Letter pubs.acs.org/journal/estlcu
Confocal Laser Scanning Microscopy-Compatible Microfluidic Membrane Flow Cell as a Nondestructive Tool for Studying Biofouling Dynamics on Forward Osmosis Membranes Manisha Mukherjee,†,‡ Nishanth V. Menon,§ Xin Liu,∥ Yuejun Kang,⊥ and Bin Cao*,†,‡ †
School of Civil and Environmental Engineering, Nanyang Technological University, Singapore Singapore Centre for Environmental Life Sciences Engineering, Nanyang Technological University, Singapore § School of Chemical and Biomedical Engineering, Nanyang Technological University, Singapore ∥ Singapore Membrane Technology Centre, Nanyang Environment and Water Research Institute, Nanyang Technological University, Singapore ⊥ Institute for Clean Energy and Advanced Materials, Faculty of Materials and Energy, Southwest University, Chongqing, China ‡
S Supporting Information *
ABSTRACT: In membrane biofouling studies, quantification of biofouling is often conducted destructively and the results reflect only a snapshot of the biofouling processes. This limitation is mainly due to the lack of tools that allow us to monitor dynamics of biofouling without the need to disassemble the membrane testing systems. In this study, we developed a novel multichannel fluidic membrane biofilm flow cell that allows nondestructive, real-time monitoring of biofouling dynamics on forward osmosis (FO) membranes using confocal laser scanning microscopy. As a proof of concept, we used green fluorescent protein-tagged Shewanella oneidensis as a model organism and examined its biofilm development on membranes in FO mode. The temporal profiles of quantitative biofouling parameters such as surface coverage, biovolume, and biofilm thickness were obtained without disrupting the continuous operation of the membrane testing system. We also demonstrated the applicability of the microfluidic membrane flow cells, revealing biofouling dynamics of natural, untagged bacteria on FO membranes. The microfluidic membrane flow cell developed in this study can be readily applied to evaluate antibiofouling activities of FO membranes and allows direct comparison of biofouling dynamics between FO membranes with different surface modifications.
■
substances present in the feedwater.5 The undesirable growth of microorganisms and biofilm formation, i.e., biofouling, on membrane surfaces represents the “Achilles heel” of membrane processes for drinking water production and wastewater treatment.5 Biofouling has been shown to be the potential contributor to >45% of all membrane fouling6 and has been reported to pose a crucial threat to membrane processes.7 Therefore, the study of biofilm formation and biofilm dynamics
INTRODUCTION In recent years, membrane technology has emerged as one of the leading technologies for addressing the challenge of clean water access.1 The application of membrane filtration in water treatment facilitates the production of biologically stable and microbiologically safe drinking water via the removal of inorganic and organic compounds and microorganisms.2 However, prolonged membrane operations often result in membrane fouling, which adversely affects membrane performance.3,4 A majority of the abiotic fouling components (both inorganic and organic) in membrane processes can be removed by pretreatment; most microorganisms can be removed, too, but the remaining microorganisms are capable of proliferating with the availability of a minute amount of biodegradable © XXXX American Chemical Society
Received: June 21, 2016 Revised: July 17, 2016 Accepted: July 19, 2016
A
DOI: 10.1021/acs.estlett.6b00218 Environ. Sci. Technol. Lett. XXXX, XXX, XXX−XXX
Letter
Environmental Science & Technology Letters
Figure 1. Schematic illustration of the CLSM-compatible microfluidic flow cell for nondestructive monitoring of biofouling dynamics on waterpurifying membranes. The flow cell contains three independent channels with each channel separated into two chambers by a flat sheet membrane. The drawing is not to scale.
