This is the submitted version of the journal article: Fernandez Verdejo, David Juan; Cortés Garmendia, M. Pilar; Blánquez Cano, Paqui; [et al.]. «Enhanced dechlorination of 1,2-dichloropropane to propene in a bioelectrochemical system mediated by Dehalogenimonas». Journal of hazardous materials, Vol. 416 (Aug. 2021), art. 126234. DOI 10.1016/j.jhazmat.2021.126234 This version is available at https://ddd.uab.cat/record/273831 under the terms of the license 1 Enhanced dechlorination of 1,2-dichloropropane to propene in a bioelectrochemical 2 system mediated by Dehalogenimonas 3 David Fernández-Verdejo1, Pilar Cortés2, Paqui Blánquez1, Ernest Marco-Urrea1,*, 4 Albert Guisasola 3 5 1 6 Biorem UAB, Department of Chemical, Biological and Environmental Engineering, 7 School of Engineering, Universitat Autònoma de Barcelona, 08193, Bellaterra, 8 Barcelona, Spain. 2 9 Departament de Genètica i de Microbiologia, Facultat de BioCiències, Universitat 10 Autònoma de Barcelona, 08193 Bellaterra, Barcelona, Spain 3 11 GENOCOV, Department of Chemical, Biological and Environmental Engineering, 12 School of Engineering, Universitat Autònoma de Barcelona, 08193, Bellaterra, 13 Barcelona, Spain. 14 15 * Corresponding author: 16 Biorem UAB, Department of Chemical, Biological and Environmental Engineering, 17 School of Engineering, Universitat Autònoma de Barcelona, 08193, Bellaterra, 18 Barcelona, Spain. Email: ernest.marco@uab.cat 19 20 21 1 22 Abstract 23 Bioelectrochemical systems (BES) are promising technologies to enhance the growth of 24 organohalide-respiring bacteria and to treat chlorinated aliphatic hydrocarbons. In this 25 study, two carbon-based cathodic electrode materials, a graphite brush and a carbon cloth, 26 were used as hydrogen suppliers to couple growth of Dehalogenimonas and 27 dechlorination of 1,2-DCP to nontoxic propene in the cathode vessel. The BES with 28 graphite brush electrode consumed ~4000 µM 1,2-DCP during 110 days and exhibited a 29 degradation rate 5.6-fold higher than the maximum value obtained with the carbon cloth 30 electrode, with a cathode potential set at -0.7 V. Quantitative PCR confirmed that 31 Dehalogenimonas gene copies increased by two orders of magnitude in the graphite brush 32 BES, with an average yield of 1.2·108 ± 5·107 cells per µmol of 1,2-DCP degraded. The 33 use of a pulsed voltage operation (cathode potential set at -0.6 V for 16h and -1.1 V for 34 8h) increased the coulombic efficiency and degradation of 1,2-DCP when compared with 35 a continuous voltage operation of -1.1 V. Bacterial cell aggregates were observed in the 36 surface of the graphite brush electrodes by electron scanning microscopy, suggesting 37 biofilm formation. This study expands the range of chlorinated compounds degradable 38 and organohalide-respiring bacteria capable of growing in BES. 39 Keywords: Dehalogenimonas, 1,2-Dichloropropane, Bioelectrochemistry, Remediation, 40 Organohalide respiring bacteria. 41 42 43 44 2 45 1. Introduction 46 1,2-dichloropropane (1,2-DCP) is an halogenated organic compound used in industry 47 either as a chemical intermediate in the production of other valuable organic substances, 48 such as tetrachloroethene and tetrachloromethane or as a solvent [1]. Due to its toxicity 49 and carcinogenic effects, it has been designated as a high priority chemical substance for 50 risk evaluation by the United States Environmental Protection Agency [2]. 51 Due to accidental spills, 1,2-DCP has been frequently detected in groundwater sources. 52 The anoxic conditions typically found in contaminated groundwater limit the aerobic 53 biodegradation of 1,2-DCP [3–6]. Under anaerobic conditions, 1,2-DCP can serve as 54 terminal electron acceptor for organohalide-respiring bacteria (OHRB) belonging to the 55 genera Dehalogenimonas and Dehalococcoides mccartyi, among others [1,7–9]. The 56 reductive dechlorination reaction of 1,2-DCP for both genera involve a 57 dichloroelimination mechanism producing propene as final non-toxic product [1,7–9]. 58 Enhanced in situ anaerobic bioremediation can be an effective method of degrading 59 chlorinated compounds dissolved in groundwater [10,11], but the injection of electron 60 donors in groundwaters to stimulate OHRB growth presents some limitations (i.e. the 61 uneven distribution of the substrates, possible induction of secondary contamination or 62 competition with non-dechlorinating hydrogenotrophic bacteria). In the recent years, 63 bioelectrochemical systems (BES) have been proposed as an efficient technology to 64 promote anaerobic reductive dechlorination processes [12–18]. BES catalyse reductive 65 dechlorination reactions in the cathode by supplying electrons either by direct electron 66 transfer from the electrode surface or by indirect electron transfer via soluble redox 67 mediators or H2 generation through electrochemical dehydrogenation [19]. One of the 68 advantages of using BES over conventional enhanced in situ anaerobic bioremediation is 3 69 that the supply of electron donor, i.e. hydrogen, can be easily monitored and fine-tuned. 