Influence of Electric Fields on Biofouling of Carbonaceous Electrodes

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The influence of electric fields on biofouling of carbonaceous electrodes Soumya Pandit, Sneha Shanbhag, Meagan S Mauter, Yoram Oren, and Moshe Herzberg Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.6b06339 • Publication Date (Web): 25 Jul 2017 Downloaded from http://pubs.acs.org on July 29, 2017

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Environmental Science & Technology

The influence of electric fields on biofouling of carbonaceous electrodes

1 2 3 4 5

Soumya Pandit1, Sneha Shanbhag2, Meagan Mauter2,3, Yoram Oren1 and Moshe Herzberg1*

6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Author Affiliations: 1

Zuckerberg Institute for Water Research, Blaustein Institutes for Desert Research, Ben Gurion University of the Negev, Midreshet Ben Gurion, 84990, ISRAEL 2

Department of Civil & Environmental Engineering, Carnegie Mellon University, 5000 Forbes Ave., Pittsburgh, PA, 15213, USA

3

Department of Engineering and Public Policy, Carnegie Mellon University, 5000 Forbes Ave., Pittsburgh, PA, 15213, USA

*Author to Whom Correspondence Should Be Addressed e-mail: [email protected] phone: +972-86563520

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Abstract

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Biofouling commonly occurs on carbonaceous capacitive deionization electrodes in the

38

process of treating natural waters. Although previous work reported the effect of electric

39

fields on bacterial mortality for variety of medical and engineered applications, the effect

40

of electrode surface properties and the magnitude and polarity of applied electric fields on

41

biofilm development has not been comprehensively investigated. This paper studies the

42

formation of Pseudomonas aeruginosa biofilm on Papyex graphite (PA) and carbon

43

aerogel (CA) in presence and absence of an electric field. The experiments were

44

conducted using a two-electrode flow cell with voltage window of ±0.9 V. The carbon

45

aerogel was less susceptible to biofilm formation compared to Papyex graphite due to its

46

lower

47

properties. For both positive and negative applied potentials, we observed an inverse

48

relationship between biofilm formation and the magnitude of the applied potential. The

49

effect is particularly strong for CA electrodes, and may be a result of cumulative effects

50

between material toxicity and the stress experienced by cells at high applied

51

potentials. Under the applied potentials for both electrodes, high production of

52

endogenous reactive oxygen species (ROS) was indicative of bacterial stress. For both

53

electrodes, the elevated specific ROS activity was lowest for the open circuit potential

54

(OCP) condition, elevated when cathodically and anodically polarized, and highest for

55

the ±0.9 V cases. These high applied potentials are believed to affect the redox potential

56

across the cell membrane and disrupt redox homeostasis, thereby inhibiting bacterial

57

growth.

surface

roughness,

lower

hydrophobicity,

and

significant antimicrobial

58 59 60

Keywords: Carbon electrodes; Biofilm; Electric field; Reactive Oxygen Species; Bioelectric Effect.

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70 71


TOC ART Open circuit

- 0.9 V

+ 0.9 V

Reactive oxygen species

72 73

Working electrode (carbon aerogel/graphite)

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1.

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Introduction

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Capacitive deionization (CDI) is an energy efficient process for low salinity brackish

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water treatment, in which dissolved ions are electro-adsorbed in the electrical double

92

layer of carbonaceous electrodes.1-4 When treating natural waters, these electrodes are

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susceptible to fouling by organics, colloids, and bacteria.2 While pretreatment can remove

94

organic and colloidal foulants, even at low concentration of nutrients proliferation of

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bacteria is unavoidable under non-sterile conditions.5 Biofouling of CDI electrodes can

96

adversely affect CDI performance by reducing its electrosorption capacity,6 due to

97

reduced electrode surface area, and increasing the energy consumption of the process,

98

due to increased electrical resistance of the electrode6-8. It is critical to understand the

99

mechanisms of biofilm development on electrodes in order to reduce their adverse impact

100

on CDI processes.

101

Previous work on bacterial attachment and biofilm formation has identified key

102

surface properties and operating parameters that promote biofilm growth on diverse

103

surfaces. Surfaces decorated with charged functional groups,9 hydrophobic surfaces,10

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rough surfaces,11 and low toxicity surfaces generally promote cell attachment and growth.

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In addition, operating conditions such as hydrodynamics, aquatic chemistry,12 and applied

106

potential13-18 affect the rate of biofilm formation.

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Although the effect of an external electric field on biofilm formation has been

108

addressed for a range of electrode surfaces, most studies have focused on the effect of

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electric field on bacteria adhesion19-21 and biofilm formation22-25 from a sanitation

110

perspective.

