Bacterial cell surface properties: Role of loosely bound extracellular polymeric substances (LB-EPS)

Colloids and Surfaces B: Biointerfaces 128 (2015) 600–607

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Bacterial cell surface properties: Role of loosely bound extracellular polymeric substances (LB-EPS) Wenqiang Zhao 1 , Shanshan Yang 1 , Qiaoyun Huang, Peng Cai ∗ State Key Laboratory of Agricultural Microbiology, College of Resources and Environment, Huazhong Agricultural University, Wuhan 430070, China

a r t i c l e

i n f o

Article history: Received 26 April 2014 Received in revised form 12 September 2014 Accepted 2 March 2015 Available online 13 March 2015 Keywords: Bacteria LB-EPS Functional group Surface charge Hydrophobicity

a b s t r a c t This study investigated the effect of loosely bound extracellular polymeric substances (LB-EPS) on the comprehensive surface properties of four bacteria (Bacillus subtilis, Streptococcus suis, Escherichia coli and Pseudomonas putida). The removal of LB-EPS from bacterial surfaces by high-speed centrifugation (12,000 × g) was confirmed by SEM images. Viability tests showed that the percentages of viable cells ranged from 95.9% to 98.0%, and no significant difference was found after treatment (P > 0.05). FTIR spectra revealed the presence of phosphodiester, carboxylic, phosphate, and amino functional groups on bacteria surfaces, and the removal of LB-EPS did not alter the types of cell surface functional groups. Potentiometric titration results suggested the total site concentrations on the intact bacteria were higher than those on LB-EPS free bacteria. Most of the acidity constants (pKa ) were almost identical, except the increased pKa values of phosphodiester groups on LB-EPS free S. suis and E. coli surfaces. The electrophoretic mobilities and hydrodynamic diameters of the intact and LB-EPS free bacteria were statistically unchanged (P > 0.05), indicating LB-EPS had no influence on the net surface charges and size distribution of bacteria. However, LB-ESP could enhance cell aggregation processes. The four LB-EPS free bacteria all exhibited fewer hydrophobicity values (26.1–65.0%) as compared to the intact cells (47.4–69.3%), suggesting the removal of uncharged nonpolar compounds (e.g., carbohydrates) in LB-EPS. These findings improve our understanding of the changes in cell surface characterizations induced by LB-EPS, and have important implications for assessing the role of LB-EPS in bacterial adhesion and transport behaviors. © 2015 Elsevier B.V. All rights reserved.

1. Introduction Extracellular polymeric substances (EPS) are biopolymers resulting from bacterial active secretion, shedding of cell surface materials, cell lysis materials and adsorption of organic matters from the environment [1]. EPS are a heterogeneous mixture composed dominantly of polysaccharides and proteins, with nucleic acids and lipids as minor constituents [2,3]. Bacteria have a double-layered EPS structure of loosely bound EPS (LB-EPS) diffused from the tightly bound EPS (TB-EPS) that surround the cells [4]. A schematic representation of the distribution of LB-EPS and TB-EPS around bacteria was shown in Fig. 1. LB-EPS can be separated from the bacterial culture by speed centrifugation with TB-EPS still attached on the cells [5,6]. Methods of removing TBEPS from the cells include chemical methods such as treatment with EDTA or formaldehyde [6], or physical methods including

∗ Corresponding author. Tel.: +86 27 87671033; fax: +86 27 87280670. E-mail address: [email protected] (P. Cai). 1 These authors contributed equally to this work. http://dx.doi.org/10.1016/j.colsurfb.2015.03.017 0927-7765/© 2015 Elsevier B.V. All rights reserved.

sonication-cationic exchange resin treatment [5], or biological methods such as enzyme treatment [7]. A thorough knowledge of the role of EPS on bacterial surface properties is necessary to understand bacteria interactions with heavy metals, bacteria transport, adhesion and survival in the environments [7–10]. Recently a few studies have been performed to investigate the role of TB-EPS on cell surface properties [7,11,12]. For example, Tourney and Ngwenya [12] found that after Bacillus licheniformis S86 TB-EPS were stripped by cation exchange resin, the total acidic site concentrations of the bacteria decreased by 34.2%. Kim et al. [7] observed that after partial removal of TB-EPS on Escherichia coli O157:H7 using a proteolytic enzyme, the cell surface exhibited more negative electrophoretic mobility at ionic strength (IS) ≤1 mM. Fang et al. [8] demonstrated after removing TB-EPS of Bacillus subtilis and Pseudomonas putida by cationic exchange resin, the deprotonation constants (pKa ) values for the functional groups of the untreated and EPS-free cells were similar, while the total deprotonated site concentrations of untreated cell surfaces were significantly higher than those of EPS-free cell surfaces. Although it has been established the effect of TB-EPS on bacterial surface properties, the impact of LB-EPS has not been well examined. Actually,

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2.2. SEM imaging Intact and LB-EPS free bacteria were imaged by scanning electron microscopy (SEM) to determine the extent of EPS removal. The bacterial samples were fixed in 3% glutaraldehyde for 2 h and freeze-dried (Freezone 6, Labconco Co., USA) for 24 h [19]. The dried cells were then coated under reduced pressure with a thin layer of gold and examined using scanning electron microscopy (JSM6390/LV).