70 In addition, the utilisation of BES could increase the low cell densities obtained for OHRB 71 using conventional suspended lab culture techniques, which is one of the bottlenecks for 72 the obtention of commercial bioaugmentation cultures [20]. This could be achieved by 73 the formation of biofilms attached to high-surface carbon electrodes, that would allow a 74 higher OHRB concentration in the reactor [17]. 75 To date, the application of biocathodes for the treatment of chlorinated aliphatic 76 hydrocarbons by OHRB has been limited to the degradation of chlorinated ethenes 77 (tetrachloroethylene, trichlorethylene and 1,2-dichloroethylene) and the chlorinated 78 ethane 1,2-dichloroethane [21]. The OHRB identified playing a role in these BES systems 79 belonged to the genera Dehalococcoides mccartyi or Geobacter [12,15,21,22]. 80 The aim of this study is to explore the feasibility of performing the dechlorination of 1,2- 81 DCP to non-toxic propene by a Dehalogenimonas-containing culture in a two-chamber 82 BES separated by a cationic exchange membrane, where the growth of Dehalogenimonas 83 is driven by abiotically generated H2 in the cathode. Graphite fibre and carbon cloth were 84 selected as the cathodic electrode materials in view of its biocompatibility with 85 Dehalogenimonas sp. Both cathodes were compared in terms of energetic consumption 86 and degradation rate at the same cathode potential. This work further expands the range 87 of chlorinated compounds and OHRB that can be stimulated by using BES. 88 2. Materials and methods 89 2.1. Bioelectrochemical cells description 90 The electrochemical cells consisted of two 165 mL glass vessels separated by a cation- 91 exchange membrane (CMI-7000, Membranes International INC, USA) with 4 cm of 92 aperture diameter. In the anodic compartment, a titanium sheet was used as the electrode. 4 93 In the cathodic compartment, a graphite fiber brush (35 mm length x 30 mm diameter, 94 7.2 µm fiber diameter, Millrose Co., USA) or a carbon cloth fragment (85 mm length x 95 25 mm width x 0.673 mm thickness, Zoltek Co., USA), provided with a surface area of 96 approximately 13.2 dm2 and 0.213 dm2 respectively, were used as working electrodes. 97 Both vessels were mixed with a magnetic stirrer. A power source (Quad Potentiostat, 98 Whistonbrook Software) was used to control the voltage and the potential of the system. 99 The applied cathode potential was measured against an Ag/AgCl reference electrode 100 placed in the cathodic vessel. When controlling the voltage, a digital multimeter (Hayoue 101 DT830B) was used to ensure the cathodic potential. All the potential values mentioned 102 are V vs Standard Hydrogen Electrode (SHE) unless otherwise stated. Butyl rubber 103 septums and aluminium crimp caps were used to seal the system while providing a 104 sampling port to allow liquid extraction or addition from both cell compartments. 105 2.2. Cultivation of the Dehalogenimonas-containing culture in serum bottles 106 A 1,2-DCP-degrading Dehalogenimonas-containing culture was previously enriched 107 from sediments obtained in the Besós river (Spain) [1]. The genome of the strain 108 contained in this culture was recently sequenced and annotated, and it was denominated 109 Dehalogenimonas alkenigignens strain BRE15M [23]. The culture was maintained in 100 110 mL glass bottles containing 65 mL of anaerobic medium previously described [24]. In 111 brief, it contained minerals, trace elements, vitamins, sodium acetate (5 mM) as a carbon 112 source, titanium (III) citrate (0.29 mM) as reducing agent and sodium bicarbonate (12 113 mM) as buffering agent. Bottles were sealed with butyl rubber septums and aluminium 114 crimp caps and were gassed with N2/CO2 (4:1 at 0.2 bar of overpressure) and H2 (0.4 bar 115 of overpressure). Cultures were periodically spiked with 1,2-DCP (500 µM, nominal 116 concentration) and transferred into fresh medium after the consumption of ~3000 µM. 5 117 2.3. Operation of BES 118 Cathode and anode chambers contained 130 mL of the anaerobic medium described 119 above. Cathodic compartments were initially spiked with 500 µM 1,2-DCP and 120 inoculated with 3 mL of an active culture of Dehalogenimonas with a density of ~9·107 121 cells·mL-1. Biotic open circuit BES without electrode were also included as controls. The 122 cathodic potentials were adjusted to -0.7 V vs Standard Hydrogen Electrode (SHE) 123 against an Ag/AgCl reference electrode (RE-1B, BAS Inc., +197 mV vs SHE) by 124 applying a potentiostatic control in both graphite brush and carbon cloth-provided 125 reactors. The BES were maintained in fed-batch mode by adding 1,2-DCP when 126 exhausted (500 or 1000 µM). 127 For the experiments carried out with BES containing graphite brush electrodes under 128 pulsed and continuous voltage operation, a cathode potential of -0.6 V and -1.1 V was 129 applied during time-periods of approximately 16 and 8 h respectively from day 0 to day 130 46. Afterwards, a cathodic potential of -1.1 V was applied until the end of the experiment 131 at day 136. In this case, a pH probe (Hach) was employed to ensure the acidity of the 132 cathodic vessel and manual additions of an anaerobic stock solution of 1 M of HCl were 133 employed to adjust pH at a value of 7. 