Parallel artificial and biological electric circuits power petroleum decontamination: The case of snorkel and cable bacteria

Journal Pre-proof Parallel artificial and biological electric circuits power petroleum decontamination: The case of snorkel and cable bacteria Ugo Marzocchi, Enza Palma, Simona Rossetti, Federico Aulenta, Alberto Scoma PII:

S0043-1354(20)30056-7

DOI:

https://doi.org/10.1016/j.watres.2020.115520

Reference:

WR 115520

To appear in:

Water Research

Received Date: 23 September 2019 Revised Date:

13 December 2019

Accepted Date: 17 January 2020

Please cite this article as: Marzocchi, U., Palma, E., Rossetti, S., Aulenta, F., Scoma, A., Parallel artificial and biological electric circuits power petroleum decontamination: The case of snorkel and cable bacteria, Water Research (2020), doi: https://doi.org/10.1016/j.watres.2020.115520. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2020 Published by Elsevier Ltd.

HC+ CO2 HC-

SO42-

H 2S

Snorkel + Cable bact.

*

Cable bact.

e-

0.10

Snorkel

e-

Control

e-

t. zero

e-

n-Alkanes (mg/gDW)

O2 + 4H+ 2H2O

0.15

*

0.05

**

0.00

1

PARALLEL ARTIFICIAL AND BIOLOGICAL ELECTRIC CIRCUITS POWER

2

PETROLEUM DECONTAMINATION: THE CASE OF SNORKEL AND CABLE

3

BACTERIA

4

Ugo Marzocchi1,2*, Enza Palma3, Simona Rossetti3, Federico Aulenta3 and Alberto Scoma4,5

5 6

1

7

Aarhus University, Aarhus, Denmark

8

2

9

Institute of Marine Biology, Ecology and Biotechnology, Napoli, Italy

Center for Electromicrobiology, Section for Microbiology, Department of Bioscience,

Integrative Marine Ecology Department, Stazione Zoologica Anton Dohrn, National

10

3

Water Research Institute (IRSA), National Research Council (CNR), Monterotondo, Italy

11

4

Section of Microbiology, Department of Bioscience, Aarhus University, Aarhus, Denmark

12

5

13

University, Aarhus, Denmark

Biological and Chemical Engineering (BCE), Department of Engineering, Aarhus

14 15

*Correspondence to: Ugo Marzocchi, Center for Electromicrobiology, Section for

16

Microbiology, Department of Bioscience, Aarhus University.

17

Ny Munkegade 114, 8000-C Aarhus, Denmark.

18

Email: [email protected]

19

Phone: +45-87154335

20 21

Keywords: Cable bacteria; snorkel; sediment; hydrocarbon; long-distance electron transport;

22

remediation.

23 24

25

ABSTRACT

26

Degradation of petroleum hydrocarbons (HC) in sediments is often limited by the

27

availability of electron acceptors. By allowing long-distance electron transport (LDET)

28

between anoxic sediments and oxic overlying water, bioelectrochemical snorkels may

29

stimulate the regeneration of sulphate in the anoxic sediment thereby accelerating petroleum

30

HC degradation. Cable bacteria can also mediate LDET between anoxic and oxic sediment

31

layers and thus theoretically stimulate petroleum HC degradation. Here, we quantitatively

32

assessed the impact of cable bacteria and snorkels on the degradation of alkanes in marine

33

sediment from Aarhus Bay (Denmark). After seven weeks, cable bacteria and snorkels

34

accelerated alkanes degradation by +24 and +25%, respectively, compared to control

35

sediment with no cable bacteria nor snorkel. The combination of snorkels and cable bacteria

36

further enhanced alkanes degradation (+46%). Higher degradation rates were sustained by

37

LDET-induced sulphide removal rather than, as initially hypothesized, sulphate regeneration.

38

Cable bacteria are thus overlooked players in the self-healing capacity of crude-oil

39

contaminated sediments, and may inspire novel remediation treatments upon hydrocarbon

40

spillage.

41 42

1. INTRODUCTION

43

Microbial oil degradation in sediments is often limited by the availability of electron

44

acceptors (e.g. Meckenstock et al. 2015). In impermeable coastal and shelf sediment, the

45

mass transport of solutes from the water is governed by slow molecular diffusion, which

46

restricts the availability of electron acceptors such as oxygen (O2) and nitrate (NO3-) to the

47

topmost millimeters (Fenchel and Jørgensen 1977; Glud 2008). Sulphate (SO42-) may

48

penetrate centimetres to meters into marine sediments and thus, despite being a less

49

favourable electron acceptor, may account for a large fraction of organic matter degradation

50

(Jørgensen 2006). In addition to diffusion from the water, SO42- can be regenerated within the

51

anoxic sediment via the re-oxidation of sulphide (H2S) and other reduced sulphur species

52

with insoluble iron (Fe) and manganese (Mn) oxides serving as electron acceptor. As these

53

minerals are ultimately formed under oxic conditions, in the absence of sediment-mixing by

54

bioturbating organisms, H2S re-oxidation is constrained to the proximity of oxic-anoxic

55

interfaces. In petroleum HC contaminated sediment, the high demand of SO42- by oil-

56

degrading microorganisms may exceed the rate of SO42- supply from the water and via H2S

57

oxidation, potentially leading to local SO42- limitation. Supply limitations and the less

58

favourable thermodynamic of SO42- may account for a long persistence of petroleum HC in

59

the anoxic compartment of marine sediments (e.g. Reddy et al. 2002; Peterson et al. 2003;

60

Culbertson et al. 2008).

