Bioresource Technology 193 (2015) 274–280

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Bioresource Technology journal homepage: www.elsevier.com/locate/biortech

Extracellular polymeric substances govern the development of biofilm and mass transfer of polycyclic aromatic hydrocarbons for improved biodegradation Yinping Zhang a,b,1, Fang Wang a,1, Xiaoshu Zhu b, Jun Zeng a, Qiguo Zhao a, Xin Jiang a,⇑ a b

State Key Laboratory of Soil and Sustainable Agriculture, Institute of Soil Science, Chinese Academy of Sciences, Nanjing 210008, China Nanjing Normal University Center for Analysis and Testing, Nanjing 210046, China

h i g h l i g h t s  The production of EPS controlled the formation of biofilm.  The biofilm enhanced biodegradation of PAHs, compared to suspended bacteria.  The EPS secreted from bioflim resulted in enhanced bioavailability of PAHs.

a r t i c l e

i n f o

Article history: Received 13 April 2015 Received in revised form 21 June 2015 Accepted 22 June 2015 Available online 27 June 2015 Keywords: Polycyclic aromatic hydrocarbons (PAHs) Extracellular polymeric substances (EPS) Biofilms Biodegradation

a b s t r a c t The hypothesis that extracellular polymeric substances (EPS) affect the formation of biofilms for subsequent enhanced biodegradation of polycyclic aromatic hydrocarbons was tested. Controlled formation of biofilms on humin particles and biodegradation of phenanthrene and pyrene were performed with bacteria and EPS-extracted bacteria of Micrococcus sp. PHE9 and Mycobacterium sp. NJS-P. Bacteria without EPS extraction developed biofilms on humin, in contrast the EPS-extracted bacteria could not attach to humin particles. In the subsequent biodegradation of phenanthrene and pyrene, the biodegradation rates by biofilms were significantly higher than those of EPS-extracted bacteria. Although, both the biofilms and EPS-extracted bacteria showed increases in EPS contents, only the EPS contents in biofilms displayed significant correlations with the biodegradation efficiencies of phenanthrene and pyrene. It is proposed that the bacterial-produced EPS was a key factor to mediate bacterial attachment to other surfaces and develop biofilms, thereby increasing the bioavailability of poorly soluble PAH for enhanced biodegradation. Ó 2015 Published by Elsevier Ltd.

1. Introduction Polycyclic aromatic hydrocarbons (PAHs) occur in various ecosystems and are causing increased environmental concern due to their toxicity to humans and ecosystems. Bioremediation is widely accepted as a green and promising new technology for the ecological recovery of PAH-contaminated sites (Vaiopoulou et al., 2015). Indeed, the degradation capacity can be enhanced by the inoculation of specialized bacterial isolates for degradation of water-soluble contaminants, such as phenol (Jemaat et al., 2014), in wastewater treatment systems. Whereas, enhanced biodegradation in large scale ⇑ Corresponding author. Tel.: +86 881195; fax: +86 25 86881000. 1

E-mail address: [email protected] (X. Jiang). Equal contribution.

http://dx.doi.org/10.1016/j.biortech.2015.06.110 0960-8524/Ó 2015 Published by Elsevier Ltd.

PAH-contaminated soil sites as a result of inoculation with PAH-degrading bacterial strains has never been demonstrated. Crucial roles for the efficiency of soil bioremediation are the survival of added bacteria and the degradation rate of contaminants (Johnsen et al., 2005), which would affect their bioavailability to the active bacteria. A general definition of bioavailability is the degree of interaction of chemicals with living organisms. Factually, in the PAH-contaminated soil, PAH would easily partition to organic materials in the soil, resulting in its poor bioavailability (Lehnik-Habrink et al., 2010). Furthermore, the inoculated planktonic bacteria might lose their degradation ability and may be barely keep alive in the soil environment because of abiotic and biotic environmental stresses (Johnsen et al., 2005). Therefore, seeking and understanding the intrinsic mechanisms between bacteria and PAHs would enable the accurate optimization of bioremediation strategies.

