Effect of biofilm parameters and extracellular polymeric substance composition on polycyclic aromatic hydrocarbon degradation

Abstract

Marine bacterial biofilms were studied under different physicochemical conditions for enhanced bioremediation of polycyclic aromatic hydrocarbons (PAHs). Molecular characterization of ten environmental isolates was done by 16S rRNA gene sequencing. The effect of different physicochemical parameters, such as pH, salt concentration, temperature, carbon source on their biofilm production capability was monitored. Various topological parameters of the biofilms such as total biomass (EPS and cells content), thickness, roughness coefficient, diffusion distance and surface to biovolume ratio were studied using a confocal scanning laser microscope (CSLM). Among the various strains studied, the total biomass was maximum for P. aeruginosa N6P6 (106.64 μm³ μm⁻²) followed by S. acidaminiphila NCW702 (26.92 μm³ μm⁻²) indicating the formation of dense biofilm. Significant negative correlation (P < 0.05) was observed between the roughness coefficient of the biofilm and PAH degradation, whereas a significant positive correlation (P < 0.05) was observed between PAH (phenanthrene and pyrene) degradation and total biomass, thickness and diffusion distance of the biofilms. PAH degradation was studied both in planktonic and biofilm modes of growth. Biofilm facilitated degradation of the two PAHs was higher than the planktonic cells. This work demonstrates that the attached phenotypes of the marine bacteria showed noticeable variation in biofilm architecture and, in turn, biodegradation of PAHs.

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Introduction

Interface ecosystems are becoming increasingly saturated with pollutants due to anthropogenic activities. Particularly, coastal sediments and near shore waters act as sinks for many types of pollutants such as hydrocarbons.¹ Among organic pollutants, polycyclic aromatic hydrocarbons (PAHs) are lethal contaminants, ubiquitously present in coastal areas.² PAHs and related compounds adversely affect marine biota due to their acute lipophilic and persistent nature. PAHs get accumulated in the lipids compounds present in the various tissues. Thus, even moderate quantity of PAH compounds can have harmful effects on aquatic organisms.^3,4 These organic compounds can undergo complex biotransformation through many metabolic routes generating a number of intermediate metabolites. In many cases, acute toxicity of some intermediates than the parent compounds have been reported in marine organisms residing in the vicinity of the contaminated sites.^5,6

Bacteria are among the first organisms to colonize surfaces when exposed to aquatic environments. Biofilms can form in almost any hydrated environment that has the proper nutrient conditions, and can develop on a wide variety of abiotic, hydrophobic and hydrophilic surfaces. The polymers secreted by the microorganisms provide structural stability to the biofilms and maintain their metabolic activity and stability.⁷ This stable and dynamic nature of biofilm makes it resistant to various environmental stressors and xenobiotics. The exopolymeric constituent of biofilm i.e., EPS can trap various organic contaminants present in the immediate milieu. EPS matrix traps the nutrients for metabolic utilization by the resident bacteria and water is efficiently retained through H-bond interactions with hydrophilic polysaccharides.^8,9 The unique mass transport property along with several other parameters makes biofilm the most suitable biological agent for the degradation of various recalcitrant organic compounds such as PAHs.^10–13 Microbial biofilm communities have many advantages over planktonic cells when used as a biocatalysts in biodegradation processes. Naturally immobilized bacterial cells in biofilms exclude the necessity of cell-immobilization as biofilm cells are embedded in the extracellular polymeric matrix.¹⁴

Biofilms support high biomass density thus maintaining optimum condition in the niche, this facilitates the mineralization process. Accordingly biofilm-mediated bioremediation process offers an efficient and cheap method over the planktonic cells which requires huge infrastructure. In order to optimize biofilm based bioremediation processes, a systematic understanding of biofilm structure, dynamics and interaction with pollutants is required. The respective properties of cells in a biofilm differ from its planktonic counterpart in various aspects such as high cell density, altered gene expression, metabolic resilience, cellular heterogeneities.¹⁵ Though, most of the bacteria have their distinct biofilm architectures and growth characteristics, still biofilm properties are modulated by various physicochemical factors such as pH, temperature, ionic strength and nutrients.¹⁶ Therefore, a detailed biofilm characterisation is warranted under different physicochemical conditions before its application for a remediation purpose. The present work investigates the effect of different physicochemical conditions such as pH, temperature, carbon source and salt concentration on biofilm growth of PAHs degrading marine bacteria isolates. The biofilm architecture of the isolates was studied using confocal scanning laser microscopy (CSLM) and scanning electron microscopy (SEM). Finally, PAHs degradation efficiency of marine bacterial isolates was investigated both in biofilm and planktonic mode of growth. To the best of our knowledge, this is the first study where a direct correlation was investigated between biofilm parameters, EPS characteristics etc., and the PAHs degradation using a number of isolates.

Experimental

Bacterial culture

Biofilm forming and PAHs degrading bacteria were isolated from Odisha state of eastern coastal region of India, by selective enrichment with phenanthrene and pyrene. The isolates were identified by 16S rRNA gene sequencing as described earlier.¹³ Ten isolates used in the present study are Paenibacillus lautus NE3B01, Pseudomonas alcaliphila NE3B02, Alcaligenes faecalis NCW-402, Stenotrophomonas acidaminiphila NCW-702, Lysinibacillus fusiformis NCW903, Sporosarcina luteola NR402, Pseudomonas mendocina NR802, Pseudomonas pseudoalcaligenes NP103, Pseudomonas sp. NP202 and Pseudomonas aeruginosa N6P6. The 16S rRNA gene sequences have been submitted to NCBI GenBank and cultures have been deposited to microbial culture collection (ESI Table S1†).

