Multiple Roles of Extracellular Polymeric ... - ACS Publications

1 downloads 8 Views 1MB Size Report
Nov 20, 2012 - and Youngwoo Seo. †,‡,*. †. Department of Civil ... industrial sectors and public services.1,2 Known as the major constituent of biofilms ... biodegradable organic matter substantially increased the free chlorine demand to ...

Article pubs.acs.org/est

Multiple Roles of Extracellular Polymeric Substances on Resistance of Biofilm and Detached Clusters Zheng Xue,† Varun Raj Sendamangalam,‡ Cyndee L. Gruden,† and Youngwoo Seo†,‡,* †

Department of Civil Engineering, University of Toledo, 3006 Nitschke Hall, Mail Stop, 307, 2801 W. Bancroft Street, Toledo, Ohio Department of Chemical and Environmental Engineering, University of Toledo, Toledo, Ohio



S Supporting Information *

ABSTRACT: In this study, multiple roles of biofilm EPS were assessed with respect to the resistance of biofilm and detached biofilm clusters to chlorine disinfection. Strains from an opportunistic pathogen, Pseudomonas aeruginosa (wild type, EPSand EPS+) with altered extracellular polymeric substances (EPS) secretion capabilities were tested. The impact of biofilm EPS quantity on disinfection was evaluated by monitoring biofilm viability, biofilm structure, removal of dissolved organic matter (DOM), and viability of detached biofilm simultaneously during chlorine disinfection. The obtained results suggested that the presence of EPS increased biofilm and detached biofilm resistance to chlorine in both presence and absence of DOM. The quantity of EPS had an effect on biofilm structure and the structural characteristics were closely related to both overall biofilm viability and the spatial distribution of viable cells within the biofilm. Additionally, the increased amount of EPS influenced selective removal of DOM with polar functional groups. However the DOM removal did not have a significant impact on the viability of biofilm cells during chlorine disinfection. Meanwhile, the viability of detached biofilm clusters, particularly the EPS overproducing strain, was significantly increased in the presence of DOM. The combined results suggested that biofilm EPS played multiple roles toward influencing the resistance of both biofilm and detached biofilm to disinfectant. biofilm life cycle.9 Biofilm detachment returns sessile cells to the bulk solution. These detached cells are redistributed and can colonize new surfaces, adversely influencing water quality and increasing public health risks. Currently, there is no robust study addressing the synergistic effects of EPS on detached biofilm clusters. In drinking water distribution systems, facilitated by the large surface area of the pipeline and the production of EPS, at least 95% of the total bacterial biomass in water distribution systems is found as biofilm.10 Therefore, studying the efficiency of disinfectant residuals is of great importance to control biofilm growth.11 Previous studies monitoring biofilm resistance to disinfection mainly focused on the viable cell reduction in total,4,12 without correlating biofilm structural properties to the spatial distribution of viable cells within biofilms. On the other hand, studies that have examined the influence of DOM on biofilm focused primarily on the physical sorption and biodegradation of DOM by biofilm13 without considering the interaction between adsorbed DOM and disinfectant residuals as well as the impact of this interaction on the growth and

1. INTRODUCTION Biofilm formation has been an issue of great concern for many industrial sectors and public services.1,2 Known as the major constituent of biofilms, extracellular polymeric substances (EPS) has many roles including enhancing bacterial adhesion, promoting structural development of biofilm, providing a protective barrier, as well as adsorbing and storing nutrients for biofilm growth. Among these roles, the protective role of EPS has been reported to significantly increase biofilm tolerance to antibiotics and disinfectants.3,4 Mechanisms of EPS protection previously reported include transport limitation of biocides through the EPS matrix, sacrificial reaction of EPS with biocides, and formation of a nutrient gradient within the biofilm resulting in complex phenotypes of cells.5 However, EPS may affect other biofilm characteristics which may also influence its resistance to disinfectants. Specifically, the biofilm structure mediated by EPS production will consequently impact the interaction between biofilms and the fluid environment.2 In addition, EPS provide adsorption sites for dissolved organic matter (DOM) as a result of their composition and influence on biofilm structure.6,7 This DOM, which is not sufficiently removed during conventional treatment processes,8 can exert a disinfectant demand. Beyond their significant influence on the resistance to disinfection, EPS may affect fate and redistribution of detached biofilm clusters, which is an essential component in © 2012 American Chemical Society

Received: Revised: Accepted: Published: 13212

July 31, 2012 November 19, 2012 November 20, 2012 November 20, 2012 dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