Y. Zhang et al. / Bioresource Technology 193 (2015) 274–280

Interestingly, the biofilms can function as a cooperative consortium, in a relatively complex and coordinated manner, to obtain benefit of the phenotypic versatility of their neighbors (Denkhaus et al., 2007), thereby showing the potential to withstand the environmental abiotic and biotic stresses. For example, the formation of biofilm has enhanced N,N-dimethylformamide degradation in the presence of secondary carbon sources (Nisha et al., 2015). In the membrane aerated biofilm reactors, the biofilm can trap fluoride during the biodegradation of 4-fluorobenzoate (Misiak et al., 2011). However, the bacterial cells may probably undergo profound changes during their transition from planktonic microorganisms to cells that are part of a complex, surface-attached biofilms. With the genetic and molecular approaches, important genes and regulatory circuits have been identified for bacterial biofilms formation and growth, and indicated that, compared to planktonic cells, the expression of genes for polysaccharide production is up-regulated in biofilms in liquid media. Further, high cellular levels of cyclic di-GMP are produced (Rigano et al., 2007), which are apparently involved in bacterial attachment and its subsequent biofilms formation by conditioning the compositions and external structures of bacterial surface extracellular polymeric substances (EPS) that are mainly composed of polysaccharides, proteins and nucleic acids. With synchrotron radiation-based fourier transformed infrared spectroscopy, a variety of active functional groups-mainly including carboxyl, phosphoric, amine, and hydroxyl groups-have been detected in the EPS of the bacterial surface. These functional groups represent potential binding sites for the absorption of toxic substances, such as Cd, Cu (Wei et al., 2011) and water soluble dye (Binupriya et al., 2010). Furthermore, the analysis of EPS compositions with the nuclear magnetic resonance technique and three-dimensional excitation-emission matrix fluorescence spectroscopy both indicate that EPS carries large quantities of aromatic structures and unsaturated fatty chains, and contains three dimensional networks (Sheng et al., 2010), which can interact with molecules containing aromatic rings. Thus, the possible role of EPS involved in the bacterial biofilm formation and the subsequent biodegradation of hydrophobic PAHs is particularly exciting in view of their potential application in the bioremediation PAHs-contaminated soil environments. However, no specific information on the dominant mechanism by the bacteria has been provided. The present study was conceived to test the hypothesis that EPS controlled the attachment of bacteria and subsequently favored mass transfer of PAHs, thereby enhancing the biodegradation efficiency. Low-molecular-weight (LMW) PAHs are acutely toxic, while high-molecular-weight (HMW) PAHs are mutagenic and carcinogenic, thus phenanthrene (PHE) and pyrene (PYR) were selected as the model LMW and HMW PAHs, respectively. Correspondingly, PHE- and PYR-degrading bacterial strains, Micrococcus sp. PHE9 (strain PHE9) and Mycobacterium sp. NJS-P (strain NJS-P) were employed to investigate the attachment and biodegradation behavior. The production of EPS was quantified and its biosorbed PHE and PYR were also analyzed to study the biodegradation progress.

2. Methods 2.1. Chemicals and materials All the chemicals used were of the highest available purity. Phenanthrene (PHE) and pyrene (PYR) were obtained from Supelco Corporation (USA, purity >95%), while n-hexane, methanol, acetone, and all other chemicals were obtained from Nanjing Chemicals Reagent (Nanjing, China). Humin was purchased from

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Beijing Kaien Company (Beijing, China). For use in the trials, humin was purified. Briefly, humin was rinsed with milli-Q water, and then dried at 108 °C for 5 h. The elemental composition (C, H, N, S) of the rinsed humin, without pH adjustment, was determined by an elemental analyzer (Vario Micro, Elementar, Germany) – using the high-temperature combustion method – as 53.32%, 2.81%, 1.12%, and 0.49% for C, H, N, and S, respectively. 2.2. Bacterial strains and preparation Micrococcus sp. PHE9 (strain PHE9) and Mycobacterium sp. NJS-P (strain NJS-P), both of which were deposited in the China Center for Type Culture Collection with No. CCTCC AB 2010362 and No. CCTCC M 2011011, respectively, were used as test bacteria. Both strains were thawed and regenerated using an autoclaved nutrient broth (NB) medium. Bacterial cells of strains PHE9 and NJS-P were harvested at the exponential phase at an optical density at 600 nm (OD600) of 0.8 and 0.6, respectively. The cells were re-suspended in phosphate buffer solution (PBS) after washing twice with PBS, with cell densities of each strain adjusted to an OD600 of 0.25, which corresponds to approximately 1.6  108 CFU mL 1. 25.0 mL of the cell suspension was used as untreated bacteria (with EPS), and another 25.0 mL of the cell suspension was used to prepare EPS-extracted bacteria (without EPS). EPS-extracted cells were achieved via the employment of cation exchange resin treatment technique as which displays no effect on rates of bacterial survival and growth described in the previous publication (Zhang et al., 2011). The treated bacteria pellets were then re-suspended in PBS to the initial volumes. The cell surface hydrophobicity was quantified using a test that measured bacterial adhesion to hydrocarbons by calculating the percentage of cells adhering to hexadecane (Rosenberg, 2006). The percentage of cells adhering to hexadecane were 35.50% for strain PHE9 and 41.27% for strain NJS-P when grown on NB medium, while the EPS-extracted cells showed much less adhesion behavior. 2.3. Biodegradation kinetic experiments To test the PAH-removing efficiency of both incubated bacteria and EPS-extracted bacteria, a batch of the experiment was set up in a series of 50-mL sterile tubes containing 0.5 mL of PHE (2000.0 mg L 1) or PYR (2000.0 mg L 1) acetone stock solution. After evaporating the acetone, 10.0 mL sterile mineral medium (MM) was added to give an initial concentration of 100.0 mg L 1. For biofilm formation, 0.1 g of humin that served as carriers was then added into each tube. Simultaneously, biofilm formation was started by inoculating 1.0 mL of untreated PHE- and PYR-degrading bacteria into corresponding tubes. An aliquot of 1.0 mL of EPS-extracted bacteria were subsequently inoculated into a series of tubes as controls, for evaluating their biofilm formation and biodegradation kinetics. The sets without inoculation were used to assess the abiotic loss of PHE and PYR. All the flasks were then shaken at 160 rpm and 28 °C in the dark. Due to the low water solubility of PHE and PYR, sacrifice sampling was conducted to ensure better determination of PHE and PYR concentrations in the whole culture. At time intervals of 12 h for PHE and over 3 days for PYR, three tubes were sacrificed for cell counting and quantification of PAHs. Another three tubes were drawn out for EPS extraction and quantification. At the end of experiment, approximately 0.1 g of humin was collected for scanning electron microscopy (SEM) observation of biofilms. Bacterial cells of strains PHE3 and NJS-P still suspended in the MM were examined at an optical density at 600 nm (OD600).