111

infections in hospital environments or to disinfect contaminated liquids

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been little research on the role of electric fields and electrical charges on the proliferation

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of microbial biofilms on electrode surfaces, and the publications that do exist offer only

114

qualitative descriptions about the mechanisms of bacteria attachment,26-28 growth,29,

115

and inactivation15, 31-33 on the electrode surface.14, 17, 34-37 Determining the mechanisms by

116

which applied potentials influence biofilm development on the electrode surface may also

117

inform a suite of solutions for controlling biofilm growth in CDI as well as in other

118

electrochemically driven water treatment processes.

In these studies the primary objective was to prevent device-related

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13-18

. There has

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While it is unclear how electric fields will influence biofilm growth on carbon

121

electrodes in CDI, previous work has documented a variety of potential mechanisms by

122

which electric fields either enhance or reduce biofilm viability. The electro-migration of

123

ions in the presence of an applied electric field may disrupt normal cellular processes,

124

including adenosine triphosphate production that depend on ion gradients across the cell

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membrane.38 Antimicrobial activity may also result from byproducts of electrolysis (e.g.

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H2O2, oxidizing radicals, chlorine), oxidation of cellular enzymes and coenzymes (e.g.

127

coenzyme A, involved in the citric acid cycle),16 damage to the integrity of the cellular

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membrane, and decreased bacterial respiration rates.39

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In addition, there is a well-documented “bioelectric effect”, by which

130

antimicrobial agents become significantly more potent in the presence of an electric

131

field40-43. Proposed mechanisms relevant to inactivation on CDI electrodes include

132

increased permeability of the cellular membrane, electrochemical production of

133

potentiating oxidants, increased ion transport through electro-osmosis, physical

134

disruption of the biofilm by electrolytically generated bubbles, and increased temperature

135

at the electrode-biofilm interface.

136

have some inherent antimicrobial activity, there may be a cumulative bacterial

137

inactivation effect from the antimicrobial properties of the electrode materials and from

138

the inactivation resulting from the applied electric field.

39

In CDI where the carbon electrode material may

139

This paper addresses the critical gap that remains in understanding the extent of P.

140

aeruginosa PAO1 biofilm formation on two distinct carbonaceous electrodes as a

141

function of electrode properties and applied electric fields. After documenting the effect

142

of electrode surface properties and the applied electric field on biofilm development, we

143

investigate potential mechanisms of action. We comprehensively characterized electrode

144

surface properties, the relative quantities of biofilm components (live cells, dead cells,

145

extracellular polymeric substances), the endogenous production of reactive oxygen

146

species (ROS) production by bacteria in response to applied potentials, and the

147

exogenous production of H2O2 at the cathode. This work contributes to the literature on

148

bacterial mechanisms of inactivation in the presence of an applied potential, as well as

149

insights into pathways for mitigating biofilm formation in CDI and other water treatment

150

electrochemical processes.



151

2. Materials and Methods

152

2.1 Electrode materials

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Carbon aerogel film (CA; Marketech International Inc.) and graphite paper

154

“PAPYEX” (PA; Mersen/Carbone Lorraine) were used as carbon electrode surfaces. The

155

CA electrode contains an embedded carbon fiber mat, which provides structural integrity

156

to the electrode but does not influence the surface properties.

157

2.2 Carbon electrode characterization

158

Hydrophobicity: The captive bubble method was used to measure the hydrophobicity

159

of the carbon electrode surfaces (OCA 20, DataPhysics).

The electrodes were

160

equilibrated in either 10 or 100 mM sodium sulfate solution for 24 hours prior to the

161

experiment. The porous structure of the CA electrode readily adsorbed the air bubble and

162

precluded our ability to measure contact angle. To circumvent this issue, we performed

163

contact angle measurements on the diamond cut surface of a carbon aerogel monolith

164

supplied by the same manufacturer of the carbon aerogel film (Marketech International

165

Inc.).

166

Zeta potential: We approximate the zeta potential of the carbon electrodes by

167

pulverizing the electrode and immersing the powder in background electrolyte solution at

168

a concentration of 25 mg/mL. The samples were allowed to settle for 24 hours, and the

169

supernatant containing 0.7-2.7 um particles was analyzed (Zeta Plus 1994, Brookhaven

170

Instrument Co., Holtsville, NY). Streaming potential measurements (SurPass, Anton

171

Paar) of the PA surface corroborated the zeta-potential measurements, but the porosity of

172

the CA electrode prevented similar comparison.

