Biofouling and Microbial Communities in Membrane Distillation and

Subscriber access provided by NEW YORK UNIV

Article

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

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Environmental Science & Technology is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 28

Environmental Science & Technology

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38

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

ACS Paragon Plus Environment


39

Abstract

40

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

42

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

43

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

44

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

45

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

46

desalination systems expose the natural microbial community to vastly different

47

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

48

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

49

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

50

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

51

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

52

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

53

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

54

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

55

Bacteria representing the orders Burkholderiales, Rhodobacterales, and Flavobacteriales were

56

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

57

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

58 59

TOC ART

60 61 1 ACS Paragon Plus Environment

Page 2 of 28

Page 3 of 28


62 63

INTRODUCTION

64

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

65

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

66

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

67

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

68

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

69

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

70

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

71

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

72

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

73

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

74

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

75

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

76

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

77

in MD systems.

78

Biofouling in MD may lead to increased temperature polarization (thermal

79

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

80

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

81

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

82

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

83

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

84

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

85

thermophilic

86

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

87

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

88

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

organisms,

including

Meithermus

hypogaeus,

Tepidimonas

sp,

and

89

There are some major differences between these membrane-based desalination

90

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

91

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

92

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

93

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


Page 4 of 28

94

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

95

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

96

higher temperatures than RO.

97

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

98

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

99

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

100

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

101

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

102

development of anti-biofouling materials and treatments.

103 104

MATERIALS AND METHODS

105

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

106

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

107

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

108

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

109

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

110

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

111

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

112

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

113

MD and RO bench-scale systems.

114

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

115

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

116

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

117

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

118

Environmental Science, Chesapeake Biological Laboratory).

119

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

Particulate and dissolved

120

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

121

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

122

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

123

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

124

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

125

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

Page 5 of 28


126

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

127

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

128

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

129

dark.

130

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

131

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

132

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

133

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

134

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

135

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

136

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

137

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

138

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

139

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

140

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

141

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

142

was observed during any of the experiments.

143

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

144

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

145

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

146

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

147

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

148

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

149

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

150

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

151

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

152

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

153

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

154

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

155

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

156

flux.

18

. RO membrane salt rejection was

157

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

158

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


159

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

160

glutaraldehyde prior to measurement of photosynthetic picoplankton and total bacteria.

161

Bacteria Abundance in Feed Reservoirs. Photosynthetic picoplankton

162

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

163

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

164

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

165

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

166

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

167

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

168

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

169

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

170

forward-scatter (a proxy of cell size22).

171

Membrane Biofilm Characterization. Biofilm formation on the MD and RO

172

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

173

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

174

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

175

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

176

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

177

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

178

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

179

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

180

biofilm viewing cell.

181

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

182

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

183

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

184

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

185

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

186

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

187

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

188

stack was performed using Otsu’s method.

5 ACS Paragon Plus Environment

Page 6 of 28

Page 7 of 28


189

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

190

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

191

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

192

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

193

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

194

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

195

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

196

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

197

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

198

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

199

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

200

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

201

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

202

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

203

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

204

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

205

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

206

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

207

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

208

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

209

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

210

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

211

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

212

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

213

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

214

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

215

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

216

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

217

weighted UniFrac distances between all samples were calculated from the trimmed

218

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

219

distance matrices.

220



221

RESULTS AND DISCUSSION

222

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

223

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

224

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

225

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

226

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

227

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

228

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

229

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

230

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

231

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

232

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

233

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

234

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

235

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

236

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

237

collection.

238

TABLE 1

239

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

240

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

241

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

242

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

243

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

244

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

245

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

246

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

247

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

248

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

249

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

250

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

251

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

252

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

253

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

Page 8 of 28

Page 9 of 28


254

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

255

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

256

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

257

desalination plant intakes would be located farther from shore.

258

FIGURE 1

259

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

260

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

261

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

262

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

263

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

264

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

265

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

266

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

267

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

268

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

269

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

270

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

271

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

272

environment (Figure 2B).