61

A bioelectrochemical snorkel is an electrically-conductive, non-polarized material

62

(e.g., graphite, platinum, stain-less steel) applied to short-circuit redox gradients (Erable et al.

63

2011; Hoareau et al. 2019). Snorkels offer a preferential route for electrons to move from

64

highly reduced to oxidized zones, where electrons can react with O2 to generate water. This

65

electron flow may occur over centimetres distances, and thus may enable microorganisms the

66

remote access to otherwise out-of-reach O2, eventually resulting in enhanced oxidation rates

67

under anaerobic conditions. The application of snorkels in petroleum HC-contaminated

68

marine sediment resulted in improved microbial HC degradation in combination with

69

increased SO42- availability, indicating that the snorkel enhanced SO42- regeneration via

70

promoting H2S oxidation on the electrode (Cruz Viggi et al. 2015). The detection of ferric

71

iron (Fe3+) minerals on the surface of snorkels applied in riverine sediments (Viggi et al.

72

2017) suggested that the re-oxidation of H2S to SO42- may also occur via reduction of Fe

73

oxides with the produced ferrous iron (Fe2+) scavenged by the snorkel. Despite the fact that

74

these studies show that snorkels can accelerate petroleum HC biodegradation in sediments,

75

the biogeochemical mechanism that controls the transfer of electrons from the HC to snorkels

76

remains to a large extent uncertain and may vary from site to site.

77

Owing to their ability to access remote electron acceptors and stimulate regeneration

78

of SO42- at depth (Risgaard-Petersen et al. 2012), cable bacteria can be considered the

79

biological analogue of a snorkel. Cable bacteria are filamentous Desulfobulbaceae able to

80

mediate electric currents thereby coupling anodic H2S oxidation at a few centimetres depth

81

with cathodic O2 or NO3- reduction close to the sediment-water interface (Nielsen et al. 2010;

82

Pfeffer et al. 2012; Marzocchi et al. 2014). Cable bacteria were reported from a wide range of

83

sediments (e.g. Risgaard-Petersen et al. 2015; Burdorf et al. 2017; Marzocchi et al. 2018)

84

including oil-contaminated sites (Müller et al. 2016; Matturro et al. 2017). Because of their

85

ability to mediate long-distance electron transport (LDET) they have been hypothesized to

86

stimulate petroleum HC degradation by functioning as snorkels. However, a cause-effect

87

relationship between the activity of cable bacteria and petroleum HC degradation has not

88

been so far demonstrated.

89

In the present study, we quantitatively assessed the impact of such biological (cable

90

bacteria) and engineered (snorkel) LDET systems, deployed alone and in combination, on the

91

degradation of petroleum HC in marine sediments from Aarhus Bay (Denmark). Moreover,

92

we comparatively investigated their respective geochemical imprint, with a focus on sulphur

93

(S) and iron (Fe) cycling and impact on the benthic microbial community.

94 95 96

2. METHODS 2.1. Sediment sampling and pre-treatment

97

Intact sediment samples were collected from Aarhus Bay (Denmark) at station M5

98

(56º06’20’’N, 10º27’48’’E; depth 30 m), which is known to host cable bacteria (Marzocchi et

99

al. 2014), using a box-corer. On site, the upper 10-12 cm of sediment were discarded to

100

exclude large shells and stones that might interfere with the microprofiling measurements

101

(see section 2.2). The underlying sulfidic sediment was sealed in airtight bags, brought to the

102

laboratory and stored at 15°C. Within a few weeks the bags were opened and the sediment

103

was sieved (mesh size 0.5 mm), homogenized, and contaminated with crude oil (Statfjord

104

oil). Sediment contamination to a final concentration of 1% (v:w) was performed as in Viggi

105

et al. (2015).

106

The contaminated sediment was then packed into 12 glass liners (4.3 x 8 cm, diam. x

107

height). Four treatments were prepared to discern the individual and combined impact of

108

cable bacteria and snorkel on sediment geochemistry, i.e., 1) negative control with no snorkel

109

and no cable bacteria (Control); Snorkel with no cable bacteria (Snorkel), Cable bacteria with

110

no snorkel (Cable bacteria); and 4) Snorkel and cable bacteria (Snorkel + Cable bacteria) (Fig

111

S1). Cable bacteria growth was inhibited in the Snorkel treatment and in the Control by

112

inserting filters (mesh size 0.2 µm, Advantec, Japan) in the sediment similarly to Pfeffer et al.

113

(2012). Compared to the original method, we applied glass-fiber instead of the polycarbonate

114

filters to avoid adsorption of oil HC to the filter material, and applied two filters (at the

115

depths of 5 and 10 mm) instead of one to further insure the inhibition of cable bacteria.

116

Snorkels were made of graphite (>99.9995% purity; Alfa Aesar, Milan, Italy) rods (6.15 x

117

120 mm, diam. x height) inserted vertically into the sediment to the depth of 7 cm. Disks (40

118

x 10 mm, diam. x height) of carbon felt (Hi-Tech carbon Co.limited, China) were connected

119

to the snorkel in the water column to maximize the cathodic surface. Three cores were

120

prepared for each treatment and for the filter control.

