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Biofouling and Microbial Communities in Membrane Distillation and Reverse Osmosis Katherine R. Zodrow, Edo Bar-Zeev, Michael J Giannetto, and Menachem Elimelech Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/es503051t • Publication Date (Web): 08 Oct 2014 Downloaded from http://pubs.acs.org on October 19, 2014

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Environmental Science & Technology

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Biofouling and Microbial Communities in Membrane Distillation and Reverse Osmosis


Revised: October 6, 2014

Katherine R. Zodrow, Edo Bar-Zeev, Michael J. Giannetto, and Menachem Elimelech *

Department of Chemical and Environmental Engineering, Yale University, New Haven, Connecticut 06520-8286

* Corresponding author: Menachem Elimelech, Email: [email protected], Phone: (203) 432-2789

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Abstract

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Membrane distillation (MD) is an emerging desalination technology that uses low-grade heat

41

to drive water vapor across a microporous hydrophobic membrane. Currently, little is known

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about the biofilms that grow on MD membranes. In this study, we use estuarine water

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collected from Long Island Sound in a bench-scale direct contact MD system to investigate

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the initial stages of biofilm formation. For comparison, we studied biofilm formation in a

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bench-scale reverse osmosis (RO) system using the same feed water. These two membrane

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desalination systems expose the natural microbial community to vastly different

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environmental conditions  high temperatures with no hydraulic pressure in MD and low

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temperature with hydraulic pressure in RO. Over the course of 4 days, we observed a steady

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decline in bacteria concentration (nearly 2 orders of magnitude) in the MD feed reservoir.

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Even with this drop in planktonic bacteria, significant biofilm formation was observed.

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Biofilm morphologies on MD and RO membranes were markedly different. MD membrane

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biofilms were heterogeneous and contained several colonies, while RO membrane biofilms,

53

although thicker, were a homogenous mat. Phylogenetic analysis using next-generation

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sequencing of 16S ribosomal DNA showed significant shifts in the microbial communities.

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Bacteria representing the orders Burkholderiales, Rhodobacterales, and Flavobacteriales were

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most abundant in the MD biofilms. Based on the results, we propose two different regimes

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for microbial community shifts and biofilm development in RO and MD systems.

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TOC ART

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INTRODUCTION

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Water and energy are two of the grand challenges of the 21st century. Many technologies

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have been developed to meet these challenges, including several novel membrane-based

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processes.1,2 One such process is membrane distillation (MD), which uses a partial vapor

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pressure difference across a water-excluding, hydrophobic microporous membrane to

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perform a separation.3–5 Membrane distillation is currently used for separations in the oil and

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gas industry,6 nutrient recovery,7 chemical separations,8 and water treatment and

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desalination.5 MD can desalinate sea water or brine at a relatively low temperature (e.g. 50 –

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80 ˚C) using solar energy or low-grade heat.9

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Several pilot scale MD plants are currently under development.5 However, like all

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developing technologies, MD faces several challenges, many of them similar to challenges

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faced by other membrane technologies. These include fouling by inorganic,10,11 organic,12 and

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biological matters.8,13,14 Several studies have investigated the effects of inorganic and organic

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fouling on MD performance;10–12 however, much less is known about the biofilms that form

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in MD systems.

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Biofouling in MD may lead to increased temperature polarization (thermal

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resistance), increased resistance to vapor flow, pore blockage, wetting, or a decrease in the

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vapor pressure driving force.10,13,14 Biofilms do form in direct contact MD with a coastal sea

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water feed,14 although little is known about the organisms that constitute those biofilms.

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However, it is known that the survival of certain species is limited by the relatively high

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temperatures employed in this process.14 Biofilms in MD with a wastewater feed were probed

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with denaturing gradient gel electrophoresis (DGGE) and found to contain several

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thermophilic

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Caldalkalibacillus uzonesis.13 This is in contrast to the organisms commonly found on RO

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membranes, including producers of glycosphingolipids, which, along with Rhodobacteraceae,

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are suspected to be some of the primary colonizers of RO membranes.15,16

organisms,

including

Meithermus

hypogaeus,

Tepidimonas

sp,

and

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There are some major differences between these membrane-based desalination

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methods. RO separates water from salt using a dense selective layer. The driving force for

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this separation is hydraulic pressure, and this process is carried out at ambient temperature.

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MD uses a different driving force — a vapor pressure difference across a microporous

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hydrophobic membrane. This hydrophobic membrane excludes water but allows water vapor 2 ACS Paragon Plus Environment


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transport across the membrane. Separation in MD occurs at the interface between the liquid

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and the vapor phase present inside the membrane. Thus, MD is intentionally carried out at

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higher temperatures than RO.

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In this study, we explore biofouling in an MD system and compare it to biofouling in

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RO. Operating MD and RO simultaneously with natural sea water feeds allows us to compare

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fouling propensity and biofilm structure in these two systems. We additionally provide the

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first analysis of the bacterial community on an MD membrane using next-generation

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sequencing. Our results expand the understanding of biofouling in MD and may assist further

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development of anti-biofouling materials and treatments.