273

FIGURE 2

274

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

275

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

276

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

277

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

278

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

279

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

280

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

281

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

282

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

283

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

284

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

285

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



286

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

287

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

288

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

289

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

290

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

291

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

292

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

293

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

294

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

295

biofouling.

296

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

297

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

298

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

299

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

300

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

301

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

302

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

303

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

304

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

305

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

306

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

307

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

308

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

309

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

310

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

311

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

312

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

313

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

314

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

315

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

317

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

318

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

Page 10 of 28

Page 11 of 28


319

Octadecabacter and Loktanella. Additional differences between the microbial communities

320

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

321

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

322

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

323

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

324

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

325

and MD systems.

326

FIGURE 3

327

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

329

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

330

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

331

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

332

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

333

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

334

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

335

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

336

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

337

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

338

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

339

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

340

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

342

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

343

autumn and winter runs (Figure S4).

344

FIGURE 4

345

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

346

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

347

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

348

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

349

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

350

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


351

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

352

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

353

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

354

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

355

observed in RO (discussed in the next section).

356

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

357

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

358

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

359

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

360

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

361

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

362

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

363

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

REFERENCES

479 480

(1)

Elimelech, M.; Phillip, W. A. The future of seawater desalination: Energy, technology, and the environment. Science 2011, 333, 712–717.

481 482

(2)

Logan, B. E.; Elimelech, M. Membrane-based processes for sustainable power generation using water. Nature 2012, 488, 313–319.

483 484

(3)

Khayet, M. Membranes and theoretical modeling of membrane distillation: A review. Adv. Colloid Interface Sci. 2011, 164, 56–88.

485 486 487

(4)

Saffarini, R. B.; Summers, E. K.; Arafat, H. A.; Lienhard V, J. H. Economic evaluation of stand-alone solar powered membrane distillation systems. Desalination 2012, 299, 55–62.

488 489 490

(5)

Saffarini, R. B.; Summers, E. K.; Arafat, H. A.; Lienhard V, J. H. Technical evaluation of stand-alone solar powered membrane distillation systems. Desalination 2012, 286, 332–341.

491 492 493 494

(6)

Shaffer, D. L.; Arias Chavez, L. H.; Ben-Sasson, M.; Romero-Vargas Castrillón, S.; Yip, N. Y.; Elimelech, M. Desalination and reuse of high-salinity shale gas produced water: Drivers, technologies, and future directions. Environ. Sci. Technol. 2013, 47, 9569–9583.

495 496 497

(7)

Xie, M.; Nghiem, L. D.; Price, W. E.; Elimelech, M. A forward osmosis-membrane distillation hybrid process for direct sewer mining : System performance and limitations. Environ. Sci. Technol. 2013, 47, 13486–13493.

498 499

(8)

Gryta, M. The assessment of microorganism growth in the membrane distillation system. Desalination 2002, 142, 79–88.

500 501 502 503

(9)

Guillen-Burrieza, E.; Thomas, R.; Mansoor, B.; Johnson, D.; Hilal, N.; Arafat, H. Effect of dry-out on the fouling of PVDF and PTFE membranes under conditions simulating intermittent seawater membrane distillation (SWMD). J. Memb. Sci. 2013, 438, 126–139.

504 505

(10)

Gryta, M. Fouling in direct contact membrane distillation process. J. Memb. Sci. 2008, 325, 383–394.

506 507

(11)

Nghiem, L. D.; Cath, T. A scaling mitigation approach during direct contact membrane distillation. Sep. Purif. Technol. 2011, 80, 315–322.

508 509

(12)

Srisurichan, S.; Jiraratananon, R.; Fane, a. G. Humic acid fouling in the membrane distillation process. Desalination 2005, 174, 63–72.