121 122

2.2. Sediment incubation and sampling procedures

123

All sediment cores were incubated for seven weeks in the same aquarium containing

124

artificial seawater (salinity 30%. Red Sea Salts, Red Sea Fish Pharm Ltd, Eilat, Israel). The

125

water was kept gently stirred by an aquarium pump. Depth microprofiles of O2, H2S, pH, and

126

redox potential were measured at the beginning and at the end of the incubation period in all

127

cores. In addition, microprofiles were measured weekly (except for week 6) in all treatments

128

except for those with filters to avoid filter damage. Sediment was sampled at the beginning

129

(two days after contamination) and at the end of the incubation. For sampling, sediment cores

130

were removed from the aquaria and the sediment was extracted from the liners under N2

131

atmosphere and homogenised. Approx. 1 g. of sediment was snap-frozen (-80°C) for later 16s

132

rRNA extraction. Samples for fluorescence in situ hybridization (FISH) for cable bacteria

133

detection were collected from the sediment and from the surface of snorkels as in (Matturro

134

et al. 2017). Four to seven grams of sediment were collected and stored at -20°C for later

135

analysis of the content of n-alkanes in the C9-C31 interval, hereon referred to as alkanes. The

136

rest of the sediment was centrifuged (8000 r.p.m x 15 min.) and the supernatant was sampled

137

(1.5 mL) for SO42- determination. Additional 0.8 mL of supernatant were filtered and

138

acidified (+0.1 µL of 6M HCl) for later Fe2+ determination. Samples for SO42-and Fe2+ were

139

stored at -20°C until further analysis. The centrifuged sediment was stored at -20°C for later

140

acid volatile sulphur (AVS) determination.

141 142

2.3. Microsensor measurements

143

High-resolution depth profiles of H2S, O2, pH, and Redox potential were recorded

144

with micro-electrodes built at Aarhus University (Revsbech and Jorgensen 1986; Revsbech

145

1989; Jeroschewski et al. 1996; Kühl and Revsbech 2001). Microprofiling procedures and

146

H2S, O2, and pH microsensors calibrations were conducted as in (Marzocchi et al. 2018). The

147

offset of the reference of the Redox sensor compared to the SHE electrode was quantified via

148

a two-point calibration procedure using quinhydrone buffers. Microprofiles were recorded at

149

100-400 µm vertical resolution and measured at approx. 2 cm distance from the snorkel. A

150

reference electrode (REF201 Red Rod electrode; Radiometer Analytical, Denmark) was used

151

for Redox and pH measurements. Total hydrogen sulphide (ΣH2S = [H2S] + [HS-] + [S2-])

152

concentrations were calculated at each depth from the measured H2S and pH values

153

(Jeroschewski et al. 1996).

154 155

2.4. Chemical analysis

156

Sulphate concentrations in the sediment porewater were determined by Ion

157

Chromatography (Dionex IC-2500, Thermo Fisher Scientific). Ferrous iron was determined

158

using the Ferrozine method (Stookey 1970). Acid Volatile Sulphides (AVS) were determined

159

on a wet sample after acidification as in Yücel et al. (2010).

160

Quantification of alkanes in sediment samples was performed by GC-MS. In brief,

161

sediment samples (approximately 10 g) were air dried overnight and extracted with a Thermo

162

Scientific ASE (DIONEX ASE 150) using a dichloromethane (DCM): hexane (1:9 v/v)

163

mixture at 100 °C and a system pressure of 1500 psi. The extract was evaporated to a final

164

volume of 5 mL under a gentle nitrogen stream. A sample of the extract (1 µL) was then

165

injected (in pulsed split-less mode) into a GC-MS (Perkin Elmer Clarus 680/600; column:

166

HP-5 MS (Agilent) 30 m, ID 0.25 mm, 0.25 mm film thickness; carrier gas: helium at 1

167

mL/min; injector temperature: 280 °C; oven temperature program: initial Temp 40 °C, 18

168

°C/min to 250 °C, 10 °C/min to 280 °C, hold for 17 min; MS-scan 30-600, 2-32 min).

169

Quantification of alkanes was performed by means of external standards (C8-C40 Alkanes

170

Calibration Standard, Sigma-Aldrich).

171 172

2.5. DNA extraction, 16S rRNA gene sequencing, and FISH analysis

173

DNA was extracted from ~1 g of wet sediment and from the material scraped from

174

the graphite rods surface with DNeasy PowerSoil Kit (QIAGEN, Italy) following the

175

manufacturer’s instructions. Extracted DNA was amplified in a first PCR with the primer pair

176

27F (5’-AGAGTTTGATCCTGGCTCAG-3’) and 534R (5’-ATTACCGCGGCTGCTGG-3’)

177

targeting the region V1-V3 of bacterial 16S rRNA gene. Reactions were set up in 25 µL

178

volumes containing 15 ng of DNA, 0.5 µM primers and 1X Phusion High-Fidelity PCR

179

Master Mix (Thermo Fisher Scientific, Waltham, MA USA). PCR settings are detailed in

180

(Matturro et al. 2017). The amplicon libraries were purified using the Agencourt® AMpure

181

XP bead protocol (Beckmann Coulter, USA). Sequencing libraries were prepared from the

182

purified amplicon libraries using a second PCR as described in Matturro et al. (2017). The

183

amplicon libraries were purified using the Agencourt® AMpure XP bead protocol

184

(Beckmann Coulter, USA). The purified libraries were pooled in equimolar concentrations

185

and diluted to 4 nM. The samples were paired end sequenced (2x301bp) on a MiSeq platform

186

(Illumina) using a MiSeq Reagent kit v3, 600 cycles (Illumina, USA) following the standard

187

guidelines for preparing and loading samples. 10% Phix control library was spiked in to

188

overcome low complexity issue often observed with amplicon samples. The bioinformatic

189

processing was performed as detailed in (Crognale et al. 2019).

190

FISH was performed according to previously published protocols (Pernthaler et al.

191

2001) as described in (Marzocchi et al. 2018). Cable bacteria were detected with probe

192

DSB706 (Loy et al. 2002), targeting most Desulfobulbaceae (Pfeffer et al. 2012).

