Polyelectrolyte and Silver Nanoparticle Modification of Microfiltration

Feb 13, 2012 - Microfiltration Membranes To Mitigate Organic and Bacterial Fouling ... quality. Microfiltration membranes, in particular, have shown p...
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Polyelectrolyte and Silver Nanoparticle Modification of Microfiltration Membranes To Mitigate Organic and Bacterial Fouling Fatou Diagne,† Ramamoorthy Malaisamy,† Vic Boddie,‡ R. David Holbrook,§ Broderick Eribo,‡ and Kimberly L. Jones*,†

Environ. Sci. Technol. 2012.46:4025-4033. Downloaded from pubs.acs.org by UNIV OF THE SUNSHINE COAST on 06/27/18. For personal use only.

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Department of Civil and Environmental Engineering and ‡Department of Biology, Howard University, 2300 Sixth Street Northwest, Washington, D.C. 20059, United States § Chemical Science and Technology Laboratory, National Institute of Standards and Technology, Gaithersburg, Maryland 20899, United States ABSTRACT: Membrane fouling remains one of the most problematic issues surrounding membrane use in water and wastewater treatment applications. Organic and biological fouling contribute to irreversible fouling and flux decline in these processes. The aim of this study was to reduce both organic and biological fouling by modifying the surface of commercially available poly(ether sulfone) (PES) membranes using the polyelectrolyte multilayer modification method with poly(styrenesulfonate) (PSS), poly(diallyldimethylammonium chloride) (PDADMAC), and silver nanoparticles (nanoAg) integrated onto the surface as stable, thin (15 nm) films. PSS increases the hydrophilicity of the membrane and increases the negative surface charge, while integration of nanoAg into the top PSS layer imparts biocidal characteristics to the modified surface. Fouling was simulated by filtering aqueous solutions of humic acid (5 and 20 mg L−1), a suspension of Escherichia coli (106 colony-forming units (CFU) mL−1), and a mixture of both foulants through unmodified and modified PES membranes under batch conditions. Filtration and cleaning studies confirmed that the modification significantly reduced organic and biological fouling.

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INTRODUCTION Membrane applications in water and wastewater treatment have increased steadily over the past few years due to improved membrane design, smaller footprint, and reliable effluent quality. Microfiltration membranes, in particular, have shown promise as a cost-effective alternative to conventional water treatment, as pretreatment for nanofiltration (NF) and reverse osmosis (RO), and as key components in membrane bioreactors. Despite this potential, organic fouling and biological fouling remain problematic for membranes. Fouling increases operating and replacement costs and compromises membrane performance. The abundance of natural organic matter (NOM) in natural waters makes organic fouling of membrane processes in water treatment inevitable; biofouling is considered more problematic since bacteria can reproduce at the membrane surface, create biofilms, and cause additional fouling.1 Approaches to limit biofouling aim to prevent or delay one or many of the steps necessary for biofilm growth: adhesion, reproduction, and proliferation. Several methods to mitigate fouling focus on altering the surface charge or increasing the hydrophilicity of the surface to discourage attachment and adsorption of foulants onto the surface.2−6 Since initial stages of fouling involve membrane− foulant interactions, membrane surface modification methods © 2012 American Chemical Society

have been identified as promising methods to reduce fouling. Promising surface modification methods include polymerization,6−10 polyelectrolyte adsorption, and plasma modification.11 First introduced by Decher et al.,12 polyelectrolyte multilayer modification (PEM), an alternate electrostatic deposition of oppositely charged polyelectrolytes, is a reproducible, versatile method that allows control over the surface charge, pore size, hydrophilicity, film thickness, and ionic charge of membranes. Multilayers are formed on the membrane surfaces by charge reversal and overcompensation, with the final polyelectrolyte layer determining the charge of the multilayer films.9,13−15 While PEM capped with negatively charged poly(styrenesulfonate) (PSS) addresses organic fouling by increasing electrostatic repulsion near the surface, most polyelectrolytes do not directly prevent bacterial growth on the membrane surface. Therefore, silver nanoparticles (nanoAg) are incorporated into the top PSS layer of the PEM-modified Received: Revised: Accepted: Published: 4025