134 2.4. Coulombic efficiency and energy consumption 135 The coulombic efficiency (CE) for each degradation experiment at the cathode was 136 calculated using equation 1: 2·V·F·[1,2−DCP]deg 137 CE = (1) ∫ I(t)dt 138 where 2 is the number of electrons required to dechlorinate a molecule of 1,2-DCP to 139 propene, V is the cathodic liquid volume (L), F is Faraday’s constant (96485 A·s·mol-1), 6 140 [1,2-DCP]deg is the degraded concentration of 1,2-DCP (M) and ∫I(t)dt is the integration 141 of the measured intensity throughout the selected experimental time (A·s). 142 Additionally, the energetic input (EI) required for each experiment was calculated as 143 follows: VA·∫ I(t)dt 144 EI = (2) V·[1,2−DCP]deg 145 Where VA is the voltage applied in the system (V). 146 2.5. Analytical methods 147 1,2-DCP and propene were measured by static headspace gas chromatography. Liquid 148 samples (1 mL) were taken from the cathode and were transferred to 10 mL sealed vials 149 containing 5.5 mL deionized water. The vials were placed in a headspace sampler 150 (Agilent 7964) and heated to 85 ºC for 15 min. Automatically, 1 mL headspace gas sample 151 from the vials was injected into an Agilent 6890N gas chromatograph provided with an 152 Agilent DB-624 column (30 m × 0.32 mm with 0.25 μm film thickness) and a flame 153 ionization detector following a method described elsewhere [25]. Calibration of both 154 compounds was based on aqueous standards with the same liquid and gas volumes than 155 the experimental BES. Details for the calculation of propene concentration is provided in 156 the Supplementary Information Materials and Methods section. 157 2.6. Microbiological analysis 158 Liquid samples (3 mL) were obtained from the cathode at different time points during the 159 experiments and centrifuged at 9000 g for 20 minutes. The supernatant was removed and 160 the DNA was extracted using a NucleoSpin Tissue kit (Machery Nagel) following the 161 manufacturer instructions and eluted in a final volume of 50 µL. 7 162 Quantitative hydrolysis probes based real-time PCR (qPCR) was performed with a 163 Lightcycler 480 instrument (LC480; Roche) in order to monitor the increase of 164 Dehalogenimonas cells in the BES cultures. The qPCR used a set of primers (Dhgm478F 165 and Dhgm536R) and a TaqMan probe (Dhgm500Probe) which were used in a previous 166 study and specifically targeted the 16S gene from the genus Dehalogenimonas (Table S1) 167 [26]. Each qPCR assay was performed in a total volume of 20 µL and containing 1x 168 LightCycler 480 Probes Master Mix (Roche Diagnostics), 0.4 µM final concentration of 169 each primer and TaqMan probe, ultrapure water and 2 µL of the extracted DNA sample. 170 The qPCR cycle conditions were: pre-incubation at 95oC for 10 min, followed by 50 171 cycles of 15 s at 95 °C, 1 min at 56 °C and 1 second at 72 oC (fluorescence detection). 172 Standard curve for quantifying the gene copies obtained in the assays was created as 173 following. A synthetic sequence corresponding to the whole 16S gene of 174 Dehalogenimonas alkenigignens (NR_109657.1) and containing the amplified region 175 was purchased (ATG biosynthetics). The gene sequence of interest was amplified by 176 conventional PCR using the previously described primers and the amplicon (59 pb) 177 presence was checked with a 2% agarose electrophoresis gel. The PCR product was 178 purified using an NZYGelpure kit (Nzytech) following the manufacturer instructions, and 179 was ligated to a pGEMT vector (Promega) and transformed into Escherichia coli DH5α 180 following standard procedures. The presence of the cloned sequence in the transformed 181 vector was assessed by sequencing (Macrogen). 182 Plasmid DNA was prepared using an NZY Miniprep kit (Nzytech) according to the 183 manufacturer instructions and it was diluted in 10-fold dilutions ranging from 103 to 109 184 gene copies per reaction mix and used as template for qPCR. The quantification cycle 185 (Cq) obtained for each dilution was plotted against the logarithm of the gene copy 186 numbers in the template. Each dilution was tested by triplicate. The number of copies of 8 187 the cloned sequence in plasmid DNA was calculated with the formula used by Ritalahti 188 et al. [27]. Also, the 16S gene copies per mL of culture sample were determined from the 189 standard curve (Fig. S1) following a previously described equation [28]. All the samples 190 were quantified in triplicate. 191 Due to the significant similarities in the amplified region between the genus 192 Dehalococcoides and Dehalogenimonas, two negative controls using a DNA extraction 193 from a Dehalococcoides mccartyi-containing culture and water instead of experimental 194 sample were used to assess the specificity of the amplification. 