2.3. Cell viability tests

Fig. 1. Distribution of LB-EPS and TB-EPS surrounding bacteria.

TB-EPS and LB-EPS have different functions in bacterial aggregation behaviors. It was reported by Eboigbodin and Biggs [13] that the addition of LB-EPS to E. coli MG1655 resulted in aggregation of the cells. However, the TB-EPS did not contribute to aggregation process. Therefore, it is necessary to clarify the role of LB-EPS in the surface properties of bacteria. This study was designed to investigate the effect of LB-EPS on surface properties of different species of bacteria. To achieve this purpose, high-speed centrifugation was employed to remove the LB-EPS on the outer surface of the bacteria cells. Four types of bacteria were used: B. subtilis (Gram-positive), S. suis (Grampositive), E. coli (Gram-negative) and P. putida (Gram-negative). Surface properties of untreated cells and high-speed treated cells were characterized and compared via Live/Dead BacLight Viability Kits, Fourier transform infrared (FTIR) spectroscopy, potentiometric titration, electrophoretic mobility and hydrophobicity analysis. 2. Materials and methods 2.1. Bacterial cultivation and LB-EPS removal B. subtilis 168, P. putida X4 and E. coli WH09 were cultivated in Luria broth medium (5.0 g L−1 yeast extract, 10.0 g L−1 tryptone and 5.0 g L−1 NaCl) at 28 ◦ C and 180 rpm for 24 h, 18 h and 18 h, respectively. S. suis SC05 strain was grown statically in Tryptone Soy Broth medium (17 g L−1 pancreatic digest of casein, 3 g L−1 papaic digest of soybean, 2.5 g L−1 dextrose, 5 g L−1 sodium chloride and 2.5 g L−1 dipotassium phosphate) at 37 ◦ C for 10 h [14]. Both growth media were sterilized in an autoclave at 121 ◦ C for 30 min before use. In the standard case all cultures were harvested at early stationary phase by centrifugation at 3220 × g for 10 min, and then rinsed three times with ultrapure water to remove residual growth medium. This low centrifugation speed has been utilized as a model procedure to separate intact bacteria from the growth media, and they most closely resembled unmanipulated microorganisms in the cell surface parameters [15,16]. For the removal of LB-EPS, the cell suspension was centrifuged at a higher speed (12,000 × g, 10 min) following the same rinsing protocol [17,18]. Our preliminary experiments showed that if the centrifugation speed is below 10,000 × g, much fewer LBEPS amounts were collected in the supernatants. On the other hand, if the speed was very high (e.g., 20,000 × g), some bacterial cells would die or lyse. Hence, this centrifugation speed was chosen for LB-EPS removal. The intact and LB-EPS free bacteria were then re-suspended in 10 mM KNO3 for subsequent characterization. Total LB-EPS, polysaccharide, and protein contents were assayed by ethanol precipitation, phenol–sulfuric acid, and bicinchoninic acid method (Boisynthesis Co., Ltd., Beijing), respectively [2].

Viability tests were performed to examine the integrity of the cell membrane after exposure to high speed centrifugation, and the tests were conducted in triplicate (unadjusted pH 5.5–5.8). The viability of cells was evaluated using Live/Dead BacLight Bacterial Viability Kits, which was widely used to enumerate viable bacteria. Experimental details have been well described in previous studies [20–22]. Bacterial suspension (∼108 cells mL−1 ) was stained with a dye solution consisting of 40 ␮L Live/Dead BacLight stain (L-7012, Molecular Probes, Eugene, OR) in 2 mL of 10 mM KNO3 for 15 min. Then 100 ␮L of bacterial suspension was observed by fluorescence microscopy (IX-70, Olympus), and 20 images were taken for each cell type. Bacteria with intact cell membranes stain fluorescent green (considered to be viable), whereas bacteria with damaged membranes stain fluorescent red (considered to be dead) [20].

2.4. Fourier transform infrared (FTIR) spectroscopy FTIR spectroscopy has been used to characterize bacterial cell wall chemistry and to identify functional groups of macromolecules on the cell surface [23–25]. FTIR spectra of the intact and LBEPS free bacteria were obtained using a Fourier transform infrared spectrometer (Nicolet AVAR 330, USA). The infrared spectra were recorded over the range of 4000 cm−1 to 400 cm−1 , and each sample was scanned 256 times at a resolution of 4 cm−1 . Sample disks were made from 5 mg of bacterial cells encapsulated in 150 mg of KBr. This ratio resulted in better-resolved IR spectra than the 1:100 typically recommended ratio [26].

2.5. Potentiometric titration Potentiometric titrations were conducted using an automatic potentiometric titrator (Metrohm titrator 836) according to the procedures described by Yee et al. [27]. The washed bacteria were resuspended in 40 mL of 10 mM KNO3 solution at final concentrations of 2.250–5.508 mg dry weight mL−1 . The bacterial suspension was then placed in a sealed titration vessel and purged with N2 gas until the system maintained a constant pH for a period of 10 min. The suspension was initially acidified to approximately pH 2.5 using 100 mM HNO3 and equilibrated for 40 min, then titrated up to pH 10.0 with 100 mM NaOH. At each titration step, a stability of 0.1 mV S−1 was attained before the addition of the next dripping of titrant. Blank titrations were performed using 10 mM KNO3 and each titration was conducted in triplicate. In order to elucidate the effect of LB-EPS on proton-active binding sites on bacteria, a nonelectrostatic approach was used to calculate the acidity constant and the concentration of each site on intact and LB-EPS free bacteria. The deprotonation of a functional group can be represented by the following generic reaction [28]: R − AH0 → R − A− + H+

(1)

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where R is the bacterium to which the functional group type A is attached. The acidity constant Ka for reaction (1) can be expressed as:

 Ka =

R−A



−



aH+

R − AH0



(2)

where [R − A− ] and [R − AH0 ] represent the concentration of deprotonated and protonated sites, respectively, and aH+ represents the activity of protons in the bulk solution. Titration data were modeled using the data optimization program Protofit 2.1 to obtain site concentrations and acidity constants (pKa ) for proton-active functional groups present on the bacterial surface [29].