173

Surface roughness: Surface roughness and topography of CA and PA electrodes were

174

analyzed using a Nanoscope IIID MultiMode Atomic Force Microscope (Veeco-DI),

175

which utilizes a NP-K cantilever (Veeco-DI) with a spring constant of 0.12 N/m in

176

contact mode. The scans were performed in air at three different resolutions: 1 µm2, 25

177

µm2 and 400 µm2. Roughness indices were estimated using the method of Root-Mean-

178

Square for the Z-plane at a resolution of 1 µm2.

179

Metabolic analysis: Reduction of XTT ((2, 3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-

180

2H-tetrazolium-5-carboxanilide)) to formazan is a colorimetric assay for assessing cell 6 ACS Paragon Plus Environment

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metabolic activity, which is correlated with cell viability44,

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antimicrobial activity of the following concentrations of suspended CA and PA carbon

183

particles: 0, 4.2, 42 and 420 NTU (Nephelometric Turbidity Units; used to measure

184

particulate content in terms of turbidity) in 10% Luria Bertani (LB) broth containing

185

planktonic P. aeruginosa PAO1 in the semi-log growth phase at OD600nm 0.5 or 1.0. The

186

XTT assay was prepared by mixing 2.5 µL of XTT solution (1 mg/mL of XTT (Thermo

187

Fisher Scientific) in LB) with 1 mL of an electron carrier solution containing N-methyl

188

dibenzopyrazine methyl sulfate (PMS; AppliChem, Darmstadt, Germany) (0.3 mg/mL of

189

PMS in phosphate buffered saline (PBS; Sigma Aldrich)).

190

45

. We evaluated the

The assay was performed by incubating 25 µL of XTT/PMS mixture with 50 µL of

191

bacterial cells and 50 µL of carbon electrode particles for 2 hours at 37°C.

The

192

suspension was then centrifuged as 13,000 RPM for 2 minutes. The supernatant was

193

collected and absorbance at 450 nm was immediately measured (Infinite 200 PRO,

194

Tecani Control).

195

An additional set of control experiments was performed in the absence of bacteria to

196

ensure that XTT does not adsorb to carbon particles and that the carbon particles do not

197

oxidize XTT to formazan. A control solution was prepared by combining 25 µL of

198

XTT/PMS solution with 100 µL of carbon electrode particle suspensions. A calibration

199

curve for the reduction of XTT to formazan by P. aeruginosa PAO1 is provided in Figure

200

SI-1.

201

2.3 Flow cell description

202

A FC 81‐PC transmission flow cell (BioSurface Technologies Corporation, Montana

203

USA) was customized to produce an electrochemical parallel plate flow-cell (Figure 1and

204

SI-2). The flow-cell was operated in pseudo-reference mode with the ITO electrode

205

acting both as a counter and the reference electrode. The absence of the proper reference

206

electrode (Ag/AgCl or sat’d calomel) was to avoid possible introduction of toxic metals

207

from reference electrodes to the bacterial culture. The voltage windows for the CA, PA,

208

and ITO electrodes were determined using cyclic voltammetry (CV) as described in SI-3.

209 210 211



212 213

214 215

Figure 1. A schematic view of the customized flow cell. The scheme of a full

216

experimental set-up is provided in SI-4.

217 218

2.4 Biofilm formation experiments

219

The biofilm formation experiments were conducted as follows. The flow cell was

220

sterilized with 70% ethanol for 30 minutes and thoroughly rinsed with sterile double

221

distilled water for 60 minutes. Three independent biofilm experiments were initiated from

222

a stationary phase overnight culture of P. aeruginosa PAO1.46 Each culture originates

223

from one distinct bacterial colony. After 8.5 hours of incubation at 30°C and 150 rpm,

224

the liquid culture was diluted 100x in LB broth and incubated overnight. We washed 80

225

mL of overnight culture in 100 mM Na2SO4 three times and adjusted the optical density

226

to OD600nm= 0.1. We injected the washed suspension to the flow cell at a rate of 2

227

mL/min (shear rate of 27 s‐1) for 40 minutes.

228

After the bacterial deposition phase, we injected a bacterial growth media for 12 or 36

229

hours at 2 mL/min while polarizing the electrodes. The growth media consisted of 1.0

230

g/L Bacto Tryptone (Becton, Dickinson and Company), 0.5 g/L yeast extract (Becton,


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Dickinson and Company), and 100 mM of Na2SO4 (Frutarom) and adjusted to pH 7.0 ±

232

0.1 M Na2SO4 was selected as the background electrolyte to avoid the potential for

233

electrochemical oxidation of Cl- to free dissolved chlorine on the anode. The pH of the

234

electrolyte effluent from the flow cell was continuously monitored using a benchtop pH

235

meter (S20 Seven Easy™ pH, Mettler Toledo). The pH microenvironment of the

236

electrode surface was checked using pH test paper (Hydrion brilliant pH strip) at the

237

conclusion of each experiment.

