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Biofouling in forward osmosis and reverse osmosis: Measurements and mechanisms
Author’s Accepted Manuscript Biofouling in forward osmosis and reverse osmosis: Measurements and mechanisms Sarah E. Kwan, Edo Bar-Zeev, Menachem Elimelech www.elsevier.com/locate/memsci
PII: DOI: Reference:
S0376-7388(15)30055-7 http://dx.doi.org/10.1016/j.memsci.2015.07.027 MEMSCI13846
To appear in: Journal of Membrane Science Received date: 2 December 2014 Revised date: 13 July 2015 Accepted date: 14 July 2015 Cite this article as: Sarah E. Kwan, Edo Bar-Zeev and Menachem Elimelech, Biofouling in forward osmosis and reverse osmosis: Measurements and m e c h a n i s m s , Journal of Membrane Science, http://dx.doi.org/10.1016/j.memsci.2015.07.027 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Biofouling in forward osmosis and reverse osmosis: Measurements and mechanisms
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Journal of Membrane Science
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Revised: July 12, 2015
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Sarah E. Kwan+, Edo Bar-Zeev+, and Menachem Elimelech*
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Department of Chemical and Environmental Engineering, Yale University, P.O. Box 208286 New Haven, Connecticut 06520-8286, USA
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+
Contribute equally to this work
* Corresponding author: Menachem Elimelech, Tel: +1 (203) 432-2789; Fax: +1 (203) 432-4387 email: [email protected]
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Abstract
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We have investigated biofouling behavior in forward osmosis (FO) and reverse osmosis (RO)
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membrane systems. Analysis of these two systems was done via comparison of biofilm structure
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and membrane permeate water flux decline. Experiments were performed using a model
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wastewater solution inoculated with Pseudomonas aeruginosa in laboratory-scale FO and RO
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test cells. For a meaningful comparison, biofouling runs were carried out using the same thin-
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film composite polyamide FO membrane, identical hydrodynamic conditions, and an initial
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permeate water flux of 19 L m-2 hr-1. Water flux decline after 600 mL of cumulative permeate
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volume (or ~17-18 hours of fouling) was significantly lower in FO (~10%) compared to RO
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(~30%). FO biofilms grew in a loosely organized thick layer, with eminent mushroom-shaped
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structures, while RO biofilms grew in tightly organized mats encased in larger amounts of
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extracellular polymeric substances (EPS) per cell. The more compact biofilms in RO induced
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greater biofilm-enhanced osmotic pressure and hydraulic resistance to water flow compared to
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FO, which resulted in higher flux decline. We attribute the differences in biofouling behaviors in
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FO and RO to the different driving forces: osmotic pressure in FO and hydraulic pressure in RO.
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Keywords: Biofouling; Forward osmosis; Reverse osmosis; Compaction; Hydraulic pressure
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Graphic abstract
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1. Introduction
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Membrane-based technologies play an important role in various separation processes at the
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water-energy nexus [1,2]. Forward osmosis (FO) is an emerging membrane technology within
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this nexus that has potential applications in desalination [3], wastewater treatment/reuse [4], and
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treatment of various industrial wastes [3–5]. However, the performance of FO, like all
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membrane-based processes, is hampered by biofouling, an obstruction that increases operational
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costs and shortens membrane life.
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Membrane fouling can be classified into four main categories organic, inorganic,
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colloidal, and microbial (biofouling) with biofouling being the most ubiquitous and
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recalcitrant [6,7]. Membrane biofouling is manifested as a complex sessile microbial community
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permanently attached to the surface by gel-like, self-produced extracellular polymeric substances
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(EPS) [8–10]. This microbial cake layer features a multicellular architecture of live and dead
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cells encased in EPS, which is primarily composed of polysaccharides and proteins [11]. Once
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established, biofilms are notoriously resistant to removal by treatment with oxidizing agents,
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biocides, or antibiotics due to the protection provided by the EPS matrix [11].
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Previous FO fouling studies have focused on organic [12–14] and colloidal fouling [15,16].
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These studies revealed that fouling in FO is less detrimental to membrane performance than in
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reverse osmosis (RO) [3,13,17]. Furthermore, in addition to diminished adverse fouling effects,
3
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fouling in FO was found to be more reversible than in RO [15]. Studies have attributed the
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differences in the impact of fouling on membrane performance in FO compared to RO to lack of
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applied hydraulic pressure in FO [3,13,17].
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Our understanding of biofouling propensity and characteristics in FO is limited [18,19],
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especially in comparison to RO. To date, only one study compared biofilms developed in FO and
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RO systems [20]. Biofouling in FO was found to form a loose film and induced less permeate
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water flux decline than RO [20]. However, the mechanisms responsible for the differences in
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biofilm characteristics and the resulting effect on FO performance compared to RO are still not
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well understood.
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In this paper, we report a direct comparison of biofouling behavior in FO and RO
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configurations, conducted by analyzing permeate water flux decline and relating system
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performance to biofilm characteristics. Our results shed new light on biofilm structural
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composition in both FO and RO systems and the relationship between permeate water flux
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decline and biofouling characteristics. Moreover, we provide insights into the mechanisms of
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biofouling in osmotically driven FO and hydraulic-pressure driven RO systems.
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2. Materials and methods
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2.1. Bacteria strain and growth conditions
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A Pseudomonas aeruginosa (ATCC® 27853TM) monoculture was used as a model biofilm
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strain in all biofouling experiments. Approximately 250 mL of Luria-Bertani (LB, BD
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Biosciences) broth were inoculated with a fresh single colony of P. aeruginosa and allowed to
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grow overnight in a shaking incubator at 37 C. Sterile, model wastewater was freshly prepared
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for each biofouling run, as described elsewhere [21]. Wastewater was prepared by supplementing
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deionized (DI) water (Nano Pure II, Barnstead, Dubuque, IA) with 8.0 mM NaCl, 0.15 mM
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MgSO4, 0.5 mM NaHCO3, 0.2 mM CaCl2, 0.2 mM KH2PO4, 0.4 mM NH4Cl, and 0.6 mM
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Na3C6H5O7 (sodium citrate) as a carbon source. All components were individually dissolved in
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~60 mL of DI water and sterilized in an autoclave, then added to sterile DI water to prepare a 10
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L wastewater solution batch. The final solution pH and conductivity were 7.6 ± 0.5 and 1142 ±
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50 μS cm-1, respectively, with a calculated ionic strength of 15.9 mM.
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2.2. Thin-film composite membrane 4
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A commercial thin film composite (TFC) FO membrane (Hydration Technology Innovations,
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Albany, OR, USA) was used as a model membrane for biofouling experiments in both FO and
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RO. The membrane was received as a flat sheet and stored at 4 C in DI water. Prior to use, the
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membrane was wetted in a 25% isopropyl alcohol solution for 20 min, and then rinsed for 10
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minutes in DI water three times. Intrinsic membrane transport properties were determined by
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using the method of Tiraferri et al. [22], resulting in water permeability coefficient, A, of 1.08 L
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m-2 hr-1 bar-1, salt (NaCl) permeability coefficient, B, of 0.603 L m-2 hr-1, and structural
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parameter, S, of 419 μm. Reverse salt flux through the membrane was experimentally determined
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according to the procedure outlined by Yong et al. [23].
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2.3. Membrane crossflow test units
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All FO and RO biofouling experiments were carried out at fixed hydrodynamic operating
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conditions, with a crossflow velocity of 8.5 cm s-1 and an initial permeate water flux of 19 ± 1 L
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m-2 hr-1. For both RO and FO, the inner membrane cell dimensions were 7.7 cm × 2.6 cm × 0.3
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cm (length, width, and height, respectively). Under these conditions, the resulting cell Reynolds
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number (Re) was 460 and the mass transfer coefficient (k) was 1.46 × 10-5 m s-1. Initial feed
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volume was 10 L with a constant feed temperature of 25 C. No permeate was recycled in either
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the FO or RO setup.