195 2.7. Cell growth kinetic calculations 196 The maximum specific growth rates (µ, d-1) were calculated by selecting the exponential 197 growth phase period according to the cell concentrations of Dehalogenimonas obtained 198 by qPCR and using the following equation: 1 dX 199 µ= · (3) X dt 200 where X is the cell concentration in the bioelectrochemical reactor (Dehalogenimonas 201 cells·mL-1). 202 The biomass growth yield (Y, cells generated / µmol 1,2-DCP degraded) was calculated 203 as it follows: Xgen 204 Y= (4) 1,2−DCPdeg 205 Where Xgen is the increase of Dehalogenimonas cells and 1,2-DCPdeg are the µmol of 1,2- 206 DCP degraded during a certain time period. 207 The maximum specific 1,2-DCP degradation rate by Dehalogenimonas (q) can be 208 calculated by applying µ and Y as it is shown in the following equation: µ 209 q= (5) Y 9 210 211 2.8. Scanning electron microscopy 212 Field emission-scanning electron microscopy was performed with liquid samples of the 213 culture contained in the cathode chamber and with cathodic electrode material at the end 214 of each working period of BES. Liquid samples (5 mL) were filtered through 0.2 µm 215 pore-size polycarbonate filters to collect the cells in suspension. Both kind of samples 216 were fixed with a 2.5 % glutaraldehyde solution and treated as previously described [29]. 217 Imaging of the samples was done with a Zeiss EVO-MA10 field emission scanning 218 electrode microscope at the Servei de Microscopia (Universitat Autònoma de Barcelona). 219 3. Results and discussion 220 3.1. Effect of electrode material on 1,2-DCP dechlorination 221 BES containing graphite fibre brush and carbon cloth cathodes were poised at -0.7 V of 222 cathode potential and operated during 110 days with periodical amendments of 1,2-DCP 223 when consumed (Figure 1A). The first dose of 1,2-DCP was completely dechlorinated to 224 propene in 37 days regardless of the electrode. However, the subsequent amendments of 225 1,2-DCP were consumed at increasing rates over time in BES containing graphite fibre 226 brush whereas dechlorination of 1,2-DCP in BES containing carbon cloth proceeded 227 slower (Fig. 1A). 228 During this time-span, graphite fiber brush-containing cells were capable of depleting 229 seven 1,2-DCP doses, while the carbon cloth-containing cell was only capable of 230 degrading three, which resulted in a final accumulated degradation of 3930 ± 605 and 231 1442 µM of 1,2-DCP, respectively (Fig. 1B). The production of propene was monitored 232 in the BES containing graphite fibre brush because exhibited an enhanced 1,2-DCP 233 dechlorination performance. As shown in Fig. 1B, the moles of 1,2-DCP degraded 10 234 stoichiometrically agree with the propene generated. At the end of the experiment (110 235 days), propene accumulated was 4039 ± 425 (102.7% of the 1,2-DCP consumed), 236 demonstrating that full dechlorination of 1,2-DCP was achieved during the process and 237 discarding the presence of other undesired and potentially harmful by-products. Biotic 238 controls that consisted of opened circuit system without electrode required 75 days to 239 degrade 540 µM of 1,2-DCP (Figure S2), showing that 1,2-DCP degradation notably 240 decreased in absence of a continuous supply of electrochemically generated hydrogen 241 from an electrode. 242 Degradation of 1,2-DCP was coupled to an increase in the number of Dehalogenimonas 243 16SrDNA copies in BES containing graphite fibre brush. The consumption of the four 244 first amendments of 1,2-DCP led to an increase of Dehalogenimonas 16SrDNA gene 245 copies by over two orders of magnitude (Figure 1C). The concentration of 246 Dehalogenimonas 16S rDNA genes during the conversion of the following two 1,2-DCP 247 doses continued increasing but at lesser extent, obtaining a final cell density of 2.1·108 ± 248 8·107 cells/mL Dehalogenimonas (assuming one 16S rRNA gene per genome). This value 249 fits well with cell densities reported for organohalide-respiring bacteria, which rarely 250 grow above 108 cells/mL [30]. Relative to the biotic controls, the consumption of a single 251 dose of 540 µM 1,2-DCP in 75 days was consistently accompanied by a modest growth 252 from 1.3·106 ± 3·105 to 2.8·106 ± 8·105 (Figure S2). 253 The 1,2-DCP degradation rates obtained in the graphite brush experiment increased 254 during the first four contaminant amendments from 9.84 ± 1.50 µM·d-1 to 89.18 ± 6.39 255 µM·d-1 and afterwards exhibited stable values during three consecutive 1,2-DCP 256 amendments (Figure 2A). The degradation rate reached its maximum at the sixth addition 257 (98.24 ± 82.27 µM·d-1). The maximum degradation rate in the carbon cloth BES (16 258 µM·d-1) was 5.6-fold lower than that obtained in the graphite brush BES. Both graphite 11 259 and carbon cloth electrodes showed higher degradation rates than biotic controls (10.95 260 ± 1.38 µM·d-1) indicating that direct transfer of H2 from electrochemical dehydrogenation 261 promoted the growth of Dehalogenimonas and enhanced 1,2-DCP degradation. 