Effect of physicochemical conditions on biofilm growth

A single colony of each isolate was inoculated in Luria Bertani¹⁷ broth (Himedia, India) and incubated at 37 °C overnight. The overnight grown cultures were checked for culture purity and diluted to 1 : 100 in LB broth. One ml of the diluted culture was transferred to a glass tube and incubated under static condition at 37 °C for 48 h in a sterile moist chamber. After 48 h, the tubes were washed twice with phosphate buffer saline (PBS), air dried and stained with 1 ml of 0.2% crystal violet for 5 min. Thereafter, the tubes were washed twice with water, air dried and destained with 1 ml of 95% ethanol for 30 min. After 30 min, 200 μl of the destained solution was transferred to a 96-well microtiter plate in triplicate. The biofilm growth was quantified in terms of absorbance at 595 nm, measured using Plate Reader (Victor X3, PerkinElmer, USA). Biofilm growth was quantified in the presence of fructose, glucose, xylose, sucrose, arabinose, rhamnose and raffinose in Basal salt medium (BSM) (Himedia, India). 1% concentration of each sugar was individually provided as sole carbon source.

Ten selected marine bacterial isolates were studied for biofilm formation under various physiological conditions like ionic strength, temperature and pH. Ionic strength test was carried out with NaCl in LB broth at a concentration of 0–6%. LB broth with pH range of 6 to 9 was used to study the effect of pH. Effect of temperature was studied at 28 °C and 37 °C.

Characterization of biofilm architecture by scanning electron microscopy (SEM) and confocal scanning laser microscopy (CSLM)

Biofilm architecture was studied employing SEM and CSLM. The samples were prepared following Mangwani et al.¹³ Briefly, the overnight grown culture of the isolate was diluted to 1 : 100 in LB broth. Three ml aliquot of this suspension was transferred to 6-well plate with glass slide (1 × 1 cm for SEM and 2 × 3 cm for CSLM study). The plates were incubated at 37 °C under static condition for 48 h, thereafter the glass slide for SEM study was fixed with 2.5% glutaraldehyde (for 12 h at 4 °C) and 1% aqueous solution of tannic acid. The glass slide was dehydrated using 30%, 70% and 100% alcohol and air dried.¹⁸ Biofilm slide was then coated with platinum and observed under SEM (Jeol T-330 Scanning Electron Microscope, Germany).

The biofilm slides prepared for CSLM studies was stained with SYTO9 (Invitrogen, USA) and concanavalin A-tetramethylrhodamine isothiocyanate [ConA-TRITC (Invitrogen, USA)] and biofilm architecture was studied following.¹⁹ A cover slip was placed over the stained biofilm and mounted upside down over the objective lens of CSLM (TCS-SP2-AOBS) equipped with DM IRE 2 inverted microscope (Leica Microsystems, Hessen, Wetzlar, Germany). Water immersed 63× objective lens with 1.2 NA was used to image the biofilm. About 10 image stacks were collected randomly from different points in order to acquire statistically significant data. Biofilm parameters (average thickness, maximum thickness, total biomass, diffusion distance surface to biovolume ratio, and roughness coefficient) were quantified by the analysis of image-stacks using COMSTAT.^20,21

Extraction and characterization of biofilm associated EPS

Biofilm associated EPS was extracted by chilled ethanol following Mangwani et al.¹⁹ An IR spectrum of the dried EPS KBr pellet was measured using Fourier-transform infrared (FTIR) spectrophotometer (PerkinElmer, USA). Total carbohydrate and protein content of the extracted EPS was estimated by Bradford method²² and phenol sulphuric acid method²³ respectively.

Emulsification assay was carried out with biofilm grown on glass beads of diameter 2 mm (in 3 ml LB broth) in a glass tube. After 48 h, free planktonic cells were carefully aspirated from the tubes and the biofilm grown over the glass beads was rinsed with sterile PBS. The glass beads were gently vortexed to detach biofilm and 3 ml of LB broth was added and mixed gently. The EPS and cells were separated by centrifugation at 6000 rpm for 15 min and 2 ml of the aqueous upper layer was transferred to a test tube. 1 ml of n-octane was added to the tube followed by vortexing for 2 min. After 24 h, total height of liquid layer and emulsion layer was measured. 1% SDS solution in LB broth was used as positive control and LB broth was used as negative control. Emulsifying activity was expressed as the percentage of the total height occupied by the emulsion in the test tube. The following mathematical equation was used to calculate emulsification index.²⁴

Emulsification index (E₂₄%) = (height of the emulsion layer/total height of liquid phase) × 100

PAHs bioremediation study

Phenanthrene and pyrene degradation was studied using biofilm and planktonic culture of the isolates following Mangwani et al.¹⁹ (details included in ESI file†).

Statistical analysis

Data are expressed as mean ± standard deviation (SD) of the triplicate experimental data. A two tailed student's t-test was used to determine the differences in biofilm formation under different physiological conditions. One-way ANOVA was also calculated between the groups to evaluate the effect of physiological conditions on biofilm growth. P ≤ 0.05 was considered statistically significant. Biofilm parameters obtained from CSLM were expressed as mean ± standard error (SE). Correlation coefficient was calculated to evaluate the relationship between different biofilm topological parameters.

