This is the accepted version of the article: Trueba Santiso, Alba María; Fernandez Verdejo, David Juan; Marco Rius, Irene; [et al.]. «Interspecies interaction and effect of co-contaminants in an anaerobic dichloromethane-degrading culture». Chemosphere, Vol. 240 (Feb. 2020), art. 124877. DOI 10.1016/j.chemosphere.2019.124877 This version is avaible at https://ddd.uab.cat/record/212957 under the terms of the license 1 Interspecies interaction and effect of co-contaminants 2 in an anaerobic dichloromethane-degrading culture 3 Alba Trueba-Santisoa, David Fernández-Verdejoa, Irene Marco-Riusb, Jesica M. Soder- 4 Walza, Oriol Casabellaa, Teresa Vicenta, Ernest Marco-Urreaa,*. a 5 Departament d'Enginyeria Química, Biològica i Ambiental, Universitat Autònoma de 6 Barcelona (UAB), Carrer de les Sitges s/n, 08193 Cerdanyola del Vallès, Spain. b 7 Institute for Bioengineering of Catalonia. Parc Científic de Barcelona. Edifici Clúster 8 c/ Baldiri Reixac 10-12, 08028 Barcelona, Spain. 9 10 *Corresponding author 11 Departament d’Enginyeria Química, Biològica i Ambiental, Universitat Autònoma de 12 Barcelona (UAB), c/ de les Sitges s/n, 08193 Cerdanyola del Vallès, Spain. E-mail: 13 ernest.marco@uab.cat, Phone: +34 935812694. 14 15 1 16 ABSTRACT 17 An anaerobic stable mixed culture dominated by bacteria belonging to the genera 18 Dehalobacterium, Acetobacterium, Desulfovibrio, and Wolinella was used as a model to 19 study the microbial interactions during DCM degradation. Physiological studies 20 indicated that DCM was degraded in this mixed culture at least in a three-step process: 21 i) fermentation of DCM to acetate and formate, ii) formate oxidation to CO2 and H2, and 22 iii) H2/CO2 reductive acetogenesis. The 16S rRNA gene sequencing of cultures enriched 23 with formate or H2 showed that Desulfovibrio was the dominant population followed by 24 Acetobacterium, but sequences representing Dehalobacterium were only present in 25 cultures amended with DCM. Nuclear magnetic resonance analyses confirmed that 13 26 acetate produced from C-labeled DCM was marked at the methyl ([2-13C]acetate), 27 carboxyl ([1-13C]acetate), and both ([1,2-13C]acetate) positions, which is in accordance 28 to acetate formed by both direct DCM fermentation and H2/CO2 acetogenesis. The 29 inhibitory effect of ten different co-contaminants frequently detected in groundwaters 30 on DCM degradation was also investigated. Complete inhibition of DCM degradation 31 was observed when chloroform, perfluorooctanesulfonic acid, and diuron were added at 32 838, 400, and 107 µM, respectively. However, the inhibited cultures recovered the 33 DCM degradation capability when transferred to fresh medium without co- 34 contaminants. Findings derived from this work are of significant relevance to provide a 35 better understanding of the synergistic interactions among bacteria to accomplish DCM 36 degradation as well as to predict the effect of co-contaminants during anaerobic DCM 37 bioremediation in groundwater. 38 KEYWORDS: 39 Dehalobacterium; dichloromethane; co-contaminants; inhibition; bioremediation. 40 2 41 1. INTRODUCTION 42 Dichloromethane (DCM) is a frequently detected groundwater contaminant that 43 typically occurs in mixtures in either disposal sites or domestic well samples [1,2], and 44 it is ranked 11th (out of 33) on the European Union priority list of pollutants [3]. 45 Biodegradation of DCM has been described in the literature under both oxic and anoxic 46 conditions [4], and it has become apparent that bacteria play a key role in the natural 47 attenuation of sites impacted with DCM [5]. Most of the described DCM-degrading 48 bacteria are aerobic methylotrophic [4], but in contaminated aquifers oxygen is oftenly 49 very limited. Under anoxic conditions, Dehalobacterium, Dehalobacter, and 50 Candidatus Dichloromethanomonas are the only known genera capable of utilizing 51 DCM as growth substrate [6,7,8]. Dehalobacterium formicoaceticum is the only 52 reported isolate [6], and previous studies suggested that it utilizes DCM as sole growth 53 substrate and ferments this compound to acetate and formate [9]. Detailed physiological 54 studies with Candidatus Dichloromethanomonas elyuquensis and Dehalobacter- 55 containing cultures have shown that DCM degradation requires syntrophic partners (i.e. 56 hydrogenotrophic homoacetogens and methanogens) present in the mixed culture, 57 probably to prevent H2 from reaching inhibitory levels [10,11]. The reasons why H2 58 exerts and inhibitory effect on DCM degradation are still unclear, but Chen et al. [11] 59 hypothesized that it could be provoked by formate accumulation inside the cell of 60 Candidatus Dichloromethanomonas elyuquensis because this strain does not contain 61 formate transporters and formate conversion to CO2 and H2 is impeded at high 62 concentrations of H2. 