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2.3.1. FO crossflow test unit
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A custom-made laboratory-scale FO test unit was used for FO biofouling experiments (Fig.
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S1 of Supplementary data). Feed and draw crossflow rates were held constant using two gear
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pumps with Micropump® A-Mount cavity style pump heads (Cole Palmer). Retentate was
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recycled back into the feed reservoir. Both solutions were held at a constant temperature by a
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Thermo Scientific (Heslab, RTE 7) chiller attached to a water bath (Thermo Scientific, MX-OG
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11B). An initial permeate water flux was attained using a 1.3 M (NaCl) draw solution. Permeate
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water flux was monitored throughout the experiment by placing the draw solution reservoir on a
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digital scale (Denver Instrument, P4002) interfaced with a data acquisition system (PC
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interfaced).
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2.3.2. RO crossflow test unit
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A custom made laboratory-scale RO test unit was used for RO biofouling experiments (Fig.
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S2 of Supplementary data). The hydraulic pressure of the system was held constant at 14.5 ± 0.7
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bar (210 ± 10 psi) throughout the entire experiment by a high-pressure pump (Hydra-Cell,
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Wanner Engineering Inc.). Feedwater temperature was controlled by a chiller unit (Neslab, RTE-
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111), and the feed tank was further isolated in a fabricated refrigerator for better temperature
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control. A digital flow meter (Optiflow 1000 flowmeter, Humonics, CA) was interfaced with a
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data acquisition system (PC interfaced) to acquire real-time permeate water flux. Retentate was
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recycled back into the feed reservoir, while permeate was discarded.
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2.4. Biofouling tests and protocols
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A biofouling protocol was designed to allow adequate biofilm development on the membrane
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surface in both the FO and RO crossflow test units.
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2.4.1. System cleaning procedure
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At the beginning of each experiment, the FO and RO units were thoroughly cleaned. Initially,
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15% bleach was circulated through the system for 30 minutes, followed by two DI water rinses
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for 10 minutes each. Next, 5 mM EDTA solution at pH 11 was recirculated in the test unit for 30
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minutes, followed by two 10 minute DI water rinses. Finally, the units were cleaned by
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recirculating 95% ethanol for one hour. All product residues were then thoroughly rinsed out of
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the system by circulating DI water through the unit for 10 minutes, three times. The membrane
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was inserted into the units following the cleaning protocol.
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2.4.2. FO biofouling procedure
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Following the cleaning procedure, the system was stabilized at a zero flux, with membrane in
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place and sterilized DI water in both the feed and draw reservoirs. The draw DI water was then
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replaced with 2 L of 1.3 M NaCl solution and allowed to stabilize for approximately one hour.
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Once the desired initial water flux was obtained (19 ± 1 L m-2 hr-1), the feed solution was
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replaced with 10 L of sterile wastewater and temperature was allowed to restabilize. The system
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was run for 24 hours to obtain an accurate background baseline. At the conclusion of the
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background run, both the feed and draw were replaced with 10 L of fresh, sterile wastewater and
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2 L of 1.3 M NaCl, respectively. Late exponential stage P. aeruginosa (OD600 of 0.6-1.0) was 6
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centrifuged for 15 minutes at 4,000 rpm and 4 C to remove the LB broth. The bacteria were then
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resuspended in 10 mL of sterile wastewater by vortexing for 30 seconds. The feed reservoir was
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inoculated with P. aeruginosa to achieve an initial bacterial concentration of ~2 × 106 cells mL-1.
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Water subsamples of the inoculated feed solution were taken throughout the experiment to
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monitor pH, conductivity, and bacterial concentration (via plate counts and OD600
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measurements). Permeate water flux was measured continuously throughout the experiment.
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Baseline data was then used to correct the water flux for the dilution of draw solution and
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reduction of the osmotic driving force during the biofouling run, as we outlined elsewhere [12].
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2.4.3. RO biofouling procedure
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Following the previously described disinfection procedure, the membrane was compacted at
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20.7 bar (300 psi) with sterile DI water until a stable permeate water flux was obtained, taking
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approximately 12-14 hours. Following membrane compaction, pressure was adjusted down to
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14.5 ± 0.7 bar (210 ± 10 psi) and the system left to run for three hours to obtain a stable permeate
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water flux (19 ± 1 L m-2 hr-1). The pressure, crossflow, and temperature were maintained
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throughout both the background and biofouling experiments. After attaining a stable water flux,
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the DI feed solution was replaced with 10 L of sterile wastewater. The system temperature was
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allowed to restabilize and a background baseline run was conducted for 24 hours. At the
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completion of the background baseline, the feed solution was inoculated with P. aeruginosa
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using the same procedure as described for FO biofouling. The feed solution was monitored for
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pH, conductivity, and bacterial concentration throughout the experiment.
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2.5. Biofilm characterization
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At the end of each experiment, the biofouled membrane coupon was removed from the RO
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and FO cells and dissected into two subsamples for confocal laser scanning microscope (CLSM)
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and total organic carbon (TOC) analysis. CLSM analysis was done directly after each run.
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Coupon samples for TOC analysis were stored in separate, sterile conical centrifuge tubes at -80
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C for future analysis.
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2.5.1. CLSM analysis
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A central section from the biofouled membrane was cut into two 1 cm × 1 cm coupons. Each
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coupon was rinsed three times with sterile wastewater to remove planktonic and loosely adhered
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bacteria. After rinsing, the biofilm was stained with SYTO® 9 and propidium iodide (PI)
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according to the reagent manual (LIVE/DEAD® BacLightTM, Invitrogen, USA) to allow
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identification of live and dead cells (green and red, respectively). Simultaneously, the biofilm
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samples were stained with 30 µL of 50 μM concanavalin A (Con A, Alexa Flour® 633,
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Invitrogen, USA) to identify polysaccharides (blue) as part of the biofilm EPS. All staining was
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conducted over 30 minutes in the dark. Unbound stain was then removed with three final sterile
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wastewater rinses. Stained coupons were then mounted in an unconfined biofilm characterization
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chamber, as described by Bar-Zeev et al. [21]. 8
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CLSM images of the stained coupons were captured using an inverted CLSM (Zeiss LSM 510,
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Carl Zeiss, Inc.) outfitted with a Plan-Apochromat 20 × 0.8 numerical aperture objective. At least
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seven Z stack random fields (635 μm × 635 μm) were collected, with a slice thickness of 2.14
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µm using ZEN® (Carl Zeiss, Inc.), to obtain representative areas of each biofilm. SYTO® 9, PI,
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and Con A were excited with 488 nm argon, 561 nm diode-pumped solid state, and 633 nm
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helium-neon lasers, respectively. Image analysis for each set of data was conducted using Auto-
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PHLIP-ML (http://sourceforge.net/projects/phlip/), ImageJ software (http://rsbweb.nih.gov), and
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MATLAB® (The Mathworks™, Inc.). Biovolume was determined for polysaccharides-EPS, live,
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and dead bacterial cells. Total biovolume was calculated by summing together the biovolume of
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each component, and total biofilm thickness was measured on Zen® by averaging three different
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thickness measurements from each CLSM image collected for both FO and RO biofilms.
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Orthogonal CLSM biofilm images were reconstructed using Zen imaging software (Carl Zeiss,
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Inc.). Vertical pore sizes were determined with ZEN-Ortho software by measuring 3 pore widths,
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using different biofilm side view sections, from each CLSM biofilm (at least 7 CLSM images
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were taken for each FO and RO run).
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2.5.2. TOC measurements
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For biofilm TOC analysis, a central 2 cm × 1 cm subsample was dissected from each of the
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pre-stored membrane coupons and placed into separate, clean 25 mL glass vials with 24 mL of
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fresh DI water. Biofilm was removed from the membrane coupons via bath sonication for 1.5
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hours. TOC analysis was then conducted with a TOC analyzer (Shimadzu TOC-V CSH)
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according to manufacturer’s operating instructions. TOC concentrations were normalized
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according to the membrane coupon sample size cut (TOC mass × membrane area-1).
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2.6. Statistical analysis.
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Confocal and TOC data were displayed as means with corresponding standard deviations. A
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student t-test with paired two-tailed distribution was applied to all confocal biovolume data
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(n=11-16). Data were obtained from two biologically different experiments with 7-9 areas
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analyzed from each experiment, a total of 16 FO images and 16 RO images. Data were
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considered statically significant if P < 0.01. Permeate flux data were displayed as a running 16-
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minute average for all runs. 9
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3. Results and discussion
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3.1. Influence of biofouling on forward osmosis and reverse osmosis water flux
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Permeate water flux decline was determined for two independent forward osmosis (FO) and
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reverse osmosis (RO) biofouling experiments, which were terminated after ~24 hours. To
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compare biofouling behaviors, experimental conditions in FO and RO were kept identical,
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including type of membrane used, initial permeate water flux, crossflow velocity, solution
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composition, channel dimensions, and initial concentration of P. aeruginosa. A marked
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difference in permeate water flux behavior in FO and RO was observed (Fig. 1). During
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biofouling experiments, a permeate water flux decline of ~10% was observed in the two FO runs
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(at 600 mL cumulative permeate volume, corresponding to ~18 hours). A more substantial
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decrease in water flux (~30%) was observed in the two RO runs (at 600 mL cumulative permeate
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volume, corresponding to ~17 hours). Our results were generally consistent with water flux
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decline behavior due to biofouling, observed in FO and RO [20].
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FIGURE 1
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Previously, sodium ions have been shown to disrupt calcium ion bridging in the fouling layer
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of alginate (a model polysaccharide), potentially reducing FO fouling with polysaccharides
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[15,22,23]. Since EPS is a major component of the biofouling layer formed in our FO
243
experiments, we have surmised that reverse diffusion of sodium ions from the NaCl draw
244
solution may have disrupted the biofilm structure, thus reducing the impact of the biofilm on
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permeate water flux.
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An additional FO biofouling experiment was conducted at the same initial water flux using
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MgCl2 (~1.6 M) as a draw solution to determine whether the lower biofouling propensity of FO
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in comparison to RO can be attributed to reverse salt flux of sodium ions through the membrane
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[15,22,23]. Magnesium chloride, as a divalent salt, had a much lower measured reverse
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permeation rate (73.89 ± 23.88 mmol m-2 hr-1) than NaCl (166.74 ± 23.97 mmol m-2 hr-1),
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thereby serving as a suitable draw solution to assess the impact of reverse permeation of sodium
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ions on EPS and biofouling characteristics. As observed, permeate water flux decline using
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MgCl2 draw solution was similar to FO biofouling runs with NaCl as a draw solution (Fig. S3 of
10
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Supplementary data). These results indicate that sodium ion permeation through reverse
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diffusion in FO did not disrupt the biofilm structure and was not the cause for the reduced water
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flux decline in FO.
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In RO systems, hydraulic pressure (∆P) is applied on the feed side to produce permeate
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water flux through the semi-permeable membrane. This application of ∆P caused bacteria in the
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feed solution to be deposited and subsequently compacted on the membrane, thus affecting
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permeate water flux behavior. We propose that biofilm compaction hindered membrane water
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permeate flux via two co-occurring factors. First, it is possible that biofilm enhanced osmotic
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pressure due to solutes accumulating at the membrane surface was intensified in the compacted
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biofilms compared to looser ones [7,24]. Second, the biofilm hydraulic resistance increased
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primarily due to compaction of the EPS-hydrogel [25,26].
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In FO systems, an osmotic pressure difference (∆π) draws permeate water flux across the
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semi-permeable membrane. As no ∆P was applied in FO, no biofilm compaction occurred. We
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suggest that hydraulic resistance in the thicker FO biofilm was negligible due to its loose
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conformation. Moreover, the sparse and channeled FO biofilm structure could account for
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enhanced back diffusion of the solutes to the bulk solution, thus lessening concentration
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polarization within the biofilm.
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Our observations led us to conclude that the differences in FO and RO water flux decline
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behavior must be attributed to changes in biofilm characteristics. Because the FO and RO
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biofouling experiments were operated under identical conditions, the differences in biofilm
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characteristics are directly related to the disparate driving forces of the two systems.
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3.2. Biofilm characteristics in FO and RO systems
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Marked differences in membrane structure as well as biofilm dimensions, architecture, and
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composition (“live” cells, “dead” cells, and polysaccharide-EPS) were observed in RO and FO as
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a result of the two different driving forces, ∆P and ∆π, respectively (Fig. 2). In FO, the
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membrane structure was similar to its initial condition (i.e., before a biofouling run), featuring a
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finger-like structure, as previously demonstrated for TFC membranes with a polysulfone support
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layer [27,28]. The FO biofilm exhibited a loosely organized thick layer (~50 µm), with eminent
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mushroom-shaped structures (~77 µm) (Fig. 2A), encased in a mild polysaccharide matrix, as 11
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evident from the cells/polysaccharide-EPS biovolume ratios (1.85 and 0.80 for FO and RO,
284
respectively; Table 1). These distended mushroom structures with high three-dimensional
285
complexity developed more readily in FO due to the absence of applied ∆P. The sparse biofilm
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organization was also characterized by large diameter vertical pores and cavities, captured within
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the biofilm mat and in between the mushroom-like structures (Table 1).
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FIGURE 2
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TABLE 1
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We posit that the open biofilm structure promoted nutrient and oxygen-rich feedwater to pass
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throughout the entire biofilm structure. These favorable FO conditions likely encouraged cell
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growth over EPS secretion, resulting in a high cells to polysaccharide-EPS biovolume ratio, as
293
seen in Table 1 (0.72 and 1.00 for NaCl and MgCl2 runs, respectively) [29,30]. Biofilm
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morphological characteristics that developed in the FO system were similar to “classic” biofilms
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captured on various unpressurized, impermeable surfaces [10,11,31]. Nevertheless, biofilm
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development in FO was distinctly organized in three layers: a polysaccharide matrix
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encapsulating a thick live layer, with an explicit dead bacterial mat attached to the membrane
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surface (Fig. 2A). We further suggest that cells attached to the surface of the FO membrane may
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have been subjected to a saline stress, caused by reverse NaCl flux from the high-salinity draw
300
solution [17,32].
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Biofilm dimensions, architecture, and composition were also measured for the MgCl2 FO run
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(Table 1). These results were used to determine whether biofilm changes between FO and RO
303
could be attributed to reverse sodium ion permeation. The MgCl2 biofilm was a porous, loosely
304
organized, thick layer (~50 μm), much like the NaCl FO biofilm (Fig. S4 of Supplementary
305
data). Live, dead, and polysaccharide-EPS biovolume compositions were also close to those
306
observed in the NaCl experiments (36%, 32%, and 32%, respectively).
307
In contrast to FO runs, RO membranes were deformed in a weave-like pattern, a result of
308
membrane compaction over the embedded supporting woven mesh due to ∆P of ~14.5 bar (Fig.
309
2B, SEM image). Consequently, the biofilms that developed in RO displayed a distinct
310
configuration that closely followed the shape of the membrane surface (Fig. 2B, top view). A
311
closer, enlarged view revealed a biofilm with a tight cell organization (~29 μm thick), embedded 12
312
in a polysaccharide layer (cells/polysaccharide-EPS biovolume ratio of 0.80, Table 1) that
313
undulated in response to the membrane surface (Fig. 2B, side view). Often, peaks of the biofilm
314
were thicker than the troughs (29 ± 4 μm and 17 ± 3 μm, respectively) and were composed
315
mainly of live cells, whereas most of the cells within the troughs appeared dead (Fig. 2B).
316
Average biovolumes (μm3 μm-2) and corresponding ratios for live cells, dead cells, and
317
polysaccharide-EPS are shown in Fig. 3 and Table 1. Live and dead biofilm components
318
developed in FO were both significantly larger than RO, as evidenced by their respective total
319
biovolumes of 71 ± 10 and 44 ± 5 μm3 μm-2, respectively. We note that total biovolumes were
320
calculated as the sum of live, dead, and polysaccharide-EPS biovolumes. Live to dead cell
321
biovolume ratios for FO and RO (0.72 and 0.52, respectively) shed light on the viability of the
322
biofilm (Table 1). We suggest that a similar number of live and dead cells were present in the FO
323
biofilm, while significantly more dead cells were present in RO (Table 1), likely a result of the
324
compact cell configuration in the RO biofilms.
325
FIGURE 3
326 327
3.3. Biofilm compaction in FO and RO systems
328
Previous studies have reported that the biofilm EPS matrix is responsible for increasing
329
hydraulic resistance in RO membrane biofouling [17,25,26]. Biofilms on solid surfaces are
330
typically loosely organized with many horizontal channels and mushroom-shaped structures
331
[33]. However, for RO biofouling under ∆P, the biofilms appear to compact into tight, film-like
332
mats (Fig. 2B), as previously shown for various RO systems [7,34].
333
To assess biofilm compaction, the biofilm densities were indirectly estimated by dividing
334
TOC biomass by the total biovolume. Average TOC biomasses for the entire biofilm matrix were
335
similar in FO (1.08 ± 0.12 pg μm-2) and RO (1.46 ± 0.30 pg μm-2) (Table 1). However, due to
336
significant differences in average biovolumes, biofilm densities for FO and RO were 0.015 pg
337
TOC μm-3 and 0.033 pg TOC μm-3, respectively. Taken together, it is clear that the pressurized
338
environment in RO is responsible for development of highly compacted biofilms, compared to
339
those developed in osmotically-driven FO systems (Fig. 2, Table 1). 13
340
However, the mechanisms that control biofilm structures in forward and reverse osmosis
341
systems are poorly understood. The initial permeate water flux was similar in both FO and RO
342
experiments; therefore, permeate flux was not responsible for the differences in biofilm
343
structures. We posit that the application of ∆P in RO caused a pressure gradient to be generated
344
across the biofilm thickness as well as across the membrane. This pressure gradient caused the
345
overall biofilm structure to rearrange into a tight organization, with collapsed channel and pores.
346
Conversely, FO biofilms, which are not exposed to ∆P, do not experience this compaction. As a
347
result, biofilms developed in FO appear as sparse, thick films with horizontal and vertical
348
channels, akin to the architecture of biofilms that are grown over impermeable surfaces [33].
349 350
4. Concluding remarks
351
Our results indicate that FO biofilms have little effect on permeate water flux compared to
352
RO. We conclude that this observation can be attributed to the loose, open structure of FO
353
biofilms. Such a biofilm structure imposes low hydraulic resistance to water flow and low
354
concentration polarization at the membrane surface due to enhanced back transport of solutes to
355
the bulk solution. Conversely, RO biofilms, subjected to hydraulic pressure (∆P), cause severe
356
permeate water flux decline, due to biofilm compression induced by ∆P that leads to a compact
357
biofilm structure. We surmise that the compaction of the biofilm into a tight film-like mat added
358
hydraulic resistance and possibly intensified biofilm-enhanced osmotic pressure, both of which
359
contributed to the overall water flux decline. Taken together, the results highlight FO biofilms
360
advantages over RO in membrane cleaning and treatment of high-fouling potential feedwaters.
361 362
Acknowledgments
363
This research was made possible by the National Science Foundation Graduate Research
364
Fellowship to Sarah Kwan (Grant No. DGE-1122492), and the support through the postdoctoral
365
fellowship to Dr. Edo Bar-Zeev provided by the United States-Israel Binational Agricultural
366
Research and Development (BARD) fund. We also extend our thanks Laura H. Aria Chavez and
367
Tony Straub for technical assistance using the Shimadzu TOC-V CSH and membrane
368
characterization, respectively. We would like to thank Dr. Joseph Wolenski from the Molecular,
369
Cellular, and Developmental Biology Department at Yale University for technical assistance 14
370
using the CLSM. Lastly, facilities use was supported by YINQE and NSF MRSEC DMR
371
1119826.
372
15
373
References
374 375 376
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466 467
18
468 469 470 471
19
Normalized Water Flux (Jw/Jw0)
472
1.2 1.0
FO
0.8
RO
0.6 0.4 0.2 0.0 0
100 200 300 400 500 600
Permeate Volume (mL) 473 474 475 476
Fig. 1. Normalized permeate water flux decline (Jw/Jw0) during biofouling with P. aeruginosa of
477
duplicate FO and RO biofouling experiments. Experimental conditions were the following:
478
initial water flux 19 ± 1 L m-2 hr-1, cross velocity 8.5 cm s-1, initial cell concentration 2 × 106
479
cells mL-1, pH 7.6 ± 0.5, and model wastewater medium with initial ionic strength of 15.9 mM
480
and conductivity of 1,142 ± 50 μS.
481
20
482
483 484 485
Fig. 2. CLSM orthogonal view of P. aeruginosa biofilm structures developed on (A) FO and (B)
486
RO membranes (SEM image) after biofouling for 24 hours. The RO biofouling weave
487
morphology followed the deformed membrane structure, as observed in the cross-section SEM
488
images. Top insets are matching enlargements of the biofilm layer in side view. Biofilms were
489
stained with Con A (blue), SYTO® 9 (green), and PI (red) dyes specific for polysaccharide-EPS,
490
live cells, and dead cells, respectively.
491
21
Biovolume (m3/m2)
492 40 35 30 25 20 15 10 5 0
Live
Dead
Poly.
Biofilm Components FO
RO
493 494 495
Fig. 3. Comparison of FO and RO biovolumes of P. aeruginosa biofouling layer components
496
observed by CLSM for live cells, dead cells, polysaccharide-EPS (denoted as Poly.), shown in
497
green, red, and blue, respectively. Asterisk above bar indicates measurement between FO and
498
RO was significantly different (P < 0.01). Error bars indicate one standard deviation (n = 11-16).
499 500
Table 1. Comparison of biofilm component characteristics between FO and RO membrane
501
systems represented as percentages and ratios for polysaccharide-EPS, dead cells, live cells, and
502
cells (summation of live and dead cells). Parameter
Units
Total thickness (μm) Ave Pore diameter (μm) TOC
µm µm pg μm-2
Polysaccharide- EPS Dead cells
% %
Forward Forward Osmosis Osmosis (NaCl draw) (MgCl2 draw) 56 ± 9a 65 ± 5a 12 ± 4 11 ± 4 1.08 ± 0.12 1.12 Percent Biovolume Composition 35 36 38 32 22
Reverse Osmosis 29 ± 7a 5±2 1.46 ± 0.30 56 29b, d
Live cells
%
27
Live cells/Dead cells V/V Cells/Polysaccharide-EPS V/V Density pg TOC μm-3
0.72 1.85 0.015
32 Biovolume Ratios 1.00 1.78 0.011
15c 0.52 0.80 0.033
503
a
Indicates significant difference between FO and RO total thickness (P < 0.01)
504
b
Indicates significant difference between polysaccharide-EPS and dead cell measurement (P < 0.01)
505
c
Indicates significant difference between polysaccharide-EPS and live cell measurement (P < 0.01)
506
d
Indicates significant difference between live and dead cell measurement (P < 0.01)
507 508 509
HIGHLIGHTS We compare biofouling behaviors in FO and RO membrane systems
511 512
Water flux decline was significantly more pronounced in RO than FO Biofilms in FO were thick, loosely organized, and with mushroom shaped structures
513
Biofilms in RO were tightly organized due to the applied hydraulic pressure
514
Differences in biofouling are due to differences in driving forces in FO and RO
510
515
23
PII: DOI: Reference:
S0376-7388(15)30055-7 http://dx.doi.org/10.1016/j.memsci.2015.07.027 MEMSCI13846
To appear in: Journal of Membrane Science Received date: 2 December 2014 Revised date: 13 July 2015 Accepted date: 14 July 2015 Cite this article as: Sarah E. Kwan, Edo Bar-Zeev and Menachem Elimelech, Biofouling in forward osmosis and reverse osmosis: Measurements and m e c h a n i s m s , Journal of Membrane Science, http://dx.doi.org/10.1016/j.memsci.2015.07.027 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Biofouling in forward osmosis and reverse osmosis: Measurements and mechanisms
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Journal of Membrane Science
12 13
Revised: July 12, 2015
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Sarah E. Kwan+, Edo Bar-Zeev+, and Menachem Elimelech*
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Department of Chemical and Environmental Engineering, Yale University, P.O. Box 208286 New Haven, Connecticut 06520-8286, USA
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+
Contribute equally to this work
* Corresponding author: Menachem Elimelech, Tel: +1 (203) 432-2789; Fax: +1 (203) 432-4387 email: [email protected]
1
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Abstract
34
We have investigated biofouling behavior in forward osmosis (FO) and reverse osmosis (RO)
35
membrane systems. Analysis of these two systems was done via comparison of biofilm structure
36
and membrane permeate water flux decline. Experiments were performed using a model
37
wastewater solution inoculated with Pseudomonas aeruginosa in laboratory-scale FO and RO
38
test cells. For a meaningful comparison, biofouling runs were carried out using the same thin-
39
film composite polyamide FO membrane, identical hydrodynamic conditions, and an initial
40
permeate water flux of 19 L m-2 hr-1. Water flux decline after 600 mL of cumulative permeate
41
volume (or ~17-18 hours of fouling) was significantly lower in FO (~10%) compared to RO
42
(~30%). FO biofilms grew in a loosely organized thick layer, with eminent mushroom-shaped
43
structures, while RO biofilms grew in tightly organized mats encased in larger amounts of
44
extracellular polymeric substances (EPS) per cell. The more compact biofilms in RO induced
45
greater biofilm-enhanced osmotic pressure and hydraulic resistance to water flow compared to
46
FO, which resulted in higher flux decline. We attribute the differences in biofouling behaviors in
47
FO and RO to the different driving forces: osmotic pressure in FO and hydraulic pressure in RO.
48
Keywords: Biofouling; Forward osmosis; Reverse osmosis; Compaction; Hydraulic pressure
49
2
Graphic abstract
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51 52 53 54
1. Introduction
55
Membrane-based technologies play an important role in various separation processes at the
56
water-energy nexus [1,2]. Forward osmosis (FO) is an emerging membrane technology within
57
this nexus that has potential applications in desalination [3], wastewater treatment/reuse [4], and
58
treatment of various industrial wastes [3–5]. However, the performance of FO, like all
59
membrane-based processes, is hampered by biofouling, an obstruction that increases operational
60
costs and shortens membrane life.
61
Membrane fouling can be classified into four main categories organic, inorganic,
62
colloidal, and microbial (biofouling) with biofouling being the most ubiquitous and
63
recalcitrant [6,7]. Membrane biofouling is manifested as a complex sessile microbial community
64
permanently attached to the surface by gel-like, self-produced extracellular polymeric substances
65
(EPS) [8–10]. This microbial cake layer features a multicellular architecture of live and dead
66
cells encased in EPS, which is primarily composed of polysaccharides and proteins [11]. Once
67
established, biofilms are notoriously resistant to removal by treatment with oxidizing agents,
68
biocides, or antibiotics due to the protection provided by the EPS matrix [11].
69
Previous FO fouling studies have focused on organic [12–14] and colloidal fouling [15,16].
70
These studies revealed that fouling in FO is less detrimental to membrane performance than in
71
reverse osmosis (RO) [3,13,17]. Furthermore, in addition to diminished adverse fouling effects,
3
72
fouling in FO was found to be more reversible than in RO [15]. Studies have attributed the
73
differences in the impact of fouling on membrane performance in FO compared to RO to lack of
74
applied hydraulic pressure in FO [3,13,17].
75
Our understanding of biofouling propensity and characteristics in FO is limited [18,19],
76
especially in comparison to RO. To date, only one study compared biofilms developed in FO and
77
RO systems [20]. Biofouling in FO was found to form a loose film and induced less permeate
78
water flux decline than RO [20]. However, the mechanisms responsible for the differences in
79
biofilm characteristics and the resulting effect on FO performance compared to RO are still not
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well understood.
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In this paper, we report a direct comparison of biofouling behavior in FO and RO
82
configurations, conducted by analyzing permeate water flux decline and relating system
83
performance to biofilm characteristics. Our results shed new light on biofilm structural
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composition in both FO and RO systems and the relationship between permeate water flux
85
decline and biofouling characteristics. Moreover, we provide insights into the mechanisms of
86
biofouling in osmotically driven FO and hydraulic-pressure driven RO systems.
87 88
2. Materials and methods
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2.1. Bacteria strain and growth conditions
90
A Pseudomonas aeruginosa (ATCC® 27853TM) monoculture was used as a model biofilm
91
strain in all biofouling experiments. Approximately 250 mL of Luria-Bertani (LB, BD
92
Biosciences) broth were inoculated with a fresh single colony of P. aeruginosa and allowed to
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grow overnight in a shaking incubator at 37 C. Sterile, model wastewater was freshly prepared
94
for each biofouling run, as described elsewhere [21]. Wastewater was prepared by supplementing
95
deionized (DI) water (Nano Pure II, Barnstead, Dubuque, IA) with 8.0 mM NaCl, 0.15 mM
96
MgSO4, 0.5 mM NaHCO3, 0.2 mM CaCl2, 0.2 mM KH2PO4, 0.4 mM NH4Cl, and 0.6 mM
97
Na3C6H5O7 (sodium citrate) as a carbon source. All components were individually dissolved in
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~60 mL of DI water and sterilized in an autoclave, then added to sterile DI water to prepare a 10
99
L wastewater solution batch. The final solution pH and conductivity were 7.6 ± 0.5 and 1142 ±
100
50 μS cm-1, respectively, with a calculated ionic strength of 15.9 mM.
101
2.2. Thin-film composite membrane 4
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A commercial thin film composite (TFC) FO membrane (Hydration Technology Innovations,
103
Albany, OR, USA) was used as a model membrane for biofouling experiments in both FO and
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RO. The membrane was received as a flat sheet and stored at 4 C in DI water. Prior to use, the
105
membrane was wetted in a 25% isopropyl alcohol solution for 20 min, and then rinsed for 10
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minutes in DI water three times. Intrinsic membrane transport properties were determined by
107
using the method of Tiraferri et al. [22], resulting in water permeability coefficient, A, of 1.08 L
108
m-2 hr-1 bar-1, salt (NaCl) permeability coefficient, B, of 0.603 L m-2 hr-1, and structural
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parameter, S, of 419 μm. Reverse salt flux through the membrane was experimentally determined
110
according to the procedure outlined by Yong et al. [23].
111
2.3. Membrane crossflow test units
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All FO and RO biofouling experiments were carried out at fixed hydrodynamic operating
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conditions, with a crossflow velocity of 8.5 cm s-1 and an initial permeate water flux of 19 ± 1 L
114
m-2 hr-1. For both RO and FO, the inner membrane cell dimensions were 7.7 cm × 2.6 cm × 0.3
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cm (length, width, and height, respectively). Under these conditions, the resulting cell Reynolds
116
number (Re) was 460 and the mass transfer coefficient (k) was 1.46 × 10-5 m s-1. Initial feed
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volume was 10 L with a constant feed temperature of 25 C. No permeate was recycled in either
118
the FO or RO setup.
119
2.3.1. FO crossflow test unit
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A custom-made laboratory-scale FO test unit was used for FO biofouling experiments (Fig.
121
S1 of Supplementary data). Feed and draw crossflow rates were held constant using two gear
122
pumps with Micropump® A-Mount cavity style pump heads (Cole Palmer). Retentate was
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recycled back into the feed reservoir. Both solutions were held at a constant temperature by a
124
Thermo Scientific (Heslab, RTE 7) chiller attached to a water bath (Thermo Scientific, MX-OG
125
11B). An initial permeate water flux was attained using a 1.3 M (NaCl) draw solution. Permeate
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water flux was monitored throughout the experiment by placing the draw solution reservoir on a
127
digital scale (Denver Instrument, P4002) interfaced with a data acquisition system (PC
128
interfaced).
129
2.3.2. RO crossflow test unit
5
130
A custom made laboratory-scale RO test unit was used for RO biofouling experiments (Fig.
131
S2 of Supplementary data). The hydraulic pressure of the system was held constant at 14.5 ± 0.7
132
bar (210 ± 10 psi) throughout the entire experiment by a high-pressure pump (Hydra-Cell,
133
Wanner Engineering Inc.). Feedwater temperature was controlled by a chiller unit (Neslab, RTE-
134
111), and the feed tank was further isolated in a fabricated refrigerator for better temperature
135
control. A digital flow meter (Optiflow 1000 flowmeter, Humonics, CA) was interfaced with a
136
data acquisition system (PC interfaced) to acquire real-time permeate water flux. Retentate was
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recycled back into the feed reservoir, while permeate was discarded.
138
2.4. Biofouling tests and protocols
139
A biofouling protocol was designed to allow adequate biofilm development on the membrane
140
surface in both the FO and RO crossflow test units.
141
2.4.1. System cleaning procedure
142
At the beginning of each experiment, the FO and RO units were thoroughly cleaned. Initially,
143
15% bleach was circulated through the system for 30 minutes, followed by two DI water rinses
144
for 10 minutes each. Next, 5 mM EDTA solution at pH 11 was recirculated in the test unit for 30
145
minutes, followed by two 10 minute DI water rinses. Finally, the units were cleaned by
146
recirculating 95% ethanol for one hour. All product residues were then thoroughly rinsed out of
147
the system by circulating DI water through the unit for 10 minutes, three times. The membrane
148
was inserted into the units following the cleaning protocol.
149
2.4.2. FO biofouling procedure
150
Following the cleaning procedure, the system was stabilized at a zero flux, with membrane in
151
place and sterilized DI water in both the feed and draw reservoirs. The draw DI water was then
152
replaced with 2 L of 1.3 M NaCl solution and allowed to stabilize for approximately one hour.
153
Once the desired initial water flux was obtained (19 ± 1 L m-2 hr-1), the feed solution was
154
replaced with 10 L of sterile wastewater and temperature was allowed to restabilize. The system
155
was run for 24 hours to obtain an accurate background baseline. At the conclusion of the
156
background run, both the feed and draw were replaced with 10 L of fresh, sterile wastewater and
157
2 L of 1.3 M NaCl, respectively. Late exponential stage P. aeruginosa (OD600 of 0.6-1.0) was 6
158
centrifuged for 15 minutes at 4,000 rpm and 4 C to remove the LB broth. The bacteria were then
159
resuspended in 10 mL of sterile wastewater by vortexing for 30 seconds. The feed reservoir was
160
inoculated with P. aeruginosa to achieve an initial bacterial concentration of ~2 × 106 cells mL-1.
161
Water subsamples of the inoculated feed solution were taken throughout the experiment to
162
monitor pH, conductivity, and bacterial concentration (via plate counts and OD600
163
measurements). Permeate water flux was measured continuously throughout the experiment.
164
Baseline data was then used to correct the water flux for the dilution of draw solution and
165
reduction of the osmotic driving force during the biofouling run, as we outlined elsewhere [12].
166
7
167
2.4.3. RO biofouling procedure
168
Following the previously described disinfection procedure, the membrane was compacted at
169
20.7 bar (300 psi) with sterile DI water until a stable permeate water flux was obtained, taking
170
approximately 12-14 hours. Following membrane compaction, pressure was adjusted down to
171
14.5 ± 0.7 bar (210 ± 10 psi) and the system left to run for three hours to obtain a stable permeate
172
water flux (19 ± 1 L m-2 hr-1). The pressure, crossflow, and temperature were maintained
173
throughout both the background and biofouling experiments. After attaining a stable water flux,
174
the DI feed solution was replaced with 10 L of sterile wastewater. The system temperature was
175
allowed to restabilize and a background baseline run was conducted for 24 hours. At the
176
completion of the background baseline, the feed solution was inoculated with P. aeruginosa
177
using the same procedure as described for FO biofouling. The feed solution was monitored for
178
pH, conductivity, and bacterial concentration throughout the experiment.
179
2.5. Biofilm characterization
180
At the end of each experiment, the biofouled membrane coupon was removed from the RO
181
and FO cells and dissected into two subsamples for confocal laser scanning microscope (CLSM)
182
and total organic carbon (TOC) analysis. CLSM analysis was done directly after each run.
183
Coupon samples for TOC analysis were stored in separate, sterile conical centrifuge tubes at -80
184
C for future analysis.
185
2.5.1. CLSM analysis
186
A central section from the biofouled membrane was cut into two 1 cm × 1 cm coupons. Each
187
coupon was rinsed three times with sterile wastewater to remove planktonic and loosely adhered
188
bacteria. After rinsing, the biofilm was stained with SYTO® 9 and propidium iodide (PI)
189
according to the reagent manual (LIVE/DEAD® BacLightTM, Invitrogen, USA) to allow
190
identification of live and dead cells (green and red, respectively). Simultaneously, the biofilm
191
samples were stained with 30 µL of 50 μM concanavalin A (Con A, Alexa Flour® 633,
192
Invitrogen, USA) to identify polysaccharides (blue) as part of the biofilm EPS. All staining was
193
conducted over 30 minutes in the dark. Unbound stain was then removed with three final sterile
194
wastewater rinses. Stained coupons were then mounted in an unconfined biofilm characterization
195
chamber, as described by Bar-Zeev et al. [21]. 8
196
CLSM images of the stained coupons were captured using an inverted CLSM (Zeiss LSM 510,
197
Carl Zeiss, Inc.) outfitted with a Plan-Apochromat 20 × 0.8 numerical aperture objective. At least
198
seven Z stack random fields (635 μm × 635 μm) were collected, with a slice thickness of 2.14
199
µm using ZEN® (Carl Zeiss, Inc.), to obtain representative areas of each biofilm. SYTO® 9, PI,
200
and Con A were excited with 488 nm argon, 561 nm diode-pumped solid state, and 633 nm
201
helium-neon lasers, respectively. Image analysis for each set of data was conducted using Auto-
202
PHLIP-ML (http://sourceforge.net/projects/phlip/), ImageJ software (http://rsbweb.nih.gov), and
203
MATLAB® (The Mathworks™, Inc.). Biovolume was determined for polysaccharides-EPS, live,
204
and dead bacterial cells. Total biovolume was calculated by summing together the biovolume of
205
each component, and total biofilm thickness was measured on Zen® by averaging three different
206
thickness measurements from each CLSM image collected for both FO and RO biofilms.
207
Orthogonal CLSM biofilm images were reconstructed using Zen imaging software (Carl Zeiss,
208
Inc.). Vertical pore sizes were determined with ZEN-Ortho software by measuring 3 pore widths,
209
using different biofilm side view sections, from each CLSM biofilm (at least 7 CLSM images
210
were taken for each FO and RO run).
211
2.5.2. TOC measurements
212
For biofilm TOC analysis, a central 2 cm × 1 cm subsample was dissected from each of the
213
pre-stored membrane coupons and placed into separate, clean 25 mL glass vials with 24 mL of
214
fresh DI water. Biofilm was removed from the membrane coupons via bath sonication for 1.5
215
hours. TOC analysis was then conducted with a TOC analyzer (Shimadzu TOC-V CSH)
216
according to manufacturer’s operating instructions. TOC concentrations were normalized
217
according to the membrane coupon sample size cut (TOC mass × membrane area-1).
218
2.6. Statistical analysis.
219
Confocal and TOC data were displayed as means with corresponding standard deviations. A
220
student t-test with paired two-tailed distribution was applied to all confocal biovolume data
221
(n=11-16). Data were obtained from two biologically different experiments with 7-9 areas
222
analyzed from each experiment, a total of 16 FO images and 16 RO images. Data were
223
considered statically significant if P < 0.01. Permeate flux data were displayed as a running 16-
224
minute average for all runs. 9
225 226
3. Results and discussion
227
3.1. Influence of biofouling on forward osmosis and reverse osmosis water flux
228
Permeate water flux decline was determined for two independent forward osmosis (FO) and
229
reverse osmosis (RO) biofouling experiments, which were terminated after ~24 hours. To
230
compare biofouling behaviors, experimental conditions in FO and RO were kept identical,
231
including type of membrane used, initial permeate water flux, crossflow velocity, solution
232
composition, channel dimensions, and initial concentration of P. aeruginosa. A marked
233
difference in permeate water flux behavior in FO and RO was observed (Fig. 1). During
234
biofouling experiments, a permeate water flux decline of ~10% was observed in the two FO runs
235
(at 600 mL cumulative permeate volume, corresponding to ~18 hours). A more substantial
236
decrease in water flux (~30%) was observed in the two RO runs (at 600 mL cumulative permeate
237
volume, corresponding to ~17 hours). Our results were generally consistent with water flux
238
decline behavior due to biofouling, observed in FO and RO [20].
239
FIGURE 1
240
Previously, sodium ions have been shown to disrupt calcium ion bridging in the fouling layer
241
of alginate (a model polysaccharide), potentially reducing FO fouling with polysaccharides
242
[15,22,23]. Since EPS is a major component of the biofouling layer formed in our FO
243
experiments, we have surmised that reverse diffusion of sodium ions from the NaCl draw
244
solution may have disrupted the biofilm structure, thus reducing the impact of the biofilm on
245
permeate water flux.
246
An additional FO biofouling experiment was conducted at the same initial water flux using
247
MgCl2 (~1.6 M) as a draw solution to determine whether the lower biofouling propensity of FO
248
in comparison to RO can be attributed to reverse salt flux of sodium ions through the membrane
249
[15,22,23]. Magnesium chloride, as a divalent salt, had a much lower measured reverse
250
permeation rate (73.89 ± 23.88 mmol m-2 hr-1) than NaCl (166.74 ± 23.97 mmol m-2 hr-1),
251
thereby serving as a suitable draw solution to assess the impact of reverse permeation of sodium
252
ions on EPS and biofouling characteristics. As observed, permeate water flux decline using
253
MgCl2 draw solution was similar to FO biofouling runs with NaCl as a draw solution (Fig. S3 of
10
254
Supplementary data). These results indicate that sodium ion permeation through reverse
255
diffusion in FO did not disrupt the biofilm structure and was not the cause for the reduced water
256
flux decline in FO.
257
In RO systems, hydraulic pressure (∆P) is applied on the feed side to produce permeate
258
water flux through the semi-permeable membrane. This application of ∆P caused bacteria in the
259
feed solution to be deposited and subsequently compacted on the membrane, thus affecting
260
permeate water flux behavior. We propose that biofilm compaction hindered membrane water
261
permeate flux via two co-occurring factors. First, it is possible that biofilm enhanced osmotic
262
pressure due to solutes accumulating at the membrane surface was intensified in the compacted
263
biofilms compared to looser ones [7,24]. Second, the biofilm hydraulic resistance increased
264
primarily due to compaction of the EPS-hydrogel [25,26].
265
In FO systems, an osmotic pressure difference (∆π) draws permeate water flux across the
266
semi-permeable membrane. As no ∆P was applied in FO, no biofilm compaction occurred. We
267
suggest that hydraulic resistance in the thicker FO biofilm was negligible due to its loose
268
conformation. Moreover, the sparse and channeled FO biofilm structure could account for
269
enhanced back diffusion of the solutes to the bulk solution, thus lessening concentration
270
polarization within the biofilm.
271
Our observations led us to conclude that the differences in FO and RO water flux decline
272
behavior must be attributed to changes in biofilm characteristics. Because the FO and RO
273
biofouling experiments were operated under identical conditions, the differences in biofilm
274
characteristics are directly related to the disparate driving forces of the two systems.
275
3.2. Biofilm characteristics in FO and RO systems
276
Marked differences in membrane structure as well as biofilm dimensions, architecture, and
277
composition (“live” cells, “dead” cells, and polysaccharide-EPS) were observed in RO and FO as
278
a result of the two different driving forces, ∆P and ∆π, respectively (Fig. 2). In FO, the
279
membrane structure was similar to its initial condition (i.e., before a biofouling run), featuring a
280
finger-like structure, as previously demonstrated for TFC membranes with a polysulfone support
281
layer [27,28]. The FO biofilm exhibited a loosely organized thick layer (~50 µm), with eminent
282
mushroom-shaped structures (~77 µm) (Fig. 2A), encased in a mild polysaccharide matrix, as 11
283
evident from the cells/polysaccharide-EPS biovolume ratios (1.85 and 0.80 for FO and RO,
284
respectively; Table 1). These distended mushroom structures with high three-dimensional
285
complexity developed more readily in FO due to the absence of applied ∆P. The sparse biofilm
286
organization was also characterized by large diameter vertical pores and cavities, captured within
287
the biofilm mat and in between the mushroom-like structures (Table 1).
288
FIGURE 2
289
TABLE 1
290
We posit that the open biofilm structure promoted nutrient and oxygen-rich feedwater to pass
291
throughout the entire biofilm structure. These favorable FO conditions likely encouraged cell
292
growth over EPS secretion, resulting in a high cells to polysaccharide-EPS biovolume ratio, as
293
seen in Table 1 (0.72 and 1.00 for NaCl and MgCl2 runs, respectively) [29,30]. Biofilm
294
morphological characteristics that developed in the FO system were similar to “classic” biofilms
295
captured on various unpressurized, impermeable surfaces [10,11,31]. Nevertheless, biofilm
296
development in FO was distinctly organized in three layers: a polysaccharide matrix
297
encapsulating a thick live layer, with an explicit dead bacterial mat attached to the membrane
298
surface (Fig. 2A). We further suggest that cells attached to the surface of the FO membrane may
299
have been subjected to a saline stress, caused by reverse NaCl flux from the high-salinity draw
300
solution [17,32].
301
Biofilm dimensions, architecture, and composition were also measured for the MgCl2 FO run
302
(Table 1). These results were used to determine whether biofilm changes between FO and RO
303
could be attributed to reverse sodium ion permeation. The MgCl2 biofilm was a porous, loosely
304
organized, thick layer (~50 μm), much like the NaCl FO biofilm (Fig. S4 of Supplementary
305
data). Live, dead, and polysaccharide-EPS biovolume compositions were also close to those
306
observed in the NaCl experiments (36%, 32%, and 32%, respectively).
307
In contrast to FO runs, RO membranes were deformed in a weave-like pattern, a result of
308
membrane compaction over the embedded supporting woven mesh due to ∆P of ~14.5 bar (Fig.
309
2B, SEM image). Consequently, the biofilms that developed in RO displayed a distinct
310
configuration that closely followed the shape of the membrane surface (Fig. 2B, top view). A
311
closer, enlarged view revealed a biofilm with a tight cell organization (~29 μm thick), embedded 12
312
in a polysaccharide layer (cells/polysaccharide-EPS biovolume ratio of 0.80, Table 1) that
313
undulated in response to the membrane surface (Fig. 2B, side view). Often, peaks of the biofilm
314
were thicker than the troughs (29 ± 4 μm and 17 ± 3 μm, respectively) and were composed
315
mainly of live cells, whereas most of the cells within the troughs appeared dead (Fig. 2B).