Results

Effect of temperature on biofilm growth of marine isolates

Biofilm growth was quantified after 48 h at 28 °C and 37 °C (Fig. S1†). Maximum biofilm growth was observed at 37 °C by most of the isolates. Biofilm growth of Paenibacillus lautus NE3B01, Lysinibacillus fusiformis NCW903 and Pseudomonas alcaliphila NE3B02 was not affected by changes in temperature. At both the temperatures, maximum biofilm growth was observed in P. aeruginosa N6P6 followed by P. pseudoalcaligenes NP103 and P. mendocina NR802.

Effect of sugars on biofilm growth

Effect of different sugars (fructose, xylose, raffinose, arabinose, sucrose, rhamnose and glucose) on biofilm growth was studied by glass tube assay. All the isolates were able to form biofilm in the presence of glucose, as carbon source. However, they showed diverse response to the presence of other sugars. Among all the marine isolates, P. aeruginosa N6P6 and P. mendocina NR802 were able to form biofilm in the presence of all the tested sugars. Result signifies that biofilm growth of marine isolates was significantly affected by the presence of different type of sugars in comparison to LB (P < 0.05; one way ANOVA followed by Tukey's HSD test). Among the all tested sugars, maximum biofilm growth was observed in presence of arabinose and sucrose in P. mendocina NR802 and P. aeruginosa N6P6 respectively, but the increase was non-significant when compared with other sugars (P > 0.05; Tukey's HSD test) (Fig. 1). Except P. lautus NE3B01 and S. acidaminiphila NCW702, biofilm growth of other isolates significantly increased in LB medium as compared to the tested sugars (P < 0.05; Tukey's HSD test). The most significant effect of tested sugars was observed on S. acidaminiphila NCW702 biofilm (P < 0.05; Tukey HSD test). The biofilm growth of S. acidaminiphila NCW702 increased significantly in the presence of rhamnose as compared to the other tested sugars (P < 0.05; Tukey HSD test). The biofilm growth in presence of glucose increased significantly in P. lautus NE3B01 and S. acidaminiphila NCW702 (P < 0.05; Tukey HSD test). Significant decrease in biofilm growth of P. lautus NE3B01 and S. acidaminiphila NCW702 was observed when fructose was used as carbon source as compared to other sugars (P < 0.05; Tukey HSD test) and it was found to be least preferred carbon source.

Fig. 1 Effect of different sugars on biofilm growth over glass surface quantified in terms of A₅₉₅ after crystal violet staining. Data are expressed as mean ± SD.

Effect of pH on biofilm growth of all isolates

The PAHs utilizing bacteria isolated from different marine samples showed diverse response to pH shifts (Fig. 2). Biofilm growth of P. lautus NE3B01, L. fusiformis NCW903, S. luteola NR402, P. alcaliphila NE3B02 and P. mendocina NR802 remained unaffected by changes in pH of the medium (P > 0.05; Tukey HSD test). Maximum biofilm growth was observed at pH 7 for A. faecalis NCW402 and S. acidaminiphila NCW702 (Fig. 2). A. faecalis NCW402 biofilm growth was found to be sensitive towards alkaline pH and a significant decline in biofilm growth was observed at pH 8 and pH 9 (P < 0.05; Tukey HSD test). Maximum biofilm growth was observed at pH 6 for P. pseudoalcaligenes NP103 and Pseudomonas sp. NP202. In P. aeruginosa N6P6, pH 7 was found to be optimum for biofilm growth. Biofilm growth of Pseudomonas sp. NP202 was significantly affected by change in pH of growth medium (P < 0.05; Tukey HSD test).

Fig. 2 Effect of pH on biofilm growth of marine bacterial isolates. Data are expressed as mean ± SD of triplicate samples.

Effect of NaCl on biofilm growth and cell attachment to the surface

Different concentrations of NaCl (0–6%) was used to estimate the ionic strength of the biofilm. Fig. S2† shows that Na⁺ concentration was found to affect significantly the development of biofilm and surface attachment potential of marine isolates (P < 0.05; one way ANOVA, Tukey HSD test). S. acidaminiphila NCW702 and P. aeruginosa N6P6 were able to form luxuriant biofilm at 2% NaCl and cell surface attachment was observed even at 4% NaCl. Absence of Na⁺ in medium showed negative effect on cell attachment to the surface and biofilm growth was also limited. At high salt concentration (>4%), inhibitory effect on both biofilm growth and cell attachment was observed.

Scanning electron microscopic studies of bacterial biofilms on glass surface

The cell morphology, aggregation and arrangement within biofilm were studied by scanning electron microscopy (Fig. 3). Biofilm of P. lautus NE3B01 was composed of numerous pleomorphic cells attached to the surface, lacking dense EPS or compressed aggregation (Fig. 3a). Biofilm of P. alcaliphila NE3B02 was composed of rod shaped cells attached together (Fig. 3b). However, total coverage and cell density was less as compared to P. lautus NE3B01. Biofilms of A. faecalis NCW402 and S. acidaminiphila NCW702 (Fig. 3c and d) were composed of rod shaped cells embedded in a dense matrix of EPS. Scanning electron micrographs of L. fusiformis NCW903 and S. luteola NR402 showed long rods attached to the surface surrounded by EPS, but cell aggregation was lacking between cells (Fig. 3e and f). P. mendocina NR802, P. pseudoalcaligenes NP103, Pseudomonas sp. NP202 and P. aeruginosa N6P6 scanning electron micrographs revealed the presence of cell aggregates and dense biomass encrusted in EPS matrix (Fig. 3g–j).

