental uptake rate and concentrate the dissolved organic compounds to make them available for microbial growth (Singh et al. 2011).

Chemical composition analysis by the phenol–sulphuric acid method and Bradford assay revealed carbohydrates to be the principal constituent of EPS. High carbohydrate content in the EPS makes EPS viscous. Sutherland (2016) confirmed the extracellular polysaccharide as the major component of the matrix. Furthermore, ATR-FTIR spectral analysis revealed the functional nature of EPS, together with chemical composition, type of glycosides linkage, and branching of polysaccharides that influence the complete structure of polysaccharides. The broad stretching peak at 3461 cm⁻¹ indicated the presence of hydroxyl groups (O–H) and bound water molecules that make EPS water-soluble. Kavita et al. (2014) also reported the broad stretch peak of the hydroxyl group in the region 3200–3600 cm⁻¹ in a marine bacterium Oceanobacillus iheyensis. The absorption peak at 2924 cm⁻¹ was attributed to the stretching vibration of aliphatic alkane (C–H) in carbohydrates (Boukhelata et al. 2019). The absorption peak at 2360 cm⁻¹ might be due to CO₂ adsorption or amine group (Kavita et al. 2014). The stretching peak of amide I at 1643 cm⁻¹ and amide II at 1544 cm⁻¹ is the characteristic of protein (Carrión et al. 2015). The absorption peak at 1642 cm⁻¹ was attributed to the carbonyl stretching of acetamido groups in the N-acetylated sugars in Bacillus megaterium RB-05 (Chowdhury et al. 2011). A similar C-O stretching peak of amide I and N–H stretching of amide II was also observed in EPS from Pseudomonas sp. ID1(Carrión et al. 2015). According to Christensen (1989), most of the amino sugars found in the bacterial EPS are usually N-acetylated. The absorption peak observed at 1405 cm⁻¹ depicted carboxylate groups that serve as the binding site for divalent cations. Additionally, both carboxyl and hydroxyl groups are responsible for viscous polysaccharide formation, essential for emulsification activity (Singh et al. 2011). The stretching peak at 1232 cm⁻¹ indicated amino sugars in the polysaccharide similar to B. megaterium RB-05 (Chowdhury et al. 2011). The characteristic peak at 1079 cm⁻¹ represented the O-acetyl ester linkage bond of uronic acid specifying the presence of alginate-like polysaccharides also observed in Bacillus, and Klebsiella (Kavita et al. 2014). The presence of acetyl group makes EPS hydrophobic to a certain extent, which adds to their emulsifying ability (Mata et al. 2006). The peak in the region 800–926 cm⁻¹ is the characteristic of polysaccharides resembling the polysaccharide region found in the EPS of Pseudomonas and Acidobacteria. The peak indicated β-glycosidic linkage between sugar monomers. This spectral domain is considered the EPS fingerprint region (Kielak et al. 2017). The absorption peak at 682 cm⁻¹ was assigned to the alkyl-halide that signifies the bacterial origin from the marine environment. Kavita et al. (2014) and Sardari et al. (2017) reported alkyl-halide in the EPS of Oceanobacillus iheyensis and Rhodothermus marinus. A variety of sugars, functional groups, and peptidic moieties suggested the functional role of EPS in the varied biotechnological application. The hydrophobic and hydrophilic nature of EPS makes them suitable stabilizers for the kinetic stability of the emulsion.

Phase identification of EPS was analyzed using XRD. The sharp thin peak observed in the XRD profile represents the crystalline portion of the EPS, while the broad peaks correspond to the amorphous part of the EPS (Mishra et al. 2011). The crystallinity of EPS is determined by the ratio between the sharp narrow diffraction peaks and broad peaks. From the XRD pattern, it appears that crystalline peaks were superimposed in the amorphous phase of the EPS with CI_XRD = 0.336. EPS of Leuconostoc lactis also exhibited amorphous nature with CI_XRD = 0.334 (Saravanan and Shetty 2016). Thus, a polymer can be considered amorphous with little crystallinity. The XRD profile and d-spacing are useful for studying the physical characteristics of a polymer that dictate about the phase of the EPS. Moreover, the crystalline domains act as a reinforcing grid that allows for TGA and DSC analysis over a wide range of temperatures (Angelaalincy et al. 2017). The structural characterization determines the physio-chemical properties of EPS, which impacts its physiological function.