mL/min) at room temperature (22 ± 2 °C) (SI Figure S2).28 Two carboys were used as nutrient media and chlorine supply reservoirs, respectively. All feeds were delivered using a multichannel peristaltic pump (ISMATEC, Glattbrugg, Switzerland). Flow cells, tubing and solutions were sterilized prior to each experiment. The flow cell channels were aseptically inoculated with bacterial suspension and incubated 2 h without flow for initial bacterial attachment. Then, flow was introduced and gradually increased to 0.2 mL/min. Biofilm was grown six days to reach structural maturity, which was defined as a steady state in detached cell counts in the biofilm effluent.29 For each flowcell, one channel was used as a control, while disinfectant was applied to the other. Chlorine concentration was maintained at 0.5 mg/L at the flow cell inlet throughout the disinfection process. For DOM experiments, media containing DOM was mixed with chlorine in a bubble trap (HRT = 25 min) before entering the flow cells. Chlorine feed concentration was adjusted to compensate for the rapid chlorine demand exerted by DOM (20% more chlorine demand than no DOM condition) at the point of mixing,30 so inlet chlorine concentration could be maintained at 0.5 mg/L. Flowcell effluent was collected every 30 min for 2 h during the disinfection process and quenched with 0.1 M sodium thiosulfate before further analysis. The total disinfection time was chosen based on the chlorine decay kinetic with biomass in water distribution systems.31 For each bacterial strain, the experiment was repeated more than three times (min: 3 times; max: 5 times). 2.4. Confocal Laser Scanning Microscopy and Image Analysis. The biofilm content on glass slides was discriminated using BacLight LIVE/DEAD staining kit (Molecular Probes Inc.) to differentiate live and dead cells. Extracellular polysaccharides content in the biofilm were visualized by applying Alexa 633 conjugated concanavalin A (ConA-Alexa 633). This stain specifically targets the polysaccharides (Dglucose and D-mannose residues) that are major EPS components of P. aeruginosa.32 A mixture of stains was prepared in CDF buffer with a final concentration of 2.5 μM each for SYTO 9 and propidium iodide (PI), as well as 200 μg/ mL of ConA.33 The mixture was then injected into flow channels and incubated in the dark for 15 min. Live SYTO 9stained cells, dead PI-stained cells, and ConA-Alexa 633 stained EPS were visualized with a Leica TCS SP5 confocal laser scanning microscope (CLSM) equipped with a 63× oil immersed objective and a 20× dry objective. For each biofilm sample slide, at least five positions were randomly selected for image acquisition and further image analysis. CLSM images were further processed using the image processing program COMSTAT to determine total biomass, alginate EPS content, and biofilm structural parameters, as defined in detail elsewhere.34 A fixed threshold and connected volume filtration were used for all image processing. Important structural parameters discussed in this study include (i) the roughness coefficienta measure of how the thickness of biofilm varies; (ii) surface area to volume ratiorepresenting the spatial complexity of biofilm structure; and (iii) diffusion distancedefined as the shortest distance from a pixel containing biomass to a pixel not containing biomass. Average diffusion distance is from the average of all pixels containing biomass. This parameter indicates the extent of void spaces in the biofilm structure. EPS content per unit area was generated from image analysis to evaluate the role of EPS on biofilm structure and biofilm resistance to disinfectant. To investigate

susceptibility of biofilm. Butterfield et al. (2002) measured organic carbon removal rate by biofilm along with biofilm growth rate and yield, suggesting that adsorbed carbon compounds may have protective role against chlorine disinfection besides providing nutrients for biofilm growth.14 Ndiongue et al. (2005) also reported that the presence of biodegradable organic matter substantially increased the free chlorine demand to control biofilm formation.15 While most previous studies have focused on biofilm disinfection and disinfectant penetration in biofilm,16 susceptibility of detached biofilm against disinfectants remains elusive. Previous research investigating biofilm detachment has quantified the frequency of detachment and size of cell clusters;17 however, resistance of detached biofilm clusters has not been explored considering competitive disinfectant consumption exerted by biofilm components and other disinfectant demanding substances under continuous flow conditions.9,18 Accordingly, examining combined roles of EPS on resistance of both biofilm and detached biofilm clusters can provide important information regarding minimal disinfectant residuals to control biofilm and prevent bacterial redistribution in water distribution systems. To address the current knowledge gap, this study assessed the multiple roles of biofilm EPS with respect to biofilm and detached biofilm resistance to a common disinfectant (chlorine). Specifically, this study investigated (i) the effect of biofilm EPS on biofilm structure, disinfectant transport and DOM removal; (ii) the role of biofilm structure on the spatial distribution of viable cells in biofilm; and (iii) the influence of DOM on susceptibility of biofilm and detached biofilm clusters during chlorine disinfection.

2. MATERIALS AND METHODS 2.1. Bacteria Culture. P. aeruginosa is an opportunistic pathogen which has been identified in water distribution systems.19−21 The EPS of P. aeruginosa is primarily composed of polysaccharides including alginate, which are considered as the major component of biofilm EPS.22 Wild-type PAO1 (WT) and two mutant strains, alginate EPS production inhibited algT(U) (EPS-) and alginate EPS overproduced mucA22 (EPS +), were selected to construct confluent biofilms (Supporting Information (SI) Figure S1). All strains were grown in onetenth strength LB broth (2.5 g/L, Difco Laboratory, Detroit, MI) at 37 °C and then harvested during the late-exponential phase (14−16 h incubation) by centrifugation at 2000g for 15 min. The cells were diluted in chlorine demand free (CDF) buffer (0.54 g Na2HPO4 and 0.88 g KH2PO4 per liter, pH 6.98) as a bacterial suspension (OD600 = 0.5 ± 0.02).23 2.2. Solution Preparation. Biofilms were cultivated in 0.02 strength LB broth to create nutrient limited growth conditions.24,25 Biofilm interaction with DOM was conducted using Suwannee River Natural Organic Matter (SR-NOM; IHSS, MN) filtered through a 0.45 μm membrane and applied at a final concentration of 2 ± 0.2 mg/L.26 Chlorine solutions were prepared by adding Clorox bleach (The Clorox Co., Oakland, CA) to autoclaved deionized water. The chlorine concentration was determined with the N, N-diethyl-pphenylenediamine (DPD) method using a DR/2700 spectrophotometer (HACH Company, Loveland, CO).27 2.3. Biofilm Cultivation and Disinfection in Flow Cell System. Biofilms were cultivated in two channel flowcell systems (BioSurface Technologies Corp., Bozeman, MT) fitted with a glass microscope slide opposing a glass coverslip (channel dimensions, 1.6 by 12.7 by 47.5 mm; flow rate =0.2 13213

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

using unpaired t test or one-way ANOVA test. The association between biofilm structural parameters, EPS content and biofilm viability was analyzed by linear regression analysis. P < 0.05 was accepted as statistically significant. All calculations were performed by using SigmaPlot (Jandel Scientific, Sausalito, CA).