Fig. 3 Scanning electron micrograph of bacterial biofilm (a) Paenibacillus lautus NE3B01 (b) P. alcaliphila NE3B02 (c) Alcaligenes faecalis NCW402 (d) Stenotrophomonas acidaminiphila NCW702 (e) Lysinibacillus fusiformis NCW903 (f) Sporosarcina luteola NR402 (g) P. mendocina NR802 (h) P. pseudoalcaligenes NP103 (i) Pseudomonas sp. NP202 (j) P. aeruginosa N6P6 (Paenibacillus lautus NE3B01 and Stenotrophomonas acidaminiphila NCW702 images are reproduced with permission).

Characterization of biofilm structure by confocal scanning laser microscopy

Biofilm formation by marine bacteria was characterized by CSLM and images were analysed using COMSTAT. Total biomass (EPS and cells), thickness, roughness coefficient, diffusion distance and surface to biovolume ratio were determined to characterize biofilm structure of marine bacteria (Fig. 4). The estimated biofilm parameters are illustrated in Table 1. Total biomass (or biovolume) was measured as biomass of two key components cells and EPS, which were measured by staining biofilm with specific fluorescent dyes, SYTO9 and ConA-TRITC.

Fig. 4 CSLM micrograph of bacterial biofilm (after 48 h of growth) stained with SYTO9 and ConA-TRITC. SYTO9 specifically stains cells whereas ConA-TRITC binds bacterial exopolysaccharides. Green and red colours indicate the presence of cells and exopolysaccharides respectively. Each CSLM image shows four sections (i) cells, (ii) EPS, (iii) superimposed image of (i) and (ii), (iv) superimposed image of (i) and (ii) with vertical cross sections showing biofilm thickness. (a) Paenibacillus lautus NE3B01 (b) P. alcaliphila NE3B02 (c) Alcaligenes faecalis NCW402 (d) Stenotrophomonas acidaminiphila NCW702 (e) Lysinibacillus fusiformis NCW903 (f) Sporosarcina luteola NR402 (g) P. mendocina NR802 (h) P. pseudoalcaligenes NP103 (i) Pseudomonas sp. NP202 (j) P. aeruginosa N6P6 (P. lautus NE3B01 and S. acidaminiphila NCW702 images are reproduced with permission).

Table 1 Topological parameters of marine bacterial biofilm (after 48 h of growth) studied by CSLM. Data are expressed as mean ± SE

Biofilm parameters	Paenibacillus lautus NE3B01	P. alcaliphila NE3B02	Alcaligenes faecalis NCW402	Stenotrophomonas acidaminiphila NCW702	Lysinibacillus fusiformis NCW903	Sporosarcina luteola NR402	P. mendocina NR802	P. pseudoalcaligenes NP103	Pseudomonas sp. NP202	P. aeruginosa N6P6
Total biomass (EPS + cell) (μm^3 μm^−2)	8.86 ± 1.84	9.75 ± 3.9	22.98 ± 5.42	26.92 ± 5.6	8.91 ± 2.23	23.24 ± 6.09	16.79 ± 3.26	15.91 ± 3.84	19.16 ± 3.42	106.64 ± 11.42
Total biomass (EPS) (μm^3 μm^−2)	3.94 ± 0.78	4.09 ± 1.73	10.49 ± 2.28	8.21 ± 2.4	4.15 ± 1.3	8.59 ± 2.9	6.03 ± 1.27	3.89 ± 1.67	5.68 ± 1.68	54.03 ± 5.01
Total biomass (cell) (μm^3 μm^−2)	4.91 ± 1.05	5.66 ± 2.1	12.49 ± 3.14	18.7 ± 3.2	4.75 ± 0.9	14.64 ± 3.18	10.75 ± 1.99	12.02 ± 2.16	13.48 ± 1.74	52.611 ± 6.4
Average thickness (μm)	9.24 ± 1.98	8.36 ± 2.28	19.49 ± 4.7	23.93 ± 2.61	11.79 ± 4.3	25.14 ± 1.91	19.73 ± 2.45	28.73 ± 3.22	26.71 ± 2.48	66.76 ± 4.11
Maximum thickness (μm)	23.57 ± 0.75	14.93 ± 2.0	30.07 ± 3.24	30.94 ± 2.24	28.51 ± 2.54	32.01 ± 0.86	22.69 ± 2.19	48.85 ± 2.37	31.42 ± 3.21	80.34 ± 3.82
Roughness coefficient	1.04 ± 0.14	0.53 ± 0.23	0.34 ± 0.27	0.24 ± 0.07	1.0 ± 0.22	0.20 ± 0.05	0.16 ± 0.06	0.47 ± 0.12	0.08 ± 0.02	0.10 ± 0.36
Maximum diffusion distance (μm)	12.94 ± 0.98	6.53 ± 1.46	6.89 ± 0.46	13.77 ± 1.46	9.50 ± 0.78	10.37 ± 1.39	8.60 ± 1.02	15.89 ± 4.09	7.20 ± 0.12	35.23 ± 6.79
Average diffusion distance (μm)	0.35 ± 0.091	0.42 ± 0.22	0.65 ± 0.17	2.68 ± 0.7	0.19 ± 0.04	1.05 ± 0.35	0.69 ± 0.16	0.79 ± 0.33	0.63 ± 0.17	4.46 ± 1.33
Surface to biovolume ratio (μm^2 μm^−3)	0.26 ± 0.04	0.29 ± 0.08	0.25 ± 0.13	1.97 ± 0.25	0.33 ± 0.07	0.13 ± 0.03	0.17 ± 0.02	0.23 ± 0.08	0.16 ± 0.03	0.053 ± 0.008