63 Halogenated compounds are often present in aquifers as complex mixtures from 64 industrial, urban or agricultural practices, and in some cases may impede the 65 accomplishment of remedial goals. In particular, chloroform (CF), chlorinated ethenes, 3 66 and chlorinated ethanes have been shown to inhibit the dechlorinating activity of 67 organohalide-respiring bacteria [12-15]. To date, publications on the inhibitory effect of 68 certain chlorinated solvents over DCM-degrading bacteria are scarce. For instance, 69 previous studies showed that l,1-dichloroethane, tetrachloroethene, and trichloroethene 70 (TCE) did not affect DCM degradation in D. formicoaceticum, while 71 tetrachloromethane, CF, and chloromethane caused an immediate and complete 72 inhibition of DCM degradation and bacterial growth [9]. A better understanding of the 73 effect of co-contaminants in enrichment cultures under laboratory conditions may 74 provide a basis to anticipate whether bioremediation is a feasible strategy to remediate 75 groundwater when multiple contaminants are present. 76 The aim of the work presented herein is two-fold. First, we attempt to examine 77 the bacterial interactions in a mixed culture during DCM degradation. We use a 78 Dehalobacterium-containing culture derived from a membrane bioreactor operating in 79 an industrial wastewater treatment plant that had been enriched with DCM for more 80 than three years [16]. Previous studies indicated that this mixed bacterial culture was 81 mainly composed by four genera (Dehalobacterium, Acetobacterium, Desulfovibrio, 82 and Wolinella) and produced acetate and formate when grown with DCM and H2 in a 83 bicarbonate-buffered medium [16]. Secondly, we assessed the effect of selected 84 halogenated contaminants on the DCM-degrading capability of this mixed culture. The 85 list of tested compounds comprises chloroalkanes, chloroalkenes, CF, acetone, 86 polyfluoroalkyl substances, as well as the halogenated pesticide diuron and its 87 transformation product 3,4-dichloroaniline (3,4-DCA). The selected compounds are 88 among the most frequently detected in contaminated groundwater, do not serve as 89 substrates for this culture [16], and are all included in the 2017 ATSDR Priority List of 90 Hazardous Substances [17], with the exception of 3,4-DCA. 4 91 2. MATERIALS AND METHODS 92 2.1 Cultivation 93 The experiments were performed with a stable DCM-degrading consortium composed 94 predominantly by bacteria from the genera Dehalobacterium, Acetobacterium, 95 Wolinella and Desulfovibrio derived from slurry samples of the membrane bioreactor of 96 an industrial wastewater treatment plant [16]. The culture was grown in 100 mL glass 97 serum bottles (microcosms) with 65 mL of reduced, anoxic, bicarbonate-buffered 98 medium (pH=7) described elsewhere but lacking acetate and H2 [16]. Briefly, this 99 medium contained DCM, yeast extract (200 mg/L), vitamins, trace elements, tungsten 100 (22.8 μM), selenium (24.2 μM), and Na2S · 9H2O and L-cysteine (0.2 mM each) as 101 reducing agents. Bicarbonate (0.01 M) and resazurin (0.25 mg/L) were used as a buffer 102 and redox indicator, respectively. The serum bottles were sealed with Teflon-coated 103 butyl rubber stoppers and aluminium crimp caps and gassed with N2 (0.4 bar 104 overpressure). The bacterial consortia was maintained active by transferring a 3% (v/v) 105 inoculum during the exponential degradation phase of 2 mM DCM doses into fresh 106 medium. The microcosms were cultivated under static conditions (25°C) in the dark. 107 Two controls were always included for each experiment in triplicate: i) abiotic controls 108 consisting of anaerobic medium plus DCM to discard its abiotic transformation, and ii) 109 biotic controls consisting of anaerobic medium plus bacterial inoculum but without 110 DCM to evaluate production of metabolites from the components of the medium. DCM 111 was replaced by either formate or H2 when indicated. 112 To elucidate the role of the bacteria present in the mixed culture in the 113 degradation of the daughter products of DCM, the changes in the bacterial community 114 structure were studied after applying the dilution-to-extinction method [18] in 12-mL 5 115 vials containing the anoxic medium described above but using either formate (420 116 µmol) or H2 (0.2 bar overpressure) as substrate in parallel cultures without DCM. After 117 two extinction series, the more diluted vial showing activity against the tested substrate 118 was selected as inoculum for serum bottle microcosms and it was amended with 119 different substrates. The procedure followed and the vials selected are depicted in Fig. 120 S1A and B. 121 2.2 Analytical methods 122 Volatile halogenated compounds and acetone were quantified by injecting a 0.5-mL 123 headspace sample into an Agilent 6890N gas chromatograph (GC) fitted with an 124 Agilent DB-624 column (30 m × 0.32 mm with 0.25 μm film thickness) and a flame 125 ionization detector as previously described [16]. Formate and acetate were measured 126 using a Dionex 3000 Ultimate high-performance liquid chromatography