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MATERIALS AND METHODS

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Water Collection and Pre-Treatment. Surface water (40 L) was collected in late autumn

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(December 2013) and winter (January 2014) from a pier in Long Island Sound at the New

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Haven Harbor near the mouths of the Quinnipiac River and Morris Creek (41˚ 14’ 47.5656”

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N, 72˚ 54’ 03.1788” W, Figure S1). Plastic carboys were rinsed 3 times with sea water prior

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to collection, and the water was stored in the dark until use (< 24 h). Pretreatment was carried

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out using a 10 µm filter (Millipore Isopore, TCTP) and a dead-end cell connected to a

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peristaltic pump (Masterflex L/S Easy-Load II). Filters were replaced, as necessary, to

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mitigate cake buildup. Pretreated water was characterized and then used as the feed water in

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MD and RO bench-scale systems.

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Sea Water Nutrient Analysis. Water samples were collected in clean 400-mL

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plastic bottles and frozen (-20 ˚C) prior to analysis. Nutrients (ammonium, nitrate+nitrite,

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particulate and dissolved nitrogen and phosphorous) were quantified using standard operating

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procedures at the Nutrient Analytical Services Laboratory (University of Maryland Center for

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Environmental Science, Chesapeake Biological Laboratory).

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samples were separated using an acid-washed GF/F filter (Millipore, 0.7 µm) filter.

Particulate and dissolved

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Bench-Scale Membrane Distillation and Reverse Osmosis. Bench-scale MD

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and RO experiments were carried out for 4 days with identical initial feed water and similar

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hydrodynamic operating conditions. Crossflow velocity was 4.3 cm s-1, with initial permeate

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(distillate) flux of 20 ± 2 L m-2 h-1. Inner membrane cell dimensions in both systems were 7.7

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cm × 2.6 cm × 0.3 cm (length × width × height). Both MD and RO systems were thoroughly

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cleaned and disinfected before each experiment by washing with 10% sodium hypochlorite, 5 3 ACS Paragon Plus Environment

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mM ethylenediaminetetraacetic acid (EDTA), 90% ethanol, and three deionized (DI) water

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rinses. 4 L (MD) and 10 L (RO) of each of the above solutions were circulated through the

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system for 1 hour. During the experiments, both the MD and RO feed tanks were kept in the

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dark.

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The direct contact MD system consisted of two closed loops — a feed and a distillate

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(Figure S2). Details of this system can be found in our previous publication.17 The cold

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distillate stream was maintained at 18.1 ± 0.37 ˚C, while the hot feed stream was set to 50.4 ±

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0.56 °C using two chillers (Cole-Parmer Polystat). In order to achieve this temperature, the

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feed reservoir was maintained at 60 °C. Thermocouples (DS18B20), connected to a

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microcontroller, were placed at the entrances and exits of the membrane module. A

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temperature drop across the membrane feed channel of 3.8 °C was observed. A supported

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PTFE membrane (Millipore, FGLP, 0.2 µm) was used as the distillation membrane. As water

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vapor condensed on the distillate side and the volume of the distillate increased, water flowed

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out of the side arm of the flask and passed through a drop counter.17 Any collected distillate

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water was then pumped through a peristaltic pump (Masterflex L/S Easy-Load II) back into

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the feed reservoir to prevent concentration of the feed. No increase in distillate conductivity

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was observed during any of the experiments.

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Reverse osmosis experiments were carried out in a custom-built, closed-loop

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system.18 Thin-film composite (TFC) polyamide RO membrane (SW30XLE, Dow Filmtec)

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coupons were used for all experiments (active area of 7.7 cm × 2.6 cm). Dry membrane

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coupons were wetted using 25% isopropanol (J.T Baker, PA, USA) for 30 minutes and

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washed 3 times (1 hour for each wash cycle) using DI water before mounting in the RO test

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cell. The membrane water permeability coefficient, A, was 3.3 L m-2 h-1 bar-1, and the salt

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(NaCl) permeability coefficient, B, was 0.184 L m-2 h-1

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98.6 ± 0.2 % during the experiments. Hydraulic pressure was held constant at 36 bar (520

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psi) by a high-pressure pump (Hydra-Cell, Wanner Engineering Inc.), yielding an initial

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water flux of 20 ± 0.3 L m-2 h. During the experiments, temperature was held constant at 25 ±

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0.2 °C using a high capacity chiller unit (Polyscience, USA). The feed tank was further

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isolated in a modified refrigerator for better temperature control. A digital flow meter

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(Humonics 1000, CA, USA) was interfaced with a PC to acquire real-time permeate water

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flux.

18

. RO membrane salt rejection was

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Feed water and permeate samples (50 mL) were routinely collected for determining

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salt rejection (based on electric conductivity). Planktonic samples (1.8 ml) were taken 4 ACS Paragon Plus Environment


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periodically from both the MD and RO feed reservoirs and frozen at -80 ˚C with 1 %

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glutaraldehyde prior to measurement of photosynthetic picoplankton and total bacteria.