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Among the ten environmental isolates studied, the total biomass was maximum for P. aeruginosa N6P6 (106.64 ± 11.42 μm³ μm⁻²) including both cells and EPS components followed by S. acidaminiphila NCW702. For other isolates the total biomass was in the range of 8–25 μm³ μm⁻². P. lautus NE3B01 and L. fusiformis NCW903 had low biomass values. Cells/EPS ratio was near to 1 for P. aeruginosa N6P6 (0.97), A. faecalis NCW402 (1.19) and L. fusiformis NCW903 (1.14) indicating equal proportion of cells and EPS. For P. lautus NE3B01 and P. alcaliphila NE3B02, cells/EPS ratio was <0.8, indicating less proportion of cells over EPS (Table 1; Fig. 6a–j). Average thickness was highest for P. aeruginosa N6P6 (66.76 ± 4.11 μm) and P. pseudoalcaligenes NP103 (28.73 ± 3.22 μm). The average thickness was least for P. lautus NE3B01 and P. alcaliphila NE3B02 (i.e. <10 μm).

Roughness coefficient (R*) showed considerable difference in biofilm structure of isolated marine bacteria. R* was >1 for Lysinibacillus fusiformis NCW903 and Paenibacillus lautus NE3B01, which reflects biofilm heterogeneity and growth as microcolonies. R* for Alcaligenes faecalis NCW402, Stenotrophomonas acidaminiphila NCW702, P. pseudoalcaligenes NP103, P. alcaliphila NE3B02, P. mendocina NR802, and Sporosarcina luteola NR402 was between 0.15 and 0.6. Whereas, in Pseudomonas sp. NP202 and P. aeruginosa N6P6 R* was <0.1.

Average diffusion distance is a measure of the distance over which nutrients and other substrate components diffuse inside biofilm and to bacteria within microcolonies. P. aeruginosa N6P6 and S. acidaminiphila NCW702 had average diffusion distance of 4.46 ± 1.33 μm and 2.68 ± 0.7 μm respectively, which was maximum among studied isolates. Surface to biovolume ratio indicates the adaptation of biofilm to the environment. The surface to volume ratio increases in order to optimize access to the limited supply of nutrients. Surface to volume ratio also indicates portion of the biofilm, exposed to the nutrient flow. S. acidaminiphila NCW702 had maximum surface to biovolume ratio of 1.97 ± 0.25 μm² μm⁻³, whereas P. aeruginosa N6P6 had least surface to biovolume ratio 0.053 ± 0.008 μm² μm⁻³.

To evaluate the relation between different biofilm topological parameters calculated using COMSTAT, correlation coefficient (r) was calculated (Table 2). From the correlation matrix, it was interpreted that there was a significant positive correlation between biomass, thickness and diffusion distance of biofilm (r > 0.9, P < 0.001).

Table 2 Correlation matrix of biofilm topological parameters and PAHs (phenanthrene and pyrene) degradation (n = 10; * P < 0.05; ** P < 0.01; *** P < 0.001)

Biofilm parameters	Total biomass (EPS + cell)	Total biomass (EPS)	Total biomass (cell)	Average thickness	Maximum thickness	Roughness coefficient	Maximum diffusion distance	Average diffusion distance	Surface to biovolume ratio	Phenanthrene degradation	Pyrene degradation
Total biomass (EPS + cell)	1
Total biomass (EPS)	0.9928***	1
Total biomass (cell)	0.9913***	0.9684***	1
Average thickness	0.9494***	0.9157***	0.9707***	1
Maximum thickness	0.8967***	0.8757**	0.9049***	0.9526***	1
Roughness coefficient	−0.4710	−0.3940	−0.5478	−0.5896	−0.3571	1
Maximum diffusion distance	0.9133***	0.9061***	0.9059***	0.8875***	0.9297***	−0.2124	1
Average diffusion distance	0.9206***	0.8828***	0.9467***	0.8784***	0.8085**	−0.5010	0.8767***	1
Surface to biovolume ratio	−0.0984	−0.1641	−0.0249	−0.1230	−0.1619	−0.0604	−0.0474	0.2810	1
Phenanthrene degradation	0.5769	0.5254	0.6237*	0.6550*	0.6405*	−0.50905	0.6020	0.6526*	0.21152	1	—
Pyrene degradation	0.4629	0.4138	0.5090	0.5888*	0.6077*	−0.4508	0.4525	0.4794	0.1312	—	1

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FTIR characterization of biofilm associated exopolymers

Composition of the functional groups of biofilm-EPS was analysed by FTIR spectroscopy. The presence of peaks corresponding to functional groups like –COC–, PO4–, –NH2 and –CH2–, in biofilm-EPS shows the presence of polysaccharides, nucleic acid, protein and lipid in EPS. The band in range of 1320–1000 cm⁻¹, assigned to uronic acids, was present in all EPS. A peak between 3200 and 2800 cm⁻¹ indicated the presence of lipids in EPS (Fig. 5). The existence of carbohydrate was indicated by –COC—stretch between 1750 and 1625 cm⁻¹. Characteristic peak of amide was observed at 1650–1600 cm⁻¹, which was assigned to stretching vibration of N–H. The absorption band in range of 1655–1680 cm⁻¹ was assigned to the strong vibration of C=O and C–N groups associated with peptide bonds. A band at 1200–900 cm⁻¹ suggested the stretching vibration of C–O–C ring of carbohydrates. The band at 1200–1240 cm⁻¹ was assigned to the phosphate groups (Fig. 5). The carbohydrate and protein content is shown in ESI Fig. S3.†

Fig. 5 FTIR spectra of biofilm associated EPS (A) P. lautus NE3B01 (black), P. alcaliphila NE3B02 (red), A. faecalis NCW402 (pink), S. acidaminiphila NCW702 (blue), L. fusiformis NCW903 (green) (B) Sporosarcina luteola NR402 (black), P. mendocina NR802 (red), P. pseudoalcaligenes NP103 (pink), Pseudomonas sp. NP202 (blue), P. aeruginosa N6P6 (green). The existence of carbohydrate was suggested by the adsorption bands at 1200–900 cm⁻¹ and 1750–1625 cm⁻¹ corresponding stretching vibration of C–O–C. Characteristic peak at 1650–1600 cm⁻¹, suggest presence of N–H. The absorption band in range of 1655–1680 cm⁻¹ is assigned to C=O and C–N groups associated with peptide bonds.