Besides chemical properties, the applicability of EPS is dependent mainly on its mechanical properties. The thermogravimetric analysis showed thermal stability of EPS with three well-differentiated steps of EPS degradation. Fagerson (1969) described that during initial temperature, increased gelatinization, and swelling is the primary event that appears first in the thermogram. A further rise in temperature leads to three thermal events: dehydration, pyrolysis, and reorganization of linkages. During these events, the breakdown of linkages in polysaccharides leads to the formation of volatile and non-volatile products and the formation of new bonds. As revealed by the TGA curve, the first step of degradation is the dehydration reaction, mainly due to the loss of bound water molecules with carboxylic acid groups of the EPS (Sajna et al. 2013; Solmaz et al. 2018). The second step, i.e., degradation, is owed to the structural breakdown and pyrolysis of saccharide moieties, leading to depolymerization (Kavita et al. 2014). In the third step, C–O, C–C bonds in the ring units break, followed by C–O, CO_2, and H₂O evaporation, causing EPS decomposition, which leads to the formation of polynuclear aromatic and aliphatic carbon structures (Parikh and Madamwar 2006; Can et al. 2019). Zamora et al. (2002) also described the same phenomenon in the EPS from Pediococcus damnosus (strain 2.6). Madorsky (1964) and Conley (1970) reviewed such a thermal decomposition process for cellulose. The present finding of the events of distinct phases of polymer degradation stands true with the observation of Parikh and Madamwar (2006), who reported similar pyrolysis patterns of EPS produced by Cyanothece sp. and xanthan with the thermal stability of 550 °C and 500 °C, respectively. The TGA curve revealed that about 12.07% of the EPS content remained after all decomposition (< 600 °C).

The high thermal stability and residue content after 600 °C might be due to the complex and heterogeneous molecular structure of EPS containing uronic acid and calcite-like crystals that prevent complete degradation (Can et al. 2019). In xanthan, the residue content of 27% after 500 °C might be due to its complex molecular structure and cations like Na⁺, K⁺, Ca²⁺, which bonded differently charged sugar moieties (Shah et al. 2000). The stability of EPS produced by P. furukawaii PPS-19 was up to 600 °C that can be termed as thermostable. The thermostable nature will probably be an asset for industrial and biotechnological applications. DSC thermogram presented both exothermic and endothermic thermal events during a thermal scan of 29 °C to 800 °C. DSC analysis suggested a significant thermal transition of EPS into a crystalline state. As the temperature increases during the DSC scan, the amorphous part of EPS becomes less viscous. When it reached a particular temperature (called crystallization temperature), molecules obtain sufficient degree of freedom to spontaneously organize themselves into a crystalline state (Singh et al. 2011). Two endothermic peaks exhibited by the DSC thermogram are polymer transition from crystalline to amorphous state (melting temperature). The possible reason for two endothermic peaks might be due to some conformational changes of the sugar molecules. The first melting peak could be related to the molded component, while the second melting peak might denote the material property of the EPS (Parikh and Madamwar 2006).