the spatial distribution of viability, data was acquired by imaging in 1 μm depth increments throughout the image acquisition process, then calculating a cell viability of live over total cell biomass. 2.5. Bacterial Enumeration and Viability. After disinfection and microscopic analysis, biofilms from both control and disinfected channels were scraped from glass slides, resuspended in CDF buffer and carefully dispersed by vortexing. Viable cells were enumerated using heterotrophic plate counting (HPC) on R2A agar plates (Difco Laboratories, Detroit, MI) in duplicate. The chlorine disinfection efficacy was evaluated using the ratio of viable bacteria in disinfected biofilm to the control biofilm value. Enumerations of viable bacterial cells in the effluent samples followed the same procedure. Viability ratio in the effluent was defined as percentage of viable cells under disinfected condition as compared to viable cells under control condition. 2.6. Flow Cytometry Analysis. While plate counting yields quantity of culturable cells, flow cytometry analysis of the detached biofilm was used to differentiate and quantify dead, injured and live cells as a function of fluorescence intensity, based on the extent of membrane damage. Data were acquired in “log” mode using a FACScalibur flow cytometer (BD Biosciences, San Jose, CA). PI and SYTO 9 were used in combination to determine membrane compromised cells and intact cells, respectively. Stains were simultaneously added at final concentrations of 0.25 μM PI and 0.17 μM of SYTO 9 to vortexed effluent samples and incubated as described above. Cell concentration was determined by comparing cell events to events from a microsphere standard of known concentration (InVitrogen, Carlsbad, CA). On the basis of negative and positive controls, flow cytometry analysis was performed on two fluorescent channels (PI and SYTO 9) to evaluate the cellular viability. Each acquired data plot was analyzed using WinMDI (J. Trotter 1993−1998) in four quadrants:35 (A) PI positively stained dead cells with a permeabilized cell membrane; (B) both PI and SYTO 9 positively stained membrane compromised cells; (C) SYTO 9 positively stained live cells with intact cell membrane; and (D) negative signals. 2.7. Dissolved Organic Matter (DOM) Analysis. During the six day biofilm cultivation, effluent samples from biofilm and abiotic flow cell channels were collected for DOM analysis. Five samples per day were collected from each flow channel. Collected effluent samples were filtered through 0.45 μm membrane filters to remove bacterial cells and cell bound EPS. The filtered samples were analyzed in triplicate using a UV spectrophotometer at 220−350 nm (UV-1800, Shimadzu) to estimate overall DOM removal as well as selective removal of DOM by biofilms.36,37 The area under the spectra between 250 and 350 nm (A250−350) was calculated. The area difference between with and without DOM conditions was used to quantify the overall DOM concentration. The 220−230 nm band is within the benzenoid (Bz) band range representing aromatic rings substituted predominantly with aliphatic functional groups. The 240−260 nm band assigned as the electron transfer (ET) band is related to the presence of polar functional groups such as hydroxyl, carboxyl and ester groups on the ring. The ratio of area under the spectra of 220−230 nm and 240− 260 nm was calculated to evaluate the comparative removal of specific fractions in DOM by biofilm. 2.8. Statistical Analysis. Data are presented as mean ± standard deviation or standard error. Differences were analyzed

3. RESULTS 3.1. Effect of EPS Content on Biofilm Structure. After six days growth, biofilm composition, structure, and viability ratio was quantitatively analyzed using CLSM (Table 1). Image Table 1. Biofilm Structural Parameters, Biomass Content and Viability Ratio (Without DOM Condition)a roughness coefficient surface area to volume ratio (μm2/ μm3) average diffusion distance (μm) alginate EPS (μm3/ μm2) total biomass (μm3/ μm2) cell viabilityb (%) − control cell viabilityb (%) − disinfected

EPS−

WT

EPS+

1.00 ± 0.26 2.31 ± 1.28

0.69 ± 0.26 1.99 ± 0.94

0.41 ± 0.41 1.15 ± 0.36

0.80 ± 0.20

0.58 ± 0.06

0.42 ± 0.13

0.12 ± 0.03

0.87 ± 0.32

1.40 ± 1.50

16.83 ± 6.26

15.43 ± 2.44

21.54 ± 5.82

76.66 ± 18.41

78.94 ± 26.89

87.21 ± 16.35

0.96 ± 0.42

10.20 ± 3.94

23.17 ± 9.47

a Values represent average ± standard error (N ≥ 15). bCell viability was expressed as the biomass of live cells (stained with SYTO 9) divided by the biomass of total cells (SYTO 9 stained live cells and PI stained dead cells) × 100%.

analysis revealed the alginate EPS content in biofilm was 0.12 ± 0.03 μm3/μm2 for EPS- biofilm, 0.87 ± 0.32 μm3/μm2 for WT biofilm, and 1.4 ± 1.5 μm3/μm2 for EPS+ biofilm. Despite clear differences in biofilm EPS content, biofilm from all strains possessed a similar level of bacterial cell biomass (18 ± 3 μm3/ μm2). However, EPS production was determined to significantly affect biofilm structural parameters, such as roughness coefficient, surface area to volume ratio, and average diffusion distance. These three parameters were found to be negatively related to the EPS amount (P < 0.05). No significant difference between control and disinfected biofilm in structural parameters, cell biomass and EPS content for all strains. 3.2. DOM Removal by Biofilm. The spectroscopic study of DOM revealed both overall and specific removal of different fractions of DOM by biofilms. DOM removal was not specifically determined but implied by difference between the influent and effluent of flowcell reactors. The area under the spectra between 250 and 350 nm (A250−350) was used to measure the overall DOM concentration (Figure 1(a)). Around 20% reduction in DOM concentration was observed from the second day through the sixth day of biofilm growth compared to the concentration at the beginning of the experiment. In general, there was no significant difference in DOM removal among the three biofilms during the 6 day biofilm growth, although DOM concentration was slightly lower in the effluent of EPS+ biofilm than the WT biofilm and the EPS- biofilm. In order to characterize the selective removal of different fractions in the DOM, area under the spectra between 220 and 230 nm (A220−230) was compared to the area between 240 and 260 nm (A240−260). In Figure 1(b), A220−230/A240−260 ratio was 13214

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

plotted as the distance from the substratum to the biofilm surface versus the cell viability determined in each slice, where each curve represents an individual strain and condition (Figure 2).