November 4, 2011 February 1, 2012 February 13, 2012 February 13, 2012 dx.doi.org/10.1021/es203945v | Environ. Sci. Technol. 2012, 46, 4025−4033

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however, the number of bilayers was limited to 1.5 to minimize the thickness and flux decline. For the nanoAg-impregnated surfaces, the polyelectrolyte solution was prepared to result in 0.02 mol L−1 PSS and 0.001 mol L−1 citrate-stabilized silver nanoparticles. The stability of the citrate-coated nanoAg suspension was confirmed for up to 134 days.28 Furthermore, due to the stabilizing effect of citrate, precipitation was not observed when the negatively charged PSS was dispersed in the nanoAg suspension. The modified membranes were then used for characterization, filtration, and silver toxicity studies. Membrane Characterization. ζ Potential. An asymmetric clamping cell (Anton Paar, Graz, Austria) attached to an electro kinetic analyzer (EKA; Brookhaven Instruments, Holtsville, NY) was employed to measure the ζ potential of the unmodified and modified membranes with 1 mmol/L KCl as electrolyte at pH 5.5 ± 0.2. The experiments were carried out as by Malaisamy et al.29 The reported values are the average of ζ potentials determined in both flow directions with at least three membranes. Membrane Morphology. Surface morphologies of the unmodified and modified membranes with and without nanoAg at various magnifications were obtained by environmental scanning electron microscopy (ESEM) combined with energydispersive X-ray spectroscopy (EDS) (Quanta 200 FEG ESEM, FEI, Hillsboro, OR). The membranes for SEM analysis were initially air-dried and frozen in liquid nitrogen. The dried surfaces were sputter-coated with gold−palladium alloy using a sputter coater to a thickness of 5 nm prior to imaging. Secondary and backscattered electron imaging was performed to illustrate surface morphology, architecture, and compositional differences, while EDS was used to verify the presence of nanoAg and aggregated Ag. Identification of Surface Functionalization. The surface chemical functionality of the unmodified and modified PES membranes was obtained with an attenuated total reflectance Fourier transform infrared (ATR-FTIR) spectrometer (Bruker Vertex 80v). Clean, dry membrane pieces were mounted on the ATR diamond crystal, and 256 IR scans were performed at a resolution of 4 cm−1 at an incident angle of 45°. The IR penetration depth for this incident angle is 2.0−4.36 μm. A baseline was obtained for each membrane, and the sample chamber was maintained under vacuum at 230 Pa to avoid interference from air and moisture. The detector was cooled with liquid nitrogen to further reduce baseline noise. Membrane Hydrophilicity. The contact angles of the membranes were measured immediately after modification to determine the change in hydrophilic/hydrophobic behavior upon modification and after 14 days to assess the stability of the modified membranes. Each contact angle measurement was conducted in triplicate; for each sample, at least 10 measurements per membrane were completed using the sessile drop method on the Phoenix 150 goniometer (Surface & ElectroOptics Corp.). The high number of experiments were completed to ensure accuracy on the highly permeable membranes. Quantification of Silver. The silver content of the nanoAgmodified membranes was determined in parallel experiments before and after pure water filtration, after 7 and 14 days by inductively couple plasma mass spectrometry (ICP-MS, Agilent) to assess the stability of the nanoAg layer. Modified membranes were digested following a modified USEPA 3050b method. The membrane was soaked in 10 mL of concentrated

