Environ. Sci. Technol. 2005, 39, 6461-6469
Direct Observation of Microbial Adhesion to Membranes† SUNNY WANG,‡ GREG GUILLEN, AND E R I C M . V . H O E K * ,§ Department of Chemical and Environmental Engineering, University of California Riverside, Riverside, California 92521
Direct microscopic observation and an interfacial force model were used to better understand and control microbial adhesion to polymeric ultrafiltration membranes. The model was used to predict a “critical flux”, below which cells deposited reversibly, and direct observation was used to visually quantify cell deposition and removal. In preliminary direct observation experiments, permeate reversal (backpulsing) was more effective than cross-flow hydrodynamics at removing deposited cells. In experiments conducted below the critical flux, no cell accumulation was observed over repeated forward-reverse filtration cycles; however, a small fraction of cells deposited irreversibly regardless of the flux, membrane, or solution chemistry. The fraction of irreversibly deposited cells was consistent with the equilibrium surface coverage attained without permeation (i.e., due to heterogeneous adsorption). Although steric forces were not invoked to establish a critical flux, when operating above the critical flux, a balance between permeation drag and steric repulsion appeared to determine the strength of adhesion of cells to membranes. Direct observation also confirmed that above the critical flux fouling occurred and pressure losses accumulated over several backpulse cycles, whereas below the critical flux there were no observable pressure losses or fouling.
Introduction Fouling is a major performance limiting factor in environmental membrane processes that causes higher operating pressures, increased chemical cleaning, and accelerated membrane replacement (1). Biological fouling (biofouling) has become a significant concern in membrane filtration processes (i.e., microfiltration, MF, and ultrafiltration, UF) used to treat municipal wastewater and alternative (brackish) waters (2). Biofouling occurs through a cascade of events including transport, deposition, adhesion, exopolymer production, growth, and proliferationsall contributing to biofilm formation (3-5). Physical and chemical factors govern the first three events, while biological factors determine the later three (6). The present study of biofouling focuses on understanding and controlling the events leading to adhesion of microbial cells to polymeric UF membranes challenged by concentrated cell suspensions. †
This paper is part of the Charles O’Melia tribute issue. * Corresponding author phone: (310)206-3735; fax: (310)206-2222; e-mail:
[email protected]. ‡ Present address: Black & Veatch, 800 Wilshire Blvd. Suite 600, Los Angeles, CA 90017. § Present address: Department of Civil and Environmental Engineering, University of California Los Angeles, 5732G Boelter Hall, Los Angeles, CA 90095. 10.1021/es050188s CCC: $30.25 Published on Web 06/24/2005
2005 American Chemical Society
In membrane filtration processes, transport and deposition of microbial particles is dominated by bulk hydrodynamic factors such as permeate convection, Brownian diffusion, inertial lift, and shear induced diffusion (6-9). Initial adhesion of particles is mediated by interfacial forces arising from physicochemical properties of cells and membranes. Interfacial forces in aquatic media are traditionally described through the classical Derjaguin, Landau, Verwey, Overbeek (DLVO) theory of colloid stability, which accounts for attractive van der Waals and repulsive electrostatic double layer interactions (10, 11). However, Lewis acid-base (12), steric (13), and interfacial hydrodynamic (14, 15) interactions also influence initial microbial adhesion membranes (1619). In recent years, the importance and roles of non-DLVO interactions for colloidal transport, microbial adhesion, and membrane fouling has been a major research focus of environmental scientists and engineers interested in interfacial phenomena (20-24). Recently, Kang et al. (25) combined classical DLVO and interfacial hydrodynamic interactions to describe initial deposition of microbial particles onto membranes. The model predicted a “critical flux” whereby DLVO repulsion was in excess of permeation drag and particles deposited reversibly. Direct microscopic observation confirmed that microbial cells readily deposited at subcritical flux conditions, but most cells quickly released under moderate laminar cross-flow when permeation ceased. These observations contradicted previous studies in which critical flux was defined as “a flux below which there is no observed flux decline with time” (26, 27) or as “a flux below which particle deposition does not occur at all” (28, 29). In this study, critical flux was predicted by an interfacial force model accounting for interfacial hydrodynamic, DLVO, and acid-base interactions between a model microbial particle and two commercial UF membranes. Deposition and removal of microbial cells at subcritical and supercritical fluxes were assessed by direct microscopic observation. Cells deposited under dead-end filtration conditions were removed by either cross-flow scouring or rapid permeate backpulsing. The impacts of permeate flux, cross-flow hydrodynamics, backpulse intensity, solution chemistry, and membrane properties on microbial cell deposition and removal were investigated. The potential effects of microscopic heterogeneities and cell surface polymers on microbial adhesion are elucidated and implications for practical environmental membrane processes are discussed.