238

Biofilm studies were done at OCP as well as at ± 0.3, ± 0.6, and ± 0.9 V constant

239

voltages applied for 36 hours between the working and the counter electrodes. Electrical

240

current vs. time was acquired using PowerSuit® (version 2.58) software (Princeton

241

Applied Research).

242

2.5 Biofilm characterization

243

Scanning electron microscopy: The biofilms were fixed,47 dehydrated,48 sputtered

244

with a 300-350 Ǻ layer of gold (HITACHI E-101 ion sputterer) at 0.1-0.01 Torr vacuum.

245

SEM images of prepared samples were obtained using a Micro FASEM FEI Quanta 200

246

scanning electron microscope with incident electron beam energy of 1 keV and working

247

distance of 6 mm.

248

Biofilm imaging and analysis: The biofilm stain solution (Molecular probes, Inc.) was

249

prepared by mixing 5 µM of SYTO 9™ (live cell stain), 3 µM propidium iodide (PI; dead

250

cell stain), and 0.1 mg/mL of Concanavalin A conjugated to Alexa Fluor 633 (binds to

251

alpha-linked mannose residues of extracellular polymeric substances (EPS)) in a PBS

252

solution (Invitrogen).

253

the dark. The stained biofilm samples were visualized using a confocal laser scanning

254

microscope (CLSM; ZeissMeta510, Carl ZEISS, Inc., USA) equipped with Zeiss dry

255

objective LCI Plan-Neo Fluar (20 x magnification and numerical aperture of 0.5).

256

Stacked images were collected from 20 random positions on each surface. Image

257

acquisition and analysis was performed as previously described.46, 50

49

Biofilms were incubated in the stain solution for 30 minutes in

258

Intracellular oxidative stress analysis: The oxidative stress probe dihydrorhodamine

259

123 (DHR 123, Sigma-Aldrich) was used to measure the intracellular oxidation potential

260

in bacteria. Intracellular ROS is quantified by measuring rhodamine 123 production due

261

to the endogenous reduction of DHR 123. It should be mentioned that DHR can be 9 ACS Paragon Plus Environment


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oxidized only in the presence of in-situ peroxidase enzymes.51 At the end of the 12 hour

263

biofilm formation experiment under different applied potentials, biofilms samples were

264

incubated in a 5.0 µg/mL DHR in PBS solution (pH 7.2) for 15 min.52 Excess DHR dye

265

was carefully washed with PBS and CLSM imaging was performed.

266

2.6 Hydrogen peroxide quantification on polarized carbon electrodes

267

Production of HP via electrochemical oxygen reduction in the flow cell was measured

268

using an Amplex Red Hydrogen Peroxide/Peroxidase Assay Kit (Molecular Probes).53, 54

269

A solution of 10% LB

270

chronoamperometry tests were performed for 30 minutes at different applied voltages: 0,

271

0.3, 0.6, 0.9 V. After 30 minutes, 50 µL of the LB solution was collected directly from

272

the flow cell, mixed with 50 µL of Amplex solution, excited at 530 nm, fluoresced at 590

273

nm, and analyzed by a multimode reader device (Infinite 200 PRO, Tecani-control).

was placed in the flow cell (batch mode) while

274 275

3. Results and Discussion

276

Fouling of electrode surfaces is a significant issue in CDI and other electrochemical

277

processes that interact directly with natural waters. In this work, we assess the extent of

278

biofouling as a function of electrode material properties and applied voltage. The

279

potential mechanisms for observed differences in biofilm formation were delineated.

280

3.1 Characterization of electrode material properties

281

Electrode hydrophobicity: Surface hydrophobicity was estimated via captive air

282

bubble contact angle measurements on pristine electrode surfaces (Table 1). The contact

283

angle of the CA electrode is higher than that of the PA electrode, implying that the PA

284

electrode is more hydrophobic. Higher surface roughness often leads to higher biofilm

285

volumes on the surface.55

286

Zeta potential of the electrode surfaces: We approximate the zeta potentials of the

287

electrode surfaces by pulverizing the electrode material and measuring the eletrophoretic

288

mobility of the resulting sub-µm particles. The Smoluchwoski equation56 was then used

289

to approximate the zeta potential over the environmentally relevant range from pH 3 to

290

pH 7. The CA electrode material has a more negative zeta potential value than the PA


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electrode material over the range of pH tested (Table 1; Figure SI-5). We attribute the

292

more negative charge of CA electrodes to a higher density of carboxylic, hydroxyl, or

293

carbonyl groups on the surface.