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Bacteria Abundance in Feed Reservoirs. Photosynthetic picoplankton

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abundance was determined using an Attune® Acoustic Focusing Flow Cytometer (Applied

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Biosystems) with a syringe based fluidic system and 488 and 405 nm lasers. Samples were

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fast-thawed at 37 ˚C, and a 1 µm bead (Polysciences) was used as a standard19. For

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heterotrophic bacteria, 300 µL aliquots of the water samples were incubated at room

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temperature with the nucleic acid stain SYTO® 9 for 10 min in the dark. A flow rate of 25

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µL min-1 was used to determine green fluorescence (520 nm) of 75 µL of sample. Taxonomic

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discrimination was made based on orange fluorescence of phycoerythrin (585 nm), red

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fluorescence of Chlorophyll a (630 nm)20, side-scatter (a proxy of cell volume21), and

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forward-scatter (a proxy of cell size22).

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Membrane Biofilm Characterization. Biofilm formation on the MD and RO

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membrane surfaces was quantified using confocal laser scanning microscopy (CLSM, Zeiss

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LSM 510, Carl Zeiss, Inc.). Biofouled membrane coupons were removed from the MD and

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RO systems and rinsed gently in sterile synthetic sea water (Instant Ocean). The biofilms

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were stained with a solution of SYTO® 9 and propidium iodide (PI) according to manual

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(LIVE/DEAD® BacLight™, Invitrogen) to identify live and dead cells. Concurrently,

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biofilm extracellular polymeric substances (EPS) were stained with 50 mM concanavalin A

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(Con A, Alexa Flour® 633, Invitrogen). All samples were stained for 40 min in the dark.

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Biofilms were rinsed again before viewing with the confocal microscope in a custom-made

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biofilm viewing cell.

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Confocal images were captured using a CLSM equipped with a Plan-Apochromat

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20×/0.8 numerical aperture objective. A minimum of seven Z stack random fields (635 µm ×

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635 µm) were collected for each sample, with a slice thickness of 2.3 µm, using ZEN® (Carl

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Zeiss, Inc.). SYTO® 9 was excited with an argon laser at 488 nm, PI was excited using a

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diode-pumped solid state (DPSS) 561 nm laser, and Con A was excited with a helium-neon

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633 nm laser. FITC, Cy3, and Cy5 filter sets were used. Biovolume was calculated using the

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COMSTAT223 plug-in for ImageJ 1.41 software24. Automatic thresholding for each image

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stack was performed using Otsu’s method.

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Contact angle measurements of control and biofouled membranes with DI water were

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carried out using a Goniometer (VCA Video Contact Angle System, AST Products, Billerica,

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MA). Ten drops of 2 µL on at least two different membrane samples were measured.

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DNA Extraction and Sequencing. Both RO and MD were carried out in a closed-

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loop system, allowing for sampling of the feed before entering the system (‘initial feed’) and

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at the end of the experiment (‘final feed’). Feed water samples (as much as 1 L) were filtered

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through a 0.22 µm Durapore membrane (Millipore GVWP) and, together with the membrane

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subsamples, frozen at -80 ˚C until DNA extraction. After thawing, biomasses from the filters

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and membrane samples were resuspended with 1 mL lysis buffer (40 mM EDTA, 50 mM tris

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pH = 8.3, and 0.75 M sucrose) in 2 mL screw-cap tubes. Nucleic acids were then extracted

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using a phenol–chloroform extraction method modified according to Massana et al.25 Paired-

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end sequencing of the extracted DNA was performed on an Illumina MiSeq platform26 by

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Research and Testing Laboratory (Lubbock, Texas). Bacterial 16S rRNA variable regions

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V1-V3 were targeted using the 28f and 519r primer pair.27

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Microbial Community Analysis. The forward and reverse reads were merged and

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denoised using the Illumina Paired-End Read Merger (PEAR28). All further analysis was

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performed with the pipeline Quantitative Insights Into Microbial Ecology (QIIME), version

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1.729. Sequences with fewer than 200 bases, quality scores lower than 25, more than 6

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homopolymers, or any ambiguous bases were removed. Remaining sequences were clustered

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into operational taxonomic units (OTUs) based on 97% similarity with the UCLUST

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algorithm,30 and the cluster seeds were chosen to represent each OTU for downstream

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analysis. These representative sequences were aligned to the 97% clustered Greengenes

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database (August 2013).31,32 Chimeras were removed with ChimeraSlayer.33

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Taxonomy was assigned to OTUs by matching the representative sequences to the

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complete August 2013 Greengenes database with the RDP Classifier at a 0.9 confidence

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level.34,35 To analyze α-diversity, all samples were first trimmed to an even size of 22,000

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sequences with a single rarefaction. Then, the trimmed dataset was rarefied 100 times from 0

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to 22,000 seqs/sample, at increments of 1,000. For β-diversity analysis, unweighted and

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weighted UniFrac distances between all samples were calculated from the trimmed

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dataset,36,37 and principle coordinates analysis (PCoA) was performed on the resulting

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distance matrices.