Materials and Methods Ultrafiltration Membranes. Membranes used in this study were referred to as MX and EW (GE Osmonics, Minnetonka, MN). According to the manufacturer, EW is a polysulfone membrane and MX membrane is a modified polyacrylonitrile. The manufacturer provided data indicated molecular weight cutoffs of 60 and 70 kDa for EW and MX, respectively, at 98% retention of polysaccharides. Apparent hydraulic resistance was determined from the slope of pressure-flux data using electrolyte solutions to account for possible electroviscous effects on membrane permeability (30). Membrane surface (zeta) potential was determined from streaming potentials (Anton Paar EKA, Brookhaven Instrument Corp., Holtsville, NY) measured at pH 8.0 ( 0.1 for 10 and 50 mM NaCl solutions as previously described (31). Contact angles of deionized water, ethylene glycol, glycerol, and diiodomethane were measured on dried membrane samples mounted on glass slides with double-sided tape using a contact angle goniometer (VCA-1000, AST Products Inc., Billerica, MA). Surface VOL. 39, NO. 17, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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tensions of the probe liquids were taken from values reported previously (32). Microbial Cell Suspensions. Kang et al. (25) previously demonstrated that Saccharomyces cerevisiae (active dry yeast, Fleischmann, Inc.) served as excellent model microbial particles for deposition and adhesion studies; hence, S. cerevisiae were selected as model microbial particles for this study. Cell suspensions were prepared using the procedure described by Kang et al. (25), which enabled direct microscopic observation by light microscopy. Suspensions were prepared in concentrations of 25, 50, and 100 mg/L (as dry yeast), which corresponded to number concentrations of 2.5, 5.0, and 10 million cells per milliliter. The size and shape of the model microbial cells were previously determined using optical microscopy, scanning electron microscopy, and a particle size analyzer (25). Yeast cell size was confirmed in this study by dynamic light scattering (BI 90, Brookhaven Instrument Corp., Holtsville, NY). Electrophoretic mobility of cells was measured in 10 and 50 mM NaCl solutions using a particle electrophoresis instrument (ZetaPALS, Brookhaven Instrument Corp., Holtsville, NY) with solution pH adjusted to 8.0 ( 0.1 by addition of ∼6 × 10-5 M Na2CO3. Measured electrophoretic mobility was converted to zeta potential by the Helmholtz-Smoluchowski equation. Contact angles of various probe liquids on filtered lawns of yeast cells were obtained from literature values reported elsewhere (33). Surface Thermodynamic Parameters. Lifshitz-van der Waals (γ LW), electron donor (γ-), and electron acceptor (γ+) components of surface tension were determined from contact angles of three probe liquids with known surface tensions following the approach described by van Oss (12). Interfacial free energies arising from Lifshitz-van der Waals and Lewis acid-base interactions were determined from the computed surface tensions by an extension of the Dupre equation. The pertinent equations and examples relevant to microbial particle adhesion in environmental systems are available in numerous reviews (16-19). Interfacial Force Model. Lifshitz-van der Waals (LW), Lewis acid-base (AB), and electrostatic double layer (EL), i.e., “extended DLVO” (12) or “XDLVO”, interactions were combined with interfacial hydrodynamic forces of crossflow lift (CL) and permeation drag (PD) in the interfacial force model. The net interfacial force (FIF) was determined from
FIF ) FPD + FCL + FLW + FAB + FEL
(1)
FPD ) -(6πµavw)φH
(2)
FCL ) -81.2(Fwµγ˘ 03)1/2a3
(3)
FLW ) 2πh02∆GIw2LWa
( )(
(
) ( )( ) ( )
h0 - h FAB ) 2π∆GIw2ABa exp λAB FEL ) 4π
)
1 5.32h 1+ 2 λLW h
zF RT
2
γc γm h a exp λEL λEL
-1
(4)
(5)
(6)
where µ is the solution viscosity, a is particle radius, vw is the permeate water velocity, φH is the hydrodynamic correction factor (14), γ˘ 0 is the wall shear rate, F is the density of water, λLW ()100 nm (34)), λAB (=0.6 nm (32)), and λEL () [3.28 × 109 xcNaCl]-1, cNaCl in moles per liter) are characteristic decay lengths for LW, AB, and EL interactions in water, h0 represents the minimum interfacial separation distance (0.157 ( 0.009 nm (12)), is the dielectric permittivity of water ()78.5‚8.854 6462
9
× 10-12 C‚V-1 m-1), γm and γc are determined from the dimensionless surface (zeta) potentials (tanh [zFζ/4RT]), and ζc and ζm are zeta potentials of the cell and membrane. Direct Microscopic Observation. A previously described flow cell was used in direct microscopic observation experiments (25). The flow cell was mounted on the stage of a phase-contrast microscope (Olympus BX-51, Japan) with a 10× microscope objective lens. Images were acquired through a CCD camera with additional 5× magnification mounted on the microscope and uploaded in real time to a laboratory PC. Raw gray-scale images were later converted into black and white images. Direct pixel enumeration provided fractional surface coverage of cells on membranes during deposition, adsorption, and backpulsing experiments. In the experimental flow cell, the cross-flow Reynolds number, and wall shear rate were determined from 2000 and 6000 times the cross-flow velocity in m‚s-1, respectively. Deposition and Adsorption Experiments. In this study, “deposition” described the process by which cells accumulated at the membrane-solution interface under force of permeation drag, and “adsorption” described the process by which cells accumulated at the membrane-solution interface without permeation. All experiments were performed in solutions with pH adjusted to 8.0 ( 0.1 by addition of 6 × 10-5 M Na2CO3 and with temperature maintained at 20 ( 2 °C. Feed solution ionic strength was either 10 or 50 mM NaCl, and microbial cell concentration was 107‚mL-1 (∼100 mg/L) unless otherwise noted. All experiments were limited to durations of about 60 min or less because it was determined that model microbial cells (as prepared) began producing significant exopolymers between 60 and 120 min after adhering to the test membranes (35). In adsorption experiments, cells were fed without permeation into the flow cell at a cross-flow velocity of 0.015 m‚s-1 until the surface coverage did not increase for at least 20 min. The steadystate surface coverage was considered the extent of heterogeneous adsorption of cells on a membrane. Most experiments were repeated a minimum of three times. Cell Removal Experiments. In deposition experiments, permeation was stopped after a certain fractional coverage was attained. Adhered cells were removed by generating a permeate backpulse (flux reversal) or by inducing a crossflow velocity. Forward-reverse filtration cycles were repeated up to three times for backpulse experiments. In adsorption experiments (no flux), cross-flow velocity was increased from the starting velocity of 0.015 m‚s-1 sequentially in 1-min intervals to 0.025, 0.05, 0.125, 0.25, 0.50, and 1.0 m‚s-1 (1.0 m‚s-1 was the highest cross-flow that could be achieved in the experimental system). In preliminary experiments, cells were nearly instantly removed; no additional cell removal occurred beyond 30 s for backpulsing or beyond 1 min for cross-flow (35).