294

of biofilm formation may be affected by surface charge, we do not expect the surface

295

charge to have a significant effect on biofilm formation after 12 or 36 hours.58

57

Though the initial conditioning and attachment stages

296

Carbon electrode surface roughness: AFM contact mode scans were carried out on

297

PA and CA. The 2D and 3D visualization of the AFM scans are provided in Figures SI-6a

298

and SI-6b, respectively. The roughness of PA was greater than CA, reflecting the

299

heterogeneity of the PA surface (Table 1).

300 301 302

Table 1: Contact angle, zeta potential, and surface topography measurements of CA and PA electrodes Parameter Conditions CA PA (10mM NaCl) 162.1 ± 6.3 140.0 ± 5.1 Contact angle 1(deg.) (100mM NaCl) 159.6 ± 4.3 140.5 ± 3.8 (pH=7) -24.9 ± 0.3 -18.8 ± 0.4 Zeta potential 2 (mV) (pH=8.5) -42.2 ± 0.34 -20.8 ± 0.2 2 5 µm resolution 4.5 ± 0.4 116 ± 30 Mean roughness (nm) 2 1 µm resolution 3.3 ± 0.5 29 ± 9.5

303

1

As measured using the captive bubble method.

304

2

As measured with 0.001 M NaCl.

305 306

Metabolic effects of carbon and graphite electrodes: To assess possible

307

antimicrobial properties of CA and PA electrodes, we analyzed the metabolic activity of

308

P. aeruginosa PAO1 upon exposure to pulverized electrode material. Metabolic tests,

309

including the reduction of tetrazolium to formazan is a more sensitive test for

310

antimicrobial activity of planktonic bacteria than the live/dead assays typically used for

311

high biomass biofilms. Each column bar in Figure 2 represents absorbance value at 450

312

nm due to production of formazan salts at different combinations of different bacterial

313

cell concentration and carbon electrode particles’ concentration. The first four column

314

bars (from the left) indicate corresponding absorbance value for formazan salts

315

generation due to reduction of XTT by bacterial cells (of 1 OD) at different

316

concentrations of carbon electrode particles . The CA electrode material exhibits greater 11 ACS Paragon Plus Environment


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inhibition of metabolic activity than the PA electrode material over the range of bacterial

318

concentrations (OD600

319

(Figure 2). The formazan concentration was near the detection limit for the control

320

experiments with electrode material, in the absence of bacteria. A second control

321

experiment ensured that tetrazolium was not physically adsorbed to the carbon materials

322

(Section SI-7). Additionally, a set of experiments was conducted to indicate any possible

323

interaction between formazan produced by P. aeruginosa PAO1 and carbon particles.

324

The obtained results indicated no interaction of produced formazan with carbon particles

325

(supplementary information, section SI-7). Hence, the observed antimicrobial activity of

326

CA was confirmed. The metabolic inhibition effects of CA electrodes may be attributed

327

to the presence of resorcinol and phenolic compounds in the structure of CA.59 These

328

antimicrobial properties of the CA surface may be intensified by synergistic stressors,

329

including the presence of electric fields, as described later in this paper.

nm

=0.5 and OD600

nm

=1) and material concentrations, R, tested

330 331 332 333 334 335

Figure 2: The effect of the different carbon electrode materials on cell viability: absorbance value at 450 nm due to production of formazan salts at different combinations of different bacterial cell concentration and carbon electrode particles’ concentration: (A) CA; (B) PA. R denotes the particle concentration where R0=420 NTU, R1= 42 NTU and R2= 4.2 NTU; B refers blank or without particle.

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3.2 Biofilm growth on carbon electrodes in the presence of an electric field

340

Electrode voltage window: The voltage windows for CA, PA, and ITO electrodes

341

were determined from CV experiments in aqueous solution of 0.1 M Na2SO4 and LB

342

supplemented with 0.1 M Na2SO4 as described in details in SI-3.

343

significant difference between the cyclic voltammograms for the two electrolytes. The

344

voltage window for CA and PA was adjusted to be between -70 mV and +250 mV while

345

the ITO was between -550 mV and +375 mV (all values are vs. Hg/Hg2Cl2, KCl sat’d).

346

As depicted in Figures S2-A and S2-B by the dashed bars, these windows are well within

347

the range where Faradaic reactions (such as water electrolysis) can be considered

348

insignificant. These ranges correspond to a maximum voltage window of -0.9V to +0.9V

349

between the working (PA or CA) and the counter (ITO) electrodes.