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RESULTS AND DISCUSSION

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Natural Sea Water Feed Characteristics. Estuarine water from Long Island Sound was

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collected near the mouths of the Quinnipiac River and Morris Creek during the autumn and

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winter (Figure S1). Before passing through the MD and RO systems, the collected sea water

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was pretreated by microfiltration (10 µm) to remove suspended particles and larger

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planktonic organisms. Key nutrients and biological characteristics of the pre-filtered feed are

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summarized in Table 1. Overall, the water collected during the autumn contained lower

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concentrations of nutrients than the water collected in the winter. Correspondingly, dissolved

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carbon, nitrogen, and phosphorus followed the traditional Redfield ratio38 (102:20:1, Table

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1). A different ratio was observed during the winter (32:6:1), in addition to increased

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concentrations of carbon, nitrogen, and phosphorous. Autumn pretreated feed water

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contained 8.9 × 105 bacteria cells mL-1 (photosynthetic picoplankton and heterotrophic

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picoplankton, Table 1). Winter pre-filtered feed water contained between 6.1 × 105 and 1.7 ×

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106 cells mL-1. Although the composition of these samples is quite different, these differences

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are not necessarily due to season. It is possible that the differences between each of these

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samples are caused by variations in the tides and river flows into the region at the time of

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collection.

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TABLE 1

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Next-generation sequencing (MiSeq Illumina26) was used to evaluate the microbial

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community structure of the initial feed water. Detailed results of the sequencing are provided

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in the SI. Despite the differences in chemical composition of the two feed water samples

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collected (Table 1), the microbial community present in this water remained similar in

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species richness, evenness, and most prevalent operational taxonomic units, or OTUs (Figure

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1A, Tables S1, S2). The most prevalent OTUs in the initial feeds represented the family

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Pelagibacteraceae and the genera Octadecabacter, Sediminicola, and Loktanella. All of these

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bacteria genera are known members of sea water microbial communities. Pelagibacteraceae,

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of the class Alphaproteobacteria, is commonly found in the world’s oceans, and it includes

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Pelagibacter ubique, one of the more abundant members of the marine bacteria

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community.39,40 The genus Octadecabacter contains known psychrophiles (organisms that

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grow and reproduce at cold temperatures), including Octadecabacter arcticus and

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Octadecabacter antarcticus.41 Given the cold temperature of the sea water feed (3-7 ˚C,

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Table 1), the presence of these bacteria is not surprising. The genus Sediminicola contains

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organisms discovered in marine sediment,42 and species from the genus Loktanella has been 7 ACS Paragon Plus Environment

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observed in sea water and beach sand.43,44 Because water samples were collected close to the

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shore, the presence of bacteria commonly found in beach sand is not surprising (Figure S1).

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In practice, the feed water microbial community may be slightly different, as seawater

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desalination plant intakes would be located farther from shore.

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FIGURE 1

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Microbial Community Dynamics in MD and RO Feed Reservoirs. MD and

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RO are two very different desalination processes, especially with respect to operating

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temperature and pressure, and the response of the natural microbial community structure

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upon entrance into each of these systems is markedly different. Total bacteria concentration

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in the initial feed in each of the reservoirs (MD and RO) ranged from 8.9 × 105 to 1.7 × 106

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cell mL-1 (Table 1) and was dominated by heterotrophic bacteria species (Figure S3). Over

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the course of the MD experiments, total bacteria decreased by ~2 orders of magnitude (Figure

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2A). Given the bacterial concentration during the experiment, we conclude that the microbial

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community shifts observed in the MD feed were a result of temperature-related bacterial

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inactivation. We suggest that the drastic feed reservoir temperature change (from 3-7 ˚C to 60

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˚C) resulted in severe heat stress and subsequent population decline. In contrast, the bacteria

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concentration in the RO feed reservoir remained relatively stable throughout the entire

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experiment, with slight fluctuations that may be due to ecological succession in the altered

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environment (Figure 2B).

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FIGURE 2

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The assigned taxonomy and the principle coordinates of the unweighted and weighted

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UniFrac distance show changes in the microbial community after 4 days in the RO and MD

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systems (Figure 1B,C). In each system, members of the most prevalent OTUs in the initial

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sea water feed decreased over time and were not abundant in the final feeds of RO or MD

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(Table S2). The most abundant members of the final RO feed water community included

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OTUs that represented the family Rhodobacteraceae and the genera Erythrobacter, Ralstonia,

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and Sediminicola (Table S3). Rhodobacteraceae, members of the class Alphaproteobacteria,

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were observed in RO membrane biofilms previously,45–47 and members of the genus

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Erythrobacter (class Alphaproteobacteria and order Sphingomonadales) were some of the

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first marine bacteria discovered to contain cell membranes with glycosphingolipids.48 It has

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been postulated that bacteria that produce glycosphingolipids are some of the first colonizers

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of RO membranes.16,49–51 The genus Ralstonia, of the Betaproteobacteria class and the order



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Burkholderiales, has also been reported to colonize RO membranes.52,53 Some species of the

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genus Ralstonia are known to degrade a wide variety of contaminants that could potentially

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be present in the feed samples, which was taken near an industrial area.54,55 Although

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members of the genus Sediminicola have not yet been reported in RO systems, members of

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its family Flavobacteria have.47,56–58 The presence of Sediminicola, a genus commonly found

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in sediment, in our system may also be due to our collection of water near the shore.42 We

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postulate that the genetic similarities observed between the feed water and RO membrane

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result from physical deposition, as previously observed.46 Moreover, it may be that the

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aqueous environment of the RO system assists in selecting the organisms responsible for RO

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biofouling.