Emulsifying property of bacterial biofilm associated EPS

The emulsification data are presented in ESI Fig. S4.† The E₂₄% of P. pseudoalcaligenes NP103 and P. aeruginosa N6P6 EPS was >40%. Whereas, E₂₄% for Lysinibacillus fusiformis NCW903, P. mendocina NR802 was >30%. The emulsification index of P. pseudoalcaligenes NP103 was more than that of 1% SDS (used as positive control).

Efficacy of biofilm and planktonic cultures on PAHs degradation

Phenanthrene degradation was studied under both planktonic and biofilm mode of growth. Initial concentration of phenanthrene was 100 mg l⁻¹. More amount of phenanthrene was degraded by biofilm mode i.e. the residual concentration of phenanthrene was less in biofilm mediated degradation (Fig. 6a) after 7 d of incubation. For P. lautus NE3B01, degradation was below 30% by both planktonic cells and biofilm mode after 7 d indicating that P. lautus NE3B01 physiological state of cells does not play any role in degradation. However, for all other isolates, biofilm culture significantly increased the degradation of phenanthrene. With biofilm culture, >50% degradation was observed in S. acidaminiphila NCW702, A. faecalis NCW402, P. mendocina NR802, P. aeruginosa N6P6 and P. pseudoalcaligenes NP103 biofilm.

Fig. 6 Residual concentration of (a) phenanthrene and (b) pyrene extracted from planktonic and biofilm cultures after 7 d of incubation (PAHs degradation data for S. acidaminiphila NCW702 are reproduced with permission from our previous report¹⁹).

The percentage degradation of phenanthrene by planktonic and biofilm cultures are illustrated in Table 2. Maximum degradation of 85.5% and 78.7% was observed for P. aeruginosa N6P6 and P. pseudoalcaligenes NP103 biofilms respectively in 7 d. Degradation of pyrene was significantly higher when biofilm culture was used. For Paenibacillus lautus NE3B01, both planktonic and biofilm culture were able to degrade about 11–12% pyrene in 7 d. Pyrene degradation was 13.93% by planktonic P. alcaliphila NE3B02 culture, whereas 6.74% of pyrene was degraded by its biofilm culture (Fig. 6b).

The efficient pyrene degradation was achieved by P. pseudoalcaligenes NP103 and P. aeruginosa N6P6. With biofilm culture of P. pseudoalcaligenes NP103, 50% degradation of pyrene was observed. Whereas, P. aeruginosa N6P6 biofilm degraded 48.69% of pyrene in 7 d. Correlation between biofilm topological parameters and PAHs degradation was calculated. A significant positive correlation was observed between PAHs (phenanthrene and pyrene degradation) and biofilm total biomass and biofilm thickness (P < 0.05). However, a negative correlation was observed between PAHs degradation and roughness coefficient (Table 2).

Discussion

Bacteria residing within biofilm live in a cooperative manner and benefit each other by forming different interlinked niches.²⁵ However, biofilm growth is influenced by a number of environmental factors. Evaluation of biofilm growth at different pH, temperature, carbohydrate and NaCl conditions indicate diverse mode of biofilm growth under different physiological conditions.²⁶ In nature, bacteria produce EPS and forms biofilm to protect themselves against unfavourable conditions. Apart from cell physiology, biofilm formation is associated with cell–cell signalling (i.e. quorum sensing) in many bacterial species.^27–30 This signalling process is largely depended upon autoinducers (AIs), which can have profound effect on biofilm architecture and functioning.³¹ Thus, a better understanding of bacterial activities in biofilms can help in designing an improvised strategy for bioremediation. In the present study, biofilms of ten PAHs utilizing marine bacteria were characterized and their PAHs degradation potential was also evaluated. Seven isolates belonged to the Proteobacteria group including γ-proteobacteria (Pseudomonas and Stenotrophomonas) and β-proteobacteria (Alcaligenes). Proteobacteria not only exist widely in the environment, but also are important in pollutant degrading and biofilm forming bacteria.^32–35 The impact of the temperature on bacterial biofilm formation is commonly strain and species dependent.¹⁷ Most marine isolates in this study showed better biofilm growth at 37 °C, however, the biofilm growth response towards temperature change significantly differed among the studied isolates. Rao studied the formation of biofilm in marine ecosystem (having different water temperature) wherein a significant increase in biofilm proliferation with increase in seawater temperature from 28.5 to 35 °C was observed.³⁶