There was no

350

SEM imaging of biofilm formation on carbon electrodes: SEM analysis was

351

performed to visualize the effects of electric field on biofilm formation (Figure 3). SEM

352

images of the CA and PA electrode surface at OCP reveal a mature biofilm after 12 hours

353

of the flow cell experiment. In contrast, we observed only sporadically deposited cells

354

and no biofilm formation on the CA and PA electrode surfaces at the extreme potentials

355

of ± 0.9V. The somewhat larger number of bacteria observed on the surface of the PA

356

may be attributed to the higher hydrophobicity and surface roughness, which act to

357

protect bacteria from shear forces (Figure 3D and 3F).

358



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Figure 3: SEM images of bacterial growth on the CA and PA electrodes at different

361

applied electric fields 12 hours into the 36 hour flow cell experiment.

362

EPS production from the cells at open circuit potential obscured the morphology of the

363

bacteria. (A) CA cathodically posed at -0.9V; (B) CA at OCP condition; (C) CA

364

anodically posed at +0.9V; and (D) PA cathodically posed at -0.9V, (E) PA at OCP

365

condition; (F) PA anodically posed at +0.9V.

After 36 hours,

366 367

Biofilm formation at different voltages: We characterize biofilm formation and

368

biofilm components on the CA and PA electrode surfaces under constant hydrodynamic

369

conditions and different applied potentials using CLSM (Figure 4). A higher biovolume

370

was observed on PA electrodes compared to the CA electrodes for the full range of

371

applied potentials, including OCP (Figure 4A).

372

roughness, and the absence of antimicrobial properties of PA electrodes provide a more

373

favorable surface for biofilm formation over CA electrodes. The differences in biofilm

374

density are visualized in Figures 4C-F and differences in biofilm morphology (SI-8) and

375

biofilm thickness (SI-9) are reported in the SI.

The higher hydrophobicity, surface


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The biovolume of dead cells exceeds that of live cells for all applied potentials except

377

OCP for both CA and PA electrodes. In addition, the total biovolume is inhibited by the

378

by the presence of an electric field, with the effect largest for high applied voltages. As

379

the applied voltage became more negative, biofilm growth decreased, with the lowest

380

biovolumes of live and dead cells observed for the -0.9 V condition. At positive applied

381

potentials, there is difference in the trends of biovolume between the CA and PA

382

electrodes. The increased biovolume observed under positive applied potentials in Figure

383

4 may be a result of the attractive electrostatic forces between negatively charged bacteria

384

and the positively charged electrode.

385

decreased to very low values at higher positive applied potentials than +0.3V, the

386

biovolume on the PA electrode was only moderately decreased. This may be a result of

387

higher CA material toxicity and the cumulative stress experienced by cells at high applied

388

potentials.

While the biovolume on the CA electrode

389

To monitor the potential of bactericidal effects by water splitting and pH change13 at

390

the extremes of the CA voltage window, we continuously monitored the pH of the

391

bacterial solution and tested the pH of the electrode surface at the conclusion of the

392

experiments using a pH strip. We observed no change in pH over the duration of the CA

393

or PA experiments for either the continuously flowing solution or the electrode surface.

394

Under the influence of an external electric field, the possibility of change in electrolyte

395

pH is further diminished due to the generation of CO2 during oxidation of carbon, while

396

carbon is used as working electrode, which results in a carbonate buffer

397

microenvironment.

398

polarized working electrode.60

This reaction is thermodynamically favorable at an anodically

399

C + 2H 2 O → CO 2 + 4H + + 4e - , E o = 0.207 V

400

Another suggested effect of external electric potential on bacterial cells is stress

401

response, which can generate free radicals that hamper normal metabolic activities and

402

disrupt cell organelles. An oxidation of the coenzyme A (CoA), an enzyme associated

403

with the synthesis of pyruvate in the citric acid respiration cycle by free radicals was

404

observed by Matsunaga et al.61 In this study, the respiratory activity of whole cells of

405

Saccharomyces cerecisiae decreased at applied potential of +0.74 V. The loss of


Eq.1


406

respiratory activity was also observed for Bacillus subtilis and Escherichia coli. The CoA

407

existing in the cell wall was electrochemically oxidized to dimeric CoA and, as a result,

408

the respiration of cells was inhibited. ROS generation by the biofilm is further

409

investigated in Section 3.3.