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The biggest difference between the RO and MD feed waters are temperature, with the

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RO feed reservoir at 25 ˚C and the MD feed reservoir at 60 ˚C. This temperature difference

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led to significant changes in the bacteria community (Figure 1B,C). ANOSIM analysis

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(excluding initial feed communities) of the unweighted UniFrac distances indicates

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significant (p = 0.017) differences between the grouped RO (biofilms and final feeds) and

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grouped MD samples (biofilms and the final feeds). Unlike the unweighted UniFrac

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distances, the weighted UniFrac distances take into account the relative abundance of each

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OTU, and the differences between the RO and MD group weighted UniFrac distances were

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not as pronounced (p = 0.087). Therefore, the rare taxa may be driving these differences.

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Major contributors to the final MD feed included members genera of the genera

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Ralstonia and Erythrobacter and the order Bacillales. The OTUs identified by Ralstonia and

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Erythrobacter are shared between the final MD and RO feeds. This similarity is curious but

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not unreasonable, as members of Ralstonia have been shown to be thermotolerant,55 as have

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close relatives to Erythrobacter.59 Members of the order Bacillales can form spores that allow

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them to survive in extreme environments. Thus, they have been found in hot springs60 and

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marine vents.61 Members of this family were also previously observed on MD membranes.62

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The differences in the taxa present in both RO and MD systems can also be seen in

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Figure 3, which shows the 15 most abundant orders (averaged over all MD samples and all

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RO samples). The orders Burkholderiales, Flavobacteriales, and Rhodobacterales are

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prevalent in MD and RO biofilms. In the MD system, Burkholderiales, including the genus

316

Ralstonia, appears to be more prevalent. The order Flavobacteriales, which makes up a

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significantly larger portion of the RO community than the MD community (p = 0.08,

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Student’s t-test), includes Sediminicola and Olleya. The order Rhodobacterales includes 9 ACS Paragon Plus Environment

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Octadecabacter and Loktanella. Additional differences between the microbial communities

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in the RO and MD systems include an increase in Rhizobiales (primarily Methylobacterium)

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and a decrease in Thiotrichales (primarily Methylophaga) in the MD system compared to RO.

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Both of these bacteria are methylotrophs which degrade single carbon species. These changes

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contribute to the differences observed in the UniFrac distances between each group (Figure

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1B,C). Thus, we see a clear divergence in the microbial communities after entering the RO

325

and MD systems.

326

FIGURE 3

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In the case of both MD and RO, a strong selection away from the bacteria naturally

328

abundant in the seawater was observed in the feed reservoirs by the end of the experiment. In

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MD, it is likely that this selection is a result of bacterial decline at high temperatures (Figure

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2A). In RO, however, stable bacteria concentrations indicate succession (Figure 2B).

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Nevertheless, species richness of MD samples (both the final feed and the biofilm) exceeded

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the richness of the RO samples (Figure 1A) and more closely resembled the richness of the

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initial feed sea water. It has been shown that species richness in an environment undergoing

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succession does not decrease until certain species are able to grow and dominate the system.63

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Thus, although bacteria are actively multiplying in the RO system, the microbial community

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in the MD system was unable to reach this point of ecological succession.

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Biofilms on MD and RO Membranes. Biofilms that developed on MD and RO

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membranes during both seasons were observed to have different architecture (Figure 4) and

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community structure (Tables 2 and S5). In general, the RO membrane was covered with

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homogeneous microbial mat (Figure 4B,D) while the MD membrane biofilm consisted of

341

several heterogeneous colonies (Figure 4A,C). Moreover, the total biofilm biovolume in MD

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was significantly lower than the total biovolume measured on RO membranes for both the

343

autumn and winter runs (Figure S4).

344

FIGURE 4

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In the autumn, biofilms formed in MD and RO contained mostly live cells with a

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relatively small amount of EPS (Figure S5). During the winter run, the overall trends in

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biofilm morphology resembled the autumn run  the MD biofilm appeared heterogeneous

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with several colonies (Figure 4C), and the RO biofilm was a relatively homogenous mat

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(Figure 4D). However, the biofilms formed on both MD and RO membranes in the winter

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contained a greater amount of dead cells (increased 3.1× in MD and 4.1× in RO) and EPS 10 ACS Paragon Plus Environment


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(increased 3.0× in MD and 4.4× in RO) (Figure S5). It is likely that these differences in

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biofilm composition were a result of different chemical or biological components in the

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initial sea water feeds. It is also likely that the different composition of the biofilm in the

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winter, especially the greater volume of EPS,64 played a role in the increased flux decline

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observed in RO (discussed in the next section).