2.6. Electrophoretic mobility and size distribution Freshly rinsed intact and LB-EPS free bacterial cells were diluted to an absorbance value of 0.200 at 600 nm. Approximately 1.0 mL of this bacterial suspension was transferred to electrophoresis cell, and the electrophoretic mobility measurements were conducted at 25 ◦ C using Zetasizer Nano (NanoZS ZEN3600, Malvern Instruments) [30]. All measurements were determined in triplicate over the ionic strength (IS) range of 0.01–100 mM KNO3 (unadjusted pH 5.5–5.8). Similar experiments were also performed in Ca(NO3 )2 solutions with concentrations of 0.01–100 mM. Size distribution (% number) and hydrodynamic diameters of bacterial cells in 1 mM KNO3 was also measured by Zetasizer Nano and analyzed by the Malvern software.

2.7. Cell aggregation test The washed intact and LB-EPS free bacteria were re-suspended in 1 mM KNO3 solution to an initial optical density of ∼0.6 (C0 ) at a wavelength of 600 nm. Then, the cell suspensions (0.4 mL) were transferred into quartz cuvette and the upper part of the suspension were measured spectroscopically (UV1102, Shanghai Tianmei Scientific Instrument Co., China) every 5 min (C). The measurements lasted for 2 h, and the obtained aggregation kinetic profiles were plotted as relative optical density (C/C0 ) vs. time.

2.8. Hydrophobicity Cell surface hydrophobicity was quantified using the MATH test, following a procedure described by Pembrey et al. [15] and Walker et al. [31]. Briefly, 1 mL of n-dodecane (laboratory grade) was added to 4-mL samples of a cell suspension (optical density of 0.200–0.210 at 600 nm) in 15 mL test tube. The test tube was vortexed at full speed for 2 min and then left to stand for 15 min to allow phase separation. The hydrophobicity value was expressed as the percentage of cells partitioned into the hydrocarbon phase. The average cell hydrophobicity was calculated using triplicate samples in 10 mM KNO3 .

Table 1 Infrared absorption band assignments and types of bacterial surface functional groups. Wave number (cm−1 )

Absorption band assignment

Functional group type

1660–1637

Stretching vibration of C O and C N (Amide I) Stretching vibration of C N and deformation vibration of N H (Amide II) CH2 bending vibrations Symmetric stretching vibration of C O Stretching vibration of PO2 − Deformation vibration of C O Vibration of C OH, C O C and C C Asymmetric stretching vibration of P O

Proteins (peptidic bond) [3]

1550–1540

1482–1454 1420–1400 1260–1220

1150–950

Proteins (peptidic bond) [3]

Proteins and lipids [10] Carboxylic groups [10] Phosphodiesters phosphate groups Carboxylic groups [32] Polysaccharides Phosphodiesters [33]

Note: The references for each functional group type were [3,10,32,33], respectively.

3. Results and discussion 3.1. Effectiveness of high-speed centrifugation treatment Table S1 displays the amounts of LB-EPS, polysaccharides and proteins removed from the bacterial surfaces after high-speed centrifugation. The four bacterial strains had roughly the same order of magnitude of LB-EPS contents ranging from 18.9 ␮g/108 cells (P. putida) to 37.0 ␮g/108 cells (B. subtilis). The LB-EPS of B. subtilis, E. coli, and P. putida mainly consisted of polysaccharides (10.0–16.2 ␮g/108 cells), with fewer proteins (2.9–9.2 ␮g/108 cells) in LB-EPS. On the contrary, S. suis exhibited much larger content of protein (21.9 ␮g/108 cells) than polysaccharide (3.2 ␮g/108 cells). The SEM images of the intact and LB-EPS free bacteria samples were shown in Fig. 2. Generally, intact bacterial cell surfaces (especially Gram-positive bacteria, B. subtilis and S. suis) had more LB-EPS molecules. For example, the intact B. subtilis cells were cross-linked by fibrous materials with long chains, forming a polymeric network (Fig. 2A). Slimy substances connecting the ovoid-shaped cells were observed on intact S. suis surfaces (Fig. 2C). After treatment with high-speed centrifugation, most of the surrounding LB-EPS disappeared and the bacterial surfaces became much smoother (Fig. 2B, D, F and H). The possible influence of high speed centrifugation on cell membrane integrity can be detected by fluorescence microscopy with LIVE/DEAD staining of the biomass. Fig. S1 shows that equal percentages of cells (95.9–98.0%) were viable between intact and LB-EPS free bacteria (P > 0.05), suggesting that the removal of LBEPS by high-speed centrifugation had no significant impact on cell membrane integrity. Therefore, the component analysis of LB-EPS, SEM images and viability test results clearly demonstrated that the high-speed centrifugation treatment can effectively remove the LBEPS material from bacteria, and did not destroy the cell membrane integrity. 3.2. FTIR spectrum

2.9. Statistical analysis In tables and figures, the data are presented as the mean ± SEM (standard error of the mean). Statistical differences between mean values were analyzed using a Student’s t test in SPSS 17.0 software (SPSS Inc., Chicago, USA). When P < 0.05, the differences are suggested to be statistically significant.