membranes to meet our overall goal of reducing both organic and biological fouling. Antibacterial materials such as silver, chitosan, quaternary ammonium groups, photocatalytic TiO2, and ethylene glycol oligomers have been incorporated into or onto surfaces to prevent bacterial growth.1,16−18 Silver particles in various forms release silver ions that damage the bacteria by producing reactive oxygen species or preventing DNA replication in the bacterial cell.16,19−24 NanoAg particles exhibit antibacterial properties in various substrates.17,25,26 Sondi et al. studied the effect of nanoAg on Escherichia coli and reported that nanoAg particles in agar plates completely inhibit microorganisms, whereas nanoAg particles in liquid medium only delay growth.23 Although Ag-based nanoparticles are among the most frequently used nanomaterials for antimicrobial applications, few studies have investigated their use for mitigation of biofouling of membranes. Zodrow et al. developed polysulfone silver-impregnated membranes with antibacterial and antiviral properties.16 Li et al. designed a multilayer film using polyelectrolytes and embedded silver ions, which have the potential to kill bacteria on contact.21 Most modification studies focus on a single type of fouling, whereas membranes typically experience several fouling mechanisms concurrently during operation. In this study, we address organic and biological fouling simultaneously by modifying commercially available poly(ether sulfone) (PES) membranes using (1) polyelectrolyte multilayers alone and (2) polyelectrolyte multilayers embedded with nanoAg particles. Model foulants were humic acid and an E. coli suspension. The performance of the modified and unmodified membranes was evaluated by filtration experiments in which PEM-modified membranes were compared to unmodified membranes in terms of permeability, surface charge (ζ potential), surface morphology, irreversible fouling resistances, and flux recovery after cleaning. Additionally, nanoAg-modified membranes were compared to silver-free membranes in terms of biological fouling potential. Resistance to biofouling was determined by fluorescence microscopy and by performing an aerobic plate count on unmodified membranes, PEM-modified membranes without nanoAg, and PEM-modified membranes with nanoAg.

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MATERIALS AND METHODS Surface Modification. The PES membrane coupon (0.1 μm pore size, GE Osmonics) was placed in a holder with the top surface exposed. The polyelectrolyte solutions were alternately deposited on the support for 3 min each with a water rinse between layers. Each layer serves as a surface primer for the next oppositely charged layer. The deposition was initiated by the negatively charged PSS solution (0.02 mol L−1, MW = 70 000, Aldrich Chemical Co.), which adheres to the membrane surface via hydrogen bonding and hydrogen− hydrogen interactions. 27 A positively charged poly(diallyldimethylammonium chloride) (PDADMAC) solution (0.02 mol L−1, MW = 100000−200000, Aldrich Chemical Co.) is then added, which adheres to the PSS layer via electrostatic interactions and van der Waals forces.13 The third and top layer was either another PSS layer or a PSS layer embedded with citrate-coated nanoAg (CREST center, Howard University), thus resulting in 1.5 bilayers. Details on the synthesis and characterization of the nanoAg particles can be found in the paper by Hai Ting.28 Additional layers would increase the negative charge of the terminal PSS layer (data not shown); 4026

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Figure 1. ESEM images of unmodified and modified membranes with and without silver nanoparticles. (A) Unmodified membranes. (B) Membranes modified with 1.5 bilayers of polyelectrolytes. (C) Membranes modified with 1.5 bilayers of polyelectrolytes and silver nanoparticles. The bottom images are combined secondary and backscattered electron images. Silver nanoparticles are depicted by false red to highlight the distribution. (D) Aggregates of silver nanoparticles shown on the membrane surface and in the vicinity of the membrane pores.

nitric acid and heated at 100 °C until the membrane was completely digested. The sample was cooled before 3 mL of hydrogen peroxide (30%) was added, and the mixture was heated at 100 °C until no effervescence was noted in the solution. After the sample was cooled, the solution was diluted to 100 mL before being filtered through a Whatman no. 41 filter paper. The permeate samples were collected every 30 min, and the silver concentration was quantified to assess leaching during filtration. Samples were acidified to pH 2 using concentrated nitric acid. Filtration Studies. A dead-end stirred cell consisting of a filtration cell (Amicon model 8050, Millipore Corp., Bedford, MA) with a total cell volume of 50 mL and an effective membrane area of 13.4 cm2 was connected to a cylinder with ultrapure grade nitrogen via a stainless steel pressure reservoir of 20 and 3 L, respectively, for the organic fouling and biofouling experiments. The pure water flux (Jo) was measured at an applied pressure of 69 kPa. The pure water was then replaced by the foulant solutions to carry out the fouling studies. The pure water flux was then measured before and after cleaning to determine the flux recovery and membrane resistances.