Figure 2. Spatial distribution of control and disinfected biofilm viability ratio in the absence of DOM. The vertical axis represents the distance from the substratum increasing to the biofilm surface, where the viability ratio (horizontal axis) was determined at each crosssection.

Figure 1. DOM removal by biofilms as a function of biofilm age. (a) Overall DOM concentration in flowcell effluent. The area under the spectra between 250 and 350 nm (A250−350) was calculated. The overall DOM concentrations are quantified by the area difference between with and without DOM conditions; (b) Selective removal of different DOM fractions by biofilms. Area under the spectra between 220 and 230 nm (A220−230) was compared to the area between 240 and 260 nm (A240−260). Data points represent mean ±1 standard deviation (N = 6). Values not followed by a common letter are statistically different from each other (P < 0.05, Tukey test).

A similar pattern in viability ratio distribution was observed for all strains in control channels, ranging from 70 to 100% viability with minimum viability located in the middle biofilm depth. For disinfected biofilm, the maximum viability ratio determined for EPS-, WT and EPS+ biofilms was less than 10%, 10−15% and up to 30%, respectively. The disinfectant impact depth, defined as the distance from the top surface of biofilm to the point where maximum viability ratio was observed, was 20−30 μm for EPS- biofilm, 15−20 μm for WT biofilm, and about 10 μm for EPS+ biofilm. Compared to the other strains, significant viability ratio decrease in EPS+ biofilm was observed only at the surface layer, and the viability ratio remained constant as depth increased into the middle section of biofilm. Among all strains, a decline in stratified viability was observed approaching the substratum. 3.4. Viability of Detached Clusters during Biofilm Disinfection. As an element of biofilm lifecycle, the viability of detached biofilm clusters was investigated using flow cytometry and HPC. HPC results indicated an increased viability ratio for detached biofilm from the EPS+ strain (Figure 3), while the EPS- strain was most susceptible to disinfection. In the absence of DOM, only clusters detached from EPS- biofilm reached and maintained greater than 2-log reduction (99% inactivation) at all sampling points. Clusters detached from WT biofilm reached 2-log reduction after 60-min disinfection. However, the EPS+ clusters maintained a higher viability ratio, reaching only 1-log reduction after 2 h biofilm disinfection. In the presence of DOM, the viability ratio increased for all strains, but was most pronounced for the EPS+ strain. Even the highly susceptible EPS- detached clusters could not reach 2-log reduction. WT and EPS+ clusters showed less than 1-log reduction. The viability ratio for WT biofilm clusters was around 3%. The viability ratio of EPS+ biofilm clusters was as high as 30% at the first sampling point and was reduced to a constant value of approximately 12% after 1 h biofilm disinfection.

increased differently by the three biofilms from day 2 to day 6 when compared to the ratio at the beginning of the experiment (P < 0.05). The increase in A220−230/A240−260 ratio suggested a tendency to selectively adsorb aromatic rings substituted with polar functional groups. Furthermore, the ratio was significantly higher for EPS producing strains (WT and EPS+) than the EPS- strain (P < 0.05), indicating that the presence of EPS may be related to selective removal of DOM molecules with hydroxyl, carboxyl and ester groups. 3.3. Biofilm Resistance to Chlorine Disinfection. To better understand EPS influence on the biofilm disinfection process, both HPC and CLSM image analysis from control and disinfected channels were interpreted to determine both the overall and spatial distribution of biofilm viability upon disinfectant exposure. The HPC results of resuspended biofilm showed the EPS+ biofilm had the highest viability ratio (34.33 ± 0.15%), followed by the WT biofilm (19.78 ± 0.18%), and the EPS- biofilm (0.67 ± 0.26%), where no significant difference in overall biofilm viability was determined considering the presence or absence of DOM (P = 0.61). The overall viability ratio was increased by 30 times in the EPS+ biofilm compared to the EPS- biofilm. In addition, the overall biofilm viability ratio was negatively correlated to the roughness coefficient and average diffusion distance (P < 0.05) (SI Figure S3). Beyond the overall biofilm survival, a spatial distribution of viable biofilm cells was analyzed to investigate disinfectant efficacy among strains using CLSM image analysis. Data were 13215

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

Figure 3. Viability ratio of detached biofilm clusters determined by HPC in the presence and absence of DOM. Dotted lines indicate 1-log and 2-log reduction. Data points represent mean ±1 standard deviation (N ≥ 12).

In addition to HPC, flow cytometry provided a measure of both cell concentration and cell viability. The detached cell concentration was found consistent for all tested strains and conditions (∼106 cells/mL). To analyze cell viability, the flow cytometry acquired a data plot of PI versus SYTO 9 fluorescence intensity and data were analyzed in four quadrants (Figure 4 (a)). Reports from previous studies have shown that the staining of bacterial cells with SYTO 9 and PI did not always produce distinct “live” and “dead” populations.38,39 The appearance of yellow or orange stained cells observed in previous study indicated an intermediate state of membrane compromised cells. In this study, the intermediate state cells were observed in quadrant B (upper right), indicating both green and red positive signal. The percentage of live, dead and membrane compromised cells were plotted in a bar graph (Figure 4 (b, c)), where a comparison of flow cytometry to HPC results revealed a similar trend in viability ratio related to EPS content of tested strains. No significant difference in viability ratio was found for the control samples when comparing the conditions of with and without DOM. However, when the disinfected samples were examined, more SYTO 9 positive signals were observed (quadrants B and C in Figure 4 (a)) under the DOM condition, indicating higher viability ratio for cells during disinfection. In the presence of DOM, the portion of dead cells in detached biofilm was reduced by 8%, 24%, and 38% for EPS-, WT, and EPS+ strains, respectively. Consequently, the live cell portion increased significantly for EPS- and WT detached biofilm; however, this increase was not as substantial for the EPS+ as the other strains. Instead, with the addition of DOM, a noticeable increase in the injured cell percentage was observed for the EPS+ detached biofilm, which was 4.7 times higher compared to the absence of DOM.