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Results and Discussion Physicochemical Properties of Microbes and Membranes. Although the surrogate microbial particles were slightly ellipsoidal with average major and minor axes of 4.9 and 4.5 µm, the spherical hydrodynamic diameter of 4.7 ( 0.2 µm (determined by light scattering) was used in all model calculations. Average values and standard deviations were determined for all measured physicochemical properties of cells and membranes (Table 1). All materials possessed significant negative charge, but streaming potential or electrophoretic mobility decreased (less negative) as ionic concentration increased. Microbial cells and MX had very low pure water contact angles (∼20 and ∼25), whereas the contact angle for EW was relatively high (∼80). The calculated solid surface tensions and zeta potentials provided additional insight into the chemical nature of cell and membrane surfaces (Table 2). For example, the LW
TABLE 1. Measured Physicochemical Properties EW MX yeastb a
I (mM)
E/∆p or U/Ea (×108)
θpure water
θglycerol
θdiiodomethane
θethylene glycol
10 50 10 50 10 50
-98.9 ( 5.9 -50.0 ( 3.6 -150 ( 3.6 -117 ( 2.9 -2.07 ( 0.15 -1.34 ( 0.07
79.0 ( 0.5
82.5 (0.4
32.2 ( 1.0
38.8 ( 5.9
24.7 ( 4.6
25.6 (0.6
12.7 ( 1.0
24.9 ( 2.1
20.7 ( 2.0
21.4 ( 1.8
45.5 ( 3.1
N/A ( N/A
E/∆p ) streaming potential (V‚Pa1-m-1); U/E ) electrophoretic mobility (m2 V-1 s-1). b Yeast cell contact angles taken from ref 33.
TABLE 2. Calculated Surface Energetic Parameters γ LW (mJ/m2)
γ+ (mJ/m2)
γ- (mJ/m2)
γAB (mJ/m2)
γTOT (mJ/m2)
I (mM)
ζ (mV)
EW
41.0 ( 0.3
0.4 ( 0.2
3.8 ( 0.5
2.5 ( 0.4
43.5 ( 0.5
MX
48.9 ( 0.2
0.9 ( 0.1
38.9 ( 2.0
12.9 ( 0.3
61.0 ( 0.3
yeast
32.1 ( 0.7
4.2 ( 0.2
44.4 ( 0.9
27.3 ( 0.7
59.4 ( 0.3
10 50 10 50 10 50
-16.0 ( 1.0 -8.1 ( 0.6 -24.2 ( 0.6 -19.0 ( 0.5 -26.5 ( 1.9 -17.1 ( 1.0
TABLE 3. Surface Free Energy and Deposition Rates ∆Gh)0 MX EW a
AB (mJ/m2)
LW (mJ/m2)
EL10mMa (mJ/m2)
EL50mMa (mJ/m2)
XDLVO10mM (mJ/m2)
XDLVO50mM (mJ/m2)
k(10mM) (µm/s)
k(50mM) (µm/s)
20.3 -4.5
-4.6 -3.5
4.6 3.1
2.3 1.0
20.3 -4.9
18.0 -7.0
1.7 2.4
2.0 2.5
∆Gh)0EL )ζmζc/λEL.
component of surface tension was rather small for the microbe compared to other, more comprehensive, studies (32, 36, 37) in which most polymer surfaces and microbial lawns exhibited γ LW close to ∼40 mJ‚m-2. The resulting cellwater-MX and cell-water-EW Hamaker constants were 3.3 × 10-21 and 4.4 × 10-21 J, respectively, which were small, but not unreasonable. Both membrane surfaces were nearly monopolar, but the MX surface exhibited strong electron donor (Lewis base) character. Microbial cells not only exhibited strong electron donor character but also possessed a significant (albeit small) electron acceptor component. This suggested AB attraction would be significant between the cell and EW, especially because of the low electron donor component of EW. MX and cells both had substantial electron donor components and fairly sizable zeta potentials, suggesting significant AB and EL repulsion. The relatively weak surface (zeta) potential of EW would result in weak electrostatic interactions with yeast cells. Average hydraulic resistances for EW and MX membranes were 5.47 × 109 and 1.06 × 109 Pa-s/m, respectively. Hydraulic resistance is an important characteristic with respect to microbial deposition and adhesion because it determines the permeate drag force on a microbial particle through the hydrodynamic correction factor, φH (14, 25). The correction factors at contact, φH(h ) 0) ) [(2/3Rma)+1.0722]1/2, for microbe-EW and microbe-MX pairs were 2930 and 1290, respectively. The interfacial drag force was ∼103 greater than in the bulk, but the greater hydraulic resistance (Rm) of EW increased the effective drag by ∼51/2 over that of MX. Interfacial free energies at contact were determined for AB, LW, and EL (at 10 and 50 mM) interactions between microbial cells and membranes (Table 3). The electrostatic double layer free energy at contact was estimated from the following group of parameters, ζcζm/λEL, giving the appropriate units of energy per unit area, mJ‚m2. The key difference between MX and EW was the AB interaction energy, which was significantly positive (hydrophilic repulsion) for the MX-microbe pair and significantly negative (hydrophobic
attraction) for the EW-microbe pair. The net interfacial free energy at 10 mM ionic strength (I), indicated as “XDLVO10mM”, is governed by the AB interaction energy because the LW and EL energies effectively cancel one another. At the higher ionic strength, the XDLVO interfacial free energy is reduced in proportion to the reduction of EL repulsion. Deposition rates on the membranes at both ionic strengths (determined from direct observation at 20 gfd forward flux) changed very little with ionic strength but were noticeably higher on EW, the more hydrophobic and less negatively charged membrane. Interfacial Forces and Critical Flux. Microbe-membrane interfacial forces were determined for dead-end filtration conditions (XDLVO + PD) and plotted in Figure 1. The model results shown in Figure 