equipped with 127 a UV detector at 210 nm, as described elsewhere [16]. Hydrogen and carbon dioxide 128 concentration was measured on 0.1-mL headspace samples using an Agilent 7820A GC 129 equipped with a thermal conductivity detector. Separation was achieved using a 130 MolSieve 5A 60/80 SS (1.82 m x 2 mm, Agilent) and a Porapak Q 60/80 UM columns 131 (1.82 m x 2 mm, Agilent). Nitrogen was the carrier gas at 138 kPa. Oven temperature 132 was held isothermal at 40 ºC, the injector temperature at 200 ºC and the detector 133 temperature at 250 ºC. Peak areas were calculated using Chromeleon software (Thermo 134 Scientific). Results were presented as either nominal concentrations or total μmoles per 135 bottle. Calibration was based on aqueous standards under the same conditions of 136 temperature, pressure and liquid and headspace volumes as those in the microcosms. 137 Nuclear magnetic resonance (NMR) analysis was performed with 13 138 Dehalobacterium-containing cultures grown with 3.6 mM of C-DCM using the same 6 139 medium described above. Samples were taken at the initial time (when DCM was 140 amended) and after DCM consumption, and 500 µL of each sample were mixed with 50 141 µL deuterium oxide and transferred to 5 mm wide NMR tubes. 1H decoupled 13C-NMR 142 was performed at 25ºC in a 500 MHz vertical-bore Bruker spectrometer using a CPTCI 143 probe (Bruker BioSpin GmBH). The acquisition parameters were the following: 144 repetition time = 1 s, acquisition time = 0.56 s, nominal flip angle = 30º, pulse width = 145 5.7 s (hard pulse), spectral width = 27.8 kHz, number of points = 32768, number of 146 scans = 30720. Data were Fourier transformed, zero-filled (2x), baseline and phase 147 corrected, and 1 Hz line broadening applied using TopSpin 4.0.6 (Bruker BioSpin 148 GmBH). To confirm the chemical shift assignment of the peaks, 6.5 mg of the NMR 149 standard 4,4-dimethyl-4-silapentane-1-sulfonic acid (DSS) were added to the samples 150 above and data acquired using the same parameters aforementioned. An additional 151 sample containing 1 M of unlabelled methanol, 1 M of unlabelled formic acid and DSS 152 in medium solution was also analysed by NMR to confirm the presence or not of those 153 compounds in the bacteria-metabolised samples. 154 2.3 DNA extraction and 16S rRNA gene amplicon sequencing 155 Cells were harvested by centrifugation (7000 × g, 40 min at 10 ºC) from 65-mL samples 156 (one microcosm) using sterile falcon tubes in an Avanti J-20 centrifuge. The pelleted 157 cells were resuspended in sterile PBS buffer and DNA was then extracted using the 158 Gentra Puregene Yeast/Bact kit (Qiagen) following the instructions from the 159 manufacturer. The DNA of each of the biological replicates was extracted and analysed 160 separately. Amplicons of the region V3-V4 for 16SrRNA genes were amplified with 161 primers S-D-Bact-0341-b-S-17/S-D-Bact-0785-a-A-21 [19] using the Illumina MiSeq 162 platform at Serveis de Genòmica i Bioinformàtica from the Universitat Autònoma de 7 163 Barcelona. Gene sequences were processed using the 16S Metagenomics workflow in 164 the MiSeq Reporter analysis software based on quality scores generated by real-time 165 analysis during the sequencing run. Quality-filtered indexed reads were demultiplexed 166 for generation of individual FASTQ files and aligned using the banded Smith– 167 Waterman method [20] of the Illumina-curated version of the Greengenes taxonomic 168 database. The output of this workflow was a classification of reads at multiple 169 taxonomic levels. To calculate relative abundance, we divided the number of sequences 170 belonging to a taxonomic level by the total number of sequences obtained from the 171 specific sample. 172 2.4 Inhibition tests 173 Each co-contaminant inhibition test consisted on two groups of experimental bottles 174 prepared in triplicates at the same time: (i) controls amended only with neat DCM and 175 (ii) parallel sets of experimental bottles amended with different concentrations of the co- 176 contaminant from acetone stock solutions plus neat DCM. Both experimental bottles 177 were inoculated with the Dehalobacterium-containing culture (3 %, v/v) at the 178 exponential phase of DCM degradation. Then, a time course experiment was performed, 179 monitoring DCM concentration daily by GC. All experimental bottles amended with co- 180 contaminants were compared to the corresponding controls amended only with neat 181 DCM. Rates were calculated from linear (R2>0.98, n≥3) DCM disappearance after the 182 lag phase period. 183 To study the functional recovery of DCM degradation after the complete 184 inhibition observed with some co-contaminants, cultures inhibited by CF were purged 185 with nitrogen to remove CF, and 3.5 mL of the inhibited cultures were transferred into 186 fresh medium spiked with neat DCM. Those cultures inhibited by other halogenated 8 187 compounds were directly transferred to fresh medium without purge. These experiments 188 were set up at least in duplicate. 