410

411 412 413 414 415 416 417 418 419 420

Figure 4: (A) Biovolume of biofilms (live and dead cells) formed on the CA and PA after 36 hour of flow cell experiment using 10% LB (Luria-Bertani) medium at different applied voltage (0 V implied open circuit condition); (B) Biovolume values of EPS at the CA and PA electrodes at different electric potentials, in flow cell experiment using 10% LB medium after 36 hour. Each error bar represents one standard deviation. IMARIS 3D images of (C) CA at applied voltage of +0.9 V; (D) PA at applied voltage of +0.9 V; (E) CA at OCP condition; (F) PA at OCP condition. The red, green, and blue clusters indicate dead cells, live cells, and EPS on electrode, respectively. In (C-F) Scale bars indicate 100 µm.

421


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422

EPS production at different voltages: EPS production is common bacterial response

423

to stress in viable cells. Thus, we expect peak EPS production at intermediate applied

424

voltages where the bacteria are stressed but still viable. We evaluated the effect of the

425

applied electric field on the production of EPS and found a slight elevation in EPS

426

production at ± 0.3 V, compared to the OCP condition (Figure 4B). EPS production was

427

reduced significantly at higher applied potential of ± 0.6 V and ± 0.9 V, likely because of

428

the low percentage of viable cells in the biofilms. The effects of electric fields on EPS

429

production by mapping, firstly at which level cell physiology is altered (transcription,

430

translation, or even post-translation levels) should be a matter for future studies.

431

Variations in EPS thickness and roughness are shown in SI-10.

432

3.3 Endogenous ROS production in biofilms grown under electric fields

433

Bacteria produce intracellular ROS during normal metabolic activity, as well as in

434

response to diverse environmental stressors. In these cases, ROS generation preserves

435

redox homeostasis inside the cell and serves a signaling function for coordinating of

436

cellular activity.51,

437

depletes antioxidants and leads to the oxidation of biomolecules, such as lipids in cellular

438

membranes, structural proteins or enzymes, and DNA.64 Previous work has documented

439

excess ROS generation in response to abrupt changes in temperature, dissolved oxygen,

440

and the presence of nanoparticles.65 We hypothesized that application of external electric

441

field will affect the redox potential across the membrane, disrupt redox homeostasis, and

442

accelerate ROS production in biofilms.

62, 63

However, at these excess concentrations, intracellular ROS

443

We evaluate the accumulation of intracellular ROS by detecting the oxidation of DHR

444

in biofilms exposed to an electric field. After 36 hours, the fitness of bacteria was too

445

low to detect DHR oxidation for the highest applied voltages on the CA electrode.

446

Therefore, we performed experiments on biofilms grown for 12 hours. We normalized

447

the DHR biovolume to the viable biomass as determined by SYTO9 in order to obtain the

448

specific ROS activity. Figure 5(A) presents the ratio of DHR stained cells to the SYTO9

449

live cells for both carbon electrodes at different applied potentials. Figure 5(B) through

450

5(E) show the qualitative analysis of ROS production using IMARIS images for CA and

451

PA at two levels of -0.9 V of applied voltage and OCP condition.



452

For both electrodes, the elevated specific ROS activity was lowest for the OCP

453

condition, elevated when cathodically and anodically polarized, and highest for the ± 0.9

454

V cases. We believe that the inhibited bacterial growth at high applied potentials is a

455

result of excess generation of ROS. The DHR fluorescence intensity was also elevated as

456

the intensity of the electric field increased compared to OCP condition (SI-11).

457 458 459 460 461 462 463 464 465 466

Figure 5: (A) The ratio of the biovolume of dihydrorhodamine 123 (DHR) stained cell to live cell (LBF) (stained with SYTO9) at different carbon electrodes under influence of different electric potentials, after 12-hours flow cell experiment using 10% LB as medium. IMARIS 3-D images for ROS generation at (B) CA at applied voltage of +0.9 V; (C) PA at applied voltage of +0.9 V; (D) CA at OCP condition; (E) PA at OCP condition. The red cluster indicated DHR stained bacterial cell on electrode. Figures B-E are perspective images 600 µm × 600 µm in size. Analysis of DHR oxidation was not performed for the ± 0.3 V conditions.

467

3.4 Effect of electrochemically generated hydrogen peroxide on biofilm formation

468

Oxygen reduction to hydrogen peroxide (HP) at the cathode may induce bactericidal

469

effects in the biofilm. In spite of the very narrow voltage window selected for this study

470

as discussed in Section 3.2, we still measured HP generation by the cathode as an attempt

471

to rule out the possibility that electrochemically generated HP was responsible for

472

reduced biofilm formation and cell viability. Due to the difficulty of measuring HP


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473

production in single pass flow conditions,53 we instead adopted the very conservative

474

approach of monitoring HP production in batch mode in the absence of bacteria.