1(a) were based on the properties of EW and cells at 10 mM and pH 8 for permeate velocities of 0.0 µm‚s-1 (XDLVO only), 9.5 µm‚s-1 (20 gfd), 19 µm‚s-1 (40 gfd), 28.5 µm‚s-1 (60 gfd), and 38 µm‚s-1 (80 gfd). In absence of permeation, the XDLVO force profiles indicated a longrange attractive (negative) force of -0.4 pN at 30 nm, followed by strong repulsion beginning at 25 nm separation. A critical flux should not exert a drag force larger than the magnitude of this XDLVO repulsion, which was +0.6 nN at its peak. In Figure 1(a), the presence of a negative (attractive) interaction force at long-range suggested that at all forward fluxes cells should accumulate near the membrane-solution interface. However, the positive (repulsive) peak force at 20, 40, and 60 gfd implied that physical adhesion should not occur, while at 80 gfd, cells were strongly attracted to EW from long-range up to the point of contact due to the excessive permeate drag force. Based on the average measured physicochemical properties of the microbial cells and EW, the critical flux was determined to be ∼3.2 × 10-5 m‚s-1 (∼70 gfd). If the hydrodynamic correction factor, φH, was not considered in the permeate drag force, the critical flux predicted for EW at 10 mM would be greater than ∼10-3 m‚s-1 or ∼104 gfd. Past experiments employing the same model microbial particles and the same membranes demVOL. 39, NO. 17, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 1. Interfacial forces between the surrogate microbial particle and EW membrane (a) with forward fluxes of 0 gfd (XDLVO forces only), 20 gfd, 40 gfd, 60 gfd, and 80 gfd and (b) with forward flux of 20 gfd considering maximum variation in measured physicochemical properties. Model predictions assumed no cross-flow, 10 mM NaCl, pH 8.0, and 20 °C. onstrated about 70-80% of cell deposition to be reversible when deposition occurred at fluxes below about 40 gfd (25, 35). Therefore, it was determined that strict interpretation of interfacial force profiles based on average physicochemical properties of cells and membranes was not appropriate, especially considering the potential deviations in physicochemical parameters determined from the macroscopic techniques employed (i.e., contact angle, streaming potential, particle electrophoresis, light scattering). The maximum and minimum physicochemical parameters were estimated as three times the standard deviation of measured values added to or subtracted from the average value. The computed maxima and minima were in excess of all experimentally observed data. The analysis did not consider variations in membrane resistance or microbial particle size because model predictions were relatively insensitive to these parameters. Sensitivity of interaction forces to the maximum deviations of zeta potentials and surface tensions for EW and microbial cells was assessed for all fluxes below 70 gfd. However, minimum, average, and maximum were plotted in Figure 1(b) for 20 gfd only. The average 20 gfd-10 mM force profile shown here was computed identically to the 20 gfd profile shown in Figure 1(a). The maximum and minimum interfacial force profiles represented the “most repulsive” and “most attractive” combinations of physicochemical parameters, respectively. A membrane was considered “macroscopically attractive” to a microbial cell if the sum of all forces was (on average) entirely negative on approach. Hence, 80 gfd was considered a “supercritical flux” because the force profile indicated macroscopic attraction. A membrane was considered “macroscopically repulsive” to a microbial cell if the minimum (least repulsive) interfacial force profile remained significantly positive upon close approach. Therefore, the sensitivity analysis implied that 20 gfd was a “subcritical flux” because even the most attractive combination of measured surface properties appeared macroscopically repulsive. However, the presence of secondary XDLVO attractive forces at long-range (predominantly due to LW attraction) virtually guaranteed some microbial adhesionseven in absence of permeation. Removal by Cross-Flow and Backpulsing. Microbial particles were deposited on EW at 80 gfd (macroscopically attractive, supercritical flux conditions) and at 20 gfd (macroscopically repulsive, subcritical flux conditions) with all other experimental conditions identical to those modeled in Figure 1. The forward filtration duration was 5 and 14 min at 80 and 20 gfd, respectively, to produce the same fractional 6464
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TABLE 4. Summary of Cross-Flow and Backpulse Removal vBP (µm.s-1)
JBP (gfd)
FBP (nN)
9.4 18.9 28.3 37.7 75.4 113.2
20 40 60 80 160 240
0.451 0.901 1.352 1.803 3.605 5.408
u0 (mm.s-1)
Re (-)
FCL (nN)
25 50 125 250 500 1000
50 100 250 500 1000 2000
0.002 0.005 0.022 0.061 0.173 0.490
a
Backpulsea Jfor ) 80 gfd (% removed) (%)
Jfor ) 20 gfd (% removed) (%)
36 49 65 82 84 90 Cross-Flow Jfor ) 80 gfd (% removed) (%)
94 95 96
Jfor ) 20 gfd (% removed) (%)
18 25 35 42 71 89
33 42 64 68 91 98
Removal values are for the first backpulse cycle only.