Fouling protocols. For the organic fouling experiments, humic acid solutions (5 and 20 mg/L concentration) were filtered for 2 h (Jf). Pure water flux of the fouled membranes was measured after the membranes were hydraulically cleaned in Milli-Q water (Jr) for about 3 min. The fouled membranes were then unloaded and chemically cleaned using 0.2 mol L−1 sodium hydroxide (NaOH). After chemical cleaning, the membranes were reloaded in the cell to measure the pure water flux (Ji). For the biological fouling experiments, a suspension of E. coli (ATCC 25922) was filtered through the cell. E. coli was cultured on brain heart infusion (BHI) agar (Remel, Lenexa, KS) and incubated at 37 °C for 48 h. The cell inoculum was suspended in sterile 0.25 mol L−1 phosphate-buffered saline (PBS) and adjusted to 106 CFU mL−1 (CFU = colony-forming units). The suspension was used to assess the bactericidal activity at the membrane surface. Following the pure water flux (Jo) measurement, the suspension was filtered through the membranes for 2 h and the flux measured (Jf). Microorganisms on the fouled membranes were allowed to grow on agar plates incubated at 37 °C for 24 h. The membranes were then hydraulically cleaned for 30 min and chemically cleaned using 4027

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Table 1. Membrane Characterizationa

200 mg L−1 sodium hypochlorite (NaOCl) for 1 h in the stirred cell. The pure water flux was measured after each cleaning step and is denoted as Jr following hydraulic cleaning and Ji following chemical cleaning. For the combined fouling experiments, a mixture of humic acid (20 mg L−1) and E. coli (106 CFU mL−1) was filtered through the cell. In these experiments, the membranes were cleaned using NaOCl followed by NaOH. The normalized flux (Jn) was calculated by dividing the steady-state flux (Jf) by the initial flux (Jo). The percent flux reduction (J%) of each membrane was computed as a function of pure water flux after fouling and cleaning (PWFf) and pure water flux before fouling (PWFo): J% =

PWFo − PWFf PWFo

ζ potential (mV) inherent resistance (1/m)

contact angle, day 1 (deg) contact angle, day 15 (deg)

unmodified

modified

nanomodified

−2.96 (0.68) 4.1 (0.15)

−52.6 (0.01) 4.8 (0.15)

−51.8 (0.38) 4.7 (0.3)

unmodified

modified, 0.5 bilayer

modified, 1.5 bilayers

25.4 (3.73)

25.3 (1.37)

24.12 (2.76)

not measured

22.1(2.30) 22 (1.27)

ζ potential of membranes taken at pH 5.4 with 0.01 M KCl (charge of the membrane surfaces), inherent resistances of the membranes before fouling, and contact angle values of the membranes (hydrophilicity) right after modification and 14 days after modification. The membranes were soaked in Milli-Q water (stability of the PEM films). Standard deviations are given in parentheses.

a

× 100 (1)