193 194

2.6. Statistical analysis

195

The effects of the factors cable bacteria and snorkel (presence/absence) on our dependent

196

variables (Alkanes, SO42-, AVS, and Fe2+) were tested in the four treatments (control, cable

197

bacteria, snorkel, cable bacteria + snorkel) by means of a two-way ANOVA test. The level of

198

significance was set to p < 0.05. Posthoc pairwise multiple comparisons (Tukey test) were

199

performed to asses difference between treatments. Statistical analyses were performed using

200

OriginPro (OriginLab).

201 202 203 204

3. RESULTS 3.1. Cable bacteria geochemical imprint and ΣH2S dynamics throughout the incubation

205

After two weeks of incubation, ΣH2S and pH depth profiles in the Cable bacteria

206

cores showed the typical signature for cable bacteria activity i.e., build-up of a ~2-cm-thick

207

zone devoid of both O2 and ΣH2S, intense proton consumption in the oxic portion of the

208

sediment (pH maxima) compatible with cathodic O2 consumption, and acidification in the

209

suboxic zone compatible with anodic H2S oxidation (Fig. 1A) (e.g. Nielsen et al. 2010;

210

Meysman et al. 2015). Snorkel + Cable bacteria cores showed analogous patterns, indicating

211

the presence of cable bacteria, although with two remarkable differences: a 40% higher

212

attenuation of the upward flux of ΣH2S (6.5 vs. 3.8 µmol m-2 h-1); and a less pronounced pH

213

maxima (Fig. 1A). This is descriptive of a contribution of the snorkel to LDET, that is, ΣH2S

214

consumption over a deeper sediment horizon and cathodic O2 reduction mainly occurring in

215

the overlying water, which did not contribute to the pH increase in the subsurface sediment.

216

Over the course of the incubation in the Snorkel + Cable bacteria treatment, the

217

sulphidic front (depth with [ΣH2S] > 1 µM) deepened monotonically throughout the seven-

218

week period, moving from 1.0 cm below the sediment-water interface at time zero to 2.5 cm

219

depth at the end of the incubation (Fig. 1B). In the Cable bacteria treatment, the ΣH2S

220

concentration showed a less linear trend (Fig. 1C). Within the first two weeks, the sulphidic

221

front moved from 1.0 to 2.15 cm. In the following two weeks, the sulphidic front remained

222

steady, however, ΣH2S accumulated in the deeper sediment layers (>2.15 cm). After four

223

weeks, the continued ΣH2S accumulation in the deeper sediment layers affected the sulphidic

224

front, which raised progressively and realigned with the depth recorded at the beginning of

225

the experiment (i.e., 0.95 cm).

226

3.2. Alkanes degradation at the end of the incubation

227

At the end of the seven-week incubation period, the concentration of alkanes in

228

controls (0.114 mg g-1 dry weight [DW]) was not significantly lower compared to the

229

beginning of the experiment (t-test, p = 0.2, n = 6) (Fig. 2). Two-way ANOVA analysis

230

indicated significant variations due to the factors Snorkel and Cable bacteria (p < 0.01). In the

231

Snorkel and in the Cable bacteria treatments the concentration of alkanes dropped by -24%

232

and -25%, respectively, compared to the Control (Tukey-test, p = 0.04). The concentration of

233

alkanes in the Snorkel + Cable bacteria treatment was significantly lower compared to

234

Controls, i.e., -54% (Tukey-test, p > 0.01) and to the Snorkel and the Cable bacteria

235

treatments (Tukey-test, p = 0.02)

236 237 238

3.3. Impact of cable bacteria and snorkel on sediment biogeochemistry at the end of the incubation

239

At the end of the incubation period, ΣH2S was detected in the Control at 1.0 cm depth

240

(Fig. 3A), with the concentration increasing to >100 µM at 2.7 cm depth. In the Cable

241

bacteria treatment, ΣH2S was measured at 1.1 cm depth and reached concentrations >100 µM

242

at 3.3 cm depth. In the Snorkel treatment ΣH2S was below detection limit up to a depth of 2.0

243

cm, and at the bottom of the profile (4.0 cm depth) ΣH2S only reached 37 µM (± 5.4 s.e.m).

244

The Snorkel + Cable bacteria treatment was the most efficient in maintaining low ΣH2S

245

levels, as ΣH2S was not detected to 2.6 cm and it only reached 14 µM (±6.6) at 4.0 cm depth.

246

In the Control, the pH decreased with depth from the overlying water value of ~8.6

247

reaching a minimum of 7.77 ±0.04, at 1.05 cm. The Snorkel treatment had a similar pattern,

248

but maintained lower values starting from a ~0.6 cm depth (minimum of 7.61 ±0.02, at 1.75

249

cm). As opposed to these treatments, those including cable bacteria showed subsurface peaks

250

(with maximum value of 8.5 at 0.2-0.3 cm depth) and reached lower minimum values i.e.,

251

7.3-7.4 (Fig. 3B). In the Snorkel + cable bacteria, the pH minimum was detected at higher

252

depth (2.4 cm) compared to the Cable bacteria treatment (1.05 cm).

253

Redox profiles mirrored those of the ΣH2S, with increasingly positive values

254

following the order Control < Cable bacteria < Snorkel < Snorkel + Cable bacteria, indicative

255

of a progressively less reduced environment when applying LDET systems alone or in

256

combination (Fig. 3C). Compared to the Control, all the three treatments resulted in a

257

tangible increase in redox potential (between +80 and +150 mV) in the 0.5-1.5 cm depth

258

interval. Deeper in the sediment, the impact of cable bacteria alone on redox potential

259

attenuated and values tended to realign to the Control, whereas in both snorkel treatments

260

values remained higher compared to Control (+50 to +60 mV, at 4 cm).