surface coverage (∼40%). Forward permeation was stopped at the specified time, and adhered cells were removed either by generating a permeate backpulse or by inducing a crossflow as described above. Fractional surface coverage data shown in Table 4 were determined by direct microscopic observation. The backpulse and cross-flow removal efficacy is reported in terms of percent removal of deposited cells. In each experiment the fractional surface coverage prior to initiating the removal mechanism was 40%; hence, 50% removal indicated 20% of the membrane surface remained covered with cells. In the top half of Table 4, the backpulse intensity was reported in terms of a velocity in µm‚s-1, vBP, a flux in gfd, JBP, and a force in nN, FBP. In the bottom half of the table, cross-flow intensity was reported in terms of the cross-flow velocity in mm‚s-1, u0, Reynolds number, Re, and lift force in nN, FCL. Cells deposited at subcritical flux conditions (20 gfd) were readily removed by low intensity backpulsing but were removed comparably only at the highest cross-flow (1.0 m‚s-1). Cells deposited at supercritical flux conditions (80 gfd) were also removed, but to a much smaller extent. In general, permeate backpulsing was better at removing cells than the highest laminar cross-flows tested. For cells deposited at subcritical flux conditions, the relative insensitivity of cell removal to permeate backpulse intensity
FIGURE 2. Removal by backpulsing (a) at subcritical flux conditions (20 gfd) and different forward filtration durations and (b) at supercritical flux conditions (80 gfd) and different backpulse intensities. Common experimental conditions were no cross-flow, 10 mM NaCl, pH 8.0 ( 0.1, and 20 ( 2 °C. suggested that few cells were strongly adhered to the membrane surface. Whereas at supercritical flux conditions, cell removal was proportional to permeate backpulse intensity. In fact, the highest backpulse intensity used exceeded the manufacturer’s recommended limits for the membrane and actually caused visible physical damage (cracks and tears) to the membrane active surface. Microbial Adhesion over Repeated Backpulse Cycles. In Figure 2, fractional surface coverage data were plotted to compare the impacts of (a) forward filtration duration (tfor ) 5 and 14 min at Jfor ) 20 gfd with Jrev ) 20, 40, and 60 gfd) and (b) forward filtration and backpulse intensity (Jfor ) 80 gfd with Jrev ) 20, 40, 60, 80, 160, and 240 gfd) over repeated backpulse cycles. Cells were almost completely removed even at the lowest backpulse intensity when deposition occurred below the critical flux (Figure 2(a)). A small fraction of cells could not be removed, but this fraction did not increase over several repeated forward-reverse filtration cycles. The same result was found over longer times as well. The amount of irreversibly bound cells was on average slightly higher for the longer exposure time. However, the fractional coverage fell within the experimental error at both forward filtration durations. At significantly longer exposure times, biological factors would become important and more irreversible adhesion would be expected (38). At supercritical forward flux conditions (Figure 2(b)) and low backpulse intensities (20, 40, 60 gfd), about 10-15% of the surface remained covered with cells after the first backpulse cycle. Subsequent forward and reverse filtration cycles showed increasing accumulation, which suggests a significant deposit layer would accumulate as the process continued. At supercritical forward flux conditions and high backpulse intensities (80, 160, 240 gfd), more cells were removed per backpulse cycle. Only two forward-reverse cycles were plotted because the high-intensity backpulses ruptured the membrane by the third backpulse cycle, but it was clear that cells would continue to accumulate if operation continued. Experimental deposition and removal data were qualitatively consistent with interaction force model predictions (Figure 1), which suggested that most cells would deposit reversibly at subcritical forward fluxes and that most cells would deposit irreversibly at supercritical forward fluxes. Removal of cells deposited at 20 gfd was not significantly affected by backpulse intensity. However, removal of cells deposited at 80 gfd was significantly different depending on the backpulse intensity employed. At the lower backpulse intensities, about 10-15% of fractional surface coverage accumulated with each forward-reverse filtration cycle. At
the higher backpulse intensities, more cells were removed, but approximately 4-8% of fractional surface coverage accumulated per filtration cycle. Other Factors Influencing Microbial Adhesion. Backpulse removal experiments were performed on EW and MX membranes at subcritical flux conditions (20 gfd) with all other experimental conditions the same as those modeled in Figure 1. Fractional surface coverage data plotted in Figure 3(a) indicate slower deposition on MX, which is consistent with previously reported results comparing EW and MX (25). Average interfacial force profiles plotted in Figure 3(b) suggested cell deposition was largely reversible for both membranes. The infinite repulsion predicted for the MX membrane at small separations (due to “hydrophilic” AB repulsion) implied that deposited cells should be completely removed, but a small fraction surface coverage remained after each backpulse cycle. Additional deposition and removal data plotted in Figure 4 were obtained using 20 gfd forward fluxes on EW with solution ionic strengths of 10 and 50 mM NaCl. Deposition and removal data are shown in Figure 4(a), while the average interfacial force profiles are provided in Figure 4(b). The model predicted attraction between yeast cells and EW up to the point of contact at 50 mM. The increase in solution ionic strength reduced the zeta potentials of cells and membranes (see Table 2) and suppressed electrical double layer repulsion. Figure 4(b) suggested macroscopic attraction; however, deposition rates at both ionic strengths were practically identical. These data were consistent with previously reported results (25), but the unexpected result was that the fractional coverage remaining after each forwardreverse filtration cycle were practically identical. On the Potential Role of Microscopic Heterogeneities. Microscopic heterogeneities have been implicated in many deposition and adhesion studies (39-41). We envisioned the role of microscopic heterogeneities to be similar to that of specific interactions in cellular- or enzyme-substrate systems, which usually are only possible after the more prevalent macroscopic repulsion has been overcome (33). For example, it is well-known that hydrophilic proteins adsorb (albeit to a moderate degree) onto macroscopically repulsive surfaces such as glass (12, 42). In such cases, macroscopic repulsion can be locally overcome by a microscopic attraction between, for example, a protein chain with a small radius of curvature and a discrete attractor site on the glass surface that has a positive charge, perhaps due to specific adsorption of a cation. Further, numerical simulations have shown that nanometer-scale membrane surface roughness may significantly reduce macroscopic DLVO repulsion and create locally VOL. 39, NO. 17, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 3. Impact of membrane properties on (a) cell removal by backpulsing and (b) interfacial forces. Cells were deposited with no cross-flow at 20 gfd onto EW and MX at 10 mM NaCl, pH 8.0 ( 0.1, and 20 ( 2 °C.
FIGURE 4. Impact of solution ionic strength on (a) cell removal by backpulsing and (b) interfacial forces. Cells were deposited with no cross-flow at 20 gfd onto EW at 10 and 50 mM NaCl, pH 8.0 ( 0.1, and 20 ( 2 °C. Error bars omitted to avoid cluttering the figure.