Figure 4. Detached biofilm viability quantified using flow cytometry. (a) flow cytomery data in dot plot: Quadrant A) PI positive signal (dead cells); Quadrant B) PI and SYTO 9 positive signal (injured cells); Quadrant C) SYTO 9 positive signal (live cells); Quadrant D) negative signal. (b) percentage of live, dead and injured cells in the detached biofilm in the absence of DOM; (c) percentage of live, dead and injured cells in the detached biofilm in the presence of DOM. Notations in x-axis of figures (b) and (c): C stand for control; D stand for disinfected. Data points represent mean ±1 standard error (N ≥ 24).

4. DISCUSSION 4.1. Influence of EPS on Biofilm Structure and Biofilm Resistance during Chlorine Disinfection. In this study, we observed a relationship among alginate EPS content, biofilm structure and biofilm resistance to chlorine disinfection. EPS production has been reported as a primary cause of complexity in biofilm architecture.40 Biofilms formed by an alginate overproducing P. aeruginosa strain exhibited enhanced microcolony formation and greater structural maturity.41 In this study, overall biofilm thickness was around 30−40 μm, which

was similar to what Hentzer, et al. (2001) reported. The EPS+ biofilm had a lower biofilm roughness coefficient, surface area to volume ratio and average diffusion distance when compared to the WT biofilm and EPS- biofilm. Overall, biofilm viability was greatly enhanced by increased EPS content in biofilm structure, which confirms a previous study showing that a 13216

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

its extreme complexity, low concentration and high polarity.47 In the water treatment industry, the specific UV absorbance (SUVA) measurement at 254 nm is commonly used to assess the concentration of DOM in water. However, monitoring light absorbance at only 254 nm, indicating certain activated aromatic rings in DOM, fails to interpret vast and important information embedded in the rest of the spectra data.36 In this study, the UV254 measurement of the filtered effluent mostly showed a minimal difference (approximately 2−5%) between strains and the result was inconsistent in some independent replicates. In order to monitor DOM removal by biofilm with minimized interference, the area under the spectra between 250 and 350 nm was used as a surrogate for concentration of DOM in water.37 The integrated results covering a broader spectra range compensates for the potential random error in the absorbance determined at a single wavelength due to the heterogeneous DOM composition. The results show a continuous removal of DOM from the feed solution during biofilm formation for all three strains (Figure 1(a)). The UV absorbance provides not only a measure of overall DOM concentration, but also can be indicative of characteristics of DOM fractions. A220−230/A240−260 ratio was used to evaluate the proportion of nonpolar and polar compounds in DOM (Figure 1(b)). Specifically, a high A220−230/A240−260 ratio indicates that aromatic rings are substituted predominantly with aliphatic functional groups while a low ratio indicates that the aromatic rings are highly substituted with polar groups, such as hydroxyl, carboxyl, carbonyl, and ester groups. The A220−230/A240−260 ratio results indicated a tendency to selectively remove polar DOM molecules by EPS producing biofilms. Our previous study investigating DOM biosorption by UV inactivated planktonic Pseudomonas strains indicated that adsorption equilibrium was reached within 30 min and EPS quantity was directly related to DOM biosorption,48 suggesting DOM biosorption is limited by the available adsorption sites on cell biomass. The difference in DOM retention by biofilm among the three strains was not significant. However, the difference among strains was larger during the biofilm expansion stage (second and third day) compared to that during initial phase and maturation stage44 (Figure 1(a)). The finding could be largely related to DOM biosorption as the biodegradable fraction constitutes a small portion of DOM.26 During the initial phase of biofilm formation, due to limited adsorption sites, a difference in biosorption among strains was not significant. In the biofilm expansion stage, cell growth, and division is accelerated, producing more EPS, which provides adsorption sites to retain DOM, as well as increasing the complexity of biofilm structure. Consequently, the EPS+ strain had a higher DOM retention than the other strains. Upon formation of ample cell and EPS biomass, the biofilm matrix is built up, structural complexity is formed, and, the biofilm enters into the maturation stage. During the maturation stage, the rate to create new biosorption sites was subject to both growth rate and detachment rate, where large cluster detachment happened more frequently than in the initial stage and expansion stage. In this case, the available adsorption sites were mostly saturated thus the difference in DOM retention among strains disappeared. 4.3. Influence of DOM on Biofilm and Detached Biofilm Susceptibility. Cell biomass detached from biofilm is thought to display a transitional phenotype between attached and planktonic cells. The detached biofilm encompasses a wide size distribution, including single cells and clusters comprised of hundreds of cells.9 These large clusters may provide protection