Fig. 3 shows FTIR spectrum for the intact and LB-EPS free bacteria, and the main sharp stretching frequencies and the corresponding biological molecules are summarized in Table 1. The peak assignments of both intact and LB-EPS free bacteria were as follows [3,10,32,33]: near 1652 cm−1 (C O and C N stretching in Amide I); 1543 cm−1 (C N stretching vibration and N H

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Fig. 2. Scanning electron microscope (SEM) images for (A) intact B. subtilis, (B) LB-EPS free B. subtilis, (C) intact S. suis, (D) LB-EPS free S. suis, (E) intact E. coli, (F) LB-EPS free E. coli, (G) intact P. putida, and (H) LB-EPS free P. putida.

deformation vibration of Amide II); 1462 cm−1 (CH2 bending vibrations in proteins and lipids); 1396 cm−1 (C O symmetric stretching of carboxylic groups); near 1231 cm−1 (PO2 − stretching from phosphodiesters and phosphate groups); a broad peak near 1077 cm−1 (C OH, C O C and C C vibrations in polysaccharides). The FTIR spectra revealed the presence of phosphodiester, carboxylic, phosphate, and amino functional groups on bacterial surfaces. Cell surface macromolecules (include LB-EPS) are composed of a combination of lipid, protein, and carbohydrate compounds. No obvious

difference in the peak positions was found between the spectra of intact and LB-EPS free cells. It implied that the overall composition of LB-EPS outside the cell surfaces may be comparable to that of the intact cell walls [10]. LB-EPS has no effects on the type of the bacterial surface functional groups. This finding was consistent with previous reports that the FTIR spectra of untreated and TB-EPS free B. subtilis or P. putida cells were similar [8,34]. The FTIR experiments of LB-EPS removed from the cell surfaces were also performed. As indicated in Fig. S2, the FTIR spectra of LB-EPS removed from the

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A

1543

1462 1396

1800

1600

1400

1231

1077

1200

1000

1542 1458 1386

1800

-1

D

Absorbance

Absorbance 1800

1542

1600

1400

1200

1200

1000

Intact P. putida LB-EPS free P. putida

1653

1652

1239

1400

1076

Wavenumber (cm )

Intact E. coli LB-EPS free E. coli

1455 1395

1600

1237

-1

Wavenumber (cm )

C

Intact S. suis LB-EPS free S. suis

1652

Absorbance

1652

Absorbance

B

Intact B. subtilis LB-EPS free B. subtilis

1082

1000

-1

Wavenumber (cm )

1800

1543

1456 1393

1600

1400

1238

1200

1083

1000

-1

Wavenumber (cm )

Fig. 3. FTIR spectra of intact and LB-EPS free (A) B. subtilis, (B) S. suis, (C) E. coli, and (D) P. putida. The IR spectra were collected over the range of 1800 cm−1 to 1000 cm−1 with a resolution of 4 cm−1 .

cell surfaces displayed similar peak positions as those of intact/LBEPS free bacteria, with only minor shift of these absorption peaks (<15 cm−1 ). This result further confirmed that the function group types in LB-EPS and cell walls were similar. 3.3. Potentiometric titration Potentiometric titrations can determine the acidity constants and site concentrations of functional groups on bacterial cell walls [35]. As shown in Fig. S3, all samples exhibited substantial buffering capacity over a wide pH range. The shapes of the titration curves for the intact and LB-EPS free cells were comparable, suggesting that the types of functional groups were unchanging between the intact and LB-EPS free cells. The pKa and site concentration values were compiled in Table 2, in which a 4-site model (discrete deprotonation reaction associated with each site type) provided the best fit to the experimental data for all bacterial samples. The pKa values between 2 and 6 are ascribed to carboxyl groups, around 3.0 to phosphodiester groups, 5.6–7.2 to phosphoryl groups and approximately 10 to amine or hydroxyl groups [12,36–38]. As presented in Table 2, the four distinct types of organic acid functional groups that are displayed on intact and LB-EPS free bacterial surfaces likely corresponded to phosphodiester (site 1), carboxyl (site 2), phosphate (site 3), and amine or hydroxyl sites (site 4), respectively. Tourney et al. [12] also reported a 4-site model with phosphodiester (pKa 3.3–3.4), carboxylic (pKa 5.3–5.4), phosphoryl (pKa 7.4–7.5) and hydroxyl/amine (pKa 9.9–10.1) type groups for both intact and TB-EPS free B. licheniformis S-86 cell. Most of the pKa values were similar for intact and TB-EPS free bacteria (P > 0.05), indicating almost the same proton-active functional groups were present on the LB-EPS fraction as on the bulk cell wall. FTIR spectra in Fig. 3 further confirmed this result that no obvious difference in the absorption peaks was observed between the intact and LB-EPS free cells. However, the pKa values of phosphodiester groups on intact S. suis (2.44) and E. coli (2.57) were found to be