510 511 512

(13)

Goh, S.; Zhang, Q.; Zhang, J.; McDougald, D.; Krantz, W. B.; Liu, Y.; Fane, A. G. Impact of a biofouling layer on the vapor pressure driving force and performance of a membrane distillation process. J. Memb. Sci. 2013, 1–13.

513 514

(14)

Krivorot, M.; Kushmaro, a.; Oren, Y.; Gilron, J. Factors affecting biofilm formation and biofouling in membrane distillation of seawater. J. Memb. Sci. 2011, 376, 15–24.

515 516 517

(15)

Elifantz, H.; Horn, G.; Ayon, M.; Cohen, Y.; Minz, D. Rhodobacteraceae are the key members of the microbial community of the initial biofilm formed in Eastern Mediterranean coastal seawater. FEMS Microbiol. Ecol. 2013, 85, 348–357.

518 519

(16)

Bereschenko, L. A.; Heilig, G. H. J.; Nederlof, M. M.; van Loosdrecht, M. C. M.; Stams, A. J. M.; Euverink, G. J. W. Molecular characterization of the bacterial


Page 16 of 28

Page 17 of 28


520 521

communities in the different compartments of a full-scale reverse-osmosis water purification plant. Appl. Environ. Microbiol. 2008, 74, 5297–5304.

522 523 524 525

(17)

Zodrow, K. R.; Coulter, V. H.; Shaulsky, E.; Elimelech, M. Low flow data logger in membrane distillation : An interdisciplinary laboratory in process control. In Interdisciplinary Engineering Design Education Conference (IEDEC); Santa Clara, CA, USA, 2014; pp. 70–73.

526 527 528

(18)

Bar-Zeev, E.; Elimelech, M. Reverse osmosis biofilm dispersal by osmotic backflushing: Cleaning via substratum perforation. Environ. Sci. Technol. Lett. 2014, 1, 162–166.

529 530

(19)

Stambler, N. Light and picophytoplankton in the Gulf of Eilat (Aqaba). J. Geophys. Res. 2006, 111, C11009.

531 532

(20)

Vaulot, D.; Marie, D. Diel variability of photosynthetic picoplankton in the equatorial Pacific. J. Geophys. Res. 1999, 104, 3297–3310.

533 534 535

(21)

Simon, N.; Barlow, R. G.; Marie, D.; Partensky, F.; Vaulot, D. Characterization of oceanic photosynthetic picoeukaryotes by flow cytometry. J. Phycol 1994, 30, 935– 942.

536 537 538

(22)

Robertson, B. R.; Button, D. K.; Koch, A. L. Determination of the biomasses of small bacteria at low concentrations in a mixture of species with forward light scatter measurements by flow cytometry. Appl. Environ. Microbiol. 1998, 64, 3900–3909.

539 540 541

(23)

Heydorn, A.; Nielsen, A. T.; Hentzer, M.; Sternberg, C.; Givskov, M.; Ersbøll, B. K.; Molin, S. Quantification of biofilm structures by the novel computer program COMSTAT. Microbiology 2000, 146, 2395–2407.

542 543

(24)

Abràmoff, M. D.; Hospitals, I.; Magalhães, P. J.; Abràmoff, M. Image Processing with ImageJ. Biophotonics 2004, 11, 36–42.

544 545 546

(25)

Massana, R.; Murray, A. E.; Preston, C. M.; DeLong, E. F. Vertical distribution and phylogenetic characterization of marine planktonic Archaea in the Santa Barbara Channel. Appl. Environ. Microbiol. 1997, 63, 50–56.

547 548 549 550

(26)

Caporaso, J. G.; Lauber, C. L.; Walters, W. a; Berg-Lyons, D.; Huntley, J.; Fierer, N.; Owens, S. M.; Betley, J.; Fraser, L.; Bauer, M.; et al. Ultra-high-throughput microbial community analysis on the Illumina HiSeq and MiSeq platforms. ISME J. 2012, 6, 1621–1624.