261

The presence of the snorkel resulted in higher concentration of SO42- compared to the

262

treatments without it (two-way ANOVA, p < 0.01) (Fig. 3D). In the Snorkel treatment, SO42-

263

increased only slightly compared to the Control, however the difference was not significant

264

(Tukey-test, p = 0.39). The Snorkel + Cable bacteria treatment showed the highest

265

concentration of SO42- and the difference with the controls resulted significant (Tukey test, p

266

< 0.05). Such an increase in SO42- was coupled to a net decrease in AVS (Snorkel + Cables

267

vs. Control, Tukey-test, p < 0.05) and release of Fe2+ (Snorkel + Cables vs. Control, Tukey-

268

test, p < 0.05), confirming an ongoing FeS dissolution and H2S re-oxidation to SO42-.

269

Sulphate, AVS and Fe2+ concentrations in the Snorkel treatment with no cable bacteria,

270

showed trends analogous to the above described ones, although of an attenuated intensity

271

compared to the Snorkel + cable bacteria treatment. Only Fe2+ concentration was

272

significantly higher in the Snorkel treatment compared to the Control (Tukey-test p < 0.05).

273

The SO42- concentration was lower in the Cable bacteria treatment compared to all other

274

treatments (Tukey-test p < 0.05) confirming that their activity was decreasing after seven

275

weeks of incubation (Fig. 1C). No net increase in Fe2+ was recorded in the Cable bacteria

276

treatment compared to the Control, however, cable bacteria were the only significant factor

277

determining the decrease in AVS (two-way ANOVA p <0.01). The presence of cable bacteria

278

significantly decreased the AVS concentration (two-way ANOVA, p < 0.01) regardless of the

279

presence of the snorkel. The AVS concentrations decreased in the order Control > Snorkel >

280

Cable bacteria > Snorkel + Cable bacteria. This trend mirrored that for pH minima (i.e.,

281

lowest AVS concentration observed with lowest pH, Fig. 4B), suggesting that FeS dissolution

282

was likely driven by acidic-forming anodic oxidations mediated by cable bacteria.

283 284

3.4. Impact of cable bacteria and snorkel on bacterial communities

285

At the end of the incubation, cable bacteria were detected in the Cable bacteria and

286

Snorkel + Cable bacteria treatments at relative abundances of 0.1%, but were undetected in

287

the cores with filters, i.e., Control and Snorkel treatment. FISH imaging in both cable bacteria

288

treatments confirmed their typical filamentous form, that allows them to stretch across the

289

sediment redox zonation to perform LDET (Fig S2). Bacterial phyla composition in sediment

290

cores did not substantially differed as a result of different treatments. Chloroflexi,

291

Deltaproteobacteria, Epsilobacterarcheota, Firmicutes and Alphaproteobacteria represented

292

about 60 to 76% of the entire community in all test (Table S1). The 26 most abundant taxa

293

were common to all treatments and represented about 50 to 70% of the entire microbial

294

community of the sediment (Table 1A). The most enriched taxa in all treatments belonged to

295

the phylum Chloroflexi, namely the families Dehalococcoidia AB-539-J10 and

296

Anaerolineaceae (≥8%, Table 1A), but their exact metabolism remains uncertain (Wasmund

297

et al. 2014; Matturro et al. 2017). However, more than half of such abundant taxa (14 out of

298

26) referred to microorganisms well known for using S species either as electron donor (in

299

the form of H2S, thiosulphate [S2O32-], sulphite [SO32-] or elemental sulphur [S8]; i.e.,

300

Sulfurovum, Sulfurimonas, Desulfocapsa, Arcobacter, Thiomicrorhabdus, Sulfitobacter,

301

Magnetovibrio and Candidatus Tenderia (Janssen et al. 1996; Inagaki et al. 2004; Ivanova et

302

al. 2004; Takai et al. 2006; Bazylinski et al. 2013; Eddie et al. 2016; Boden et al. 2017) or

303

electron acceptor (in the form of SO42- or S8; i.e., Desulfatiglans, Desulfuromusa, SEEP-

304

SRB1 and MSBL7, unclassified Desulfobulbaceae, Fusibacter (Liesack and Finster 1994;

305

Ravot et al. 1999; Schreiber et al. 2010; Pachiadaki et al. 2014; Suzuki et al. 2014).

306

Irrespective of the snorkel treatment (Snorkel or Snorkel + Cable bacteria), few taxa

307

selectively enriched on the graphite rod’s surface compared to the sediment, namely

308

Arcobacter,

309

Pseudomonas, uncultured Ardenticatenales, unclassified Bradymonadales, unclassified

310

Rhodobacteraceae, and unclassified Desulfuromonadales (log2 fold change [log2fc] ≥1.05,

311

relative abundance on rods ≥1.2%; Table 1B). This core group represented ~50% of the 16S

312

rRNA gene sequences on the snorkels surface in both Snorkel treatments. On the contrary,

313

some of the most enriched taxa in sediments were poorly represented on graphite rods, e.g.,

314

uncultured Anaerolineaceae, Dehalococcoidia AB-539-J10, Sulfurovum and Desulfatiglans

315

(log2fc ≤-1.7; relative abundance on rods ≤2.5%; Table 1B). A few taxa were only detected

316

on rods and not in sediments (i.e. Leucobacter, Rhodococcus, Hydrogenophaga). Among

317

these Leucobacter, Rhodococcus appeared to be more abundant in the Snorkel + Cable

318

Bacteria treatment.