mucoid strain produces a biofilm with high disinfectant resistance.12 The presence of EPS as well as biofilm structural characteristics induced by EPS appeared to impact biofilm viability. After initial cell attachment, microcolony formation with EPS overproduction may result from daughter cells originating from the attached bacterium enveloped in the EPS matrix and held in closer proximity to the parent cell.40 The presence of EPS reduced the void area between adjacent cells, which results in lower surface area to volume ratio and lower diffusion distance, hence reduced disinfectant transport into biofilm. Previous research showed that the gel-like EPS matrix may protect the deeper layers of cells from antibiotics or disinfectants by permitting limited diffusion of these chemicals into biofilm.42 Besides the slow penetration of biocides, other hypotheses for biofilm resistance were discussed, such as altered microenvironment with nutrient and oxygen depletion or the emergence of resistant phenotypes.43 These hypotheses predicted increased resistance and survival of bacterial cells near the substratum. On the contrary, an unexpected spatial distribution pattern of viability ratio was observed in this study (Figure 2). Excluding the EPS- biofilm, which was almost fully disinfected throughout the thickness, the viability ratio distribution within WT and EPS+ biofilm begins near 0% at the biofilm surface then gradually increased with depth inside biofilm. In the middle section, a maximum viability ratio was reached; however, the viability ratio in the bottom section of biofilm began to decrease when approaching the substratum. The pattern of this viability ratio distribution was similar to the distribution of biomass occupying each horizontal cross-section of biofilm (SI Figure S4). In all three strains, the middle section of biofilm had a more compact structure, possessing a higher density of biomass, when compared to the top and bottom sections. This more compacted interior structure may retard disinfectant transport into the biofilm and may also be associated with low metabolic activity, oxygen and nutrient limitation, which promote tolerance against disinfectants.2 The bottom section of all three biofilms had a lower portion occupied by biomass when compared to their middle section. Overall, this distribution pattern could be related to the emergence of a mushroom shaped biofilm structure.44,45 The reduced biomass density in the bottom section, resulting from greater void space near the substratum, appeared to promote disinfectant transport and inactivation efficiency. 4.2. Influence of EPS on DOM Retention by Biofilm. The presence of EPS promotes retention of dissolved or colloidal organic substances by providing adsorption sites. Dissolved organic matter is one of the organic substances found in the water distribution system and is known to exert disinfectant demand. DOM, which is not completely removed by conventional water treatment processes, is known to contribute to biofilm formation and bacterial regrowth in the water distribution systems.1,26 However, reduction of organic carbon levels to inhibit regrowth is very costly and complex. With conventional water treatment processes, 2−5 mg/L effluent organic carbon concentration has been commonly reported by water utilities.46 In this study, a moderate DOM concentration (TOC = 2 ± 0.2 mg/L) was chosen to simulate a distribution system. The DOM applied in this study was prepared from Suwannee River NOM, which contains many components with varied physical and chemical properties. DOM is wellknown to provide a challenge for analytical investigation, due to 13217

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

however, the viability of detached biofilm significantly increased in the presence of DOM, especially for the EPS overproducing strain. The combined results revealed that multiple roles of biofilm EPS synergistically influence resistance of both biofilm and detached clusters leading to a higher resistance against disinfection practices. The results obtained in this study can provide valuable insight regarding both the removal of DOM and the concentration of residual chlorine required to facilitate biofilm inactivation and prevent regrowth in the distribution system.

for embedded cells and shield them from disinfectant exposure.18 Another aspect providing greater tolerance for detached clusters is the presence of EPS, which have been known to sacrificially react with oxidative disinfectants, such as chlorine.2 In our previous study, P. aeruginosa EPS showed very rapid reactions with chlorine, yielding a reaction rate constant nearly 10-fold greater than that of DOM-chlorine reaction.49 When more chlorine is consumed by the EPS, less will be available to inactivate cells both in the biofilm and in the detached clusters. It can be assumed that EPS content in the detached clusters reflects the EPS content in biofilm matrix, thus the biofilm detached from the EPS+ biofilm will presumably possess more EPS than biofilm detached from the EPS- strain. In addition to the intrinsic protective role of EPS against disinfectant, the DOM both retained in biomass and suspended in the bulk solution exerts demand on available chlorine in the system, thus these reactions further reduced available chlorine for biomass disinfection.26 Interestingly, the viability test for biofilm showed that there was no significant effect of DOM on overall biofilm viability during chlorine disinfection for all strains. However, the tolerance of detached clusters was significantly enhanced under the presence of DOM, especially the EPS+ strain. The low retention of DOM in the biofilm in comparison to the total biomass quantity indicates that adsorbed DOM did not significantly influence biofilm composition within strains. During the disinfection process, chlorine reaches the biofilm surface, gradually being consumed by biofilm components (bacterial cells, EPS, and adsorbed organic substances) during penetration into deeper biofilm layers. Even though the EPS+ biofilm accumulated more DOM after six day cultivation, the effect of DOM during disinfection was distributed through the entire biofilm matrix, which limits its effect on total chlorine demand when compared to the total biomass demand. On the other hand, unlike the limited disinfectant transport distance in biofilms, the detached clusters have a much higher exposure area to volume ratio. For the detached biofilm clusters, chlorine may react and be consumed by both retained and bulk DOM immediately with much smaller penetration distance as compared to biofilm. In this case, the disinfectant demand of DOM exerts a significant impact on the viability of detached clusters. Noticeably, the proportion of membrane injured cells in detached clusters greatly increased, especially for the EPS+ strain. These injured cells had partially compromised cell membrane after disinfection, as opposed to dead cells with complete cell membrane damage. Under hospitable conditions, injured cells may recover and recolonize downstream. Considering the possibility of cell revival, the viability ratio of the detached clusters could become even higher than what was measured. In this study, synergistic influence of biofilm EPS was assessed in respect to resistance of both biofilm and detached clusters to chlorine disinfection. The obtained results suggested that (i) the presence of EPS increased both biofilm and detached cluster resistance to chlorine in the presence and absence of DOM; (ii) EPS production had an influence on biofilm structure; Some structural characteristics, such as surface roughness, surface area to volume ratio and average diffusion distance, were closely related to both overall biofilm viability and the spatial distribution of viable cells within biofilm; and (iii) the DOM adsorption did not show significant impact on biofilm viability against chlorine disinfection;



ASSOCIATED CONTENT

S Supporting Information *

P. aeruginosa strains on agar plates and under SEM (Figure S1); Flow cell system set up (Figure S2); Relationship between biofilm structural parameters and viability ratio (Figure S3); Spatial distribution of biomass through horizontal cross sections (Figure S4); Information regarding mutant strain construction, SEM and CLSM imaging. This material is available free of charge via the Internet at http://pubs.acs.org.



AUTHOR INFORMATION

Corresponding Author

*Phone: (419) 530-8131; fax: (419) 530-8116; e-mail: [email protected] Notes

The authors declare no competing financial interest.