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2.4. Scanning electron microscopic observation of bacterial biofilms According to the method reported by Johnsen and Karlson (2004), the biofilms on humin were firstly fixed with 2.5% glutaraldehyde in PBS for 2 h. Secondly, the bacteria were gradually dehydrated with 30%, 50%, 70%, and 90% of ethanol for 10 min, respectively. Finally, the samples were critical point-dried (CPD, Emitech k850, U.K.). The biofilms attached to humin surfaces were coated with gold and observed with a scanning electron microscope (SEM, LEO1530vp, Germany) at an acceleration voltage of 8 kV. 2.5. EPS extraction and determination To investigate the different contributions of EPS excreted around the bacterial cell surfaces on the biodegradation of PHE and PYR, EPS produced by both suspended bacteria and biofilms had to be extracted and monitored. The EPS were extracted using a mild method and a harsh method in sequence, that is, the oscillation-ultrasound method followed by the cation exchange resin technique (Wang et al., 2009). Briefly, the tubes were ultra-sonicated (40 W, 21 KHz) for 2 min and then centrifuged at 4024g for 30 min at 4 °C. The organic compounds in the supernatant were regarded as the loosely bound EPS. After that, the residues left in the centrifuge tube were collected and re-suspended again in milli-Q water to the initial volumes, and transferred to a sterilized extraction beaker for further tightly bound EPS extraction. Firstly, 1.0 g cation exchange resin (CER, 001  7, sodium form, Sanxing, China), which was rinsed and soaked in 8% NaCl for 5 h prior to use, was added to the extraction beaker. The whole suspension was then stirred at 400 rpm for 2 h at 4 °C, and allowed to settle for 5 min so as to separate the CER. Afterward, the upper suspension was centrifuged at 4024g for 30 min at 4 °C, and the supernatant was taken to be the tightly bound EPS. Finally, both loosely bound and tightly bound EPS were combined and filtered through sterilized 0.22 lm cellulose acetate filters to remove particles and bacterial cells, and used as the EPS fraction for further biochemical component analysis and PAHs quantification. Polysaccharide content in the EPS was analyzed using a colorimeter with the Phenol–sulfuric acid method using dextran T70 (Pharmacia, Switzerland) as the standard (Dubois et al., 1956). The content of protein and humic acid in the EPS solutions was determined by the modified Lowry method (Frolund et al., 1995), with bovine albumin serum (Amresco, USA) and humic acid as respective standards. The content of DNA in the EPS was determined by the diphenylamine colorimetric method with 2-deoxy-D-ribose (DNA, AMRESCO, USA) as the standard (Sun et al., 1999). 2.6. Extraction of residual PAHs and HPLC analysis PHE and PYR in the MM and on the EPS was extracted by liquid– liquid extraction with an equivalent volume of n-hexane to MM. PHE and PYR adsorbed to humin and biofilm was extracted for three cycles with n-hexane:acetone (4:1, v/v) mixture at 100 °C and 1500 psi by accelerated solvent extraction (ASE200 Dionex, USA). The extracts were purified with silica-gel columns according to the protocols described in a previous report (Zhang et al., 2012). The analysis of PHE and PYR were performed using a high performance liquid chromatography system (Shimadzu, LC-20A, Japan) equipped with a fluorescence detector and a supelco PAHs special chromatographic column (250  4.6, 5 lm, Supelco, USA). The excitation and emission wavelengths were 250 nm and 365 nm for PHE, and 320 nm and 380 nm for PYR, respectively. The mobile phase was acetonitrile and milli-Q water (9:1, v/v) with a flow rate

of 1.50 mL min 1. The column temperature was held at 40 °C for a 9-min program. 2.7. Data analysis and quality control Spiking recovery experiments of PHE and PYR were conducted at concentrations of 100.0 mg kg 1 in the humin, 10.0 mg L 1 in the MM and in the EPS solutions, respectively. Sampling and analysis were performed using the same procedure as above, and the average recoveries for three replicates ranged from 85.31% to 94.81%. PAH bound to EPS was calculated as follows: C = Cd S, where C is the concentration that bound to EPS (mg L 1), Cd is its determined concentration, S is the solubility of PAH in the aqueous. Linear regression was used for determination of PHE and PYE biodegradation rates by suspended bacteria and biofilms. All the data were analyzed with SPSS 13.0. Duncan’s Multiple Range Test performed at significant level of P < 0.05.