Biofilm growth of marine bacteria in the presence of different sugars showed that, in general glucose and sucrose are the most preferred sugars for biofilm growth. For P. mendocina NR802 maximum biofilm growth was observed in the presence of arabinose. Whereas, poor biofilm growth was observed in the presence of fructose. Arabinose has been reported to induce biofilm growth and EPS synthesis in marine bacterium Vibrio fischeri.³⁷ In Escherichia coli fructose inhibits biofilm formation by suppression of curli production and AI-2 quorum sensing system.³⁸ Whereas, in Staphylococcus aureus a decrease in surface hydrophobicity and interference in cell attachment to the surface in presence of fructose was observed.³⁹ The presence of sugars can significantly affect the EPS synthesis in marine bacteria.⁴⁰ Biofilm forming marine bacteria Bacillus licheniformis and Halomonas sp. have been reported to produce higher EPS in presence of glucose as carbon source.^41,42 However, Ko et al. reported increase in EPS synthesis by marine bacterium Hahella chejuensis when sucrose was supplemented as carbon source.⁴³

Many physical and chemical parameters affect biofilm growth and development. Although bacterial biofilms are highly resistant to pH variations under natural conditions, still pH of the growth medium affects biofilm growth in vitro.^44,45 In this study, biofilms of P. aeruginosa N6P6 and P. pseudoalcaligenes NP103 showed luxuriant growth (A₅₉₅ > 0.6) at all the tested pH. Earlier reports showed that P. aeruginosa can form dense biofilm at wide pH range,^46–48 however in our study optimum pH for P. aeruginosa N6P6 and P. pseudoalcaligenes NP103 were found to be 7.0 and 6.0 respectively. To adapt pH stress during planktonic mode of growth, bacteria modulates their protein expression pattern however, in the case of biofilms, pH stress considerably affect the synthesis of EPS.^49–51 Few reports showed enhanced EPS production in response to higher pH i.e. alkaline pH in the case of Bacillus polymyxa⁵² and a marine bacterium Halomonas sp. biofilm.⁵³ Thus, it can be concluded that ideal pH can enhance biofilm growth by increasing EPS synthesis. Another factor that plays an important role in biofilm formation is Na⁺ concentration. Marine bacteria have highly specific requirement of Na⁺ for growth. Optimum level of Na⁺ is necessary for cells aggregation and attachment to the surface in marine bacteria.^17,54 In this study, S. acidaminiphila NCW702 and P. aeruginosa N6P6 both were able to form biofilm at various concentrations of Na⁺ and the absence of Na⁺ caused decline in biofilm growth of P. aeruginosa N6P6 biofilm growth.

Biofilm formation and the structure is a characteristic of species,⁵⁵ which was also observed in the present study. The bacterial biofilms have many structural elements and spatial variations which is attributed to cells aggregation and EPS components.^56–58 Staining of marine biofilms with SYTO9 and ConA-TRITC revealed the total biomass and EPS composition of the biofilms. SYTO9 specifically stains the live cells, whereas ConA binding is indicative of the presence of glycoproteins and polysaccharides i.e. EPS component in the biofilm.¹³ Cells/EPS ratio for biofilm was maximum for P. pseudoalcaligenes NP103 (3.08) and this ratio was >1.5 for S. acidaminiphila NCW702, S. luteola NR402, P. mendocina NR802 and Pseudomonas sp. NP202 indicating abundance of actively growing cells similar to the observations of Xiao et al.⁵⁹ Total biomass (or biovolume) represents the overall volume of the biofilm, and provides an estimate of the biomass in the biofilm. Total biomass was maximum for P. aeruginosa N6P6 followed by S. acidaminiphila NCW702. Patrauchan et al. studied the biofilm growth of a marine bacterium Pseudoalteromonas sp.⁶⁰ They proposed that increase in total biomass of biofilm is associated with the establishment of thicker microcolonies, indicating uniform colonization over the substratum.^20,61 S. acidaminiphila NCW702 had maximum surface to biovolume ratio, whereas P. aeruginosa N6P6 had least surface to biovolume ratio. Lower surface to biovolume ratio indicates presence of small cell clusters attached to the substratum. Single cells and small cell clusters naturally have a higher surface to volume ratio than larger micro-colonies.²⁰

A significant positive correlation between biomass, thickness and diffusion distance of biofilms was observed among the marine bacteria isolates (Table 2). Similar observations were reported by Mueller et al.,⁶² and Martín-Rodríguez et al.,¹⁷ in phototrophic and Shewanella biofilms respectively. Whereas, a non-significant moderate negative correlation of biofilm roughness coefficient was observed between biomass, thickness and diffusion distance (P > 0.05). Interestingly, it was observed from correlation matrix that surface to volume ratio has no significant relation with biofilm biomass, thickness, diffusion distance and roughness coefficient (P > 0.05). In the present study, marked variations in the biofilm topological parameters were noticed among the isolates. Analysis of the CSLM data revealed that different biomolecules present in the biofilms and its structural parameters are characteristic of each bacterial species.⁵⁸ Biofilm with high R* indicates the ability of biofilm growth through micro-colonies.²¹ R* was maximum for L. fusiformis NCW903 followed by P. lautus NE3B01. Nutrient concentration can considerably affect biofilm spatial arrangement and EPS content. Srinandan et al.,⁶³ studied the effect of nutrients on biofilm formation of a denitrifying bacterium Paracoccus sp. The average thickness, roughness coefficient and biovolume of the Paracoccus sp. biofilm showed an increase in the presence of succinate, Mg²⁺ and Ca²⁺ due to increase in EPS synthesis. Patrauchan et al., also reported increase in the total EPS content of marine bacterium Pseudoalteromonas sp. biofilm in presence of Ca²⁺.⁶⁰ Various components of biofilm associated EPS have structural, sorptive and nutritive properties.⁶⁴ EPS helps in cell–cell adhesion, formation of flocs, water retention, and protection from desiccation. It also helps in sorption of nutrients from the surrounding milieu.⁶⁴