ACKNOWLEDGMENTS



REFERENCES

We acknowledge Dr. Daniel Hassett for providing bacterial culture, Christopher Hessler for manuscript preparation and UT Imaging Center. This project is supported by National Science Foundation (Award Number: CBET0933288).

(1) Camper, A. K. Coliform regrowth and biofilm accumulation in drinking water systemsA review. Biofouling Biocorros. Ind. Water Syst. 1994, 91−105. (2) Wingender, J.; Neu, R. T.; Flemming, H.-C. Microbial Extracellular Polymeric Substances: Characterization, Structure and Function, 1st ed.; Springer: Berlin, 1999. (3) Gilbert, P.; Das, J.; Foley, I. Biofilm susceptibility to antimicrobials. Adv. Dent. Res. 1997, 11 (1), 160−7. (4) Haas, C. N.; Joffe, J.; Heath, M.; Jacangelo, J.; Anmangandla, U. Predicting disinfection performance in continuous flow systems from batch disinfection kinetics. Water Sci. Technol. 1998, 38 (6), 171−179. (5) Stewart, P. S. Mechanisms of antibiotic resistance in bacterial biofilms. Int. J. Med. Microbiol. 2002, 292 (2), 107−113. (6) Lechevallier, M. W.; Schulz, W.; Lee, R. G. Bacterial nutrients in drinking-water. Appl. Environ. Microbiol. 1991, 57 (3), 857−862. (7) Herzberg, M.; Kang, S.; Elimelech, M. Role of extracellular polymeric substances (EPS) in biofouling of reverse osmosis membranes. Environ. Sci. Technol. 2009, 43 (12), 4393−4398. (8) Hem, L. J.; Efraimsen, H. Assimilable organic carbon in molecular weight fractions of natural organic matter. Water Res. 2001, 35 (4), 1106−1110. (9) Behnke, S.; Parker, A. E.; Woodall, D.; Camper, A. K. Comparing the chlorine disinfection of detached biofilm clusters with those of sessile biofilms and planktonic cells in single and dual species cultures. Appl. Environ. Microbiol. 2011, 77 (20), 7176−7184. (10) Hu, J.; Yu, B.; Feng, Y.; Tan, X.; Ong, S.; Ng, W.; Hoe, W. Investigation into biofilms in a local drinking water distribution system. Biofilms 2005, 2 (1), 19−25.

13218

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Environmental Science & Technology

Article

(31) Lu, W.; Kiéné, L.; Lévi, Y. Chlorine demand of biofilms in water distribution systems. Water Res. 1999, 33 (3), 827−835. (32) Strathmann, M.; Wingender, J.; Flemming, H.-C. Application of fluoroscently labelled lectins for the visualization and biochemical characterization of polysaccharides in biofilms of Pseudomonas aeruginosa. J. Microbiol. Methods 2002, 50, 237−248. (33) Harrison, J. J.; Ceri, H.; Yerly, J.; Stremick, C. A.; Hu, Y.; Martinuzzi, R.; Turner, R. J. The use of microscopy and threedimensional visualization to evaluate the structure of microbial biofilms cultivated in the Calgary Biofilm Device. Biol. Proced. Online 2006, 8 (1), 194−215. (34) Heydorn, A.; Nielsen, T. A.; Hentzer, M.; Sternberg, C.; Givskov, M.; Ersboll, K. B.; Molin, S. Quantification of biofilm structures by the novel program COMSTAT. Microbiology 2000, 146, 2395−2407. (35) Gregori, J. G.; Ragheb, K.; Robinson, J. P. A Tutorial of WINMDI Version 2.8., 2004; pp 156−156 (36) Korshin, G. V.; Li, C. W.; Benjamin, M. M. Monitoring the properties of natural organic matter through UV spectroscopy: A consistent theory. Water Res. 1997, 31 (7), 1787−1795. (37) Wang, G. S.; Hsieh, S. T. Monitoring natural organic matter in water with scanning spectrophotometer. Environ. Int. 2001, 26 (4), 205−212. (38) Berney, M.; Hammes, F.; Bosshard, F.; Weilenmann, H. U.; Egli, T. Assessment and interpretation of bacterial viability by using the LIVE/DEAD BacLight kit in combination with flow cytometry. Appl. Environ. Microbiol. 2007, 73 (10), 3283−3290. (39) Stocks, S. M. Mechanism and use of the commercially available viability stain, BacLight. Cytometry, Part A 2004, 61A (2), 189−195. (40) Hentzer, M.; Teitzel, G. M.; Balzer, G. J.; Heydorn, A.; Molin, S.; Givskov, M.; Parsek, M. R. Alginate overproduction affects Pseudomonas aeruginosa biofilm structure and function. J. Bacteriol. 2001, 183 (18), 5395−5401. (41) Davies, D. G.; Parsek, M. R.; Pearson, J. P.; Iglewski, B. H.; Costerton, J. W.; Greenberg, E. P. The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 1998, 280 (5361), 295−298. (42) Gordon, C. A.; Hodges, N. A.; Marriott, C. Antibiotic interaction and diffusion through alginate and exopolysaccharide of cystic fibrosis-derived Pseudomonas aeruginosa. J. Antimicrob. Chemother. 1988, 22 (5), 667−674. (43) Stewart, P. S.; William Costerton, J. Antibiotic resistance of bacteria in biofilms. Lancet 2001, 358 (9276), 135−138. (44) Stoodley, P.; Sauer, K.; Davies, D. G.; Costerton, J. W. Biofilm as complex differentiated communities. Annu. Rev. Microbiol. 2002, 56 (1), 187−209. (45) Klausen, M.; Aaes-Jørgensen, A.; Molin, S.; Tolker-Nielsen, T. Involvement of bacterial migration in the development of complex multicellular structures in Pseudomonas aeruginosa biofilms. Mol. Microbiol. 2003, 50 (1), 61−68. (46) USEPA. Control of Biofilm Growth in Drinking Water Distribution Systems; National Service Center for Environmental Publications (NSCEP), 1992 (47) Sleighter, R. L.; Hatcher, P. G. The application of electrospray ionization coupled to ultrahigh resolution mass spectrometry for the molecular characterization of natural organic matter. J. Mass Spectrom. 2007, 42 (5), 559−574. (48) Wang, Z.; Hessler, C. M.; Xue, Z.; Seo, Y. The role of extracellular polymeric substances on the sorption of natural organic matter. Water Res. 2012, 46 (4), 1052−1060. (49) Xue, Z.; Hessler, C. M.; Panmanee, W.; Hassett, D. J.; Seo, Y. Pseudomonas aeruginosa inactivation mechanism is affected by capsular extracellular polymeric substance reactivity withchlorine and monochloramine. FEMS Microbiol. Ecol. 2012, DOI: 10.1111/j.15746941.2012.01453.x (accessed August 8 2012).