NanoAg Toxicity Experiments. Biological activity experiments were conducted with a smaller feed volume (3 mL) to directly assess the impact of silver on the E. coli. The smaller feed volume has a resulting lower bacterial deposition, allowing for more accurate plate counting. Membranes were fouled by depositing either 3 mL of 106 CFU mL−1 E. coli or 3 mL of a mixture of 3 mL of 106 CFU mL−1 E. coli and 5 mg L−1 humic acid onto the membrane. An aerobic plate count (APC) was performed to determine the viable cell count of E. coli on all membranes after exposure. Each membrane was suspended in 99 mL of 0.25 mol L−1 PBS and agitated using a Stomacher laboratory blender (80, Steward, U.K.) for 2 min to desorb the bacterial cells. A serial dilution was performed, and the diluent was plated on plate count agar (PCA) (Benton and Dickinson, Sparks, MD) and then incubated at 37 °C for 48 h. After incubation, the number of colonies formed was determined to establish the concentration of viable E. coli cells following the fouling experiments. Following agitation in the Stomacher, the membranes were individually plated on PCA to determine if any viable cells remained attached at the surface. All experiments were performed in triplicate.

shielded by hydration.26,30 For the PEM-modified membranes, the terminal PSS layer increases the negative surface charge of the membrane with a ζ potential of −52.6 mV; the addition of silver nanoparticles did not significantly affect the ζ potential of the nanomodified membranes, which was −51.8 mV. The increase in negative charge after PEM modification increases electrostatic repulsion of negatively charged species, including organic material at environmentally relevant pH values. Validation of Surface Modification by Functional Group Identification. The FTIR spectra of the unmodified and modified membrane surfaces taken on an ATR diamond crystal are shown in Figure 2. The spectrum of the PEM-modified

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RESULTS AND DISCUSSION Membrane Characterization. Membrane Morphology. There was no significant difference between the surface pore morphology of the virgin membrane and that of the modified membrane (Figure 1 A,B). Changes in surface pore size were minimized by limiting the modification film to 15 nm (1.5 bilayers). The nanoAg particles were well dispersed on the surface (Figure 1C), and the average particle size of the nanoAg was confirmed by ESEM to be approximately 50 nm, which corresponds to the nanoparticle size in the original citratestabilized solution. The integrity of the membrane pore structure and the nanoparticles remained intact after the modification. A few nanoAg aggregates were observed on the surface of the membrane (Figure 1D). Surface Charge. The PES membrane has two functional groups: sulfonyl and phenyl.26 Oxygen in the sulfonyl group participates in hydrogen bonding, and the phenyl groups undergo hydrophobic interactions with the polyelectrolyte, allowing the initial layer of PSS to create stable interactions with the PES. The mean ζ potential values were derived for PEM-modified and unmodified membranes (Table 1). The unmodified membranes had a slightly negative ζ potential of −2.96 mV because the membrane’s PES structure may be

Figure 2. FTIR spectra of the unmodified and modified membranes with and without nanoparticles. Emphasis is on the sulfonic stretch which appears on all the modified membranes (as indicated by arrows). Spectra are stacked to emphasize the peaks.

membranes has a peak at 1028 cm−1 which corresponds to the symmetric sulfonic stretch of PSS.30 The presence of this peak confirms the chemical changes imparted to the membranes by the deposition of the polyelectrolytes, specifically the top PSS layer. Hydrophilicity. Contact angle measurements confirmed the increased hydrophilicity of the PEM-modified surface. The water contact angle of the PEM-modified membranes was 14% (±6%) lower than that of the unmodified membranes; the contact angle of the membranes continued to decrease as increasing layers of PSS were deposited on the membrane 4028

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Figure 3. Flux decline of the unmodified and modified membranes at different concentrations of foulants. The flux decline represents the difference between original pure water flux (before fouling) and final pure water flux (after fouling and cleaning).