262 The maximum specific growth rate constant and the growth yield of Dehalogenimonas 263 was 0.105 ± 0.005 d-1 and 1.2·108 ± 5·107 cells per µmol of 1,2-DCP degraded, 264 respectively, calculated from values obtained from the exponential growth phase of BES 265 with graphite brush in the 1,2-DCP amendments 2 to 5. By applying Eq. (5), we obtained 266 a specific 1,2-DCP degradation rate of 1.0·10-9 ± 5·10-10 µmol of 1,2-DCP degraded per 267 cell and day during the whole exponential growth phase of Dehalogenimonas. The growth 268 yield of Dehalogenimonas in BES was one order of magnitude higher than those reported 269 for Dehalogenimonas lykanthroporepellens strain BL-DC-9 grown with 1,2-DCP [31] 270 but one order of magnitude lower than growth yields of Dehalogenimonas grown with 271 chlorinated ethenes [26]. 272 The current intensities recorded during the operation of BES (Figure S3) were used to 273 calculate the coulombic efficiencies (CEs) and the energy inputs for each experiment 274 (Figure 2B). The graphite fibre brush cells, which reached higher degradation rates than 275 those with carbon cloth, presented low CEs, ranging from 0.73 ± 0.9% in the first 276 amendment, to 4.91 ± 4.02% in the amendment that was consumed faster. At the same 277 time, the energy input per mole of contaminant degraded was inversely correlated to the 278 CEs, obtaining lower values when the degradation rate was higher, and ranging from 1.91 279 ± 0.23 kWh·mol-1 to 0.31 ± 0.02 kWh·mol-1. On the other hand, the carbon cloth BES, 280 despite presenting lower degradation rates, showed higher CEs and lower energetic 281 consumptions per mole of contaminant degraded, obtaining values at its final 1,2-DCP 282 amendment of 19.05% and 0.069 kWh·mol-1. This increase in the electrical efficiency 283 and consumption values were caused by the consistently lower intensities recorded during 12 284 the operation of the cells with carbon cloth as cathode compared to the graphite brush 285 containing ones. 286 3.2. Pulsed and continuous voltage operation of BES 287 The graphite brush was selected for further BES experiments. To increase the low CEs 288 observed in the previous experiments and increase the efficiency of the process while 289 maintaining a high degradation rate, two voltage operational modes were tested in a BES 290 for 140 days (Figure 3A). First, two different cathodic potentials were alternated during 291 the first 46 days: -0.6 V during 16 h and -1.1 V during 8 h. Then, a continuous cathodic 292 voltage of -1.1 V was applied during the next 89 days (Figure 3B). This operation enabled 293 the successful consumption of 10 mM 1,2-DCP in 140 d (Figure 4A). The maximum 294 degradation rate during the pulsed current period was achieved from day 17 to 43 and it 295 was fitted to a linear regression (Figure 4B), obtaining a value of 255.9 µM of 1,2-DCP 296 degraded per day. The same methodology was applied to the interval of day 58 to 135, 297 which corresponded to the continuous current application, and an 86.2% lower 298 degradation rate (33.4 µM·d-1) was obtained. In the abovementioned periods, the CE was 299 16.60% and 0.86% for the pulsed and continuous voltage operation, respectively. The CE 300 of the pulsed operation was slightly lower than the 19.05% obtained in the carbon cloth 301 containing BES but with a 16-fold degradation rate. This high CE is coupled with a low 302 energetic input of 0.076 kWh·mol-1, which is also similar to the values obtained in the 303 carbon cloth cells. On the other hand, when applying a continuous voltage, the decrease 304 of the degradation rate and the higher intensities applied drastically hindered CE, being a 305 95% lower than that obtained when applying a pulsed voltage. Also, when applying a 306 continuous voltage to achieve a cathode potential of -1.1 V, the required energetic input 307 increased 33-fold (2.499 kWh·mol-1). Applying a pulsed voltage not only reduced the 308 electric energy input but also stimulated the degrading activity of Dehalogenimonas. 13 309 Therefore, the pulsed voltage operation can be a suitable strategy in a hypothetical scale- 310 up of the process to reduce the energy input in a large-scale process while maintaining 311 the high degradation rates required for the sustainability of the process. 312 3.3. Scanning electron microscopy 313 Scanning electron microscope (SEM) was applied to uncover cell distribution and cell 314 morphology in the cathodic electrodes and liquid samples at the end of each operation. 315 Two main cell morphologies comprising irregular-shaped cocci and bacillus were clearly 316 identified in the filtered suspension cultures (Figure 5A, C and E). These cocci and 317 bacillus morphologies are in agreement with the presence of Dehalogenimonas and 318 Desulfovibrio, the two predominant bacterial genus in this culture [32]. Also, several 319 aggregates were observed on the graphite brush (Figure 5B and F) and, in less extension, 320 on carbon cloth electrode (Figure 5D). The formation of cell aggregates is common in co- 321 cultures with obligate syntrophic interactions, which facilitates the cell-to-cell contacts 322 and enhance metabolite fluxes between species. Desulfovibrio has been frequently 323 detected in co-cultures and tri-cultures with organohalide-respiring bacteria, and in some 324 cases its presence is required to proceed with the dechlorination [33,34]. The role of 325 Desulfovibrio is not fully understood but some studies suggest that might provide organic 326 cofactors (i.e. corrinoids) to organohalide-respiring bacteria, which are essential for the 327 growth of Dehalogenimonas [35,36]. The micrographs obtained in our study suggest that 328 the surface of the graphite brush electrode can provide suitable characteristics to attach 329 aggregates of Dehalogenimonas and Desulfovibrio, especially considering that it has been 330 previously reported the capability of Desulfovibrio populations of synthetizing 331 extracellular polymeric substances (EPS) [37], which are required for a biofilm 332 formation. 