(11) LeChevallier, M. W.; Welch, N. J.; Smith, D. B. Full-scale studies of factors related to coliform regrowth in drinking water. Appl. Environ. Microbiol. 1996, 62 (7), 2201−2211. (12) Stewart, P. S.; Raquepas, J. B. Implications of reaction-diffusion theory for the disinfection of microbial biofilms by reactive antimicrobial agents. Chem. Eng. Sci. 1995, 50 (19), 3099−3104. (13) Carlson, G.; Silverstein, J. A. Effect of molecular size and charge on biofilm sorption of organic matter. Water Res. 1998, 32 (5), 1580− 1592. (14) Butterfield, P. W.; Camper, A. K.; Ellis, B. D.; Jones, W. L. Chlorination of model drinking water biofilm: Implications for growth and organic carbon removal. Water Res. 2002, 36 (17), 4391−4405. (15) Ndiongue, S.; Huck, P.; Slawson, R. Effects of temperature and biodegradable organic matter on control of biofilms by free chlorine in a model drinking water distribution system. Water Res. 2005, 39 (6), 953−964. (16) Walters, M. C.; Roe, F.; Bugnicourt, A.; Franklin, M. J.; Stewart, P. S. Contributions of antibiotic penetration, oxygen limitation, and low metabolic activity to tolerance of Pseudomonas aeruginosa biofilms to ciprofloxacin and tobramycin. Antimicrob. Agents Chemother. 2003, 47 (1), 317−323. (17) Wilson, S.; Hamilton, M. A.; Hamilton, G. C.; Schumann, M. R.; Stoodley, P. Statistical quantification of detachment rates and size distributions of cell clumps from wild-type (PAO1) and cell signaling mutant (JP1) Pseudomonas aeruginosa biofilms. Appl. Environ. Microbiol. 2004, 70 (10), 5847−5852. (18) Fux, C. A.; Wilson, S.; Stoodley, P. Detachment characteristics and oxacillin resistance of Staphyloccocus aureus biofilm emboli in an in vitro catheter infection model. J. Bacteriol. 2004, 186 (14), 4486− 4491. (19) Hardalo, C.; Edberg, S. C. Pseudomonas aeruginosa: Assessment of risk from drinking water. Crit. Rev. Microbiol. 1997, 23 (1), 47−75. (20) Mena, K. D.; Gerba, C. P. Risk assessment of Pseudomonas aeruginosa in water. Rev. Environ. Contam. Toxicol. 2009, 201, 71−115. (21) Rusin, P. A.; Rose, J. B.; Haas, C. N.; Gerba, C. P. Risk assessment of opportunistic bacterial pathogens in drinking water. Rev. Environ. Contam. Toxicol. 1997, 57−83. (22) Liu, Y.; Li, J.; Qiu, X.; Burda, C. Bactericidal activity of nitrogendoped metal oxide nanocatalysts and the influence of bacterial extracellular polymeric substances (EPS). J. Photochem. Photobiol., A 2007, 190 (1), 94−100. (23) Thurston-Enriquez, J. A.; Haas, C. N.; Jacangelo, J.; Gerba, C. P. Chlorine inactivation of adenovirus type 40 and feline calicivirus. Appl. Environ. Microbiol. 2003, 69 (7), 3979−3985. (24) Purevdorj, B.; Costerton, J. W.; Stoodley, P. Influence of hydrodynamics and cell signaling on the structure and behavior of Pseudomonas aeruginosa biofilms. Appl. Environ. Microbiol. 2002, 68 (9), 4457−4464. (25) Delille, A.; Quilès, F.; Humbert, F. In situ monitoring of the nascent Pseudomonas f luorescens biofilm response to variations in the dissolved organic carbon level in low-nutrient water by attenuated total reflectance-fourier transform infrared spectroscopy. Appl. Environ. Microbiol. 2007, 73 (18), 5782−5788. (26) Croue, J. P.; Korshin, G. V.; Benjamin, M. M. Characterization of Natural Organic Matter in Drinking Water; American Water Works Association Research Foundation: Denver, 1999. (27) Walters, G. L. Hach Water Analysis Handbook; Hach Company: Loveland, CO, 1989. (28) Rogers, S. S.; van der Walle, C.; Waigh, T. A. Microrheology of bacterial biofilms in vitro: Staphylococcus aureus and Pseudomonas aeruginosa. Langmuir 2008, 24 (23), 13549−13555. (29) Sauer, K.; Camper, A. K.; Ehrlich, G. D.; Costerton, J. W.; Davies, D. G. Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 2002, 184 (4), 1140− 1154. (30) Gang, D.; Clevenger, T. E.; Banerji, S. K. Relationship of chlorine decay and THMs formation to NOM size. J. Hazard. Mater. 2003, 96 (1), 1−12. 13219

dx.doi.org/10.1021/es3031165 | Environ. Sci. Technol. 2012, 46, 13212−13219

Suggest Documents