3. Results 3.1. Biofilms grown on humin Representative views of the biofilms for untreated bacteria with EPS and EPS-extracted bacteria attached to humin are observed (Supplementary Fig. S1). For the untreated cells with EPS, clear biofilms developed for both strains (Fig. S1a and c), while in the control setup, only a very small number of cells of strain PHE9 and NJS-P after EPS extraction were attached to humin surfaces (Fig. S1b and d), showing that EPS sustained biofilm development. Furthermore, the production of EPS was observed in the biofilms, but barely any EPS was present in the EPS-extracted bacteria, indicating the formation of biofilms was associated with EPS production. Additionally, the untreated cells in the supernatants decreased to approximately 3.4  106 CFU mL 1, but those of the EPS-extracted cells displayed different trends, with an increase of cell numbers from about 1.5  107 to 3.7  107 and 3.9  107 CFU mL 1 for strain PHE and strain NJS-P, respectively, which further confirmed the development of biofilms. 3.2. PAH-degrading activity by biofilms and EPS-extracted bacteria The degradation of PHE and PYR by biofilms, compared to suspended EPS-extracted bacteria, showed different kinetics (Fig. 1). In the sterile control, the PHE and PYR concentrations decreased only a little during the whole incubation period. In contrast, in the inoculated sets with EPS-extracted bacteria, PHE and PYR decreased continuously. Approximately 66% of PHE was degraded in 3 days, and about 58% of PYR was degraded in 18 days, corresponding to approximately 24.38 and 3.24 mg L 1 day 1, respectively (Fig. 1, Table 1). However, with the inoculation of biofilms, more than 90% of PHE and PYR on humin and biofilms were reduced during the incubation period, and the degradation rates by biofilms were approximately 35.18 and 5.08 mg L 1 day 1 for PHE and PYR, respectively. The results showed that the biodegradation rates by biofilms were significantly higher than those with EPS-extracted bacteria, implying that the inoculation of biofilms significantly enhanced biodegradation efficiencies of PHE and PYR. PHE and PYR were further discovered in the extracted EPS solutions (Fig. 2), and the concentrations bound to EPS increased significantly for both EPS-extracted bacteria and biofilms in the growth period. Apparently, EPS from biofilms bound more PHE and PYR, with its values reaching 6.97 mg L 1 and 8.63 mg L 1, than EPS-extracted bacteria. Thereafter, both concentrations decreased dramatically in the later incubation period due to biodegradation.

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100

100

80

80

-1 Pyrene (mg L )

-1 Phenanthrene (mg L )

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60 40 Control Biofilms EPS-extracted bacteria

20 0

0.0 0 .5

1.0

1.5 2.0 Time (d)

60 40 20 0

2.5

3.0

0

5

10 Time (d)

15

Fig. 1. Biodegradation kinetics of phenanthrene and pyrene by biofilms development on humin and EPS-extracted bacteria.

Table 1 Kinetic parameters for biodegradation of phenanthrene (PHE) and pyrene (PYR) by extracellular polymeric substances (EPS)-extracted bacteria and biofilms of Micrococcus sp. PHE9 and Mycobacterium sp. NJS-P. Bacteria

PAHs

Inoculation method

Degradation efficacy (%)

Degradation rate (mg L 1 day 1)

Micrococcus sp. PHE9

PHE

Biofilms EPSextracted bacteria

98.71 ± 1.53a 66.02 ± 4.35b

35.18 ± 2.57a 24.38 ± 2.58b

Mycobacterium sp. NJS-P

PYR

Biofilms EPSextracted bacteria

90.67 ± 4.51a 57.67 ± 11.5b

5.08 ± 0.08c 3.24 ± 0.23d

The significant differences, within one column, were less than 0.05.

The results indicated that EPS secreted from bacterial surface led to the accumulation of PAHs. 3.3. EPS production characters corresponding to biodegradation The produced proteins, polysaccharides and DNA outside the cell of strain PHE9 and NJS-P displayed different characters. The concentrations of DNA were rather low, less than 0.04 mg L 1, and accounted for only a small portion in the EPS components (less than 5%), while proteins and polysaccharides were the major biochemical components of the EPS, with a proportion of over 95%. Further, the dynamics of the two main components of EPS, protein and polysaccharide, were monitored (Fig. 3). The production of proteins and polysaccharides increased dramatically for EPS-extracted bacteria of strain PHE9 in the first day of incubation, ranging from 0 to 5.47 mg L 1, and from 0 to 0.70 mg L 1, respectively, followed by a plateau in the subsequent period (Fig. 3a). Whereas, the values for strain NJS-P increased steadily during the