reduction due to the PEM modification. The film thicknesses were verified by ellipsometric measurements (data not shown). Despite efforts to minimize the thickness of the modified layer, the initial pure water flux (Jo) of PEM-modified membranes was 14% lower than that of the unmodified membranes due to a 14% higher inherent membrane resistance (Table 1). Lower initial flux in modified membranes can be offset by decreased fouling and flux decline, which would result in higher recovery and more efficient overall performance. Flux Decline with Organic Foulant. The term “flux decline” is used in this study to describe the difference between the original pure water flux and the pure water flux after fouling and chemical cleaning (Figure 3). Organic fouling experiments were conducted at a relatively low organic concentration (5 mg L−1) and a high organic concentration (20 mg L−1). The concentration of humic material in the feed solution did not have a significant effect on the overall flux decline, so the results reported are averages of both conditions. The flux decline of the PEM membranes without nanoAg was 8%, the flux decline for the PEM membranes with nanoAg was 16%, and the flux decline of the unmodified membranes was 23%. Although the initial flux decline was higher for the humic acid feed concentration of 20 mg L−1 when compared to the humic acid feed concentration of 5 mg L−1, the steady-state flux was comparable in both cases. The hydrophilic, charged surface of the modified membrane hinders the attachment of the hydrophobic humic acid due to electrostatic repulsion, resulting in a loose humic acid layer on the membrane surface. The membrane surface changes imparted on the membranes during modification reduce the hydrophobic/hydrophobic interactions between the native PES surface and the humic substances in solution, further reducing humic acid attachment. The nanoAg at the surface as evidenced by ESEM (Figure 1) might explain the slightly lower flux of the membranes modified with nanoAg compared to membranes modified without silver. The aggregated nanoAg (50 nm diameter per particle) may

surface (Table 1). The contact angle remained stable over 14 days. Leaching of Ag. The nanoAg-modified membranes contained 0.04 g of Ag/g of membrane upon initial modification; the surface concentration decreased to 0.02 g of Ag/g of membrane after 150 min of filtration. The silver detected in the permeate stream over 150 min was 5 μg L−1 on average, which is lower than the silver maximum contaminants limit (MCL; 100 μg L−1) mandated by the World Health Organization (WHO) and the U.S. Environmental Protection Agency (USEPA). For water and wastewater treatment applications, the nanoAg membrane coating should be stable enough to ensure that the permeate stream is not enriched with Ag+ or nanoAg particles. The final silver concentration (0.02 g of Ag/g of membrane) remained constant after 7 and 14 days of soaking in Milli-Q water, implying that the rate of silver leaching stabilizes after an initial depletion period. The pattern of leaching observed in this study has been documented for polysulfone membranes16 and other matrixes.31 When determining a method for introducing nanoAg particles into a membrane structure, one may either deposit the silver throughout the entire thickness of the membrane or deposit specifically near the active surface. The integration of the nanoAg at the membrane surface has advantages over dispersing silver throughout the entire membrane thickness and over using silver ions in solution. One of the advantages is that surface integration of nanoAg concentrates the available silver in the surface layer, making the silver more available for biocidal activity. Furthermore, the nanoAg layer can be regenerated when needed. Filtration Studies. Membrane Flux. One potential disadvantage of surface modification is that the additional thickness imparted by the modification film may significantly increase the membrane resistance, decreasing flux.5,32 In this study, the thickness of each polyelectrolyte multilayer was limited to 5 nm/layer (15 nm total) to minimize the flux 4029

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Figure 4. Membrane resistances for the fouling experiments. Foulants were 5 and 20 ppm humic acid for the organic fouling and 106 CFU mL−1 E. coli for biofouling. Inherent resistances were measured before any fouling. Irreversible resistances were measured after fouling and cleaning. Hydraulic and chemical cleaning were done with 0.2 M NaOH (organic fouling) and 200 ppm NaOCl (biofouling).