Magnetovibrio,

Sulfitobacter,

Desulfuromusa,

Sneathiella,

Marinicella,

319 320 321

4. DISCUSSION 4.1. Cable bacteria

322

This study presents the first evidence of a cause-effect relationship between cable

323

bacteria and enhanced microbial oil degradation. Microbial oxidation of alkanes proceeds

324

through the beta-oxidation pathway of fatty acids. None of the genes encoding for beta-

325

oxidation-related enzymes are present in sequenced cable bacterial genomes (Kjeldsen et al.

326

2019). As such, a direct engagement of cable bacteria in alkanes degradation is presently

327

ruled out.

328

H2S oxidation by cable bacteria is expected to increase the concentration of SO42- in

329

the porewater (Risgaard-Petersen et al. 2012). Increase in SO42- availability due to cable

330

bacteria activity has been hypothesized to stimulate toluene degradation in aquifer-sediments

331

locally depleted in SO42- (Müller et al. 2016). On the contrary, our data indicate a net

332

decrease in SO42- concentration in cores with cable bacteria at the end of the incubation, such

333

a decrease however was marginal as SO42- concentration remained above 15 mM. Cable

334

bacteria expected benefit on petroleum HC-oxidizing, SO42--reducing microorganisms

335

appears instead to reside in their capacity to scavenge H2S. Colonization of the cores by cable

336

bacteria in fact resulted in H2S depletion in the top two cm of sediments within the first two

337

weeks of incubation, followed by slow H2S re-accumulation later in the incubation (Fig 1C).

338

This temporal evolution mirrors cable bacteria population dynamics (peak at two-three weeks

339

and later decaying) previously reported in sediments from the same station not contaminated

340

with petroleum HC (Schauer et al. 2014), confirming their predominant role in controlling

341

H2S removal in the surface sediment. Moreover, besides removing H2S generated from SO42-

342

reduction, the metabolism of cable bacteria favours the dissolution and consequent re-

343

oxidation of H2S from the FeS mineral pool (Risgaard-Petersen et al. 2012). Increased FeS

344

dissolution in the Cable bacteria treatments was indicated by the significant depletion of

345

AVS. In principle, H2S scavenging by cable bacteria could affect the energetics of SO42--

346

driven anaerobic oxidation of petroleum HC to CO2. Thermodynamic calculations, however,

347

shows that under environmental conditions that are representative of the herein described

348

experiments, product removal would only have a marginal effect on the Gibbs free energy of

349

the reaction, which remained largely exergonic even at H2S concentrations exceeding 1 mM

350

(Fig. S3). Accordingly, it is more likely that H2S removal enhanced oil degradation (by SO42-

351

-reducing bacteria) via relieving toxicity and/or inhibitory effects induced by this metabolite

352

(Reis et al. 1992).

353

Snorkel

354

The application of the snorkel accelerated alkane degradation by 24% compared to the

355

Control. This occurred within only seven weeks (49 days), a much shorter time with respect

356

to previous reports, i.e., 175 days (Cruz Viggi et al. 2015; Viggi et al. 2017). Differently from

357

previous studies, where the incubations were conducted under stagnant water, our incubation

358

was performed under gentle stirring, suggesting a more rapid impact of the snorkel under

359

environmental-relevant conditions where advection contributes to the transport of O2 to the

360

cathode.

361

In the present set up (sediment volume: ~30 cm3; ray around the graphite rod: 2.5 cm)

362

the concentration of alkanes (preferentially bound to sediment particles) and insoluble

363

minerals (AVS) decreased substantially compared to control sediment (Fig. 2 & 4E),

364

indicating that the snorkel impacts an area well beyond its immediate proximity. Sediment

365

microprofiles describe an impact on solutes at ≥2 cm distance from the snorkel. Such distance

366

is compatible with molecular diffusion within the experiment period (e.g., ~1.3 cm d-1 for HS-

367

, Boudreau 1997), suggesting that the enhanced alkanes biodegradation was presumably

368

mediated by the alteration of the redox chemistry of solutes in the sediment.

369

The impact of the snorkel on petroleum HC degradation has been previously related

370

to its ability to promote SO42- reduction coupled to petroleum HC oxidation or, potentially, to

371

the oxidation of petroleum HC fermentation products such as H2 and acetate (Matturro et al.

372

2017; Viggi et al. 2017). Stimulation of SO42- reduction may be due to enhanced oxidation of

373

H2S to SO42-/S8 directly on the rod (Cruz Viggi et al. 2015), or coupled to FeOOH reduction

374

with consequent re-oxidation of Fe2+ on the electrode (Viggi et al. 2017). We indeed detected

375

a net production of SO42- when applying snorkels, however, as discussed above (section 4.1),

376

microbial kinetics may not be impacted in marine sediments carrying high SO42-

377

concentrations. As for cable bacteria, the snorkel induced a remarkable depletion of H2S.

378

Removal of H2S has been previously shown to stimulate SO42--reducing bacteria (Reis et al.

379

1992) and scavenging of H2S by a bioanode (poised at +300 mV vs. SHE) was proposed to

380

stimulate SO42--driven degradation of toluene (Daghio et al. 2016). Thus, as discussed for

381

cable bacteria, H2S removal rather than SO42- recycling should account for the enhanced

382

microbial oil degradation observed with snorkels. Further to removing H2S, the snorkel could

383

have also contributed to the removal of petroleum HC fermentation products, namely acetate

384

and hydrogen, with this reaction being catalysed by electroactive bacteria using the portion of

385

the electrode that is buried in the sediment as the terminal electron acceptor. While the

386

removal of H2S has a limited impact on the Gibbs free energy of the direct petroleum HC

387

oxidation to CO2 by SO42--reducing bacteria, thermodynamic modelling indicates that

388

maintaining low acetate and H2 concentrations in the sediment is likely to promote the

389

upstream fermentation of petroleum HC (Fig. S4). Collectively, these calculations suggest

390

that, in principle, the snorkel may enhance petroleum HC degradation also by removing

391

fermentation products and in turn serving as a “solid” syntrophic partner in the fermentative

392

degradation process. As cable bacteria can use propionate (Vasquez-Cardenas et al. 2015)

393

and plausibly other small chain organic substrates as carbon source, a similar mechanism

394

could be expected in sediment colonized by cable bacteria.