18 days of incubation, reaching 8.87 and 2.50 mg L 1, respectively (Fig. 3c). However, the total contents of EPS on biofilms (Fig. 3b and d) were higher than those of EPS-extracted bacteria (Fig. 3a and c), with the produced proteins and polysaccharides reaching 85.87 and 13.60 mg L 1, respectively. In fact, with PHE as the substrate, the ratio of proteins to polysaccharides for EPS-extracted bacteria of strain PHE9 decreased from 9.3 to 7.9, but that of biofilms ranged from 5.4 to 5.9. On the other hand, in the presence of PYR, the values for the proteins to polysaccharides ratio decreased from 6.4 to 3.5 for EPS-extracted bacteria of strain NJS-P, and it decreased from 7.2 to 6.3 for biofilms. Additionally, with the observation of SEM, the biofilm structures were rather constant at end of the experiment (Fig. S2), and a large number of excreted EPS were observed to be accumulated on the surfaces of biofilms for both strains. Multivariate regression was further performed to relate biodegradability of the biofilms and EPS-extracted bacteria (Fig. 4). After EPS extraction, the biodegradation capacity was significantly impacted, and it could be observed that the quantity of protein and polysaccharide produced from EPS-extracted bacteria of strain PHE9 has no significant obvious relations to PHE degradation rates (R2 6 0.64 for protein, R2 6 0.67 for polysaccharide, P > 0.05, Fig. 4a). To some extent, the production of EPS of strain NJS-P was correlated with the biodegradation of PYR (R2 6 0.93 for protein, R2 6 0.88 for polysaccharide, P < 0.05, Fig. 4c), though the biodegradation rates of PYR were much slower than that of PHE. Compared to EPS-extracted bacteria, the production of EPS demonstrated much better correlations with PAH degradation for biofilms that formed by strain PHE9 (R2 P 0.98 for protein, R2 P 0.96 for polysaccharide, P < 0.01, Fig. 4b) and by strain NJS-P (R2 P 0.98 for protein, R2 P 0.94 for polysaccharide, P < 0.01, Fig. 4d), respectively. These results indicated that the production of EPS components favored the biofilms formation for biodegradation of PAHs.

EPS-extracted bacteria Biofilms

10 Pyrene (mg L-1)

Phenanthrene (mg L-1)

8 6 4 2

8 6 4 2

0

0 0.0

0.5

1.0

1.5 2.0 Time (d)

2.5

3.0

0

5

10 Time (d)

15

Fig. 2. Bioaccumulation of phenanthrene and pyrene in the layer of EPS secreted from EPS-extracted bacteria and biofilms.

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Y. Zhang et al. / Bioresource Technology 193 (2015) 274–280 EPS-extracted bacteria

-1

-1

Concentrations (mg L )

a

8 Concentrations (mg L )

Biofilms

6 4 2

b

80 60

20

0 0.0

0.5

1.0

1.5

2.0

2.5

3.0

0.0

0 .5

1.0

Time (d)

1.5

2.0

2.5

3.0

Time (d)

c

8

Concentrations (mg L-1 )

Concentrations (mg L-1)

10

6 4 2

d

80 60

Proteins Polysaccharides

20

0 0

5

10

15

20

0

5

Time (d)

10

15

20

Time (d)

Fig. 3. The production of proteins and polysaccharides for EPS-extracted bacteria of Micrococcus sp. PHE9 (a), biofilms of Micrococcus sp. PHE9 (b), EPS-extracted bacteria of Mycobacterium sp. NJS-P (c), and biofilms of Mycobacterium sp. NJS-P (d).

EPS-extracted bacteria

Concentrations (mg L-1)

a

6 4 2

Concentrations (mg L-1)

Biofilms

8

b

80 60 40 20

0 60 80 100 PHE concentrations (mg L-1)

Concentrations (mg L-1)

10

c

8 6 4 2

0

20 40 60 80 100 PHE concentrations (mg L-1)

d Concentrations (mg L-1)

40

80 60 Proteins polysaccharides

40 20

0 40

60

80

100

PYR concentrations (mg L-1)

0

20

40

60

80

100

PYR concentrations (mg L-1)

Fig. 4. The correlations of proteins and polysaccharides production with PHE biodegradation rates by EPS-extracted bacteria of Micrococcus sp. PHE9 (a) and biofilms of Micrococcus sp. PHE9 (b), and the correlations of proteins and polysaccharides production with PYR biodegradation rates by EPS-extracted bacteria of Mycobacterium sp. NJS-P (c) and biofilms of Mycobacterium sp. NJS-P (d).

4. Discussion 4.1. Biofilm formation depends on the production of EPS Bacteria with EPS can attach to humin particles as over 95% of bacteria without EPS-extraction attached to humin particles while

those with EPS-extracted ones showed poor attachment and colonization on humin surfaces (Fig. S1). Furthermore, attachment to biotic and abiotic surfaces would stimulate EPS synthesis during biofilms formation (Jayasinghearachchi and Seneviratne, 2006), the secreted surface-active EPS therefore could be directly observed on the biofilms surfaces with SEM technique for strain