about the cake layer composition.17 These limitations may be addressed by using CLSM imaging in combination with chemical dyes for biofilms. However, flow cells that permit nondestructive, CLSM-based monitoring of biofouling dynamics on membranes are lacking. Microfluidic technology has been previously used for the development of sensors compatible with optical detection tools to nondestructively monitor water quality in real time.18 The incorporation of membranes into microfluidic designs has been reported for various applications, including organ-on-a-chip models,19 sample preparations for biosensors,20 characterization of particle deposition, and the contribution of glycolipids in the initial stage of bacterial attachment on membranes in filtration flow cells using scanning electron microscopy and fluorescence microcopy, respectively.21,22 The objective of this study was to develop a microfluidic flow cell that is compatible with direct observation using CLSM and allows nondestructive assessment of biofouling dynamics on forward osmosis (FO) membranes. Specifically, we designed and fabricated a three-channel flow cell with each channel separated by a flat sheet membrane into two chambers to house concurrent flows of different solutions, which allows for the evaluation of three membrane samples concurrently in forward osmosis (FO) mode. We used Shewanella oneidensis as a model organism and examined its biofilm development on a FO membrane. The temporal profiles of quantitative biofouling
in membrane processes is essential for the control and mitigation of membrane fouling. Devices developed for the purpose of biofilm studies are mainly designed as continuous flow-through chambers. The most commonly used flow chamber is a three-channel flow cell, which is configured to be assembled and disassembled with ease.8 Moreover, this flow chamber allows direct observation of biofilm structures under hydrodynamic conditions using confocal laser scanning microscopy (CLSM).8,9 This threechannel flow chamber as well as other similar biofilm flow cells10 has been widely applied in studies of biofilm biology. However, all these commercial CLSM-compatible flow cells contain a single chamber on one side of the substratum, in most cases, a piece of glass slide. On the other hand, most previous studies of membrane biofouling were conducted destructively by disassembling the membrane testing systems and dissecting the membranes for microscopic observation to obtain a snapshot of the biofouling process,11,12 which is, however, highly dynamic because of the intrinsic heterogeneous and dynamic nature of microbial biofilms.13,14 Recently, several techniques based on surface enhanced raman spectroscopy (SERS), stereomicroscopy, and optical coherence tomography (OCT) have been developed to nondestructively monitor biofouling dynamics, which might create artifacts during sample preparation due to dehydration,15 limit us to the study of early stage attachment,16 and is not capable of obtaining information B
DOI: 10.1021/acs.estlett.6b00218 Environ. Sci. Technol. Lett. XXXX, XXX, XXX−XXX
Letter
Environmental Science & Technology Letters
Figure 2. Biofouling dynamics of GFP-tagged S. oneidensis on the FO membrane in the microfluidic membrane flow cells. (A) CLSM images of biofilms formed on the FO membrane at different time points (24, 36, 48, 60, 72, 84, 96, and 108 h). CV represents the cumulative permeate volume corresponding to each time point. Each CLSM image contains one top-down view (x−y plane) and two side views (x−z and y−z planes). The scale bar represents 20 μm. (B) Biofilm biovolume (expressed as volume per unit base area) and (C) biofilm thickness at different time points. Experiments were conducted in triplicate, and for each replicate, three representative images were acquired and quantified. The error bar represents the standard deviation.
outlets for the flows in chambers on both sides of the membrane. The two PDMS layers and the membrane between them were sealed using a UV curable adhesive (NOA81, Norland Products Inc.). FO Membrane and Setup. The commercial FO membrane, a thin film composite (TFC) supported on poly(ether sulfone) (PES), was obtained from Hydration Technology Inc. (Hydrawell Filter, HTI). The thickness of the membrane varied from 95 to 125 μm. Figure S1 illustrates the schematic of CLSM-compatible FO setup used in this study. Three individual membrane coupons with an effective area of 160 mm2 for each were embedded in the different channels of the flow cell. Cross flow was maintained for both the feed solution and draw solution using a Masterflex peristaltic pump (Cole-Parmer). The fully assembled membrane flow cell was sterilized with recirculation of 70% ethanol for 1 h. The flow cell setup was further flushed with autoclaved deionized water to remove ethanol from the system before commencing the biofouling experiment. The autoclaved water was pumped through the flow cell setup for ∼2 h (∼300 flow channel volumes). The FO water flux was monitored by measuring the weight changes of the feed solution at regular time points with a precision balance (Metler-Toledo). To maintain the osmotic pressure gradient, the feed solution and draw solution (500 mL each) were dosed every 12 h with fresh feed and draw solution, respectively, to ensure that the exchange of liquid between the two chambers did not interfere with the measurement of
parameters such as surface coverage, biovolume, and biofilm thickness were obtained without disrupting the continuous operation of the FO membrane testing system.