without silver reject both humic acid and E. coli on the basis of electrostatic interactions and size. The PEM-modified membranes suffered a smaller flux decline of 0.41% (without nanoAg) and 8.9% (with nanoAg) when compared to the unmodified membranes (23.6%). The increased negative ζ potential of the PEM-modified membranes imparts an electrostatic energy barrier to prevent attachment of negatively charged contaminants (humic acid). The presence of nanoAg, which is very bioavailable since it is located near the surface, prevents the growth, proliferation, and accumulation of E. coli during biofouling and combined fouling. Membrane Resistance. Although the number of bilayers was limited to 1.5 (15 nm), they increased the inherent membrane resistance enough to result in lower initial clean water flux and permeability (Figure 4); the irreversible resistances of the PEMmodified membranes, however, were lower than those of the unmodified membranes at the end of the experiments. The PEM-modified membranes (no nanoAg) had an irreversible resistance of 0.4 m−1, the PEM-modified membranes (with nanoAg) had an irreversible resistance of 0.9 m−1, and the unmodified membrane had an irreversible resistance of 1.2 m−1 (Figure 4). The higher irreversible resistance of the unmodified membrane is due to stronger hydrophobic−hydrophobic interactions and attachments between the humic acid and the membrane surface.11 The irreversible resistances of the modified membranes were much lower during biofouling than organic fouling (Figure 4), primarily due to the cake layer fouling present with E. coli. Cake layer deposition is easier to clean than internal fouling; therefore, the irreversible fouling is minimal for the E. coli foulant. Conversely, humic acid, which is polydisperse and ranges in size from 0.5 to 3.3 nm depending on the pH,34 can travel into the membrane pores. Therefore, the irreversible portion of organic fouling is larger than that of biological fouling (Figure 4). The hydrophilic modified membranes prevent initial bacterial attachment, resulting in a lower irreversible resistance. The average irreversible resistance of the unmodified membranes during combined fouling was

partially block some of the membrane pores (100 nm diameter), reducing the flux through the membrane. Furthermore, Ag+ released from the nanoAg may slightly decrease the electrostatic energy barrier if the silver in solution complexes with the humic material. Researchers have shown that humic acid may influence the interactions of some nanoparticles in solutions.33 Potential interactions between humic material and silver in solution may have resulted in a more pronounced flux decline for membranes with nanoparticles than membranes modified without silver nanoparticles during organic and combined fouling. However, both types of modified membranes had a lower flux decline than unmodified membranes during organic, biological, and combined fouling. Flux Decline with Bacterial Foulant. The flux decline during biofouling was 0.3% for membranes without nanoAg and 3.7% for the membranes with nanoAg, while the unmodified membrane flux decline was 12.2% (Figure 3). E. coli is roughly 0.5 μm, which prevents it from entering the membrane pores; thus, bacterial fouling is expected to be a surface phenomenon and should result in a reversible cake layer unless the first layer is strongly bound to the surface. For the modified membranes, the increased negative charge and hydrophilicity will prevent attachment of the bacteria on the membrane surface, thereby reducing the irreversible fouling potential of the modified membranes. The reversible deposition of bacteria is therefore easily removed from the modified membrane surface as shown by the flux decline and irreversible resistance results (Figures 3 and 4). Flux Decline with Combined (Organic and Bacterial) Foulants. Flux decline of the unmodified membrane during combined fouling was 23.6%, which is similar to the flux decline during organic fouling for the unmodified (neutrally charged) membrane. Fouling using both contaminants is dominated by organic fouling and influenced by the hydrophobic interactions between the surface and the humic acid. The modified membranes (without silver) experienced 0.41% flux decline in the combined foulant experiments, similar to the flux decline for biofouling experiments (Figure 3). Modified membranes 4030

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Table 2. Biological Analysis Results of the Fouled Membranes Consisting of Plate Counts, Fluorescence Microscopy Images, and Visual Observationa

a

The plate counts are averages of over nine measurements. Microscopy images shown here are representative of other pictures taken. Membrane pictures are visual evidence of the bacterial growth after membrane desorption. In biological analysis, the foulant was 106 CFU/mL E. coli; in bioorganic analysis, the foulant was 106CFU/mL E. coli and 5 mg/L humic acid.