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PHE9 and strain NJS-P (Fig. S1), and EPS is a factor to control the biofilms formation of bacteria. It is well known that bacteria are negatively charged on their surface (Ayala-Torres et al., 2014), which could generate a repulsive force between neighboring cells that prevent their close contact. However, the EPS biopolymers on the cell surface, mainly including proteins and polysaccharides, may cause steric stabilization and, more importantly, steric destabilization. This would then bring about additional interactions which may offset the electrostatic repulsive forces between cells and facilitate cell attraction and attachment, thereby developing biofilms. This could be confirmed by the report that glycoprotein-rich EPS produced by Acinetobacter venetianus strain VE-C3 favored its attachment at the diesel-water interface even when the interactions force between surfaces is repulsive (Baldi et al., 2003). Thereby, EPS produced during the process of degradation of the substrates favored biofilms formation for both strain PHE9 and NJS-P. Furthermore, since a large proportion of humin is mainly formed from parent microbial-derived materials biopolymers, such as peptidoglycan (Simpson et al., 2007), the interactions between EPS biopolymers and humin particles can be attractive, and the excreted extracellular proteins on bacterial surface act as ‘‘adhesive polymers’’ and easily convert the humin surface into a region of gel-like nature, which form a microenvironment around the cells helping biofilms development. Additionally, as suggested by Rodrigues et al. (2005), the ratio of proteins to polysaccharides can be used to determine the structure and solidity of biofilms. In the experiment, the values of proteins to polysaccharides ratios were higher than 5, which indicated that homogenous, smooth and fragile biofilms developed for both untreated bacteria. In contrast, the EPS-extracted bacteria should directly interact with water molecules and humin particles, where the repulsive force dominates, resulting in difficulties of biofilms development and fluctuant ratios of proteins to polysaccharides. Thus, the bacteria self-secreted EPS is essential to effective cell attachment and biofilms development on humin. 4.2. Development of biofilms enhanced biodegradation of PHE and PYR Compared to inoculation of EPS-extracted bacteria, the inoculation of bacteria without EPS-extraction significantly enhanced biodegradation of both PHE and PYR (Fig. 1 and Table 1). EPS form a microenvironment around the bacterial cell surface helping them to avoid direct contact of PHE and PYR molecules to bacterial cell surfaces conferring protection from hydrophobicity damage. Simultaneously, EPS favored bacteria attachment to humin surfaces and formation of biofilms (Fig. S1), which might permit a higher bacterial growth rate. Furthermore, as suggested by Tribelli et al. (2012), in hydrocarbon contaminated micro-aerobic environments, the protein for alkane monooxygenase could be preferentially expressed at high cell densities, making strain PHE9 and strain NJS-P more active in biofilms than that suspended in aqueous phase. Additionally, the biofilms could increase bioavailability by enlarging the interfacial area between bacteria and the hydrophobic carbon substrate, thereby accelerating PHE and PYR biodegradation in biofilms. It could be proposed that the degradation of PHE and PYR by biofilms via the research of degradation dynamics, compared to suspended EPS-extracted bacteria and suspended bacteria with EPS, has more significance in practice. It is worth noting that EPS in strain PHE9 and strain NJS-P played an important role in the bioaccumulation of hydrocarbon compounds of PHE and PYR. Apolar groups from protein, such as aromatic lipids (Denkhaus et al., 2007), are present in EPS, and hydrophobic high-molecular-weight polysaccharides are distributed across the entire bacterial surface (Neu, 1996), which can offer the sites that the hydrophobic PHE and PYR dissolved in aqueous phase can be bound to, thus PHE and PYR were

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accumulated in the layer of EPS (Fig. 2), and the accumulated amount of PHE and PYR increased with the production of EPS. Whereas, the rapid decrease of PHE and PYR in the layer of EPS after a period time of biodegradation, suggested that the desorption of PHE and PYR from EPS probably occurred, and the biofilms played a role in the utilization of the accumulated compounds. Furthermore, EPS contains large amounts of cationic groups and anionic groups (Gomes et al., 2014), which have the capability to interact with nutrient element. In such a manner, EPS could act as sorbents to facilitate the PHE and PYR mass transfer from aqueous phase to bacterial cells as well as favor the acquisition of water and nutrients from aqueous phase, therefore the production of EPS from biofilms increased microbial activity. Correspondingly, the parameters for the biodegradation of PHE and PYR by biofilms were much more closely correlated with the amount of proteins and polysaccharides than those of the EPS-extracted bacteria (Fig. 4). Additionally, the amount of PHE and PYR biodegraded by strain PHE9 and strain NJS-P is dependent on a combination of physicochemical and biological factors (Simarro et al., 2012). In this study, the physicochemical characteristics of the PHE and PYR are classified as hydrophobic organic pollutants with high octanol–water partition coefficients, and PYR is much more difficult to be dissolved. Thus despite of the fact that the strain NJS-P produced much more EPS (Fig. 3), PYR were biodegraded much more slowly than PHE (Fig. 1).

4.3. Coincident increase of EPS production during the biodegradation process Usually, the bacteria respond to harsh environmental conditions by releasing EPS from the bacterial cell surface (Colica et al., 2014). In the experiment, PHE and PYR are hydrophobic, both of which are poorly accessible to microorganisms. Whereas, the production of EPS can favor the bacteria that when grown on hydrophobic compounds form hydrophobic cell surface and develop the ability to utilize the hydrophobic compounds in the aqueous phase (Bredholt et al., 2002), which would lead to the enhanced bioavailability of PAHs to microorganisms (Fig. 1). Thus, the production of EPS increased for both EPS-extracted bacteria and biofilms in the experiment. Nevertheless, the contents of proteins and polysaccharides produced by EPS-extracted bacteria of strain PHE9 did not change after one day (Fig. 3a). This could be explained by the reason that, after one day of adjustment of its extracellular composition and components of EPS, the bacterial cell surface could adapt to the hydrophobic environment and mass transfer of PHE from aqueous phase to intracellular cells. Furthermore, the amount of synthesized EPS mainly depends on the bioavailability of carbon substrates (Johnsen and Karlson, 2004). PYR displays lower water solubility and lower bioavailability compared to PHE, thus strain NJS-P produced more proteins and polysaccharides than strain PHE9 (Fig. 3) for conditions where its self-surface properties accelerate PYR biodegradation, which agreed with a former report (Johnsen and Karlson, 2004). Simultaneously, after a period of adjustment of its extracellular composition and components of EPS, the bacterial cell surface could adapt to the hydrophobic environment and mass transfer of PAHs from aqueous phase to intracellular cells and the trends for EPS increase correspondingly changed. Additionally, as indicated by Wolfaardt et al. (1995), proteins and polysaccharides can improve the efficiency of biofilms in trapping nutrients by hydrolyzation high-molecular-weight organic matter into a more easily utilizable form before it enters the cell, thus increased production of proteins and polysaccharides of both biofilms could adsorb PHE, PYR and their intermediate products for enhanced their bioaccessibility.