The findings of FTIR spectroscopy indicated the presence of amide groups in the biofilm associated EPS which was also reported by Beech et al.⁶⁵ Similarly, the presence of C–O–C ring of carbohydrates was also observed which was earlier reported by Braissant et al.⁶⁶ Most of the homopolysaccharides and heteropolysaccharides found in a biofilm resemble general polysaccharide components.⁶⁷ Presence of phosphate group in FTIR spectra, illustrated the existence of DNA in EPS.⁶⁸ The polyanionic nature of EPS was revealed in the FTIR study which exhibits electrostatic properties and draws various contaminants in the vicinity of biofilm enhancing the bioremediation of pollutants.^69,70 EPS components increase the solubility and binding of hydrophobic compounds, thus, play a crucial role in wastewater treatment and biodegradation.^12,71 EPS is also imparted with emulsifying property, which enhances the solubility of hydrocarbons.^72,73 The components of EPS i.e. polysaccharides, proteins and lipids are potential bioemulsifiers suitable for industrial and environmental application.⁷⁴ In a recent study, on P. aeruginosa it was reported that the EPS can affect both biofilm performance and PAH degradation process.^75,76 Enhancing EPS production by any means (using exogenous AIs, modulating QS or biofilm growth) can affect cell–cell interaction and cell surface properties of biofilm forming cells. The bacterium also degrades PAH via catechol pathway. Exopolysaccharides can be highly diverse, even among the bacterial strains of a single species. For instance, P. aeruginosa produces at least three kinds of exopolysaccharide; alginate, Pel and Psl.^25,77 Although exopolysaccharides are essential components of biofilm, several surface associated proteins, which play crucial role in biofilm development have been studied in bacteria such as Enterococcus faecalis, Staphylococcus aureus, Salmonella enteric, Pseudomonas fluorescens, P. putida.^78–81 The carbohydrate and protein content is illustrated in ESI Fig. 4.† Carbohydrate content was maximum in biofilm associated EPS of P. aeruginosa N6P6, whereas total protein content was higher in S. acidaminiphila NCW702 biofilm-EPS. Bacteria such as Halomonas eurihalina, Enterobacter cloacae and Gordonia alkanivorans were reported to produce EPS with emulsifying property.⁸² The results showed in this study state that E₂₄% of P. pseudoalcaligenes NP103, P. aeruginosa N6P6, L. fusiformis NCW903 and P. mendocina NR802 were comparatively higher than emulsification index of P. oleovorans and Pseudoalteromonas sp. reported earlier.^83,84 However, the emulsifying property of biofilm-EPS was less when compared to the exopolysaccharide synthesized by marine bacteria Halomonas eurihalina and Gordonia alkanivorans.^79,82 Biofilm (or associated EPS) are natural emulsifying agents and important tools for remediation of hydrocarbons.⁸⁵ Thus, the high emulsifying properties of biofilm associated EPS of P. pseudoalcaligenes NP103 and P. aeruginosa N6P6 show promising application of these EPS producing biofilm forming cultures for PAHs degradation (ESI Fig. 4†).

Phenanthrene degradation by all marine bacteria isolates was found to be significantly more in both planktonic as well as biofilm modes as compared to the degradation potential of Neptunomonas naphthovorans isolated from marine sediments by Li and Chen.⁸⁶ The bacterium N. naphthovorans was reported to degrade 30% of phenanthrene (at an initial concentration 2 mg l⁻¹) in 5 d. Whereas in the present study, P. aeruginosa N6P6 biofilm was able to degrade 85.5% of phenanthrene in 7 d, which was also higher than the biosurfactant producing bacterium Bordetella petrii reported by Reddy et al.⁸⁷ The pyrene degradation rate differed significantly among the various marine bacterial isolates studied (one way ANOVA, P < 0.05). Pyrene degradation potential of the tested marine bacterial isolates was higher as compared to previous findings on Ochrobactrum sp. and Pseudomonas sp.^88,89 Due to poor solubility, the metabolism of PAHs is often restricted by the mass transfer to the aqueous phase. The efficient degradation of phenanthrene and pyrene by marine bacterial biofilms as compared to their planktonic counterparts, showed the potential of sessile bacterial community in the bioremediation technology. The effectiveness of biofilm mediated PAHs degradation rate has been reported previously.⁹⁰ Production of metabolites and expression of PAHs degrading genes were also found to be higher in biofilms.¹⁹

Conclusions

The present study showed that physico-chemical parameters such as temperature, pH, ionic strength and carbon source are key determinants for biofilm growth in marine bacteria. Although in natural environment, biofilm growth is largely governed by environmental factors, however, in vitro conditions, the physiological and nutrient parameters significantly increase the development of biofilm. A significant difference was observed in biofilm growth condition, topological parameters and EPS composition among the various isolates, with significant positive or negative correlation with PAHs degradation. The adherence of bacteria with substratum and spatial arrangement of cells within biofilms varied among bacteria and familiarity on these aspects paved way in the study of biofilm mediated bioremediation of PAHs.

Acknowledgements

The authors would like to acknowledge the authorities of NIT, Rourkela and BARC facilities of IGCAR, Kalpakkam for providing research facilities. Financial supports received from Department of Biotechnology, Ministry of Science and Technology, Government of India on the research projects on bioremediation by biofilm forming marine bacteria are gratefully acknowledged.

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Footnotes

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra12824f

‡ Equal contributors.

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