significantly (P < 0.05) different from those on LB-EPS free S. suis (3.22) and E. coli (3.35). It revealed that high-speed centrifugation could alter the acidity constants, and the phosphodiester groups on S. suis/E. coli LB-EPS had different proton dissociation properties from the cell walls. Intact cell surfaces have a greater number of dissociable groups than the LB-EPS free cells (Table 2). The total site concentrations on intact B. subtilis, S. suis, E. coli and P. putida surface were 3.72 × 10−3 mol g−1 , 2.70 × 10−3 mol g−1 , 2.63 × 10−3 mol g−1 and 1.91 × 10−3 mol g−1 , respectively. After LB-EPS removed by high-speed centrifugation, the total site concentrations of bacteria decreased remarkably by 22.6%, 37.0%, 36.5% and 29.8%, respectively, for B. subtilis, S. suis, E. coli and P. putida. The majority of site concentrations for each functional group (sites 1–4) on LBEPS free bacteria were also fewer than intact cells. These results are attributed to the presence of a number of negatively charged functional groups on the LB-EPS molecules [2,39]. Nevertheless, decreasing trends of site concentrations were not always observed for all the four groups after high-speed centrifugation. For instance, the concentration of amine/hydroxyl group (0.58 × 10−3 mol g−1 ) on LB-EPS free S. suis were higher than that on intact S. suis (0.49 × 10−3 mol g−1 ). LBS-EPS free P. putida also possessed larger phosphate group concentration (increased by 0.05 × 10−3 mol g−1 ) than intact P. putida. Two possible mechanisms may be involved in this phenomenon. First, the macromolecules (e.g., polysaccharides and proteins) on cell surfaces possessed complex conformational structures. The OH− may not be able to interact with some of the acidic groups inside the polymeric layers on intact cell surfaces. However, after the removal of LB-EPS, the polymer chains and steric structures of macromolecules were disrupted. Some more functional groups may be exposed, leading to the increase of surface site concentrations. Second, the short titration time (1.5 h) may not be sufficient to allow all the acidic groups participating in the potentiometric titration tests. It may take a longer time for all the protons inside the surface macromolecules of intact cells to dissociate from the functional groups and transfer into the solution. Therefore, some site concentrations increased with the treatment

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Table 2 Acidity constants (pKa ) and site concentrations [site] of surface functional groups for the intact and LB-EPS free bacteria. Bacterium

pKa 1

pKa 2

pKa 3

pKa 4

[Site 1]a

[Site 2]

[Site 3]

[Site 4]

Total [site]

Intact B. subtilis LB-EPS free B. subtilis Intact S. suis LB-EPS free S. suis Intact E. coli LB-EPS free E. coli Intact P. putida LB-EPS free P. putida

2.88 ± 0.21 3.08 ± 0.41 2.44 ± 0.01 3.32 ± 0.04 2.57 ± 0.01 3.35 ± 0.05 3.43 ± 0.01 3.30 ± 0.07

4.74 ± 0.06 4.46 ± 0.64 4.73 ± 0.33 5.47 ± 0.09 5.71 ± 0.02 5.76 ± 0.04 4.98 ± 0.02 4.50 ± 0.01

6.81 ± 0.01 6.97 ± 0.30 7.06 ± 0.57 7.40 ± 0.07 7.18 ± 0.01 7.22 ± 0.11 7.27 ± 0.11 7.28 ± 0.04

9.18 ± 0.04 9.44 ± 0.04 9.30 ± 0.30 9.41 ± 0.04 9.21 ± 0.01 9.46 ± 0.29 9.64 ± 0.02 9.13 ± 0.04

0.89 ± 0.19 0.65 ± 0.13 1.29 ± 0.01 0.26 ± 0.02 1.29 ± 0.27 0.46 ± 0.01 0.33 ± 0.01 0.28 ± 0.01

0.72 ± 0.03 0.53 ± 0.14 0.55 ± 0.15 0.60 ± 0.07 0.51 ± 0.01 0.40 ± 0.01 0.46 ± 0.01 0.35 ± 0.01

0.70 ± 0.01 0.52 ± 0.09 0.37 ± 0.06 0.26 ± 0.01 0.28 ± 0.02 0.26 ± 0.01 0.22 ± 0.01 0.27 ± 0.01

1.41 ± 0.09 1.17 ± 0.01 0.49 ± 0.13 0.58 ± 0.01 0.56 ± 0.01 0.55 ± 0.18 0.90 ± 0.01 0.44 ± 0.01

3.72 ± 0.14 2.88 ± 0.11 2.70 ± 0.10 1.70 ± 0.08 2.63 ± 0.29 1.67 ± 0.19 1.91 ± 0.01 1.34 ± 0.01

a Site concentrations in 10−3 mol g−1 normalized to dry mass of bacteria. [Site 1], [site 2], [site 3], and [site 4] stand for the concentrations of phosphodiester, carboxyl, phosphate, and amine/hydroxyl sites, respectively.