EPS produced by P. furukawaii PPS-19 indicated emulsification ability by forming stable emulsions with aliphatic (n-hexane) and aromatic (toluene) hydrocarbons, as well as with hydrocarbon mixtures (petrol and diesel). An emulsion is a stabilized colloidal system where two immiscible liquids (dispersed and continuous phases) are held in suspension. Generally, the emulsion is stabilized using an emulsifier (EPS). Emulsification of dispersed phase within a continuous phase increases the total contact surface area between the two phases. Emulsifying activity is an important functional property of bacterial EPS. The emulsion stabilization mechanism relies on the accumulation of EPS at the oil–water interface in the form of a compact layer. EPS gets effectively and irreversibly adsorbed at the interface, making emulsion extremely stable to coalescence and flocculation (Wongkongkatep et al. 2012). It is evident from the ATR-FTIR analyses that the EPS produced by P. furukawaii PPS-19 possess protein and polysaccharide moieties that play an essential role in the emulsification activity. The proteins bind to the petroleum hydrocarbon through electrostatic or hydrophobic-hydrophobic interaction while reducing the end of the polysaccharides attach to the protein through covalent bonds. The polysaccharide-protein complex produces stable oil–water emulsion (Uzoigwe et al. 2015). Bramhachari et al. (2007) and Llamas et al. (2010) demonstrated that the emulsifying activity of the EPS is due to the existence of hydrophobic groups such as peptidic moieties, ester-linked acetyl groups, and deoxysugars. Proteins being the hydrophobic part, get adsorbed at the interface, consequently decreasing interfacial tension and increasing interfacial elasticity. Polysaccharides are hydrophilic and tend to remain dispersed within the aqueous phase. The inherent thermodynamic and kinetic stability is achieved when the interfacial tension between the dispersed and continuous phases decreases and gets dropped to the point where the entropy of mixing exceeds the interfacial free energy (Gernon et al. 2009). The emulsification activity of EPS is determined by its strength in holding the emulsion for a certain time period. The present study revealed that the higher the molecular weight of the substrate, the more effective EPS was in stabilizing the emulsion. Mixtures containing aromatic and aliphatic hydrocarbons (oils) were excellent substrates for emulsification. However, pure aliphatic and aromatic hydrocarbons were slightly emulsified. Navon-Venezia et al. (1995) and Gudiña et al. (2015) observed that emulsifier from Acinetobacter radioresistens and Paenibacillus strain could poorly emulsify pentane, hexane, cyclohexane, heptane, and iso-octane whereas C₁₀–C₁₈ compounds and hydrocarbon mixtures were efficiently emulsified. Emulsification of petrol and diesel can be applied for the removal of oil from the marine environment. Petrol and diesel contain various molecules such as paraffin, olefins, naphtha, saturated and aromatic compounds, thus could serve as model substrates for studying hydrocarbon biodegradation. Emulsification of toluene and hexane is also essential for industrial wastewater treatment. The emulsification ability of the EPS to emulsify lipids is the desired property with great potential for bioremediation of oil polluted marine sites (Camacho-Chab et al. 2013). The stability of the EPS solution at 85 °C temperature indicated that the marine bacterial EPS are thermostable as bioemulsifier. Similar thermostability of bioemulsifier was also found in Acidobacteria at 70 °C and 90 °C (Kielak et al. 2017). The stability of EPS concerning temperature and time exposure established its potential for applications involving extreme marine environmental surroundings (Toren et al. 2001).

The biodegradation of crude oil in the microcosm was used to evaluate the potential of EPS in the degradation of crude oil. To determine the oil loss caused by physico-chemical factors like air, dissolved oxygen, pH, and temperature, one control set containing only 2% (v/v) crude oil was conducted. Negligible degradation caused by physico-chemical factors was observed. In contrast, in the presence of P. furukawaii PPS-19, the degradation rate increased significantly by 67.83%, indicating the role of microbial mechanism in the degradation of crude oil. However, in the presence of P. furukawaii PPS-19 and EPS complex, the degradation rate was the highest (89.52%), suggesting the role of EPS in increasing the bacterial cell surface hydrophobicity, resulting in increased bacterial adherence to the surface of the oil droplets. EPS disperses the oil into small droplets leading to an increased interfacial area of the oil available to the bacteria (Das et al. 2008). EPS alone had no role in the degradation of crude oil, suggesting that EPS only acts as an associative agent for changing the cell surface characteristics of P. furukawaii PPS-19. Gong et al. (2015) also reported similar results with the bacteria-chitosan complex. Bacillus cereus degraded 65% n-tetradecane while the degradation rate increased to 92% with the bacteria-chitosan complex.

In this study, P. furukawaii PPS-19 showed strong biofilm-forming potential in the presence of 2% (v/v) crude oil at the interface that exhibited the potential of EPS in providing resistance to oil dispersion. Biofilm thickness shown in the confocal micrograph confirmed the robust biofilm formation by P. furukawaii PPS-19. The specialized function of biofilm components provides a three-dimensional structure. Biofilm formation at the oil–water interface, as revealed by SEM, indicated the potential of EPS in providing stability to biofilm. The 3D-architecture of extracted EPS exposing tiny pores makes them water-soluble. The presence of various functional groups by FTIR spectrum showed the structural complexity of the EPS indicating its functional role as bioemulsifier. XRD pattern showed that EPS was amorphous. TGA and DSC data suggested the thermostability of EPS that is important for industrial application. The characterization study demonstrated the potential of EPS for polymer reinforcement. The main property of the EPS is the ability to form a thermostable emulsion, enabling it an efficient emulsifier than commercial polymer xanthan. These biopolymer properties could lead to increased kinetic stability of the emulsion under undesirable conditions. Besides, the addition of the EPS increased the degradability of petroleum hydrocarbon by the marine bacterium P. furukawaii PPS-19. The findings of this study indicate that the EPS could be exploited for application in environmental biotechnology and bioremediation of oil polluted sites.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

SD designed the experimental plan, analyzed and interpreted the data and finalized the manuscript. Vandana executed the experimental work, generated the data and drafted the manuscript.

Declarations

Conflict of interest

The authors declare no conflict of interest.