1.18 m−1, which is very similar to the irreversible resistance during organic fouling. The membranes experience both internal fouling and cake layer deposition. However, the average irreversible resistance for the modified membrane without silver was 0.017 m−1, which is very similar to the irreversible resistance during biofouling only. For modified membranes, electrostatic repulsion reduces fouling as discussed previously; additionally, the cake layer formed by the bacteria provides an additional steric barrier that prevents the humic acid from entering the membrane pores. Thus, the internal fouling of the membranes by humic acid is reduced for the combined fouling experiments. As evidenced by the filtration data (Figures 3 and 4), the modified membranes show lower irreversible resistance for organic, bacterial, and combined fouling experiments. Antibacterial Studies. Membranes modified with nanoAg exhibited zero cell growth, even when the desorbed membranes were replated on nutrient agar. All other membranes (unmodified and PEM-modified without nanoAg) showed significant bacterial growth (Table 2), demonstrating that the nanoAg imparted bactericidal activity to these membranes. The nanoAg particles interact directly with the first layer of E. coli deposited at the surface of the membranes, subjecting them to its toxic effects. NanoAg at the surface also emits silver ions as evidenced by the ICP-MS results from the membrane permeate. The main mechanism for the antimicrobial activity of silver species is believed to be the release and subsequent bacterial uptake of silver ions. Released silver ions damage the

bacteria upon contact, produce reactive oxygen species, or prevent DNA replication in the bacterial cell. Silver ions, upon penetrating the bacterial cell, degrade the polysaccharide molecules, accumulate in “pits”, increasing the bacteria’s membrane permeability, and inactivate the respiratory enzyme.16,20−23,25 Fluorescence microscopy images give further evidence of the bactericidal properties of the nanoAg toward E. coli (Table 2). The images do not allow for a quantitative measurement of the live/dead cells; however, the microscopic images showed a significantly higher percentage of dead cells (red) on the nanoAg-modified membranes as opposed to the silver-free membranes, which had more live cells (green). The effect of humic acid on the toxicity of nanoAg was assessed using the silver-free membranes (unmodified and PEMmodified) as controls. Following toxicity studies with a combined feed solution consisting of humic acid and E. coli, no growth was measured on the nanoAg membrane, leading to the conclusion that humic acid does not have a significant effect on the biocidal properties of nanoAg under the physicochemical conditions of this study. Others have found that natural organic matter does influence the toxicity of nanoAg in solution when using a 10 mg L−1 humic acid concentration.33,35 However, in this study, for the toxicity experiments, 5 mg L−1 humic acid was used (note that the higher concentration of 20 mg L−1 humic acid was only used for the fouling experiments, not the toxicity studies). Also, it is important to note that the nanoAg particles in this study are fixed on a solid substrate 4031

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rather than suspended in solution, which prevents the humic acid from coating the entire surface of the nanoparticle. Significance of the Results. Membranes modified using the PEM techniques exhibited lower biological and organic fouling (lower flux decline and irreversible resistances) when compared to an unmodified PES membrane. Modified membranes that contained nanoAg exhibited antibactericidal properties while retaining the increased charge and hydrophilicity. These membranes would be useful for organic and biological fouling prevention during water and wastewater treatment. Fouling is inevitable; however, reducing fouling of membranes will make membrane processes more sustainable, easier to maintain, and more economical.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]; phone: (202) 806-4807; fax: (202) 806-5271. Notes

Any opinions, findings, conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the NSF or the EPA. This work has not been subjected to EPA review, and no official endorsement should be inferred. The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This material is based upon work supported by the National Science Foundation (NSF) and the Environmental Protection Agency (EPA) under NSF Cooperative Agreement EF0830093, Center for the Environmental Implications of NanoTechnology (CEINT). Additionally, we thank Andy Hai Ting and the CREST laboratories at Howard University for the synthesis of the silver nanoparticles and Dr. Seokjoon Kwon from the department of Civil Engineering at the University of Maryland, Baltimore County (UMBC) for his assistance using the ICP/MS instrument. We thank Dr. Delina Lyon, Research Associate at Howard University, for sharing her expertise on the microbial growth.

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