of LB-EPS removal. A recent study also found that the removal of TB-EPS eventually increased the concentrations of four groups on B. subtilis [34]. This contradiction (increase or decrease of surface groups after LB-EPS removal) possibly resulted from the different cell wall structures of the four bacterial strains. 3.4. Electrophoretic mobility The electrophoretic mobilities (EM) of bacteria cells measured in varying KNO3 and Ca(NO3 )2 solutions are shown in Fig. 4 and Fig. S4, respectively. The absolute EM values of the four bacteria decreased with the increase of electrolyte concentrations from 0.01 mM to 100 mM, which was ascribed to the electrostatic interactions of the electrolyte ions with the ionized surface functional groups (charge screening) and double layer compression outside the cell surfaces [40,41]. The EM values were determined to be virtually identical (P > 0.05) for all the intact and LB-EPS free cells, indicating LB-EPS had no significant influence on cell surface charges regardless of solution IS and ionic valence. It also suggested that LB-EPS may not be able to significantly affect the electrostatic interaction between bacteria and solid particle. EM is a reflection of the electrical potential existing at the electrokinetic shear plane, which is located in close proximity to the outermembrane

surface [42]. Titration data showed that the acidic functional groups (negatively charged) on cell surfaces were removed or exposed after LB-EPS removal, and the corresponding local surface charge densities were supposed to be disturbed. However, all the EM values remained statistically unchanged across the IS range (P > 0.05). This apparent discrepancy in cell charge results can be explained by the differences in measurement techniques. Titration captures the density of most dissociable functional groups on the membrane surface, cell wall and within the polymeric macromolecule layers [43]. Dissimilarly, EM is a macroscopic parameter that represents the net or average electrokinetic properties of the cell, which is not sensitive to small local-scale charge variations on the cell surface [31]. This parameter does not reflect the subtle differences in charge heterogeneity on the surfaces of intact and LB-EPS free cells. Therefore, the decrease in total site concentrations may not affect the net surface charges. Investigations on the bacterial surface charges induced by EPS removal have been previously reported, but different effects were observed. For example, Long et al. [44] found that the removal of TB-EPS from cell surfaces via CER treatment did not have large influence on the zeta potentials of E. coli BL21, Pseudomonas sp. QG6, Rhodococcus sp. QL2, and B. subtilis. On the contrary, Kim et al. [7] reported the EM valuess of E. coli O157:H7 became more negative after removing TB-EPS at

Fig. 4. Electrophoretic mobility (EM) of intact and LB-EPS free (A) B. subtilis, (B) S. suis, (C) E. coli, and (D) P. putida as a function of KNO3 concentration (0.01–100 mM). Error bars are the standard deviation of three replicates.

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Hydrophobicity (%)

100

Intact bacteria LB-EPS free bacteria

80 60 40 20 0

B. subtilis

S. suis

E. coli

P. putida

Fig. 5. Hydrophobicity (%) of intact and LB-EPS free bacteria in 10 mM KNO3 . Each experiment was conducted in triplicate.

IS 0.01–100 mM. This contradiction possibly resulted from the distinct components of charged macromolecules on different bacterial surfaces. 3.5. Size distribution and cell aggregation As indicated in Fig. S5, the size distribution showed little difference between intact and LB-EPS cells. The hydrodynamic diameters were 0.95–0.96 ␮m, 0.82–0.85 ␮m, 0.90–0.91 ␮m, and 1.25–1.28 ␮m for intact/LB-EPS free B. subtilis, S. suis, E. coli, and P. putida, respectively (Table S2). The treatment of high-speed centrifugation had no significant (P > 0.05) influences on bacterial size distribution and diameters. Cell aggregation kinetics of intact and LB-EPS free bacteria are presented in Fig. S6. As the time increased, more and more cellcell aggregates formed and settled out of the suspension, which led to the reduction of the optical density of bacterial suspension (C). The C/C0 values of B. subtilis, E. coli, and P. putida were significantly (P < 0.05) higher after LB-EPS removal, indicating the intact cells aggregated faster than LB-EPS free cells. It suggests that LBEPS on bacterial surfaces could enhance the cell-cell interactions. However, the electrophoretic mobilities of bacteria showed no significant (P > 0.05) differences between intact and LB-ESP free cells (Fig. 4), which revealed that the electrostatic repulsions among cells were almost the same. This phenomenon suggested that nonelectrostatic force (e.g., polymer interaction) was probably involved in the aggregation processes of intact bacteria. Dissimilarly, the C/C0 values of S. suis were statistically unchanged (P > 0.05) over the range of time (0–120 min). The LB-ESP on S. suis surface played negligible role in the cell–cell aggregation behaviors. 3.6. Hydrophobicity The measurement of hydrophobicity allows for further comparison between the intact and LB-EPS free cells. Fig. 5 presents the percent hydrophobicity of intact and LB-EPS free cells. The hydrophobicities of the intact B. subtilis, S. suis, E. coli and P. putida were 48.7%, 47.4%, 51.9% and 69.3%, respectively, confirming that the cells were consistently hydrophobic. After the LB-EPS removal, the hydrophobicity of the four bacteria decreased to 26.1–65.0%. The range of hydrophobicity values determined in this study was comparable to that of other bacterial strains (0.4–74.1%) [43,45]. As described by Walker et al. [31], a greater amount of polar and acidic molecules (hydrophilic) exposed on the cell surfaces would result in a lower percentage of cells partitioning into the hydrocarbon. By subtracting the total site concentrations of LB-EPS free bacteria from those of intact bacteria (Table 1), the calculated acidic site concentrations on LB-EPS ranged from 0.57 × 10−3 mol g−1 to 1.00 × 10−3 mol g−1 , which were fewer than those of LB-EPS free