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An interesting feature is that both strain PHE9 and strain NJS-P synthesized more proteins but less polysaccharides with PHE and PYR as carbon substrates, respectively (Fig. 3). This is a common trait of most bacteria during hydrophobic wastewater treatment (Frolund et al., 1996), which is involved in the regulation of organic pollutants’ transport across the bacterial cell membrane and a very complex metabolic pathway (Glazyrina et al., 2010). Additionally, it opens up the possibility of using both bacteria to produce proteins with distinctive properties simply by manipulation of the cultivation conditions. 5. Conclusions In conclusion, both strain PHE9 and NJS-P with EPS developed biofilms on humin surfaces, and the biofilms further enhanced the bioavailability of poorly soluble PAHs for subsequent biodegradation. The EPS contents in biofilms showed significant correlations with the biodegradation of PAHs, therefore the excretion of EPS can enhance bacterial activity for improving PAH removal. It could be proposed that, when choosing and inoculating bacteria into a harsh environment to promote persistent organic pollutants removal, the inoculation of biofilms should be first taken into consideration. Acknowledgements This research was financially supported by the National Key Basic Research Program of China (2014CB441105), the Specific Fund for Agro-Scientific Research in the Public Interest of China (201203045), the National Natural Science Foundation of China (21277148). Special thanks to Emily Juzwiak of Department of Plant, Soil and Microbial Sciences, Michigan State University in America for her great help in language revision. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.biortech.2015.06. 110. References Ayala-Torres, C., Hernandez, N., Galeano, A., Novoa-Aponte, L., Soto, C.Y., 2014. Zeta potential as a measure of the surface charge of mycobacterial cells. Ann. Microbiol. 64, 1189–1195. Baldi, F., Pepi, M., Capone, A., della Giovampaola, C., Milanesi, C., Fani, R., Focarelli, R., 2003. Envelope glycosylation determined by lectins in microscopy sections of Acinetobacter venetianus induced by diesel fuel. Res. Microbiol. 154, 417–424. Binupriya, A.R., Sathishkumar, M., Ku, C.S., Yun, S.I., 2010. Sequestration of Reactive Blue 4 by free and immobilized Bacillus subtilis cells and its extracellular polysaccharides. Colloids Surf. B 76, 179–185. Bredholt, H., Bruheim, P., Potocky, M., Eimhjellen, K., 2002. Hydrophobicity development, alkane oxidation, and crude-oil emulsification in a Rhodococcus species. Can. J. Microbiol. 48, 295–304. Colica, G., Li, H., Rossi, F., Li, D., Liu, Y., De Philippis, R., 2014. Microbial secreted exopolysaccharides affect the hydrological behavior of induced biological soil crusts in desert sandy soils. Soil Biol. Biochem. 68, 62–70. Denkhaus, E., Meisen, S., Telgheder, U., Wingender, J., 2007. Chemical and physical methods for characterisation of biofilms. Microchim. Acta 158, 1–27. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A., Smith, F., 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28, 350–356. Frolund, B., Griebe, T., Nielsen, P.H., 1995. Enzymatic-activity in the activatedsludge floc matrix. Appl. Microbiol. Biotechnol. 43, 755–761.