189 2.5 Microscope observation 190 Aliquots (20 µL) of Dehalobacterium-containing cultures were placed in slides, covered 191 with a cover slip, and observed under a light microscope (Zeiss Axioscop) at 40 x or 192 100 x (adding immersion oil) objectives. 193 194 3. RESULTS AND DISCUSSION 195 3.1 Mass balance and fate of DCM 196 Dehalobacterium-containing cultures fed with DCM (153.9 ± 0.4 µmol) were initially 197 stablished in the abovementioned defined medium without H2 but containing CO2 from 198 the bicarbonate buffered medium. After a lag phase of 7 d, DCM degradation occurred 199 at a rate of 902 ± 10 µmol d-1 L-1, and acetate and CO2 were produced concomitantly 200 (Fig. 1). Formate and H2 were potential degradation intermediates, however they were 201 not detected (Fig. 1). After a second feeding of DCM, it was consumed without an 202 observable lag phase, indicating that bacterial degradation of DCM is a growth- 203 supporting process. Controls without DCM also produced acetate but at noticeable 204 lower concentrations, presumably derived from the fermentation of the yeast extract 205 contained in the medium (Fig. S2). Different attempts were made to remove yeast 206 extract from the medium but always resulted in a marked decrease in DCM degradation. 207 Formate was provided at two different concentrations (430.9 ± 39.6 and 964.3 ± 208 71.7 µmol) in cultures where DCM and H2 were omitted to ascertain whether formate 209 was oxidized by the microbial consortia (Fig. 2). Formate degradation was accompanied 9 210 by an increase in acetate and H2 concentration (Fig. 2, Fig. S3A). The concentration of 211 CO2 slightly increased in the initial stages, but after 4 d it was consumed coinciding 212 with a steep increase of acetate in the medium (Fig. S3A). A decrease in CO2 was not 213 observed in controls where formate was omitted (Fig. S3B), suggesting that acetate was 214 produced via CO2 reduction. 215 The microcosms amended with 7 mL of H2 (~268 µmol) consumed both H2 and 216 CO2 and produced acetate, indicating H2/CO2 reductive acetogenesis (Fig. 3). Acetate 217 was also generated in the controls without H2 but at significantly lower concentrations 218 (Fig. 3). 219 Physiological studies with the isolate D. formicoaceticum suggested that DCM is 220 transformed to methylene-tetrahydrofolate, which is funnelled into the Wood-Ljunddahl 221 pathway and disproportionated [9]. By following this pathway, two thirds of methylene- 222 tetrahydrofolate is oxidized to formate, and the remaining third is reduced to acetate in 223 combination with CO2, according to Eq. 1: 224 3CH2Cl2 + 4H2O + CO2 → 2HCOO- + CH3COO- + 9H+ + 6Cl- (Eq. 1) 225 In our study, formate was not observed as a transient product when DCM was amended 226 to Dehalobacterium-containing cultures. Similarly, there was no evidence of formate 227 production in cultures containing Candidatus Dichloromethanomonas elyuquensis when 228 fed with DCM [11]. However, the fact that formate was turned over to CO2 and H2 (Fig. 229 2) provided a basis to hypothesize that formate derived from DCM was rapidly oxidized 230 by a yet unknown bacteria from the mixed culture (Eq. 2) and evaded detection. 231 HCOO- + H+ → H2 + CO2 (Eq. 2) 10 232 In addition, production of acetate in microcosms containing either H2 or formate 233 suggested that released hydrogen served as the electron donor for CO2 reduction to 234 acetate (Eq. 3): 235 4H2 + 2CO2 → CH3COO- + 2 H2O + H+ (Eq. 3) 236 Stoichiometric analysis revealed that the electrons generated from H2 oxidation in 237 cultures growing with DCM, formate, and H2 were mostly recovered in acetate (range 238 between 65-87%) (Table S1). The balance did not precisely match the stoichiometric 239 conservation of H2 because part of the carbon derived from DCM fermentation was 240 sequestered into the biomass and not yielded acetate. 241 242 3.2 Degradation experiment using 13C-labelled DCM 243 The metabolites produced after 13C-labelled DCM degradation were analysed by NMR. 244 As observed in Fig. 4, acetate, CO2, and methanol were detected after the consumption 245 of 3.6 mM 13C-DCM. Production of acetate and CO2 had already been demonstrated by 246 chromatographic analyses, but methanol had not been detected. Methanol was 247 previously reported in D. formicoaceticum and it was considered as a side reaction 248 when DCM is uncoupled from growth [9]. 249 The metabolism of DCM fermentation following Eq. 1 implies that the formate 13 250 and the methyl group of acetate are derived entirely from C-DCM, whereas the 251 carboxyl group of acetate is formed from CO2 [9] (Fig. S4). The 13C-NMR spectrum of 13 252 the samples after C-DCM consumption displayed peaks at 26.0 ppm and 184.1 ppm, 253 corresponding to the methyl and carboxyl groups of acetate, respectively. The inserts on 254 Fig. 4B show the splitting caused by the carbon-carbon J-coupling (~52.3 Hz) in acetate 13 255 C-labeled in both methyl and carboxyl carbons ([1,2-13C]acetate). Therefore, we can 13 256 conclude that acetate derived from C-DCM degradation accumulated in form of 11 13 257 acetate labelled with C at the carboxyl ([1-13C]acetate), methyl ([2-13C]acetate), and 258 both ([1,2-13C]acetate) positions. Since data acquisition was not done on a fully-relaxed 259 state but under a steady-state condition, we cannot reliably quantify each species. 260 However, we can roughly calculate that there was a ratio [2-13C]acetate/[1,2-13C]acetate 261 ~ 2.5 and [1-13C]acetate/[1,2-13C]acetate ~ 2.1. This data suggests that two acetate 262 producing pathways could be operating in this culture. On the one hand, acetate labelled 263 solely in the methyl group ([2-13C]acetate) is consistent with 13C-DCM fermentation by 264 Dehalobacterium using unlabelled bicarbonate from the buffered medium (Eq. 1, Fig. 265 S4). This bacterium could also produce [1,2-13C]acetate if using 13 CO2 released during 13 13 266 C-formate oxidation. Hence, CO2 could be reduced to CO by CO dehydrogenase, 267 which is encoded in D. formicoaceticum genome [21], and become the carboxyl group 268 of acetate via acetyl-CoA using the Wood-Ljungdahl pathway (Fig. S4). In a previous 13 269 study, NMR analysis of products formed from C-DCM by the isolate D. 270 formicoaceticum showed that carboxyl group was unlabelled because the only source of 13 271 CO2 was the carbonate-buffered medium and C-formate was not further oxidized by 272 other bacteria to 13CO2 [9]. On the other hand, acetogenic bacteria could produce acetate 273 labelled indistinctly at C1 or C2 positions using the pool of labelled and unlabelled CO2 274 present in the medium, but unlike to Dehalobacterium, acetogens can produce acetate 275 labelled exclusively in the carboxyl group ([1-13C]acetate). 276 277 3.3. Role of bacteria in the mixed culture 278 We aimed to identify the community members responsible for the transformation of 279 DCM-degrading metabolites in this mixed culture. Hence, the approach was extinction 280 culturing by diluting the consortium using formate and H2 as sole substrates in parallel 12 281 treatments (Fig. S1). The resulting cultures were tested against DCM, formate, and H2 282 consumption, and their DNA sequenced using paired end Mi-Seq Illumina (Fig. 5). 283 No sequences representing Dehalobacterium were detected in the dataset 284 generated with DNA extracted from cultures grown with either formate or H2 (Fig. 5B- 285 D). Accordingly, these cultures were not capable of consuming DCM (data not shown) 286 corroborating that Dehalobacterium was responsible for DCM degradation. This is in 287 agreement with previous studies showing that formate alone could not support growth 288 of Candidatus Dichloromethanomonas elyuquensis and Dehalobacterium 289 formicoaceticum [9,11]. 290 When the cultures were enriched with formate, the bacterial community was 291 dominated by Desulfovibrio (69.6%) and Acetobacterium (26.7%) genera (Fig. 5B). In 292 cultures enriched with H2, the most prevalent genera was again Desulfovibrio (76.3%), 293 and Wolinella (10.2%) and Acetobacterium (7.6%) were found at lower relative 294 abundance (Fig. 5C). When this H2-enriched culture was tested for formate degradation, 295 it was readily consumed, and the genus Acetobacterium contributed up to 16.9% of the 296 total amplicons but Desulfovibrio was still the dominant population (Fig. 5D). 297 Interestingly, Desulfovibrio was also detected in previous anaerobic enrichment cultures 298 containing Dehalobacter and Candidatus Dichloromethanomonas elyuquensis capable 299 of fermenting DCM [10,11]. 300 The Wood-Ljunddahl pathway allows acetogenic bacteria to convert H2/CO2 and 301 formate via acetyl-CoA to acetate. All members of the genus Acetobacterium use H2 as 302 the electron donor for CO2 reduction to acetate, and some species can also utilize 303 formate, which is an intermediate derived from CO2 in acetate formation [22]. Due to 304 this versatility, Acetobacterium may be involved in the reactions described in Eq. 2 and 305 Eq. 3. On the other hand, the yeast extract contained in the medium is a potential source 13 306 of sulphate for the sulphate-reducing bacterium Desulfovibrio using H2 and formate as 307 electron donors. Alternatively, Desulfovibrio species have also been shown that, in the 308 absence of sulphate, they can convert CO2 and H2 to formate (CO2 + H2 → HCOO- + 309 H+) but also produce H2 from formate (HCOO- + H2O → HCO3- + H2) [23,24]. 310 Wolinella here plays a more uncertain role. It is typically described to require H2 and 311 formate as electron donor substrates to drive the reduction of various electron acceptors 312 including inorganic nitrogen compounds (i.e. nitrate, nitrite) that could be provided by 313 the yeast extract [24], and the absence of Wolinella in formate-amended cultures (Fig. 314 5B and 5D) is likely due to lower growth rates with this substrate. 315 In previous studies, DCM degradation was severely inhibited by H2 in a mixed 316 culture containing Candidatus Dichloromethanomonas elyuquensis and the presence of 317 hydrogenotrophic partners was considered mandatory to enable DCM degradation to 318 proceed [11]. Different attempts were made to study whether exogenous added H2 319 exerted an inhibition effect on DCM degradation in our mixed culture, but inhibitory H2 320 partial pressure could not be stablished because acetogenic bacteria consumed H2 321 quickly in the microcosms (data not shown). 322 323 3.4. Effect of co-contaminants on DCM degradation 324 In this study, we assessed the effects of different organochlorines as co-contaminants on 325 the activity of DCM degradation. The results obtained with the microcosms amended 326 with chloroalkanes [1,1,2-trichloroethane (1,1,2-TCA) and 1,2-dichloroethane (1,2- 327 DCA)] and chloroalkenes [TCE and cis-dichloroethylene (cis-DCE)] showed that 1,2- 328 DCA and cis-DCE did not affect the rate of DCM degradation for the range of 329 concentrations tested (100-850 µM) (Fig. 6A-B). A slight decrease on DCM 330 degradation rate was observed for TCE and 1,1,2-TCA at concentrations of 850 and ≥ 14 331 500 µM, respectively (Fig. 6C-D). Previous studies have shown that 1,2-DCA and cis- 332 DCE provoked a decrease in the dechlorination potential of Dehalococcoides [13]. In 333 agreement with our results, DCM degradation by D. formicoaceticum was not inhibited 334 by TCE at 1 mM [6]. 335 The co-contamination of groundwater with CF and DCM is a relevant scenario 336 for this study due to the existence of anaerobic bacteria belonging to the genus 337 Dehalobacter capable of transforming CF into DCM via organohalide respiration 338 [7,26]. CF is also a known strong inhibitor for many bacterial processes, such as 339 methanogenesis [27,28] and reductive dechlorination of chloroethenes [29,30] and orto- 340 chlorophenol [31]. Here, the addition of 67 and 84 µM CF did slow DCM degradation 341 but when added at 838 µM CF, no DCM degradation was observed for 90 d (Fig. 6E). 342 Egli et al. [32] pointed out that anaerobic dechlorination of chlorinated methanes is 343 frequently reported for bacteria harbouring the reductive acetyl coenzyme A (acetyl- 344 CoA) pathway that involves the coenzyme B12 as a carrier for methyl groups. This 345 pathway is utilized by D. formicoaceticum to insert CO2 into the carboxyl group of 346 acetate during DCM fermentation [9,21]. Vitamin B12 is known to stimulate the 347 reductive dechlorination of some chlorinated methanes (i.e. CF and carbon 348 tetrachloride) in microbial mixed cultures [33]. Therefore, we tested whether the 349 addition of 10 µM of vitamin B12 was effective in reactivating the DCM degradation in 350 cultures that had been exposed to 838 µM CF for 37 d, but DCM was not degraded 40 d 351 after the addition (data not shown). Likewise, DCM fermentation by D. 352 formicoaceticum and Dehalobacter-containing cultures was shown to be inhibited by 353 CF at concentrations of 1 mM and 41.8 µM, respectively [6,34]. 354 The fluorinated organic compounds perfluorooctane sulfonate (PFOS) and 355 perfluorooctanoic acid (PFOA) were amended at concentrations up to 400 and 483 µM, 15 356 respectively. The addition of 400 µM PFOS completely inhibited DCM degradation, 357 whereas PFOA at concentrations ≥ 241 µM allowed its full degradation but at 358 degradation rates lower than the controls (Fig. 6F-G). This is in contrast with the results 359 for Escherichia coli, where PFOA displayed a higher toxicity than PFOS [35]. Recently, 360 a mixture of ten polyfluoroalkyl substances (including PFOA and PFOS) was shown to 361 prevent the full reductive dechlorination of TCE in a Dehalococcoides-containing 362 culture [12]. However, the effect of individual PFOA and PFOS on Dehalococcoides sp. 363 still remains unknown. 364 We also tested the effect of the herbicide diuron and its daughter product 3,4- 365 DCA on DCM degradation. Diuron amended at 43 µM caused a marked decrease in the 366 DCM degradation rate, but when diuron was added at concentrations of 107 and 322 367 µM, DCM degradation was completely inhibited (Fig. 6H). In contrast, the addition of 368 3,4-DCA only caused a slight reduction of the DCM degradation rate at concentrations 369 above 463 µM (Fig. 6I). 370 Lastly, acetone was also tested as a co-contaminant in order to discard that the 371 described observed inhibitory effects were caused by its presence, as previous tested 372 organohalides were added in the microcosms as acetone stock solutions. Also, acetone 373 is a common groundwater contaminant, ranked 189 (out of 275) on the 2017 Priority 374 List of Hazardous Substances [17]. The concentrations of acetone tested (8-78 µM) 375 corresponded to the range added in the previous experiments. As shown in Fig. 6J, none 376 of the acetone concentrations tested had an adverse effect on DCM degradation rate. 377 3.5. Functional recovery of DCM degradation activity after inhibitors exposure 378 Dehalobacterium-containing cultures exposed for 37 d to the inhibitory CF 379 concentration of 838 µM were used as inoculum in CF-free medium and amended with 16 380 DCM. DCM degradation started after a lag phase of 4 d. DCM was depleted in all six 381 replicates 9 d later (Fig. S5A). The same procedure was repeated with inoculum 382 exposed to 838 µM CF for 90 d. In this case, microcosms exhibited an extended lag 383 phase before DCM degradation was fully consumed (Fig. S5A), probably due to the 384 decrease of viable cells caused by the longer exposition to the inhibitory concentration 385 of CF. 386 Cultures that were inhibited by 107 and 322 µM diuron for 32 d were transferred 387 in parallel to diuron-free microcosms amended with DCM. In both cases, DCM 388 degradation activity started after 7 d of lag phase, and DCM was fully degraded two 389 days later (Fig. S5B). Similarly, DCM degradation was recovered in cultures exposed to 390 400 µM PFOS for 30 d (Fig. S5C). 391 The fact that Dehalobacterium sp. are endospore-forming bacteria [21,36] could 392 explain the recovery capacities observed in this study. Endospores are known to provide 393 bacteria with a higher degree of resistance to several physical or chemical damage, such 394 as those derived from wet and dry heat, UV and gamma radiation, extreme desiccation 395 or oxidizing agents [37]. Accordingly, endospores were also observed by optical 396 microscopy in our DCM-degrading culture (data not shown). In particular, this 397 hypothesis seems a potential explanation for the recovery after PFOS inhibition, as it is 398 described to disrupt bacterial cell membranes (among other cytotoxical effects), which 399 results in cell inactivation or cell death. However, another possibility is that these 400 pollutants act as reversible enzyme inhibitors binding via weak noncovalent bonds to 401 Dehalobacterium enzymes modulating the enzyme activity by competing either directly 402 or indirectly with DCM. Further research is required to explore the nature of this 403 inhibitory responses and the recovery capacities of Dehalobacterium sp. 17 404 4. CONCLUSIONS 405 Acetate and formate have been identified as DCM degradation products using pure 406 cultures of Dehalobacterium formicoaceticum in previous studies [9]. The physiological 407 studies reported here indicated that formate oxidation is a feasible mechanism for H2 408 generation in our Dehalobacterium-containing culture, although H2 was not detected as 409 intermediate during DCM degradation, likely due to the activity of H2 scavengers (i.e. 410 acetogenic bacteria). The analysis of the bacterial community structure of cultures 411 enriched with formate and H2 revealed a strong enrichment of Desulfovibrio and 412 Acetobacterium, suggesting that both genera are involved in the transformation of the 413 daughter products of DCM degradation. The balance of acetate produced from H2 414 released during formate oxidation revealed that most of the electrons generated from H2 415 were recovered in acetate, which is in agreement with the presence of Acetobacterium in 416 the formate and H2-grown cultures. The isotopic composition of acetate produced from 13 417 C-DCM suggests that acetate can be formed directly from DCM fermentation by 418 Dehalobacterium, but also from reductive acetogenesis using CO2 and H2. These 419 observations suggest interspecies H2 transfer during DCM degradation to 420 hydrogenotrophic partners (i.e. Acetobacterium and Desulfovibrio). 421 Contaminated sites typically involve complex mixtures of contaminants, the 422 origin and fate of which are usually driven by biochemical transformations that can 423 impact microbial attenuation. The results of this study have significant implications for 424 bioremediation practices. We demonstrate that 1,1,2-TCA, 1,2-DCA, TCE, and cis- 425 DCE barely affected DCM dechlorination at concentrations up to 850 µM. Conversely, 426 complete inhibition of DCM was observed for CF, PFOS, and diuron in a dose- 427 dependent manner. However, the Dehalobacterium-containing culture recovered its 428 DCM degradation capability after the inhibition episodes had ceased, indicating that a 18 429 concentration reduction of these co-contaminants may completely detoxify DCM in 430 groundwaters. This is of special relevance for CF, because DCM can derive from the 431 organohalide respiration of CF by Dehalobacter sp, and co-contamination of both 432 compounds is usually observed in CF-impacted aquifers. The concentrations of PFOS 433 and diuron that caused complete DCM inhibition are well above to those found in 434 environmental samples, which are usually in the range of ng/L [2,38,39,40]. Therefore, 435 we conclude that PFOS and diuron could only affect DCM degradation by 436 Dehalobacterium in a worst-case scenario. Interestingly, acetate and H2 derived from 437 DCM and formate degradation in this mixed culture can serve as a carbon source and 438 electron donor, respectively, for organohalide-respiring bacteria present in the 439 contaminated site (i.e. Dehalococcoides, Dehalobacter), which would act as 440 “biostimulants” to enhance the remediation of halogenated co-contaminants. 441 ACKNOWLEDMENTS 442 This work has been supported by the Spanish Ministry of Economy and 443 Competitiveness State Research Agency (CTM2013-48545-C2-1-R and CTM2016- 444 75587-C2-1-R) co-financed by the European Union through the European Regional 445 Development Fund (ERDF). This work was partly supported by the Generalitat de 446 Catalunya (Consolidate Research Group 2017-SGR-14). 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Time course of DCM degradation in a Dehalobacterium-containing culture. 585 Symbols means µmol per bottle of DCM (■), acetate (▲), CO2 (♦), formate (▼), and 586 H2 (●). Bars indicate standard deviation for triplicate bottles. Arrows indicate addition 587 of DCM. 588 Figure 2. Time course of formate degradation and concomitant production of acetate in 589 a Dehalobacterium-containing culture. Formate was added at two different 590 concentration in parallel treatments (open symbols and solid symbols refer to the 591 treatments with 964 and 431 µmol formate, respectively). Symbols means µmol per 592 bottle of formate (inverted triangles) and acetate (triangles). Bars indicate standard 593 deviation for triplicate bottles. Arrows indicate addition of formate. 594 Figure 3. Consumption of H2 in a Dehalobacterium-containing culture. Symbols means 595 µmol per bottle of H2 (●) and acetate (▲). Production of acetate in controls without H2 596 were plotted (∆). Bars indicate standard deviation for triplicate bottles. Arrows indicate 597 addition of H2. 13 13 598 Figure 4. C-NMR spectra of the metabolic products of C-DCM in a 599 Dehalobacterium-containing culture at two different time points: immediately after the 13 13 600 addition of C-DCM (Panel A), and after C-DCM had been consumed (Panel B). 601 Peak resonances were confirmed by adding the NMR standard 4,4-dimethyl-4- 602 silapentane-1-sulfonic acid to the samples. The identity of methanol was confirmed by 603 adding 1 M of unlabeled methanol to the medium. 604 Figure 5. Relative abundance (%) of the bacterial population in the Dehalobacterium- 605 containing culture after enriched with DCM (A), formate (B), and H2 (C) based on the 26 606 16S rRNA gene amplicon sequencing. Samples enriched with H2 that were fed with 607 formate are depicted in D. The results are average of duplicate samples. The number of 608 quality filtered reads obtained from cultures grown with DCM, H2, H2+formate and 609 formate were 93316/67640, 84389/47614, 22978/79460, 64182/74340, respectively. 610 Only taxa with greater than 1% relative abundance are shown and reported to the lowest 611 taxonomic rank possible. 612 Figure 6. Effect of different concentrations of co-contaminants on the degradation rate 613 of DCM degradation in a Dehalobacterium-containing culture. Controls (C) correspond 614 to cultures that contained DCM but no co-contaminants. 1,2-DCA (1,2-dichloroethane), 615 cis-1,2-DCE (cis-1,2-dichloroethylene), TCE (trichloroethene), 1,1,2-TCA (1,1,2- 616 trichloroethane), PFOS (perfluorooctanesulfonic acid), PFOA (perfluorooctanoic acid), 617 3,4-DCA (3,4-dichloroaniline). “*” means that degradation of DCM was completely 618 inhibited. DCM was always amended at 2000 µM. Concentration in the x-axis is 619 expressed in µM of the corresponding co-contaminant. Error bars represent the standard 620 deviations of triplicate samples. 621 622 623 624 625 626 627 27 628 FIGURE 1 629 630 631 632 633 634 635 636 637 28 638 FIGURE 2 639 640 29 641 FIGURE 3 642 643 644 30 645 FIGURE 4 646 A B 647 648 649 650 651 652 653 654 31 655 FIGURE 5 2.0 3.7 A B 8.4 Desulfovibrio Dehalobacterium 12.4 Acetobacterium 26.7 Desulfovibrio Wolinella Others 53.1 Acetobacterium Others 24.1 69.6 C Desulfovibrio 10.2 5.9 D Acetobacterium 6.7 Wolinella Desulfovibrio 7.6 Acetobacterium Others 16.9 Others 76.3 76.4 656 657 658 659 660 661 662 663 664 665 666 32 667 FIGURE 6 1,2-DCA cis-1,2-DCE cis-1,2,-DCE TCE 1200 1200 1200 DCM degradation rate (µM·d-1) DCM degradation rate (µM·d-1) DCM degradation rate (µM·d-1) A 1000 B 1000 C 1000 800 800 800 600 600 600 400 400 400 200 200 200 0 0 0 C 100 250 500 850 C 100 250 500 850 C 100 500 850 1,1,2-TCA Chloroform PFOS 1200 600 1000 DCM degradation rate (µM·d-1) DCM degradation rate (µM·d-1) DCM degradation rate (µM·d-1) D E F 1000 500 800 800 400 600 600 300 400 400 200 200 200 100 0 0 * 0 * C 100 250 500 850 C 17 67 84 838 C 20 100 400 PFOA Diuron 3,4-DCA 1600 1000 1000 DCM degradation rate (µM·d-1) DCM degradation rate (µM·d-1) DCM degradation rate (µM·d-1) 1400 G H I 1200 800 800 1000 600 600 800 600 400 400 400 200 200 200 0 0 * * 0 C 121 241 483 C 43 107 322 C 62 154 463 Acetone 1200 DCM degradation rate (µM·d-1) J 1000 800 600 400 200 0 C 8 20 32 78 668 33