bacteria (1.34–2.88 × 10−3 mol g−1 ). The LB-EPS of the four bacterial types probably contained less density of charged polar groups, which led to the higher hydrophobicities of LB-EPS constituents than the LB-EPS free bacteria. It was reported that the remaining macromolecules on cell surface have higher charges per unit volume than the enzyme cleaved macromolecules in extracellular materials [7]. In addition, the hydrophobicities of intact/LB-EPS free B. subtilis and S. suis (26.1–48.7%) were smaller than E. coli and P. putida (39.4–69.3%), which corresponded to the larger amount of acidic polar groups on both B. subtilis and S. suis surfaces (1.70–3.72 × 10−3 mol g−1 ). On the other hand, FTIR spectra verified that broad peaks were observed for the LB-EPS in the supernatants at 1077–1083 cm−1 (Fig. S2), which exhibited the character of carbohydrates or carbohydrates-like substances [46]. It indicated the uncharged hydrophobic carbohydrates were present in the LB-EPS [47]. Furthermore, by analyzing the components of LB-EPS, larger amounts of polysaccharides (10.0–16.2 ␮g/108 cells) were found than proteins (2.9–9.2 ␮g/108 cells) for B. subtilis, E. coli, and P. putida. Hence, the decrease in the hydrophobicities of LB-EPS free cells can be attributed to the removal of a great number of nonpolar/hydrophobic components (e.g., C (C,H)) in LB-EPS. Such trend was also found in the literature that the hydrophobicity of Leucobacter sp. decreased by 20–50% after the extracellular macromolecules degraded by enzymes [48]. It is interesting to note that after the treatment of high-speed centrifugation, the decreasing hydrophobicity percentages of B. subtilis and S. suis (38.6–46.4%) were greater than those of E. coli and P. putida (6.2–24.1%). This result revealed that the LB-EPS in the two Gram-positive bacterial types were more hydrophobic relative to the Gram-negative cells. Besides, it should be mentioned that the higher hydrophobicity values of intact cells may increase the hydrophobic effect among cell–cell interactions, resulting in the faster aggregation speed. 4. Conclusions The high speed centrifugation treatment can effectively remove the LB-EPS from the bacteria surface, without destroying the cell membrane integrity. Surface characterization of intact and LB-EPS free bacteria indicated that LB-EPS had different influences on the cell surface properties. Specifically, the removal of LB-EPS led to a significant decrease in the total concentrations of acidic functional groups on the four bacteria, although a few groups (e.g., phosphate and amine/hydroxyl groups) slightly increased. Higher surface hydrophobicities were found for intact bacteria, suggesting LB-ESP on cell surfaces could improve their adhesion to soil/sediment particles via the larger hydrophobic interactions. LB-ESP removal had no effect on the bacterial electrophoretic mobilities, size distribution, hydrodynamic diameters as well as the functional group types on cell surfaces. Electrostatic forces originated from LB-EPS may have negligible effects on bacterial adhesion and transport behaviors in soil-water environment. Cells having more LB-EPS may tend to aggregate faster in aquatic environment, which inhibit their transport in small soil pores. This study provides detailed insight into the variation of surface properties of different intact/LBEPS free bacterial types via molecular-level and macroscopic-level characterization techniques, which is of fundamental importance to better predict the potential impact of LB-EPS on bacterial fate (e.g., aggregation, adhesion and transport) in aquatic environments. Acknowledgements This research was supported by the National Natural Science Foundation of China (41171196), Foundation for the Author of National Excellent Doctoral Dissertation of China (201066),

W. Zhao et al. / Colloids and Surfaces B: Biointerfaces 128 (2015) 600–607

Program for New Century Excellent Talents (NCET-11-0645), and the Fundamental Research Funds for the Central Universities (2011PY005). 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. colsurfb.2015.03.017. References [1] J. Wingender, T.R. Neu, H.C. Flemming, Microbial Extracellular Polymeric Substances: Characterization, Structure, and Function, Springer, Berlin, 1999. [2] Y.Y. Cao, X. Wei, P. Cai, Q.Y. Huang, X.M. Rong, W. Liang, Colloids Surf., B: Biointerfaces 83 (2011) 122. [3] A. Omoike, J. Chorover, Geochim. Cosmochim. Acta 70 (2006) 827. [4] X.Y. Li, S.F. Yang, Water Res. 41 (2007) 1022. [5] S. Comte, G. Guibaud, M. Baudu, Process Biochem. 41 (2006) 815. [6] X. Zhang, P.L. Bishop, B.K. Kinkle, Water Sci. Technol. 39 (1999) 211. [7] H.N. Kim, Y. Hong, I. Lee, S.A. Bradford, S.L. Walker, Biomacromolecules 10 (2009) 2556. [8] L.C. Fang, X. Wei, P. Cai, Q.Y. Huang, H. Chen, W. Liang, X.M. Rong, Bioresour. Technol. 102 (2011) 1137. [9] H.N. Kim, S.A. Bradford, S.L. Walker, Environ. Sci. Technol. 43 (2009) 4340. [10] M. Ueshima, B.R. Ginn, E.A. Haack, J.E.S. Szymanowski, J.B. Fein, Geochim. Cosmochim. Acta 72 (2008) 5885. [11] J. Tourney, B.T. Ngwenya, J. Colloid Interface Sci. 348 (2010) 348. [12] J. Tourney, B.T. Ngwenya, J.W.F. Mosselmans, L. Tetley, G.L. Cowie, Chem. Geol. 247 (2008) 1. [13] K.E. Eboigbodin, C.A. Biggs, Biomacromolecules 9 (2008) 686. [14] W.Q. Zhao, X. Liu, Q.Y. Huang, X.M. Rong, W. Liang, K. Dai, P. Cai, Eur. J. Soil Sci. 63 (2012) 558. [15] R.S. Pembrey, K.C. Marshall, R.P. Schneider, Appl. Environ. Microbiol. 65 (1999) 2877. [16] S.S. Tazehkand, S. Torkzaban, S.A. Bradford, S.L. Walker, J. Environ. Qual. 37 (2008) 2108. [17] G.P. Sheng, H.Q. Yu, X.Y. Li, Biotechnol. Adv. 28 (2010) 882. [18] A. Ramesh, D. Lee, J. Lai, Appl. Microbiol. Biotechnol. 74 (2007) 699.