475

We detect a maximum dissolved HP concentration of ≈ 2.4 µM for applied potentials

476

of ± 0.9 V, with HP production higher for the CA electrode as a result of its higher

477

porosity and larger surface area (Figure 6). Though the concentration of HP is likely to

478

be higher at the cathode surface than in the bulk solution, operating this system in batch

479

mode provided a higher concentration of HP compared to the case of actual biofilm

480

growth experiments performed under flow. Moreover, the detection of HP generation in

481

both anodically and cathodically polarized electrode suggests that the ITO glass has the

482

ability to reduce the dissolved oxygen into HP. The reasons for the relatively low HP

483

concentration include the buffering of the electrolyte solution (at neutral pH) and the high

484

oxidation reactivity of HP towards LB media.

485

To quantitatively assess the effects of 2.4 µM HP on biofilm formation in this study,

486

we analyzed the effect of external dosage of HP on biofilm formation at OCP (SI-12). In

487

agreement with previous studies, the application of external HP had a negligible influence

488

on biofilm formation or cell viability (SI-12).66-68 Only at concentrations an order of

489

magnitude higher than those observed in the batch mode experiments (20 µM) did we

490

observe a small difference between biovolume and cell viability. For these experiments,

491

the increased EPS production at the elevated dosage of HP (analyzed with CLSM, SI-12)

492

also suggests a physiological response of the biofilm to negate the harmful effect of

493

HP.69, 70 A thorough discussion on HP generation in the flow cell is provided in SI-13.

494

The theoretical maximum concentration of HP electrochemically generated along the

495

electrode surface was calculated and is equal to 18µM (SI-13). The experimental results

496

obtained from the external dosage of hydrogen peroxide on biofilm formation at OCP

497

condition (SI-12) as well as previous studies67-68 suggest that this range of ~20 µM of

498

HP is still sub-inhibitory for both planktonic and sessile communities of P. aeruginosa

499

PAO1. Electrochemically generated HP may have little detrimental effect on planktonic

500

bacterial cells only at the very early stage of biofilm formation immediately after

501

deposition of bacteria on the electrode surface.

502

generation in LB media, coupled with the insignificant effects of exogenous HP on

In conclusion, the low rate of HP



503

biofilm growth and viability, confirm that electrochemically produced HP is not

504

responsible for the differences in biofilm formation and cell viability observed this study.

505 506 507

Figure 6: The concentration of H2O2 (µM) generated on CA and PA carbon electrodes.

508 509

4. Environmental Implications

510

In this study we investigated the effect of surface properties and applied potentials on

511

biofilm formation on carbon and graphite electrode. The driving force behind this study is

512

the growing interest in electrochemical systems such as CDI for water treatment and

513

microbial fuel cells for energy production in which these types of electrodes are

514

extensively used. At OCP, the CA electrode was less conducive to biofilm formation

515

than the PA electrode due to its lower hydrophobicity, lower surface roughness, and

516

inhibitory metabolic effects.

517

observed an inverse relationship between biofilm formation and the magnitude of the

518

applied potential. The effect is particularly strong for CA electrodes, and may be a result

519

of cumulative effects between material toxicity and the stress experienced by cells at high

520

applied potentials.

For both positive and negative applied potentials, we


Page 20 of 26

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521

Further indication of bacterial stress is evidenced by high endogenous ROS production

522

in the presence of an applied electric field. For both electrodes, the elevated specific

523

ROS activity was lowest for the OCP condition, elevated when cathodically and

524

anodically polarized, and highest for the ± 0.9 V cases. These relatively high potentials

525

are believed to affect the redox potential across the cell membrane and disrupt redox

526

homeostasis, thereby accelerating ROS production and inhibiting bacterial growth.

527

Consistent with this observation, peak EPS production occurred at intermediate applied

528

potentials where the bacteria were stressed but still viable.

529

The observed decline in bacterial viability and biofilm growth at the highest applied

530

voltages are not a result of HP generation. The low rate of electrochemically generated

531

HP in LB media, coupled with its insignificant effect on biofilm growth and viability,

532

support this conclusion. In conclusion, this study elucidates the critical effect of applied

533

electric potentials on ROS production in biofilms and the associated decrease in biofilm

534

development.

535

ACKNOWLEDGMENT

536

This research was supported by United States – Israel Binational Science Foundation

537

under award number 2012142 and the Planning and Budgeting Committee (PBC) of the

538

Council for Higher Education for postdoctoral fellowship award.

539

Supporting Information Available

540

Data regarding the experimental setup, measurement methods , additional results and

541

explanation is supplied in the supporting information.

542

Figures S1−S16

543 544



545

References

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Influence of Electric Fields on Biofouling of Carbonaceous Electrodes

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