333 4. Conclusions 14 334 This study demonstrates for the first time the capability of growing Dehalogenimonas in 335 a BES to degrade 1,2-DCP into the non-toxic product propene. The graphite brush 336 electrodes resulted in a 5.6-fold higher degradation rates than those with carbon cloth. 337 The application of periodic pulses of voltage allowed to maintain high CE and 338 degradation rates while decreasing the energetic input required with the graphite brush 339 electrode. The use of BES allowed to obtain final Dehalogenimonas concentrations up to 340 108 16S gene copies per mL of liquid culture, making these systems a promising 341 technology to produce high density Dehalogenimonas cultures for bioaugmentation 342 purposes. Further research is required to assess the optimal operational conditions for the 343 application of this technology in a scaled-up process. 344 345 Acknowledgements 346 This work was supported by the Spanish Ministry of Science, Innovation and Universities 347 project CTM2017-91879-EXP. D. Fernández-Verdejo acknowledges a predoctoral grant 348 from UAB (PIF 2017-2018). 349 The authors declare no conflict of interest. 350 351 References 352 [1] L. Martín-González, S. Hatijah Mortan, M. Rosell, E. Parladé, M. Martínez- 353 Alonso, N. Gaju, G. Caminal, L. Adrian, E. Marco-Urrea, Stable carbon isotope 354 fractionation during 1,2-dichloropropane-to-propene transformation by an 355 enrichment culture containing Dehalogenimonas strains and a dcpA gene, Environ. 356 Sci. Technol. 49 (2015) 8666–8674. doi:10.1021/acs.est.5b00929. 15 357 [2] Environmental Protection Agency (EPA), Draft Scopes of the Risk Evaluations To 358 Be Conducted for Thirteen Chemical Substances Under the Toxic Substances 359 Control Act; Notice of Availability, 2020. 360 [3] G. Pecoraino, L. Scalici, G. Avellone, L. Ceraulo, R. Favara, E.G. Candela, M.C. 361 Provenzano, C. Scaletta, Distribution of volatile organic compounds in Sicilian 362 groundwaters analysed by head space-solid phase micro extraction coupled with 363 gas chromatography mass spectrometry (SPME/GC/MS), Water Res. 42 (2008) 364 3563–3577. doi:10.1016/j.watres.2008.07.022. 365 [4] C. Courbet, A. Rivière, S. Jeannottat, S. Rinaldi, D. Hunkeler, H. Bendjoudi, G. 366 De Marsily, Complementing approaches to demonstrate chlorinated solvent 367 biodegradation in a complex pollution plume: Mass balance, PCR and compound- 368 specific stable isotope analysis, J. Contam. Hydrol. 126 (2011) 315–329. 369 doi:10.1016/j.jconhyd.2011.08.009. 370 [5] A.J. Tesoriero, F.E. Löffler, H. Liebscher, Fate and origin of 1,2-dichloropropane 371 in an unconfined shallow aquifer, Environ. Sci. Technol. 35 (2001) 455–461. 372 doi:10.1021/es001289n. 373 [6] M. Schmitt, S. Varadhan, S. Dworatzek, J. Webb, E. Suchomel, Optimization and 374 validation of enhanced biological reduction of 1,2,3-trichloropropane in 375 groundwater, Remediation. 28 (2017) 17–25. doi:10.1002/rem.21539. 376 [7] T.A. Key, K.S. Bowman, I. Lee, J. Chun, L. Albuquerque, M.S. da Costa, F.A. 377 Rainey, W.M. Moe, Dehalogenimonas formicexedens sp. nov., a chlorinated 378 alkanerespiring bacterium isolated from contaminated groundwater, Int. J. Syst. 379 Evol. Microbiol. 67 (2017) 1366–1373. doi:10.1099/ijsem.0.001819. 16 380 [8] W.M. Moe, J. Yan, M.F. Nobre, M.S. da Costa, F.A. Rainey, Dehalogenimonas 381 lykanthroporepellens gen. nov., sp. nov., a reductively dehalogenating bacterium 382 isolated from chlorinated solvent-contaminated groundwater, Int. J. Syst. Evol. 383 Microbiol. 59 (2009) 2692–2697. doi:10.1099/ijs.0.011502-0. 384 [9] K.E. Fletcher, F.E. Löffler, H.H. Richnow, I. Nijenhuis, Stable carbon isotope 385 fractionation of 1,2-dichloropropane during dichloroelimination by 386 Dehalococcoides populations, Environ. Sci. Technol. 43 (2009) 6915–6919. 387 doi:10.1021/es900365x. 388 [10] N. Blázquez-Pallí, M. Rosell, J. Varias, M. Bosch, A. Soler, T. Vicent, E. Marco- 389 Urrea, Integrative isotopic and molecular approach for the diagnosis and 390 implementation of an efficient in-situ enhanced biological reductive dechlorination 391 of chlorinated ethenes, Water Res. 167 (2019). doi:10.1016/j.watres.2019.115106. 392 [11] J. Herrero, D. Puigserver, I. Nijenhuis, K. Kuntze, J.M. Carmona, Combined use 393 of ISCR and biostimulation techniques in incomplete processes of reductive 394 dehalogenation of chlorinated solvents, Sci. Total Environ. 