607

[19] A. Rochex, D. Lecouturier, I. Pezron, J.M. Lebeault, Appl. Microbiol. Biotechnol. 65 (2004) 727. [20] L. Boulos, M. Prevost, B. Barbeau, J. Coallier, R. Desjardins, J. Microbiol. Methods 37 (1999) 77. [21] B. Asadishad, S. Ghoshal, N. Tufenkji, Environ. Sci. Technol. 45 (2011) 8345. [22] P. Cai, Q.Y. Huang, S.L. Walker, Environ. Sci. Technol. 47 (2013) 1896. [23] H.M. Al-Qadiri, M. Lin, A.G. Cavinato, B.A. Rasco, Int. J. Food Microbiol. 111 (2006) 73. [24] K.E. Eboigbodin, J.J. Ojeda, C.A. Biggs, Langmuir 23 (2007) 6691. [25] M. Kansiz, P. Heraud, B. Wood, F. Burden, J. Beardall, D. McNaughton, Phytochemistry 52 (1999) 407. [26] L.R. Drake, S. Lin, G.D. Rayson, P.J. Jackson, Environ. Sci. Technol. 30 (1995) 110. [27] N. Yee, L.G. Benning, V.R. Phoenix, F.G. Ferris, Environ. Sci. Technol. 38 (2004) 775. [28] D. Borrok, J.B. Fein, Geochim. Cosmochim. Acta 68 (2004) 3043. [29] B.F. Turner, J.B. Fein, Comput. Geosci. 32 (2006) 1344. [30] P. Hiemenz, R. Rajagopalan, Electrophoresis and other electrokinetic phenomena, in: Principles of Colloid and Surface Chemistry, Dekker, New York, NY, 1977. [31] S.L. Walker, J.A. Redman, M. Elimelech, Environ. Sci. Technol. 39 (2005) 6405. [32] G. Guibaud, S. Comte, F. Bordas, S. Dupuy, M. Baudu, Chemosphere 59 (2005) 629. [33] X. Gao, J. Chorover, Colloids Surf., B: Biointerfaces 71 (2009) 169. [34] Z.N. Hong, W.L. Chen, X.M. Rong, P. Cai, K. Dai, Q.Y. Huang, Chem. Geol. 360 (2013) 118. [35] J.B. Fein, J.F. Boily, N. Yee, D. Gorman-Lewis, B.F. Turner, Geochim. Cosmochim. Acta 69 (2005) 1123. [36] A. Barkleit, H. Moll, G. Bernhard, Dalton Trans. 21 (2008) 2879. [37] B.T. Ngwenya, I.W. Sutherland, L. Kennedy, Appl. Geochem. 18 (2003) 527. [38] D. Borrok, B.F. Turner, J.B. Fein, Am. J. Sci. 305 (2005) 826. [39] A. Omoike, J. Chorover, Biomacromolecules 5 (2004) 1219. [40] N. Yee, D.A. Fowle, F.G. Ferris, Geochim. Cosmochim. Acta 68 (2004) 3657. [41] W.Q. Zhao, S.L. Walker, Q.Y. Huang, P. Cai, Water Res. 53 (2014) 35. [42] A.T. Poortinga, R. Bos, H.J. Busscher, Colloids Surf., B: Biointerfaces 20 (2001) 105. [43] B.Z. Haznedaroglu, H.N. Kim, S.A. Bradford, S.L. Walker, Environ. Sci. Technol. 43 (2009) 1838. [44] G. Long, P. Zhu, Y. Shen, M. Tong, Environ. Sci. Technol. 43 (2009) 2308. [45] J. Li, L. McLandsborough, Int. J. Food Microbiol. 53 (1999) 185. [46] J.P. Croué, M. Benedetti, D. Violleau, J. Leenheer, Environ. Sci. Technol. 37 (2003) 328. [47] Z. Wang, Z. Wu, S. Tang, Water Res. 43 (2009) 2504. [48] B. Xie, J. Gu, J. Lu, J. Environ. Sci. 22 (2010) 1840.