FIGURE 5. Fractional surface coverage due to heterogeneous adsorption sites for yeast cells on (a) EW and (b) MX membranes after 60 min and on (c) EW and (d) MX for increasing cross-flow velocities. Other conditions are identical to Figure 1 at 20 gfd and 10 mM. attractive sites for spherical colloids approaching polymeric membrane surfaces (43). Both physical and chemical microscopic surface heterogeneities may have played a similar role in facilitating some microbial adhesion to the membranes in this study. Therefore, heterogeneous adsorption experiments were conducted for macroscopically repulsive conditions (i.e., at 10 mM NaCl) to establish the potential impact of microscopic surface heterogeneities on microbial deposition. Fractional surface coverage data were determined by direct observation and plotted in Figure 5 for (a) EW and (b) MX. Adsorption 6466
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isotherms rapidly increased and reached equilibrium values between 1 and 4% for MX and EW. After equilibrium was reached (at ∼60 min in Figure 5), cross-flow velocity was incrementally increased in 1-min intervals from the starting value of 0.015 m‚s-1 up to 1.0 m‚s-1. The cross-flow lift force predicted for a cross-flow velocity of 0.015 m‚s-1 was about +0.9 pN, while the maximum secondary XDLVO attractive force was about -0.5 pN. Thus, the adhesion of cells at higher velocities was attributed to microscopic heterogeneities. At the highest cross-flow, the lift force was on the order of 500 pN, which was orders of
FIGURE 6. Illustration of proposed roles for interfacial hydrodynamic, extended DLVO, and steric interactions on microbial adhesion to membranes. Hairlike structures drawn on cartoon microbial particles intended to represent cell surface macromolecules. Not drawn to scale. magnitude larger than secondary attractive forces predicted for both membrane surfacesseven considering experimental deviations in macroscopic surface properties. The relatively weak adhesion force of bound cells was qualitatively consistent with locally attractive microscopic heterogeneities that act over very small surface areas. Also, the fractional coverage of irreversibly bound cells was consistent with that observed in backpulse removal experiments. At subcritical flux conditions, cells may have only physically contacted membranes via microscopic heterogeneities. Hence, once all available heterogeneous adsorption sites on the membrane surface were occupied no additional cells adhesion occurred. When operating at supercritical fluxes, cells presumably made physical contact with the membrane surface and may have become adhered by macroscopic as well as microscopic attractive interactions. On the Potential Role of Steric Repulsion. Model predictions suggested physical adhesion would occur for cells on EW at 80 gfd and 10 mM as well as at 20 gfd and 50 mM. Cells deposited above the critical flux (at 80 gfd) were strongly adhered to the membrane and were difficult to remove even when intense backpulsing was applied. However, cells deposited at 20 gfd and 50 mM were not strongly adhered to the membrane, and only the fractional surface coverage expected from heterogeneous adsorption remained even when relatively weak backpulsing was applied. The apparent disparity between model predictions and experimental results might be explained by closer investigation into the model microbial particle cell wall structure. Yeast cells, like more common aquatic bacteria (especially gram negative strains), are thought to have about 90% of their surfaces coated with polysaccharide-like macromolecules (9). Mercier-Bonin et al. (44) reported that yeast cell walls have “an internal skeletal layer and an outer protein layer”. The skeletal layer, which provides the shape and strength of the wall, consists mostly of β-glucans and chitin. The outer layer, which determines the physicochemical properties, consists of heavily glycosylated manno-proteins. Many microbial adhesion studies have implicated the potential role of steric interactions arising from the presence of cell surface macromolecules (16, 24, 45-47). Some of these studies employed variations of the (physical) steric repulsion model originally proposed by de Gennes (13), which suggests an exponential decay such as FST ) A‚exp(-h/B), where h indicates separation distance between the bare cell wall (i.e.,
the base of terminally anchored polymer chains) and the substrate surface. Application of this steric model remains limited for microbial cells because the parameters A and B relate to the density and length of cell surface macromolecules, respectively. The steric model is thus largely empirical because these parameters cannot be predicted a priori. In addition, steric interactions are not directly additive with XDLVO interactions, and the model ignores the possibility of bridging between cell surface polymers and a membrane surface, for example, via plurivalent cations such as calcium (47). Despite the apparent limitations of the best available model of steric interactions, it is clear that such interactions play a critical and poorly understood role in microbial adhesion to membranes. Following the general conceptual framework proposed by Rijnaarts et al. (47), the illustrations in Figure 6 were drawn to depict the potential role of steric interactions in microbial adhesion to membranes. The dashed lines in the force profile illustrations were drawn to represent the balance between permeate drag and XDLVO interactions. The solid lines were drawn to represent the same interactions also taking steric repulsion into account. When permeate drag was small compared to cellmembrane XDLVO repulsion (Figure 6a), an attractive (negative) force at long-range caused cells to accumulate near the membrane-solution interface, but strong EL or AB repulsion prevented cells from making physical contact with the membrane. Hence, on average, steric forces did not need to be considered. Microbial deposition at subcritical flux conditions (20 gfd, 10 mM) was consistent with this scenario. When permeate drag and XDLVO forces were of approximately equal magnitudes (Figure 6b), cell surface macromolecules likely contacted the membrane at a few discrete points but not across a broad enough interfacial area to generate strong adhesion forces. The weak adhesion of cells deposited at 20 gfd and 50 mM were conceptually consistent with this scenario. Cell-membrane contact