648 (2019) 819–829. 395 doi:10.1016/j.scitotenv.2018.08.184. 396 [12] P. Leitão, S. Rossetti, H.P.A. Nouws, A.S. Danko, M. Majone, F. Aulenta, 397 Bioelectrochemically-assisted reductive dechlorination of 1,2-dichloroethane by a 398 Dehalococcoides-enriched microbial culture, Bioresour. Technol. 195 (2015) 78– 399 82. doi:10.1016/j.biortech.2015.06.027. 400 [13] P. Leitão, S. Rossetti, A.S. Danko, H. Nouws, F. Aulenta, Enrichment of 401 Dehalococcoides mccartyi spp. from a municipal activated sludge during AQDS- 402 mediated bioelectrochemical dechlorination of 1,2-dichloroethane to ethene, 403 Bioresour. Technol. 214 (2016) 426–431. doi:10.1016/j.biortech.2016.04.129. 17 404 [14] F. Aulenta, L. Tocca, R. Verdini, P. Reale, M. Majone, Dechlorination of 405 trichloroethene in a continuous-flow bioelectrochemical reactor: Effect of cathode 406 potential on rate, selectivity, and electron transfer mechanisms, Environ. Sci. 407 Technol. 45 (2011) 8444–8451. doi:10.1021/es202262y. 408 [15] F. Aulenta, R. Verdini, M. Zeppilli, G. Zanaroli, F. Fava, S. Rossetti, M. Majone, 409 Electrochemical stimulation of microbial cis-dichloroethene (cis-DCE) oxidation 410 by an ethene-assimilating culture, N. Biotechnol. 30 (2013) 749–755. 411 doi:10.1016/j.nbt.2013.04.003. 412 [16] F. Chen, Z.L. Li, J. qi Yang, B. Liang, X.Q. Lin, J. Nan, A.J. Wang, Effects of 413 different carbon substrates on performance, microbiome community structure and 414 function for bioelectrochemical-stimulated dechlorination of tetrachloroethylene, 415 Chem. Eng. J. 352 (2018) 730–736. doi:10.1016/j.cej.2018.07.082. 416 [17] H. Wan, X. Yi, X. Liu, C. Feng, Z. Dang, C. Wei, Time-dependent bacterial 417 community and electrochemical characterizations of cathodic biofilms in the 418 surfactant-amended sediment-based bioelectrochemical reactor with enhanced 419 2,3,4,5-tetrachlorobiphenyl dechlorination, Environ. Pollut. 236 (2018) 343–354. 420 doi:10.1016/j.envpol.2018.01.048. 421 [18] D. Cecconet, F. Sabba, M. Devecseri, A. Callegari, A.G. Capodaglio, In situ 422 groundwater remediation with bioelectrochemical systems: A critical review and 423 future perspectives, Environ. Int. 137 (2020) 105550. 424 doi:10.1016/j.envint.2020.105550. 425 [19] F. Chen, Z.L. Li, A.J. Wang, Acceleration of microbial dehalorespiration with 426 electrical stimulation, in: Bioelectrochemistry Stimul. Environ. Remediat. From 427 Bioelectrorespiration to Bioelectrodegradation, Springer Singapore, Singapore, 18 428 2018: pp. 73–92. doi:10.1007/978-981-10-8542-0_4. 429 [20] R.J. Steffan, S. Vainberg, Production and handling of Dehalococcoides 430 bioaugmentation cultures, in: H.F. Stroo, A. Leeson, C.H. Ward (Eds.), 431 Bioaugmentation Groundw. Remediat., Springer New York, New York, NY, 2013: 432 pp. 89–115. doi:10.1007/978-1-4614-4115-1_1. 433 [21] F. Chen, Z. Li, J. Yang, B. Liang, C. Huang, W. Cai, J. Nan, A. Wang, Electron 434 fluxes in biocathode bioelectrochemical systems performing dechlorination of 435 chlorinated aliphatic hydrocarbons, Front. Microbiol. 9 (2018) 1–10. 436 doi:10.3389/fmicb.2018.02306. 437 [22] R. Verdini, F. Aulenta, F. De Tora, A. Lai, M. Majone, Relative contribution of set 438 cathode potential and external mass transport on TCE dechlorination in a 439 continuous-flow bioelectrochemical reactor, Chemosphere. 136 (2015) 72–78. 440 doi:10.1016/j.chemosphere.2015.03.092. 441 [23] A. Trueba-Santiso, K. Wasmund, J.M. Soder-Walz, E. Marco-Urrea, L. Adrian, 442 Genome sequence, proteome profile, and identification of a multiprotein reductive 443 dehalogenase complex in Dehalogenimonas alkenigignens strain BRE15M, J. 444 Proteome Res. 20 (2021) 613–623. doi:10.1021/acs.jproteome.0c00569. 445 [24] L. Adrian, U. Szewzyk, J. Wecke, H. Görisch, Bacterial dehalorespiration with 446 chlorinated benzenes, Nature. 408 (2000) 580–583. doi:10.1038/35046063. 447 [25] A. Trueba-Santiso, E. Parladé, M. Rosell, M. Lliros, S.H. Mortan, M. Martínez- 448 Alonso, N. Gaju, L. Martín-González, T. Vicent, E. Marco-Urrea, Molecular and 449 carbon isotopic characterization of an anaerobic stable enrichment culture 450 containing Dehalobacterium sp. during dichloromethane fermentation, Sci. Total 19 451 Environ. 581–582 (2017) 640–648. doi:10.1016/j.scitotenv.2016.12.174. 452 [26] Y. Yang, S.A. Higgins, J. Yan, B. Şimşir, K. Chourey, R. Iyer, R.L. Hettich, B. 453 Baldwin, D.M. Ogles, F.E. Löffler, Grape pomace compost harbors organohalide- 454 respiring Dehalogenimonas species with novel reductive dehalogenase genes, 455 ISME J. 11 (2017) 2767–2780. doi:10.1038/ismej.2017.127. 456 [27] K.M. Ritalahti, B.K. Amos, Y. Sung, Q. Wu, S.S. Koenigsberg, F.E. Löffler, 457 Quantitative PCR targeting 16S rRNA and reductive dehalogenase genes 458 simultaneously monitors multiple Dehalococcoides strains, Appl. Environ. 459 Microbiol. 72 (2006) 2765–2774. doi:10.1128/AEM.72.4.2765-2774.2006. 460 [28] L. Rago, Y. Ruiz, J.A. Baeza, A. Guisasola, P. Cortés, Microbial community 461 analysis in a long-term membrane-less microbial electrolysis cell with hydrogen 462 and methane production, Bioelectrochemistry. 