■
MATERIALS AND METHODS Bacterial Strain and Seed Culture. The wild-type (untagged) and green fluorescent protein (GFP)-tagged S. oneidensis23 was grown in lysogenic broth (LB) medium at room temperature. Stock cultures were maintained in LB medium with 20% glycerol at −80 °C. The seed culture was prepared by transferring a 0.5 mL stock culture to 50 mL of LB medium in a 200 mL Erlenmyer flask and incubated for 12 h at room temperature (∼25 °C) on an incubator shaker at 200 rpm. Microfluidic Flow Cell Fabrication. The microfluidic flow cell was fabricated by a two-step photolithographic procedure24 followed by soft lithography.25 The thickness of the SU-8 mold corresponding to the two chambers is approximately 180 μm, while the thickness of the membrane chamber is approximately 100 μm. Polydimethylsiloxane (PDMS, Sylgard 184, Dow Corning) was cast on the SU-8 molds, degassed, and cured for 2 h at 70 °C. The chip design (Figure 1) comprised two layers of PDMS. The widths of the chambers are 4 mm each, while that of the membrane chamber is 10 mm. The lengths of chambers A and B are 40 and 61 mm, respectively, while the length of the membrane chamber is 54 mm. Holes (diameter of ∼2 mm) were punched onto the PDMS to serve as inlets and C
DOI: 10.1021/acs.estlett.6b00218 Environ. Sci. Technol. Lett. XXXX, XXX, XXX−XXX
Letter
Environmental Science & Technology Letters
average thickness of 5.0 ± 1.0 μm (Figure 2). The biovolume and the thickness of the biofilms on the membrane surfaces then gradually increased to 39.3 ± 12.7 μm3 μm−2 and 24.1 ± 4.1 μm, respectively, at 72 h (Figure 2). As the growth of the biofilms proceeded to ∼4 days (96 h), biofilm dispersal occurred, resulting in a significant decrease (p ≤ 0.05) of the biovolume and biofilm thickness from 32.0 ± 7.5 μm3 μm−2 and 17.7 ± 2.6 μm to 17.2 ± 6.0 μm3 μm−2 and 8.5 ± 1.2 μm, respectively, at 108 h. Intriguingly, no significant difference was observed for the biofouling dynamics in terms of temporal profiles of biovolume and biofilm thickness under the tested FO operation (with cross membrane flow) or control (no cross membrane flow) conditions (Figure S3). It should be noted that the cross membrane flow in our tested conditions was low (∼0.23 cm/s). The influence of cross membrane flow at different velocities on biofouling dynamics is an interesting research topic that requires future investigations. A typical life cycle of a biofilm is a complex process comprising distinct developmental stages. It commences with the transport of microbial cells to the surfaces; adhesion, which is initially reversible followed by irreversible attachment of cells to the surfaces facilitated by EPS components at the early stage of the biofilm development; and surface-associated growth and proliferation of microorganisms into mature biofilms followed by dispersal or disassembly of cells from the mature biofilms at the late stage of the biofilm development.33 Although this typical biofilm life cycle has been observed in many different biofilm systems, it has not been reported for biofouling on FO membranes under operating conditions. In this study, using our newly developed CLSM-compatible membrane flow cell, we for the first time obtained in a nondestructive manner the dynamics of biofouling on the FO membranes. Under the FO operation condition, bacteria in the feed solution attached to the membrane surfaces as individual cells at the initial stage and developed into cell aggregates. Then the biovolume and the thickness of the biofilms gradually increased. Mature biofilms with highly heterogeneous three-dimensional structures formed on membranes at a later stage of membrane biofouling followed by biofilm dispersal, which resulted in a decrease in biofilm biovolume and thickness. In spite of addition of fresh media every 12 h, biofilm dispersal was observed at the later stage of the biofouling in this system, which could be attributed to various passive34,35 and active mechanisms36,37 that control the precise timing of dispersal of the biofilm from the mature biofilms. The biofouling dynamics observed in this study correlated well with the typical biofilm life cycle reported in other experimental systems.38,39 In addition to biofouling dynamics, the microfluidic membrane flow cell we developed also allows us to monitor the water flux crossing the FO membranes. With the addition of bacteria to the feed solution, the water flux decreased by 25% from 3.92 ± 0.21 L m−2 h−1 at the initial stage of bacterial attachment (12 h; cumulative permeate volume, 7.4 mL) to 2.94 ± 0.31 L m−2 h−1 at the later stage of biofilm development (72 h; cumulative permeate volume, 39.4 mL) (Figure S4). In contrast, in the control experiment without inoculation of bacteria, a