Frolund, B., Palmgren, R., Keiding, K., Nielsen, P.H., 1996. Extraction of extracellular polymers from activated sludge using a cation exchange resin. Water Res. 30, 1749–1758. Glazyrina, J., Materne, E.M., Dreher, T., Storm, D., Junne, S., Adams, T., Greller, G., Neubauer, P., 2010. High cell density cultivation and recombinant protein production with Escherichia coli in a rocking-motion-type bioreactor. Microb. Cell Fact. 9. Gomes, I.B., Simoes, M., Simoes, L.C., 2014. An overview on the reactors to study drinking water biofilms. Water Res. 62, 63–87. Jayasinghearachchi, H.S., Seneviratne, G., 2006. Fungal solubilization of rock phosphate is enhanced by forming fungal–rhizobial biofilms. Soil Biol. Biochem. 38, 405–408. Jemaat, Z., Suarez-Ojeda, M.E., Perez, J., Carrera, J., 2014. Partial nitritation and ocresol removal with aerobic granular biomass in a continuous airlift reactor. Water Res. 48, 354–362. Johnsen, A.R., Karlson, U., 2004. Evaluation of bacterial strategies to promote the bioavailability of polycyclic aromatic hydrocarbons. Appl. Microbiol. Biotechnol. 63, 452–459. Johnsen, A.R., Wick, L.Y., Harms, H., 2005. Principles of microbial PAH-degradation in soil. Environ. Pollut. 133, 71–84. Lehnik-Habrink, P., Hein, S., Win, T., Bremser, W., Nehls, I., 2010. Multi-residue analysis of PAH, PCB, and OCP optimized for organic matter of forest soil. J. Soils Sediments 10, 1487–1498. Misiak, K., Casey, E., Murphy, C.D., 2011. Factors influencing 4-fluorobenzoate degradation in biofilm cultures of Pseudomonas knackmussii B13. Water Res. 11, 3512–3520. Neu, T.R., 1996. Significance of bacterial surface-active compounds in interaction of bacteria with interfaces. Microbiol. Rev. 60, 151–158. Nisha, K.N., Devi, V., Varalakshmi, P., Ashokkumar, B., 2015. Biodegradation and utilization of dimethylformamide by biofilm forming Paracoccus sp. strains MKU1 and MKU2. Bioresour. Technol. 188, 9–13. Rigano, L.A., Payette, C., Brouillard, G., Marano, M.R., Abramowicz, L., Torres, P.S., Yun, M., Castagnaro, A.P., El Oirdi, M., Dufour, V., Malamud, F., Dow, J.M., Bouarab, K., Vojnov, A.A., 2007. Bacterial cyclic beta-(1,2)-glucan acts in systemic suppression of plant immune responses. Plant Cell 19, 2077–2089. Rodrigues, A.C., Wuertz, S., Brito, A.G., Melo, L.F., 2005. Fluorene and phenanthrene uptake by Pseudomonas putida ATCC 17514: kinetics and physiological aspects. Biotechnol. Bioeng. 3, 281–289. Rosenberg, M., 2006. Microbial adhesion to hydrocarbons: twenty-five years of doing MATH. FEMS Microbiol. Lett. 262, 129–134. Sheng, G.P., Yu, H.Q., Li, X.Y., 2010. Extracellular polymeric substances (EPS) of microbial aggregates in biological wastewater treatment systems: a review. Biotechnol. Adv. 28, 882–894. Simarro, R., Gonzalez, N., Fernando Bautista, L., Carmen Molina, M., Schiavi, E., 2012. Evaluation of the influence of multiple environmental factors on the biodegradation of dibenzofuran, phenanthrene, and pyrene by a bacterial consortium using an orthogonal experimental design. Water Air Soil Pollut. 223, 3437–3444. Simpson, A.J., Song, G.X., Smith, E., Lam, B., Novotny, E.H., Hayes, M.H.B., 2007. Unraveling the structural components of soil humin by use of solution-state nuclear magnetic resonance spectroscopy. Environ. Sci. Technol. 41, 876–883. Sun, Y.D., Clinkenbeard, K.D., Clarke, C., Cudd, L., Highlander, S.K., Dabo, S.M., 1999. Pasteurella haemolytica leukotoxin induced apoptosis of bovine lymphocytes involves DNA fragmentation. Vet. Microbiol. 65, 153–166. Tribelli, P.M., Di Martino, C., Lopez, N.I., Iustman, L.J.R., 2012. Biofilm lifestyle enhances diesel bioremediation and biosurfactant production in the Antarctic polyhydroxyalkanoate producer Pseudomonas extremaustralis. Biodegradation 23, 645–651. Vaiopoulou, E., Misiti, T.M., Pavlostathis, S.G., 2015. Removal and toxicity reduction of naphthenic acids by ozonation and combined ozonation–aerobic biodegradation. Bioresour. Technol. 179, 339–347. Wang, Z., Wu, Z., Tang, S., 2009. Extracellular polymeric substances (EPS) properties and their effects on membrane fouling in a submerged membrane bioreactor. Water Res. 43, 2504–2512. Wei, X., Fang, L.C., Cai, P., Huang, Q.Y., Chen, H., Liang, W., Rong, X.M., 2011. Influence of extracellular polymeric substances (EPS) on Cd adsorption by bacteria. Environ. Pollut. 159, 1369–1374. Wolfaardt, G.M., Lawrence, J.R., Robarts, R.D., Caldwell, D.E., 1995. Bioaccumulation of the herbicide diclofop in extracellular polymers and its utilization by a biofilm community during starvation. Appl. Environ. Microbiol. 1, 152–158. Zhang, Y.P., Wang, F., Bian, Y.R., Kengara, F.O., Gu, C.G., Zhao, Q.G., Jiang, X., 2012. Enhanced desorption of humin-bound phenanthrene by attached phenanthrene-degrading bacteria. Bioresour. Technol. 123, 92–97. Zhang, Y.P., Wang, F., Yang, X.L., Gu, C.G., Kengara, F.O., Hong, Q., Lv, Z.Y., Jiang, X., 2011. Extracellular polymeric substances enhanced mass transfer of polycyclic aromatic hydrocarbons in the two-liquid-phase system for biodegradation. Appl. Microbiol. Biotechnol. 90, 1063–1071.