106 (2015) 359–368. 463 doi:10.1016/j.bioelechem.2015.06.003. 464 [29] E. Blázquez, D. Gabriel, J.A. Baeza, A. Guisasola, Treatment of high-strength 465 sulfate wastewater using an autotrophic biocathode in view of elemental sulfur 466 recovery, Water Res. 105 (2016) 395–405. doi:10.1016/j.watres.2016.09.014. 467 [30] R.E. Richardson, Organohalide respiring bacteria as members of microbial 468 communities: catabolic food webs and biochemical interactions, in: L. Adrian, F. 469 E. Löffler (Eds.), Organohalide Respiring Bacteria, Springer New York, 2015: pp. 470 309-341. doi: 10.1007/978-3-662-49875-0 471 [31] J. Yan, B.A. Rash, F.A. Rainey, W.M. Moe, Isolation of novel bacteria within the 472 Chloroflexi capable of reductive dechlorination of 1,2,3-trichloropropane, 473 Environ. Microbiol. 11 (2009) 833–843. doi:10.1111/j.1462-2920.2008.01804.x. 20 474 [32] K.S. Bowman, M.F. Nobre, M.S. da Costa, F.A. Rainey, W.M. Moe, 475 Dehalogenimonas alkenigignens sp. nov., a chlorinated-alkane-dehalogenating 476 bacterium isolated from groundwater, Int. J. Syst. Evol. Microbiol. 63 (2013) 477 1492–1498. doi:10.1099/ijs.0.045054-0. 478 [33] H.D. May, G.S. Miller, B. V. Kjellerup, K.R. Sowers, Dehalorespiration with 479 polychlorinated biphenyls by an anaerobic ultramicrobacterium, Appl. Environ. 480 Microbiol. 74 (2008) 2089–2094. doi:10.1128/AEM.01450-07. 481 [34] A. Grostern, E.A. Edwards, A 1,1,1-trichloroethane-degrading anaerobic mixed 482 microbial culture enhances biotransformation of mixtures of chlorinated ethenes 483 and ethanes, Appl. Environ. Microbiol. 72 (2006) 7849–7856. 484 doi:10.1128/AEM.01269-06. 485 [35] Y. Men, H. Feil, N.C. Verberkmoes, M.B. Shah, D.R. Johnson, P.K.H. Lee, K.A. 486 West, S.H. Zinder, G.L. Andersen, L. Alvarez-Cohen, Sustainable syntrophic 487 growth of Dehalococcoides ethenogenes strain 195 with desulfovibrio vulgaris 488 hildenborough and methanobacterium congolense: Global transcriptomic and 489 proteomic analyses, ISME J. 6 (2012) 410–421. doi:10.1038/ismej.2011.111. 490 [36] J. He, V.F. Holmes, P.K.H. Lee, L. Alvarez-Cohen, Influence of vitamin B12 and 491 cocultures on the growth of Dehalococcoides isolates in defined medium, Appl. 492 Environ. Microbiol. 73 (2007) 2847–2853. doi:10.1128/AEM.02574-06. 493 [37] R. Stadler, L. Wei, W. Fürbeth, M. Grooters, A. Kuklinski, Influence of bacterial 494 exopolymers on cell adhesion of Desulfovibrio vulgaris on high alloyed steel: 495 Corrosion inhibition by extracellular polymeric substances (EPS), Mater. Corros. 496 61 (2010) 1008–1016. doi:10.1002/maco.201005819. 21 497 Figure captions 498 Figure 1. Degradation profile of 1,2-DCP in BES (Panel A), accumulated concentration 499 of 1,2-DCP degraded and propene produced (Panel B) and Dehalogenimonas 16S rDNA 500 gene copies per mL (Panel C), with different cathodic electrode materials (●: Graphite 501 brush, ■: Carbon cloth) and cathodic potential set at -0.7 V. Solid symbols and open 502 symbols refer to 1,2-DCP and propene, respectively. Numbers indicate the number of 1,2- 503 DCP amendments added in BES with graphite brush. Values plotted for BES with 504 graphite brush are average of duplicates and error bars indicate 1 standard deviation. 505 Values plotted for BES with carbon cloth are for an individual BES. 506 Figure 2. Degradation rate and growth yield of Dehalogenimonas obtained for each dose 507 of 1,2-DCP consumed in BES (Panel A) and coulombic efficiencies and energetic inputs 508 obtained during the degradation (Panel B). Dose number refer to the 1,2-DCP 509 amendments depicted in Fig. 1. 510 Figure 3. Degradation profile of 1,2-DCP in a BES containing a graphite brush as 511 electrode (Panel A) and its measured current intensities during the operation of pulsed 512 and continuous voltage (Panel B). The BES received several additions of 1,2-DCP as 513 indicated by the arrows. 514 Figure 4. 1,2-DCP consumed in a BES containing a graphite brush as electrode (panel 515 A) and linear regression of the accumulated 1,2-DCP degradation values corresponding 516 to the degradation rate when applying pulsed (solid symbols) and continuous (open 517 symbols) voltage. 518 Figure 5. Microscopic analysis of the carbon fiber brush (A and B) and carbon cloth 519 experiments (C and D) described in section 3.1 and graphite brush experiment described 22 520 in section 3.2 (E and F) at the end of the experiments. For each BES, filtered liquid 521 samples (A, C and E) and solid electrode samples (B, D and F) were analysed by SEM. 522 23 523 Figure 1 524 525 526 527 24 528 Figure 2 529 530 531 532 25 533 Figure 3 2000 A 1,2-Dichloropropane (M) 1500 1000 500 0 0 50 100 150 Time (days) 25 B Current Density (mA/m2) 20 15 10 5 0 0 50 100 150 Time (days) 534 535 536 537 538 539 540 26 541 Figure 4 1,2-Dichloropropane degraded (M) 15000 A Pulsed voltage Continuous voltage 10000 5000 0 0 50 100 150 Time (days) 1,2-Dichloropropane degraded (M) 15000 B 10000 Y = 255.9*X - 3453 Y = 33.40*X + 6684 R2 = 0.9849 R2 = 0.9854 5000 0 0 50 100 150 Time (days) 542 543 544 545 546 547 548 27 549 Figure 5 550 551 28