395

Oxidation of Fe2+ on the electrode did not appear to play an important role in H2S

396

oxidation, in sharp contrast to what previously suggested (Viggi et al. 2017). This process is

397

expected to decrease the concentration of Fe2+ in the porewater, whereas we detected a net

398

production of Fe2+ in the presence of the snorkel (Fig. 3F). The increase of Fe2+ was linked to

399

the dissolution of FeS minerals as induced by the acid forming anodic oxidation of H2S on

400

the electrode, in agreement with pH and AVS data (Fig 3B & E). Under such Fe2+ forming

401

conditions, H2S oxidation via FeOOH reduction would not be favoured, but rather inhibited.

402

The net removal of H2S in our incubation has thus to be ascribed to its oxidation on the

403

snorkel. Such difference between the two studies is likely due to the inherent geochemical

404

features of the sediment, with the electron transfer prevalently mediated by iron cycling in

405

iron oxides-rich, less sulphidic freshwater sediment; and by sulphur cycling in sulphidic, FeS

406

mineral rich, marine sediment. Overall, this data is descriptive of the snorkel as a highly

407

plastic system supporting electron scavenging along multiple pathways able to accelerate

408

petroleum HC degradation under different conditions.

409 410

4.3. Cable bacteria & Snorkel

411

When both cable bacteria and snorkel were applied together, the final alkane

412

biodegradation was the sum of what obtained with either system alone, indicating no

413

competition for reductants between the two LDET systems. Notably, the estimated anodic

414

surface area for snorkel and cable bacteria was comparable (14.8 vs. 17.7 cm2), suggestive of

415

a direct relation between anodic surface and alkane biodegradation capacity, irrespective of

416

the nature of the system applied.

417

The simultaneous presence of snorkel and cable bacteria improved the removal of

418

electrons via LDET from the deeper sediment layers as shown by the enhanced H2S removal

419

and increased redox potential compared to the treatments where the systems were test

420

individually (Fig. 3A & C). The pH, SO42-, and Fe2+ data indicate that the snorkel favoured

421

cable bacteria metabolism. Deeper pH minima (Fig. 3B) are representative of an extended

422

cable bacteria activity over a thicker sediment layer compared to the cable bacteria alone,

423

meaning a larger volume of influence of cable bacteria. The increase in the SO42- and Fe2+

424

sediment pools compared to the snorkel alone indicates that after seven weeks cable bacteria

425

were still contributing to the oxidation of reduced S species both from the dissolved and

426

mineral (FeS) pool, contrary to when they were applied alone. The mechanisms by which

427

snorkels maintain a more active cable bacteria population is unclear. Owing to snorkels, cable

428

bacteria may be forced to grow as longer filaments to keep contact with the deepening

429

sulphidic front. Alternatively, cable bacteria and snorkel may interact to complete H2S

430

oxidation. A recent genome mapping reports that cable bacteria can oxidize H2S to S8

431

followed by either S8 oxidation to SO42- or disproportionation to SO42- and H2S (Kjeldsen et

432

al. 2019). Abiotic electrochemical oxidation of H2S on the electrode is expected to produce S8

433

(Rabaey et al. 2006; Dutta et al. 2008), which could in turn, feed cable bacteria. Filamentous

434

Desulfobulbaceae were recently reported anchored to the anode of a sediment microbial fuel

435

cell (Reimers et al. 2017), suggesting that cable bacteria may also use the electrode as

436

electron acceptors. We detected a few 16S rRNA gene sequences of Ca. Electrothrix on the

437

surface of graphite rods in the Snorkel + Cables treatment (relative abundance 0.13%), but no

438

cable bacteria were observed on rod’s samples via Scanning Electron Microscopy. Further,

439

pH maxima detected below the sediment-water interface after seven weeks suggest that cable

440

bacteria maintained a direct access to O2.

441

Along with cable bacteria, the role of other microorganisms in catalyzing all the

442

oxidative steps from H2S to SO42- in the presence of snorkels remains unclear. Most of the

443

abundant taxa in our sediments are common to oil-degrading communities, e.g., Sulfurimonas

444

(Rubin-Blum et al. 2014; Tian et al. 2017), Desulfocapsa (Ramos-Padron et al. 2011; Aktas

445

et al. 2017), Magnetovibrio (Matturro et al. 2017), Arcobacter (Prabagaran et al. 2007;

446

Scoma et al. 2019); Sulfitobacter (Krolicka et al. 2017; Gontigaki 2018), and may account for

447

the observed alkanes degradation. How they interact with snorkels and cable bacteria requires

448

further investigation. A core community of 11 taxa was selectively enriched on the snorkels’

449

surface (~50% of the whole community colonizing snorkels), many of which featured a S

450

metabolism. Arcobacter is typically detected under H2S (Dronen et al. 2014; Roalkvam et al.

451

2015) or S8 (Voordouw et al. 1996) oxidizing conditions. Magnetovibrio oxidizes H2S and

452

S2O32- and can either use O2 or nitrous oxide (N2O) as electron acceptor. Sulfitobacter is

453

rather ubiquitous in marine environments and most species oxidize SO32- (Ivanova et al.