316
Average biovolumes (μm3 μm-2) and corresponding ratios for live cells, dead cells, and
317
polysaccharide-EPS are shown in Fig. 3 and Table 1. Live and dead biofilm components
318
developed in FO were both significantly larger than RO, as evidenced by their respective total
319
biovolumes of 71 ± 10 and 44 ± 5 μm3 μm-2, respectively. We note that total biovolumes were
320
calculated as the sum of live, dead, and polysaccharide-EPS biovolumes. Live to dead cell
321
biovolume ratios for FO and RO (0.72 and 0.52, respectively) shed light on the viability of the
322
biofilm (Table 1). We suggest that a similar number of live and dead cells were present in the FO
323
biofilm, while significantly more dead cells were present in RO (Table 1), likely a result of the
324
compact cell configuration in the RO biofilms.
325
FIGURE 3
326 327
3.3. Biofilm compaction in FO and RO systems
328
Previous studies have reported that the biofilm EPS matrix is responsible for increasing
329
hydraulic resistance in RO membrane biofouling [17,25,26]. Biofilms on solid surfaces are
330
typically loosely organized with many horizontal channels and mushroom-shaped structures
331
[33]. However, for RO biofouling under ∆P, the biofilms appear to compact into tight, film-like
332
mats (Fig. 2B), as previously shown for various RO systems [7,34].
333
To assess biofilm compaction, the biofilm densities were indirectly estimated by dividing
334
TOC biomass by the total biovolume. Average TOC biomasses for the entire biofilm matrix were
335
similar in FO (1.08 ± 0.12 pg μm-2) and RO (1.46 ± 0.30 pg μm-2) (Table 1). However, due to
336
significant differences in average biovolumes, biofilm densities for FO and RO were 0.015 pg
337
TOC μm-3 and 0.033 pg TOC μm-3, respectively. Taken together, it is clear that the pressurized
338
environment in RO is responsible for development of highly compacted biofilms, compared to
339
those developed in osmotically-driven FO systems (Fig. 2, Table 1). 13
340
However, the mechanisms that control biofilm structures in forward and reverse osmosis
341
systems are poorly understood. The initial permeate water flux was similar in both FO and RO
342
experiments; therefore, permeate flux was not responsible for the differences in biofilm
343
structures. We posit that the application of ∆P in RO caused a pressure gradient to be generated
344
across the biofilm thickness as well as across the membrane. This pressure gradient caused the
345
overall biofilm structure to rearrange into a tight organization, with collapsed channel and pores.
346
Conversely, FO biofilms, which are not exposed to ∆P, do not experience this compaction. As a
347
result, biofilms developed in FO appear as sparse, thick films with horizontal and vertical
348
channels, akin to the architecture of biofilms that are grown over impermeable surfaces [33].
349 350
4. Concluding remarks
351
Our results indicate that FO biofilms have little effect on permeate water flux compared to
352
RO. We conclude that this observation can be attributed to the loose, open structure of FO
353
biofilms. Such a biofilm structure imposes low hydraulic resistance to water flow and low
354
concentration polarization at the membrane surface due to enhanced back transport of solutes to
355
the bulk solution. Conversely, RO biofilms, subjected to hydraulic pressure (∆P), cause severe
356
permeate water flux decline, due to biofilm compression induced by ∆P that leads to a compact
357
biofilm structure. We surmise that the compaction of the biofilm into a tight film-like mat added
358
hydraulic resistance and possibly intensified biofilm-enhanced osmotic pressure, both of which
359
contributed to the overall water flux decline. Taken together, the results highlight FO biofilms
360
advantages over RO in membrane cleaning and treatment of high-fouling potential feedwaters.
361 362
Acknowledgments
363
This research was made possible by the National Science Foundation Graduate Research
364
Fellowship to Sarah Kwan (Grant No. DGE-1122492), and the support through the postdoctoral
365
fellowship to Dr. Edo Bar-Zeev provided by the United States-Israel Binational Agricultural
366
Research and Development (BARD) fund. We also extend our thanks Laura H. Aria Chavez and
367
Tony Straub for technical assistance using the Shimadzu TOC-V CSH and membrane
368
characterization, respectively. We would like to thank Dr. Joseph Wolenski from the Molecular,
369
Cellular, and Developmental Biology Department at Yale University for technical assistance 14
370
using the CLSM. Lastly, facilities use was supported by YINQE and NSF MRSEC DMR
371
1119826.
372
15
373
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18
468 469 470 471
19
Normalized Water Flux (Jw/Jw0)
472
1.2 1.0
FO
0.8
RO
0.6 0.4 0.2 0.0 0
100 200 300 400 500 600
Permeate Volume (mL) 473 474 475 476
Fig. 1. Normalized permeate water flux decline (Jw/Jw0) during biofouling with P. aeruginosa of
477
duplicate FO and RO biofouling experiments. Experimental conditions were the following:
478
initial water flux 19 ± 1 L m-2 hr-1, cross velocity 8.5 cm s-1, initial cell concentration 2 × 106
479
cells mL-1, pH 7.6 ± 0.5, and model wastewater medium with initial ionic strength of 15.9 mM
480
and conductivity of 1,142 ± 50 μS.
481
20
482
483 484 485
Fig. 2. CLSM orthogonal view of P. aeruginosa biofilm structures developed on (A) FO and (B)
486
RO membranes (SEM image) after biofouling for 24 hours. The RO biofouling weave
487
morphology followed the deformed membrane structure, as observed in the cross-section SEM
488
images. Top insets are matching enlargements of the biofilm layer in side view. Biofilms were
489
stained with Con A (blue), SYTO® 9 (green), and PI (red) dyes specific for polysaccharide-EPS,
490
live cells, and dead cells, respectively.
491
21
Biovolume (m3/m2)
492 40 35 30 25 20 15 10 5 0
Live
Dead
Poly.
Biofilm Components FO
RO
493 494 495
Fig. 3. Comparison of FO and RO biovolumes of P. aeruginosa biofouling layer components
496
observed by CLSM for live cells, dead cells, polysaccharide-EPS (denoted as Poly.), shown in
497
green, red, and blue, respectively. Asterisk above bar indicates measurement between FO and
498
RO was significantly different (P < 0.01). Error bars indicate one standard deviation (n = 11-16).
499 500
Table 1. Comparison of biofilm component characteristics between FO and RO membrane
501
systems represented as percentages and ratios for polysaccharide-EPS, dead cells, live cells, and
502
cells (summation of live and dead cells). Parameter
Units
Total thickness (μm) Ave Pore diameter (μm) TOC
µm µm pg μm-2
Polysaccharide- EPS Dead cells
% %
Forward Forward Osmosis Osmosis (NaCl draw) (MgCl2 draw) 56 ± 9a 65 ± 5a 12 ± 4 11 ± 4 1.08 ± 0.12 1.12 Percent Biovolume Composition 35 36 38 32 22
Reverse Osmosis 29 ± 7a 5±2 1.46 ± 0.30 56 29b, d
Live cells
%
27
Live cells/Dead cells V/V Cells/Polysaccharide-EPS V/V Density pg TOC μm-3
0.72 1.85 0.015
32 Biovolume Ratios 1.00 1.78 0.011
15c 0.52 0.80 0.033
503
a
Indicates significant difference between FO and RO total thickness (P < 0.01)
504
b
Indicates significant difference between polysaccharide-EPS and dead cell measurement (P < 0.01)
505
c
Indicates significant difference between polysaccharide-EPS and live cell measurement (P < 0.01)
506
d
Indicates significant difference between live and dead cell measurement (P < 0.01)
507 508 509
HIGHLIGHTS We compare biofouling behaviors in FO and RO membrane systems
511 512
Water flux decline was significantly more pronounced in RO than FO Biofilms in FO were thick, loosely organized, and with mushroom shaped structures
513
Biofilms in RO were tightly organized due to the applied hydraulic pressure
514
Differences in biofouling are due to differences in driving forces in FO and RO
510
515
23