551 552

(27)

Campbell, B. J.; Kirchman, D. L. Bacterial diversity, community structure and potential growth rates along an estuarine salinity gradient. ISME J. 2013, 7, 210–220.

553 554

(28)

Zhang, J.; Kobert, K.; Flouri, T.; Stamatakis, A. PEAR: a fast and accurate Illumina Paired-End reAd mergeR. Bioinformatics 2014, 30, 614–620.

555 556 557 558

(29)

Caporaso, J. G.; Kuczynski, J.; Stombaugh, J.; Bittinger, K.; Bushman, F. D.; Costello, E. K.; Fierer, N.; Peña, A. G.; Goodrich, J. K.; Gordon, J. I.; et al. QIIME allows analysis of high- throughput community sequencing data. Nat. Methods 2010, 7, 335– 336.

559 560

(30)

Edgar, R. C. Search and clustering orders of magnitude faster than BLAST. Bioinformatics 2010, 26, 2460–2461.

561 562 563

(31)

Caporaso, J. G.; Bittinger, K.; Bushman, F. D.; DeSantis, T. Z.; Andersen, G. L.; Knight, R. PyNAST: a flexible tool for aligning sequences to a template alignment. Bioinformatics 2010, 26, 266–267. 16 ACS Paragon Plus Environment


564 565 566 567

(32)

DeSantis, T. Z.; Hugenholtz, P.; Larsen, N.; Rojas, M.; Brodie, E. L.; Keller, K.; Huber, T.; Dalevi, D.; Hu, P.; Andersen, G. L. Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Appl. Environ. Microbiol. 2006, 72, 5069–5072.

568 569 570 571

(33)

Haas, B. J.; Gevers, D.; Earl, A. M.; Feldgarden, M.; Ward, D. V; Giannoukos, G.; Ciulla, D.; Tabbaa, D.; Highlander, S. K.; Sodergren, E.; et al. Chimeric 16S rRNA sequence formation and detection in Sanger and 454-pyrosequenced PCR amplicons. Genome Res. 2011, 21, 494–504.

572 573 574 575

(34)

McDonald, D.; Price, M. N.; Goodrich, J.; Nawrocki, E. P.; DeSantis, T. Z.; Probst, A.; Andersen, G. L.; Knight, R.; Hugenholtz, P. An improved Greengenes taxonomy with explicit ranks for ecological and evolutionary analyses of bacteria and archaea. ISME J. 2012, 6, 610–618.

576 577 578

(35)

Wang, Q.; Garrity, G. M.; Tiedje, J. M.; Cole, J. R. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 2007, 73, 5261–5267.

579 580

(36)

Lozupone, C.; Knight, R. UniFrac: A new phylogenetic method for comparing microbial communities. Appl. Environ. Microbiol. 2005, 71, 8228–8235.

581 582 583

(37)

Lozupone, C. A.; Hamady, M.; Kelley, S. T.; Knight, R. Quantitative and qualitative beta diversity measures lead to different insights into factors that structure microbial communities. Appl. Environ. Microbiol. 2007, 73, 1576–1585.

584 585

(38)

Redfield, A. C. The biological control of chemical factors in the environment. Am. Sci. 1958, 46, 205–221.

586 587 588

(39)

Biers, E. J.; Sun, S.; Howard, E. C. Prokaryotic genomes and diversity in surface ocean waters: interrogating the global ocean sampling metagenome. Appl. Environ. Microbiol. 2009, 75, 2221–2229.

589 590 591

(40)

Morris, R. M.; Rappe, M. S.; Connon, S. A.; Vergin, K. L.; Siebold, W. A.; Carlson, C. A.; Giovannoni, S. J. SAR11 clade dominates ocean surface bacterioplankton communities. Nature 2002, 420, 806–810.