at discrete points may also explain (in part) the weaker adhesion that occurred in heterogeneous adsorption experiments for both membranes. When permeate drag was in great excess of XDLVO forces (Figure 6c), the cell surface polymers contacted the membrane surface at numerous locations or possibly across a broad interfacial region, thus, creating a strong adhesion force. This scenario probably relates to microbial deposition VOL. 39, NO. 17, 2005 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 7. Transmembrane pressure data acquired with the direct observation flow cell for subcritical (20 gfd) and supercritical (80 gfd) forward flux conditions. In (a) cells were deposited without backpulsing and in (b) cells were deposited with backpulsing. Common experimental conditions were no cross-flow, 10 mM NaCl, pH 8.0 ( 0.1, and 20 ( 2 °C. at supercritical flux conditions (80 gfd). Established theories of contact adhesion have shown the adhesion force between two surfaces to be proportional to the total interfacial free energy (at contact) by a length indicative of the contact area (i.e., contact radius) (48, 49). Therefore, at supercritical flux conditions, the drag force imparted by permeation probably determined the extent of contact between cells and membrane and, hence, the magnitude of the adhesion force. Implications for Fouling Control in Environmental Membrane Processes. The direct observation flow cell possessed pressure transducers in feed, retentate, and permeate ports. Therefore, in all experiments transmembrane pressure was monitored and recorded in real time along with microscopic images of adhered cells. Representative transmembrane pressure data obtained at supercritical and subcritical flux conditions acquired during deposition and backpulse removal experiments on EW were plotted in Figure 7. In the absence of backpulsing (Figure 7(a)), operating pressure increased moderately over 3 h at subcritical flux operation (20 gfd), whereas there was a dramatic increase in operating pressure at supercritical flux conditions (80 gfd). However, direct observation confirmed that there was a massive amount of cell accumulation on both membranes. Based on the rate of surface coverage determined for the experimental conditions employed (40% in 14 min at 20 gfd and 40% in 5 min at 80 gfd, see Figure 2), about 4.5 layers of cells accumulated in 160 min at 20 gfd and 3.7 layers of cells accumulated in 46.5 min at 80 gfd. When backpulsing was applied (Figure 7(b)), the pressure drop associated with cell accumulation at supercritical flux conditions (80 gfd) was significantly smaller and there was no apparent pressure drop for cells deposited at subcritical flux conditions (20 gfd). The pressure data reported in Figure 7(b) correspond to the fractional surface coverage data for “14 min exposure” at 20 gfd in Figure 2(a) and “low intensity” backpulsing at 80 gfd in Figure 2(b). The pressure increase at supercritical flux (80 gfd) was consistent with the increasing cell surface coverage over the three forward-reverse filtration cycles shown. In this study, operating conditions that lead to increasing cell surface coverage over repeated forward-reverse filtration cycles were identified a priori as supercritical flux conditions. Operating conditions for which there was no observed accumulation of cells over repeated forward-reverse filtration cycles were identified a priori as subcritical flux conditions. For the model system studied, direct microscopic observation and the interfacial force model provided valuable insight into mechanisms of microbial adhesion to membranes. Although our understanding of microbial adhesion (at least 6468
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for very simple model systems) has improved, significant complexities such as steric interactions and microscopic heterogeneities remain difficult to quantify. An improved understanding of these phenomena may hold the key to developing effective means of biofouling control in practical environmental systems.
Acknowledgments Financial support for this research was provided by the California Energy Commission through the Desalination Research Innovation Partnership (DRIP), which is managed by the Metropolitan Water District of Southern California (MWDSC). The authors thank Dr. Steven Kloos at GE Osmonics for supplying samples of EW and MX membranes and Prof. Marc A. Deshusses at the University of California, Riverside for providing access to the microscope used for direct observation experiments.
Literature Cited (1) Clark, M. M.; Allgeier, S.; Amy, G.; Chellam, S.; DiGiano, F.; Elimelech, M.; Freeman, S.; Jacangelo, J.; Jones, K.; Laine, J. M.; Lozier, J.; Marinas, B.; Riley, R.; Taylor, J.; Thompson, M.; Vickers, J.; Wiesner, M.; Zander, A. Committee report: Membrane processes. J. Am. Water Work Assoc. 1998, 90, 91-105. (2) Baker, J. S.; Dudley, L. Y. Biofouling in membrane systems-A review. Desalination 1998, 118, 81-90. (3) Flemming, H. C.; Schaule, G.; Griebe, T.; Schmitt, J.; Tamachkiarowa, A. Biofouling - the Achilles heel of membrane processes. Desalination 1997, 113, 215-225. (4) Flemming, H. C. Biofouling in water systems - cases, causes and countermeasures. Appl. Microbiol. Biotechnol. 2002, 59, 629-640. (5) Ridgway, H.; Ishida, K.; Rodriguez, G.; Safarik, J.; Knoell, T.; Bold, R. In Biofilms; Academic Press Inc.: San Diego, 1999; Vol. 310, pp 463-494. (6) Characklis, W. G.; Marshall, K. G. Biofilms; John Wiley & Sons: New York, 1990. (7) Belfort, G.; Davis, R. H.; Zydney, A. L. The Behavior of Suspensions and Macromolecular Solutions in Cross-Flow Microfiltration. J. Membr. Sci. 1994, 96, 1-58. (8) Davis, R. H. Modeling of fouling of cross-flow microfiltration membranes. Sep. Purif. Methods 1992, 21, 75-126. (9) Madigan, M. T.; Martinko, J. M.; Parker, J. Brock Biology of Microorganisms, 10th ed.; Prentice-Hall: Englewood Cliffs, 2003. (10) Verwey, E. J. W.; Overbeek, J. T. G. Theory of the Stability of Lyophobic Colloids; Elsevier: Amsterdam, 1948. (11) Derjaguin, B. V.; Landau, L. D. Theory of the stability of strongly charged lyophobic sols and of the adhesion of strongly charged particles in solutions of electrolytes. Acta Physicochim. URSS 1941, 14, 733-762. (12) van Oss, C. J. Acid-Base Interfacial Interactions in AqueousMedia. Colloids Surf., A 1993, 78, 1-49. (13) de Gennes, P. G. Polymers at an interface; A simplified view. Adv. Colloid Interface Sci. 1987, 27, 189-209.