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We postulate that the large differences in hydraulic conditions (i.e. pressure) have

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dictated the overall biofilm architecture. We observe here that the overall structure of the

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biofilms formed in MD is very different from the biofilms formed in RO. Being a membrane

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process that is not driven by hydraulic pressure, MD is more similar to forward osmosis (FO)

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than RO. The heterogeneous structure of the biofilm containing larger colonies is analogous

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to biofilms previously observed in FO.65 However, although the microscopic structure of the

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biofilms formed in MD is similar to those formed in FO, it is likely that the microbial

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community members of this biofilm are quite different than the members in biofilms formed

364

in other membrane processes due to the high temperatures employed in MD (Figure 1, Table

365

2). Additionally, although we observed a ~2 orders of magnitude decline in bacteria in the

366

feed reservoir (Figure 2A), CLSM live/dead staining indicates that the bacteria in the

367

membrane biofilms are alive and produce EPS (Figures 4A,C and S5). Thus, bacteria are able

368

to live on the MD membrane surface despite a steep, temperature-related decline in bacteria

369

concentration in the feed water.

370

Many of the OTUs that we observed in the initial feed and the RO biofilms were also

371

present in the MD biofilm (Table 2). However, certain organisms favored the MD membrane

372

surface. These organisms, represented an order of magnitude more in the biofilm than the

373

MD feed, are underlined in Table 2. Many of these organisms, including Octadecabacter,

374

Pelagibacteraceae, and Loktanella, have been discussed above, but they also include

375

Vibrioniceae and Cryomorphaceae. Vibrioniceae is a common marine bacterium, and

376

members of Cryomorphaceae, despite their name, have been shown to grow at warmer

377

temperatures.66

378

TABLE 2

379

Notably, fewer thermophilic organisms were present in the MD biofilm than we had

380

initially hypothesized. It is possible that the lack of thermophilic organisms detected in the

381

MD biofilm is a result of temperature polarization in the MD membrane module, which is

382

greatly influenced by operating conditions. In these experiments, the temperature at the


Page 12 of 28

Page 13 of 28


383

channel inlet was held at ~50 °C, and by the MD cell outlet the temperature had decreased by

384

3.8 °C. If we assume that all of this heat loss occurred through the membrane, then the

385

temperature at the membrane surface during operation was calculated to be ~13 °C lower

386

than the bulk (i.e. ~37 °C) (details in the Supporting Information). Thus, provided an

387

organism can survive for a short time at ~60 °C in the feed reservoir, that organism can

388

deposit and grow on the cooler MD membrane surface. This mode of action may be very

389

important for organisms that can form protective spores, e.g. Bacillus sp.

390

Overall, during the course of these experiments, we have observed a shift in the

391

natural microbial community in the sea water feed (Figure 1B,C). Shifts occur in both RO

392

and MD, and species that can survive and thrive in each of these environments dominate the

393

microbial communities in these systems. Our 4 day experiment is a snapshot of the initial

394

biofilms that form in these systems, and the differences between these two communities

395

would likely increase with time.

396

Impact of Biofouling on MD and RO Performance. Given a similar initial

397

water flux (20 ± 2 L m-2 h-1) and cross flow velocity (4.3 cm s-1), very different permeate

398

water flux behaviors were measured throughout the course of each experiment (Figure 5). In

399

autumn, permeate water flux in MD remained stable throughout the 4 day experiment (Figure

400

5A), and a slight decline (< 5%) of permeate water flux was observed in RO (Figure 5B). In

401

contrast, winter runs exhibited a total flux decline of 50% in MD and 7% RO. The sharp

402

decline in water flux (50%) in MD occurred during the first 12 h of the experiment. MD

403

experiments were repeated twice, verifying that the severe water flux decline observed was

404

the result of the fouling propensity of the feed water. These variations between autumn and

405

winter runs most likely resulted from a significant difference in fouling propensities of the

406

natural sea water.

407

FIGURE 5

408

Our results suggest that the fouling in MD resulted in partial pore blockage by

409

biological and/or organic constituents in the feed water that were not removed by the

410

microfiltration (10 µm) pretreatment. In the winter, poorer water quality was observed, with

411

DOC increasing 25%, particulate carbon increasing 135%, and total phosphorous increasing

412

426% (Table 1), potentially leading to rapid (~12 h) flux decline. These differences in water

413

quality likely impacted bacterial growth (Table 2) and EPS production (Figures 4 and S4).

414

Additionally, the severe flux decline in the winter MD (Figure 5A) was mirrored by a greater 12 ACS Paragon Plus Environment


415

flux decline in RO (Figure 5B). However, RO flux decline in the winter was not as large or as

416

rapid as the MD flux decline, indicating that MD is more sensitive to certain changes in feed

417

water quality than RO. This observation is a result of the different structure and fouling

418

mechanisms of these membranes. RO membranes have a dense, salt-rejecting selective layer.