592 593 594

(41)

Gosink, J. J.; Herwig, R. P.; Staley, J. T. Octadecabacter arcticus gen. nov., sp. nov., and O. antarcticus, sp. nov., Nonpigmented, psychrophilic gas vacuolate bacteria from Polar sea ice and water. Syst. Appl. Microbiol. 1997, 20, 356–365.

595 596 597

(42)

Khan, S. T.; Nakagawa, Y.; Harayama, S. Sediminicola luteus gen. nov., sp. nov., A novel member of the family Flavobacteriaceae. Int. J. Syst. Evol. Microbiol. 2006, 56, 841–845.

598 599 600

(43)

Lee, J.; Jung, J.-Y.; Kim, S.; Chang, I. S.; Mitra, S. S.; Kim, I. S. Selection of the most problematic biofoulant in fouled RO membrane and the seawater intake to develop biosensors for membrane biofouling. Desalination 2009, 247, 125–136.

601 602 603

(44)

Moon, Y. G.; Seo, S. H.; Lee, S. D.; Heo, M. S. Loktanella pyoseonensis sp. nov., isolated from beach sand, and emended description of the genus Loktanella. Int. J. Syst. Evol. Microbiol. 2010, 60, 785–789.

604 605 606

(45)

Khan, M. M. T.; Stewart, P. S.; Moll, D. J.; Mickols, W. E.; Burr, M. D.; Nelson, S. E.; Camper, A. K. Assessing biofouling on polyamide reverse osmosis (RO) membrane surfaces in a laboratory system. J. Memb. Sci. 2010, 349, 429–437.


Page 18 of 28

Page 19 of 28


607 608

(46)

Khan, M. T.; de O Manes, C.-L.; Aubry, C.; Gutierrez, L.; Croue, J. P. Kinetic study of seawater reverse osmosis membrane fouling. Environ. Sci. Technol. 2013.

609 610 611

(47)

Zhang, M.; Jiang, S.; Tanuwidjaja, D.; Voutchkov, N.; Hoek, E. M. V; Cai, B. Composition and variability of biofouling organisms in seawater reverse osmosis desalination plants. Appl. Environ. Microbiol. 2011, 77, 4390–4398.

612 613

(48)

Shiba, T.; Simidu, U. Erythrobacter longus gen. nov., sp. nov., An aerobic bacterium which contains Bacteriochlorophyll a. Int. J. Syst. Bacteriol. 1982, 32, 211–217.

614 615

(49)

Lee, J.; Kim, I. S. Microbial community in seawater reverse osmosis and rapid diagnosis of membrane biofouling. Desalination 2011, 273, 118–126.

616 617 618

(50)

Bereschenko, L. A.; Stams, A. J. M.; Euverink, G. J. W.; van Loosdrecht, M. C. M. Biofilm formation on reverse osmosis membranes is initiated and dominated by Sphingomonas spp. Appl. Environ. Microbiol. 2010, 76, 2623–2632.

619 620 621

(51)

Al Ashhab, A.; Herzberg, M.; Gillor, O. Biofouling of reverse-osmosis membranes during tertiary wastewater desalination: microbial community composition. Water Res. 2014, 50, 342-349.

622 623 624

(52)

Pang, C. M.; Hong, P.; Guo, H.; Liu, W.-T. Biofilm formation characteristics of bacterial isolates retrieved from a reverse osmosis membrane. Environ. Sci. Technol. 2005, 39, 7541–7550.

625 626 627 628

(53)

Ivnitsky, H.; Katz, I.; Minz, D.; Volvovic, G.; Shimoni, E.; Kesselman, E.; Semiat, R.; Dosoretz, C. G. Bacterial community composition and structure of biofilms developing on nanofiltration membranes applied to wastewater treatment. Water Res. 2007, 41, 3924–3935.

629 630 631 632

(54)

Mergeay, M.; Monchy, Sã©.; Vallaeys, T.; Auquier, V.; Benotmane, A.; Bertin, P.; Taghavi, S.; Dunn, J.; Lelie, D.; Wattiez, R. Ralstonia metallidurans, a bacterium specifically adapted to toxic metals: towards a catalogue of metal-responsive genes. FEMS Microbiol. Rev. 2003, 27, 385–410.