(14) Goren, S. L. The hydrodynamic force resisting the approach of a sphere to a plane permeable wall. J. Colloid Interface Sci. 1973, 69, 78-85. (15) Saffman, P. G. Lift on a small sphere in a slow shear flow. J. Fluid Mech. 1965, 22, 385-400. (16) Oliveira, R. Understanding adhesion: A means for preventing fouling. Exp. Therm. Fluid Sci. 1997, 14, 316-322. (17) Hermansson, M. The DLVO theory in microbial adhesion. Colloids Surf., B 1999, 14, 105-119. (18) Bos, R.; van der Mei, H. C.; Busscher, H. J. Physico-chemistry of initial microbial adhesive interactions - its mechanisms and methods for study. FEMS Microbiol. Rev. 1999, 23, 179-230. (19) Grasso, D.; Subramaniam, K.; Butkus, M.; Strevett, K.; Bergendahl, J. A review of non-DLVO interactions in environmental colloidal systems. Rev. Environ. Sci. Biotech. 2002, 1, 17-38. (20) Bergendahl, J.; Grasso, D. Prediction of colloid detachment in a model porous media: Thermodynamics. AIChE J. 1999, 45, 475-484. (21) Bergendahl, J.; Grasso, D. Prediction of colloid detachment in a model porous media: hydrodynamics. Chem. Eng. Sci. 2000, 55, 1523-1532. (22) Brant, J. A.; Childress, A. E. Assessing short-range membranecolloid interactions using surface energetics. J. Membr. Sci. 2002, 203, 257-273. (23) Chen, G.; Strevett, K. A. Impact of surface thermodynamics on bacterial transport. Environ. Microbiol. 2001, 3, 237-245. (24) Camesano, T. A.; Logan, B. E. Probing bacterial electrosteric interactions using atomic force microscopy. Environ. Sci. Technol. 2000, 34, 3354-3362. (25) Kang, S.; Subramani, A.; Hoek, E. M. V.; Deshusses, M. D.; Matsumoto, M. R. Direct microscopic observation of biofouling in cross-flow membrane filtration: Mechanisms of deposition and release. J. Membr. Sci. 2004, 244, 151-165. (26) Field, R. W.; Wu, D.; Howell, J. A.; Gupta, B. B. Critical Flux Concept for Microfiltration Fouling. J. Membr. Sci. 1995, 100, 259-272. (27) Howell, J. A. Subcritical Flux Operation of Microfiltration. J. Membr. Sci. 1995, 107, 165-171. (28) Altmann, J.; Ripperger, S. Particle deposition and layer formation at the cross-flow microfiltration. J. Membr. Sci. 1997, 124, 119128. (29) Bacchin, P.; Aimar, P.; Sanchez, V. Model for Colloidal Fouling of Membranes. AIChE J. 1995, 41, 368-376. (30) Huisman, I. H.; Pradanos, P.; Hernandez, A. Electrokinetic characterisation of ultrafiltration membranes by streaming potential, electroviscous effect, and salt retention. J. Membr. Sci. 2000, 178, 55-64. (31) Walker, S. L.; Bhattacharjee, S.; Hoek, E. M. V.; Elimelech, M. A novel asymmetric clamping cell for measuring streaming potential of flat surfaces. Langmuir 2002, 18, 2193. (32) van Oss, C. J. Interfacial forces in aqueous media; Marcel Dekker: New York, 1994. (33) Nakari-Setala, T.; Azeredo, J.; Henriques, M.; Oliveira, R.; Teixeira, J.; Linder, M.; Penttila, M. Expression of a fungal hydrophobin in the Saccharomyces cerevisiae cell wall: Effect on cell surface
(34) (35) (36) (37) (38) (39) (40) (41) (42)
(43) (44) (45) (46) (47) (48) (49)
properties and immobilization. Appl. Environ. Microbiol. 2002, 68, 3385-3391. Elimelech, M.; Gregory, J.; Jia, X.; Williams, R. A. Particle Deposition & Aggregation: Measurement, Modeling and Simulation; Butterworth-Heinemann: Woburn, MA, 1995. Wang, S. M.S. Thesis in Chemical and Environmental Engineering, University of California, Riverside, 2004. van der Mei, H. C.; Bos, R.; Busscher, H. J. A reference guide to microbial cell surface hydrophobicity based on contact angles. Colloids Surf., B 1998, 11, 213-221. Sharma, P. K.; Rao, K. H. Analysis of different approaches for evaluation of surface energy of microbial cells by contact angle goniometry. Adv. Colloid Interface Sci. 2002, 98, 341-463. Bowen, W. R.; Lovitt, R. W.; Wright, C. J. Atomic force microscopy study of the adhesion of Saccharomyces cerevisiae. J. Colloid Interface Sci. 2001, 237, 54-61. Adamczyk, Z.; Siwek, B.; Musial, E. Latex particle adsorption at heterogeneous surfaces. Colloids Surf., A 2003, 214, 219-229. Brant, J. A.; Childress, A. E. Colloidal adhesion to hydrophilic membrane surfaces. J. Membr. Sci. 2004, 241, 235-248. Ryan, J. N.; Elimelech, M. Colloid mobilization and transport in groundwater. Colloids Surf., A 1996, 107, 1-56. van Oss, C. J.; Docoslis, A.; Wu, W.; Giese, R. F. Influence of macroscopic and microscopic interactions on kinetic rate constants: I. Role of the extended DLVO theory in determining the kinetic adsorption constant of proteins in aqueous media, using von Smoluchowski’s approach. Colloids Surf., B 1999, 14, 99-104. Hoek, E. M. V.; Bhattacharjee, S.; Elimelech, M. Effect of membrane surface roughness on colloid-membrane DLVO interactions. Langmuir 2003, 19, 4836-4847. Mercier-Bonin, M.; Ouazzani, K.; Schmitz, P.; Lorthois, S. Study of bioadhesion on a flat plate with a yeast/glass model system. J. Colloid Interface Sci. 2004, 271, 342-350. Abu-Lail, N. I.; Camesano, T. A. Role of lipopolysaccharides in the adhesion, retention, and transport of Escherichia coli JM109. Environ. Sci. Technol. 2003, 37, 2173-2183. Ong, Y. L.; Razatos, A.; Georgiou, G.; Sharma, M. M. Adhesion forces between E-coli bacteria and biomaterial surfaces. Langmuir 1999, 15, 2719-2725. Rijnaarts, H. H. M.; Norde, W.; Lyklema, J.; Zehnder, A. J. B. DLVO and steric contributions to bacterial deposition in media of different ionic strengths. Colloids Surf., B 1999, 14, 179-195. Derjaguin, B. V.; Muller, V. M.; Toporov, Y. P. Effect of Contact Deformations on Adhesion of Particles. J. Colloid Interface Sci. 1975, 53, 314-326. Johnson, K. L.; Kendall, K.; Roberts, A. D. Surface energy and the contact of elastic solids. Proc. R. Soc. London, Ser. A 1971, 324, 301-313.
Received for review January 28, 2005. Revised manuscript received May 16, 2005. Accepted May 21, 2005. ES050188S
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