419

When a biofilm forms on these membranes, it increases the hydraulic resistance to permeate

420

water flow and decreases the net driving force for water permeation through biofilm

421

enhanced osmotic pressure.64,67,68 Thus, the fouling in RO in the winter may be due to the

422

hydraulic resistance of EPS produced over time. In contrast, fouling of a porous MD

423

membrane as observed in Figure 5A is attributed to blocking of pores of the microporous

424

membrane, which increases the resistance to water vapor transport through the membrane.13

425

The significant fouling and flux decline in the winter MD run was not accompanied

426

by wetting of the hydrophobic MD membrane. When water enters the pores of the membrane,

427

the membrane no longer provides a barrier for passage of salt.62 However, in our

428

experiments, no increase in distillate water conductivity was observed. We did, however, see

429

that the formation of a biofilm on the membrane surface greatly decreased the water contact

430

angle from 134 ± 4˚ to 32 ± 6˚ (Figure S5). Thus, a hydrophilic membrane surface is not the

431

only requirement for membrane wetting. This hydrophilic film must additionally coat the

432

interior membrane pores in order to impact membrane performance.

433

Proposed Biofouling Regimes in MD and RO. As natural sea water undergoes

434

changes in the RO system (temperature, pressure, and hydrodynamics), the microbial

435

community shifts, and certain organisms that are able to survive under these conditions begin

436

to grow. The bacteria that thrive in the feed water are very similar to those that form the

437

initial biofilm on the RO membrane (Tables S3 and S5). Thus, the aquatic environment of the

438

RO process may help select the organisms responsible for RO membrane biofilms. It may

439

also be that many bacteria deposit on the RO membrane surface due to flow conditions

440

during operation. The impact of operating conditions, including pressure and permeate flow,

441

on biofilm structure in RO must be understood in order to develop successful biofouling

442

mitigating strategies.

443

The microbial community undergoes a much different transformation in the hotter

444

MD system, and there are larger differences between the microbial community in the final

445

feed and the membrane biofilm due to temperature gradients, temperature polarization, and

446

bacterial attachment. Once the sea water feed is placed in the MD system, the number of

447

bacteria decreases. Despite this decline, organisms, along with other macromolecular and 13 ACS Paragon Plus Environment

Page 14 of 28

Page 15 of 28


448

colloidal matter, can block membrane pores and decrease distillate water flux (Figure 5A). In

449

fact, the MD membrane surface may be one of the most favorable locations in our system for

450

microbial growth, as it is the coolest location on the feed side of our system. Depending on

451

the temperatures employed and temperature polarization in a given system, bacteria that are

452

able to survive for a short time at high temperatures may attach and grow on the cooler

453

membrane surface. Thus, even with a filtration pretreatment and relatively high operating

454

temperatures, biofouling may pose a problem for MD implementation.

455 456

Supporting Information

457

Location of sea water feed sample collection (Figure S1); closed-loop MD system (Figure

458

S2); cell counts in MD and RO feed reservoirs (Figure S3); biovolume in MD and RO

459

biofilms (Figure S5); contact angle measurements on pristine and biofouled MD membranes

460

(Figure S5); sequencing and quality trimming results, temperature polarization in MD,

461

observed OTUs and Shannon diversity in initial feed (Table S1); 10 most abundant OTUs in

462

initial feeds (Table S2); 10 most abundant OTUs in final RO feeds (Table S3); 10 most

463

abundant OTUs in final MD feed (Table S4); 10 most abundant OTUs in RO biofilm (Table

464

S5); 30 most abundant OTUs in MD biofilms (Table S6). This information is available free of

465

charge via the Internet at http://pubs.acs.org/.

466 467

ACKNOWLEDGEMENTS

468

This research was made possible by the National Science Foundation Graduate Research

469

Fellowship to Katherine R. Zodrow (Grant No. DGE-1122492). We also acknowledge

470

support provided to Dr. Edo Bar-Zeev by the United States-Israel Binational Agricultural

471

Research and Development (BARD) postdoctoral fellowship fund. We thank Eyal Rahav for

472

assistance with flow cytometry, and Dr. Shihong Lin for assistance with the temperature

473

polarization model. Lastly, we thank Dr. Joseph Wolenski from the Molecular, Cellular, and

474

Developmental Biology Department at Yale University for technical assistance using the

475

CLSM.

476 477



478

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673 674


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675

Table1. Sea water characteristics. Chemical composition of sea water was determined using

676

standard methods. Total bacteria was determined using flow cytometry.

Temperature

Units

Autumn

Winter

˚C

7

3

7.71

7.68

mS cm-1

49.1

43.2

g L-1

26.6

23.3

-1

1.85

2.32

-1

0.036

0.034

-1

0.0987

0.1720

-1

0.0182

0.0722

-1

0.0181

0.0776

-1

0.38

0.43

-1

0.34

0.46

-1

0.0211

0.0433

-1

0.1140

0.268

pH Conductivity Salinity Total Dissolved Carbon Ammonium Nitrate + Nitrite Total Dissolved Phosphorous Total Phosphorous Total Dissolved Nitrogen

mg-C L

mg-N L mg-N L mg-P L mg-P L

mg-N L

Total Nitrogen

mg-N L

Particulate Nitrogen

mg-N L

Particulate Carbon

mg-C L

Total Bacteria

cells mL

-1

8.9 × 10

677 678


5

6.1 – 17 × 10

5


Page 22 of 28

679

Table 2. Ten most abundant OTUs in the MD Biofilms (%). Percent composition is given,

680

along with percent composition in the final MD feed. Underlined OTUs are present at least

681

one order of magnitude more in the biofilm than the feed. Unless otherwise noted, taxonomy

682

is given at the genus level. If genus level taxonomy is not available, family (f) or order (o) is

683

given. (A = autumn, W = winter)

684

Biofilm (A)

Biofilm (W)

Final Feed (A)

Final Feed (W)

Ralstonia

5.7

34.4

44.7

22.3

Octadecabacter

17.1

4.6

0.1

0.3

Pelagibacteraceae (f)

7.5

6.2

0.0

0.0

Loktanella

6.1

4.8

0.9

0.6

Sediminicola

7.5

3.0

0.0

2.4

Vibrionaceae (f)

8.4

1.3

0.5

0.0

Rhodobacteraceae (f)

4.8

3.9

0.4

5.7

Cryomorphaceae (f)

4.7

2.2

0.1

0.0

Flavobacteriaceae (f)

4.2

1.9

0.0

1.5

Bacillales (o)

5.3

0.4

23.4

0.0

Sum

71.1

62.8

70.2

32.7

685 686


Page 23 of 28


A Observed OTUs

3000

2000

1000 Initial Feed MD RO

5000 10000 15000 20000

Sequences per Sample B PC2 (14%)

0.4

0.2

0.0

-0.2 -0.4

-0.2

0.0

0.2

0.4

PC1 (16.9%)

PC2 (21.3%)

0.4

C

Initial Feed Final MD Feed RO Biofilm

MD Biofilm Final RO Feed

0.2

0.0

-0.2 -0.2

687

0.0

0.2

0.4

PC1 (48.6%)

688

Figure 1. Alpha- and Beta- diversity of microbial communities in this study. (A) Rarefaction

689

curves with all MD samples, including final feed and biofilms (‘MD’) and all RO samples

690

(‘RO’). Curves represent means and standard deviations of all samples in a group. (B)



691

Unweighted, and (C) weighted principle coordinate plot (B) of UniFrac distance for all

692

samples.

693


Page 24 of 28

Page 25 of 28


B) RO

A) MD

Cells mL-1

106

105

104

Autumn Winter

Autumn Winter 103

0

20

40

60

80

100

0

Time (h)

694

20

40

60

80

100

Time (h)

695 696 697

Figure 2. Concentration of cells (photosynthetic picoplankton and total bacteria) in feed

698

reservoirs during (A) MD and (B) RO. 1.8 mL samples were taken from the feed reservoirs

699

throughout each experiment, frozen at -80 ˚C in glutaraldehyde, and measured using flow

700

cytometry.

701



Page 26 of 28

MD RO

0.4 0.3 0.2

*

0.1

Acidimicrobiales

Campylobacterales

Alphaproteobacteria (c)

Rickettsiales

Vibrionales

Rhizobiales

Alteromonadales

Bacillales

Pseudomonadales

Sphingomonadales

Rhodobacterales

Flavobacteriales

Burkholderiales

Oceanospirillales

*

0.0

Thiotrichales

Fraction of Total OTUs

0.5

702 703 704 705

Figure 3. Comparison between top 15 OTUs in MD and RO samples (biofilm and final feed).

706

Taxa are grouped according to order. Significant differences between RO and MD samples

707

are marked with an asterisk (p < 0.05). Additional orders of interest are underlined.

708 709 710


Page 27 of 28


711 712

Figure 4. CLSM orthagonal views of biofilms formed in membrane distillation (A, autumn;

713

C, winter) and reverse osmosis (B, autumn; D, winter). All sizes are in µm. The left inset in

714

(A) is in a higher horizontal plane than the main orthogonal image.

715 26 ACS Paragon Plus Environment


Page 28 of 28

716 717

A) MD

B) RO

Normalized Flux

1.2 1.0 0.8 0.6 0.4

Autumn Winter

Autumn Winter

0.2 0 718

25

50

75

0

Time (h)

25

50

75

Time (h)

719 720 721

Figure 5. Normalized permeate flux during (A) MD and (B) RO experiments. Initial flux was

722

20 ± 2 L m-2 h-1. Crossflow velocity was 4.3 cm s-1. RO and MD feed temperature were

723

maintained at 25 and 50 ˚C, respectively.

724


Biofouling and Microbial Communities in ... - ACS Publications

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