633 634 635

(55)

Lee, S.-K.; Lee, S. B. Isolation and characterization of a thermotolerant bacterium Ralstonia sp. strain PHS1 that degrades benzene, toluene, ethylbenzene, and o -xylene. Appl. Microbiol. Biotechnol. 2001, 56, 270–275.

636 637 638

(56)

Mansouri, J.; Harrisson, S.; Chen, V. Strategies for controlling biofouling in membrane filtration systems: challenges and opportunities. J. Mater. Chem. 2010, 20, 4567.

639 640 641 642

(57)

Ridgway, H. F.; Kelly, A.; Justice, C.; Olson, B. H. Microbial fouling of reverseosmosis membranes used in advanced wastewater treatment technology: chemical, bacteriological, and ultrastructural analyses. Appl. Environ. Microbiol. 1983, 45, 1066–1084.

643 644

(58)

Baker, J.; Dudley, L. Biofouling in membrane systems - A review. Desalination 1998, 118, 81–89.

645 646 647 648

(59)

Yurkov, V. V; Krieger, S.; Stackebrandt, E.; Beatty, J. T. Citromicrobium bathyomarinum, a novel aerobic bacterium isolated from deep-sea hydrothermal vent plume waters that contains photosynthetic pigment-protein complexes. J. Bacteriol. 1999, 181, 4517–4525.

649 650

(60)

Baker, G. C.; Gaffar, S.; Cowan, D. a; Suharto, a R. Bacterial community analysis of Indonesian hot springs. FEMS Microbiol. Lett. 2001, 200, 103–109. 18 ACS Paragon Plus Environment


651 652 653

(61)

Caccamo, D.; Maugeri, T. L.; Gugliandolo, C. Identification of thermophilic and marine bacilli from shallow thermal vents by restriction analysis of their amplified 16S rDNA. J. Appl. Microbiol. 2001, 91, 520–524.

654 655 656

(62)

Goh, S.; Zhang, J.; Liu, Y.; Fane, A. G. Fouling and wetting in membrane distillation (MD) and MD-bioreactor (MDBR) for wastewater reclamation. Desalination 2012, 1– 9.

657 658

(63)

Jackson, C. R. Changes in community properties during microbial succession. OIKOS 2003, 2, 444–448.

659 660 661

(64)

Herzberg, M.; Kang, S.; Elimelech, M. Role of extracellular polymeric substances (EPS) in biofouling of reverse osmosis membranes. Environ. Sci. Technol. 2009, 43, 4393–4398.

662 663

(65)

Yoon, H.; Baek, Y.; Yu, J.; Yoon, J. Biofouling occurrence process and its control in the forward osmosis. Desalination 2013, 325, 30–36.

664 665 666 667

(66)

Lee, D.-H.; Choi, E.-K.; Moon, S.-R.; Ahn, S.; Lee, Y. S.; Jung, J. S.; Jeon, C. O.; Whang, K.-S.; Kahng, H.-Y. Wandonia haliotis gen. nov., sp. nov., a marine bacterium of the family Cryomorphaceae, phylum Bacteroidetes. Int. J. Syst. Evol. Microbiol. 2010, 60, 510–514.

668 669 670

(67)

Hoek, E. M. V; Elimelech, M. Cake-enhanced concentration polarization: A new fouling mechanism for salt-rejecting membranes. Environ. Sci. Technol. 2003, 37, 5581–5588.

671 672

(68)

Herzberg, M.; Elimelech, M. Biofouling of reverse osmosis membranes : Role of biofilm-enhanced osmotic pressure. J. Memb. Sci. 2007, 295, 11–20.

673 674


Page 20 of 28

Page 21 of 28


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 Membrane Distillation and

Recommend Documents