454

2004). Desulfuromusa typically reduces S8 to H2S, but is unable to use SO42-, S2O32- or SO32-

455

as

456

Desulfuromonadales are widely diverse taxa which are deeply involved in S cycling (Pujalte

457

et al. 2014; Reguera and Kashefi 2019). With the exception of Sneathiella, Marinicella and

458

the recently proposed Bradymonadales, all other taxa enriched on snorkels were detected on

459

anodes as biofilms or axenic cultures, i.e., Arcobacter (Fedorovich et al. 2009; Song et al.

460

2017), Magnetovibrio (Matturro et al. 2017), Sulfitobacter (Parot et al. 2011; Erable et al.

461

2017), Desulfuromusa (Liu et al. 2007; Carmona-Martinez et al. 2015), Ardicantena

462

maritima, type strain of the Ardicantenales (Kawaichi et al. 2018), Pseudomonas (Rabaey et

463

al. 2004), Rhodobacteraceae (Matturro et al. 2017), Desulfuromonadales (Reguera and

464

Kashefi 2019). The selective enrichment on bioelectrochemical snorkels of these taxa is thus

465

suggestive of a syntrophy in the frame of SO42--reducing, oil-degrading metabolism taking

466

advantage of LDET for the removal (oxidation) of reduced S species.

electron

acceptor

(Liesack

and

Finster

1994).

The

Rhodobacteraceae

and

467 468

5. CONCLUSIONS

469

The present data indicate that cable bacteria support microbial petroleum HC-

470

oxidizing, SO42--reducing microorganisms by alleviating the accumulation of a toxic product

471

(i.e., H2S) both from the dissolved and mineral (FeS) pool. Cable bacteria thus pull microbial

472

oil metabolism towards a faster oxidation, accelerating sediments decontamination. Their

473

impacts appears limited in time (peak at 2-4 weeks) and space (1.5 cm depth), however their

474

widespread distribution, potentially high cell number (i.e., 4 x 108 cell cm-3, see Schauer e al.

475

2014), and substantial impact on the top sediment (the one initially most impacted by

476

petroleum HC deposition after a spill) are suggestive of a previously unrecognized role in

477

petroleum HC decontamination in the environment.

478

Alkane degradation was highest in the co-presence of cable bacteria and the snorkel.

479

In the presence of the snorkel, cable bacteria expand their volume of influence and possibly

480

their life-span. Thus, our data indicate that application of a non-polarized electrode (i.e., the

481

snorkel) to oil-contaminated marine sediments did not simply mediate the removal of

482

negative charges convoyed via diffusion of redox species to the rod. Rather, it facilitated the

483

activity of specific microbial members, particularly cable bacteria, comprehensively doubling

484

the alkane biodegradation capacity of the whole microbial community. Unveiling the

485

mechanistic interaction of these microorganisms with electroactive materials is of primary

486

importance in the bioremediation field.

487 488

ACKNOWLEDGEMENTS

489

We are thankful to Lars Borregaard-Pedersen for the microsensors construction and technical

490

support throughout the project, to Karina Bomholt Oest for laboratory assistance, and to

491

Alexander Michaud and Katja Laufer for help with the AVS analysis. Results incorporated in

492

this study have received funding from the European Union’s Horizon 2020 research and

493

innovation programme under the Marie Sklodowska-Curie grant agreement No 656385

494

awarded to UM; and the Danish National Research Foundation (grant DNRF104).

495 496

COMPETING INTERESTS

497

The authors declare no competing interests in relation to this work.

498

499

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500 501 502 503 504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550

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FIGURES AND FIGURE LEGENDS

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Figure 1. Panel A: Sediment microprofiles of pH, and ΣH2S after two weeks of incubation in

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the treatment with no filters (red) and snorkel without filters (black). Data are shown as mean

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± s.e.m. (n = 3). Dotted line in the ΣH2S plot shows the microprofile measured at experiment

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start (time zero). White boxes show the oxic zone of the sediment ([O2] > 0.5 µM). Panel B

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and C show the variation of the porewater ΣH2S concentration over the seven-week

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incubation period in treatments Snorkel + Cable bacteria and Cable bacteria, respectively. In

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the treatments with filters (i.e., Control and Snorkel), ΣH2S profiles were only measured after

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seven weeks (Fig. 3A) to avoid filter damage during the incubation.

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Figure 2. Concentration of n-alkanes (C9 - C31) at the start of the experiment (time zero) and

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at the end of the incubation in the four treatments. Data are shown as mean ± s.e.m. (n = 3).

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Figure 3. ΣH2S, pH, and Redox potential sediment depth microprofiles in the four treatments

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at the end of the seven-week incubation (panels A, B, C). Average SO42-, AVS, and Fe2+,

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concentrations in the sediment depth interval 1 – 6 cm at experiment start (time zero) and

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after seven weeks of incubation in the four treatments (panels D, E, F). All data are shown as

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mean ± s.e.m. (n = 3).

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Table 1. Most abundant taxa in: A) marine sediments and B) graphite rods, seven weeks after

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contamination with Fjord oil (1% w:v) in the control and the three treatments. The red-blue heatmap

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indicates log2 fold changes, with blue being an increase and red a decrease.

HIGHLIGHTS •

Cable bacteria stimulate alkane oxidation by SO42- reducers in marine sediment.

•

The stimulation is linked to the removal of H2S rather than to the regeneration of SO42-.

•

Rates of alkane removal are highest in the co-presence of cable bacteria and snorkel.

•

The snorkel expands the volume of influence of cable bacteria in the sediment.

Declaration of interests ☒ The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. ☐The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: