Benzene Degradation by Variovorax sp within a Coal-tar Contaminated Groundwater Microbial Community

24 Investigations of environmental microbial communities are crucial for the discovery of 25 populations capable of degrading hazardous compounds and may lead to improved 26 bioremediation strategies. The goal of this study was to identify microorganisms responsible for 27 aerobic benzene degradation in coal-tar contaminated groundwater. Benzene degradation was 28 monitored in laboratory incubations of well waters using gas chromatography mass spectrometry 29 (GC/MS). Stable isotope probing (SIP) experiments using C-benzene allowed us to obtain C30 labled community DNA; from this, 16S rRNA clone libraries identified γand β-Proteobacteria 31 as the active benzene metabolizing microbial populations. Subsequent cultivation experiments 32 yielded nine bacterial isolates that were capable of growing in the presence of benzene; five were 33 confirmed in laboratory culture to grow on benzene. The isolated benzene-degrading organisms 34 were genotypically similar (>97% 16S rRNA gene nucleotide identity) to the organisms 35 identified in SIP experiments. One isolate, Variovorax MAK3, was further investigated for 36 expression of a putative aromatic ring-hydroxylating dioxygenase (RHD) hypothesized to be 37 involved in benzene degradation. Microcosm experiments using Variovorax MAK3 revealed a 38 10-fold increase in RHD (Vapar_5383) expression--establishing this gene’s link to benzene 39 degradation. Furthermore, addition of Variovorax MAK3 to microcosms prepared from site 40 waters accelerated community benzene degradation and showed a corresponding increase in 41 RHD gene expression. In microcosms using uninoculated groundwater, qPCR assays (16S rRNA 42 and RDH genes) showed Variovorax to be present and responsive to added benzene. These data 43 demonstrate how the convergence of cultivation-dependent and cultivation-independent 44 techniques can boost understanding of active populations and functional genes in complex 45 benzene-degrading microbial communities. 46 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom IMPORTANCE: Benzene is a human carcinogen whose presence in contaminated groundwater 47 drives environmental cleanup efforts. Although aerobic biodegradation of benzene has long been 48 established, knowing the identity of microorganisms in complex naturally-occurring microbial 49 communities that are responsible for benzene biodegradation has evaded scientific inquiry for 50 many decades. Here we apply a molecular biology technique known as stable isotopic probing 51 (SIP) to the microbial community residing in contaminated groundwater samples to discover the 52 identity of community members active in benzene biodegradation. We complemented this 53 approach by isolating and growing in the laboratory a bacterium representative of the bacteria 54 found using SIP. Further characterization of the isolated bacterium allowed us to track 55 expression of a key gene that attacks benzene both in pure cultures of the bacterium and in the 56 naturally-occurring groundwater microbial community. This work advances information about 57 how to document microbial processes, especially populations and genes, that contribute to 58 bioremediation. 59 60 INTRODUCTION 61 Microbial communities play a globally significant role metabolizing a diverse array of 62 compounds in Earth’s ecosystems (1, 2). Achieving a more robust understanding of key 63 microbial players and their metabolic processes is constrained by the complexity of microbial 64 communities and by methodological limitations in environmental microbiology, particularly with 65 traditional cultivation-dependent approaches (3, 4). Regarding the carbon cycle, knowing the 66 identity of microbial population responsible for metabolizing specific compounds has major 67 implications for bioremediation and predicting ecosystem responses to ongoing anthropogenic 68 release of pollutants such as hydrocarbons to terrestrial and aquatic environments (5). Despite the 69 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom importance of microbial communities in contaminant degradation, much remains to be explored 70 about the vast taxonomic and functional genetic diversity of bioremediation agents (4, 6). 71 Aerobic hydrocarbon degradation is one of the most well described microbially-mediated 72 processes in the environment (7–9). Numerous studies have applied assays for molecular 73 biomarker approaches to contaminated sites, with the objective of exploring the diversity of 74 biodegradation genes and the naturally-occurring populations involved in metabolizing 75 hydrocarbons (10–16). In addition, cultivation efforts have successfully isolated hydrocarbon76 degrading organisms from habitats including soil, ground water, marine waters, and deep sea 77 sediments (12, 17–22). Yet despite many studies successfully reporting new information about 78 the genetic basis (e.g., dioxygenase) of hydrocarbon metabolism, much remains to be learned 79 about both the identity of populations responsible for biodegradation and the diversity of 80 biodegradation genes (4, 10, 14, 16). 81 Linking the responsible agents to specific metabolic processes in the environment 82 remains difficult, even in well described processes of aerobic aromatic hydrocarbon degradation 83 (4, 8, 23, 24). Stable Isotope probing (SIP) is a promising tool to target and identify active 84 populations within microbial communities without the need for cultivation (25–28). SIP studies 85 dose substrates labeled with heavy isotopes (i.e. C, N) into mixed microbial populations. 86 Then assimilative metabolic processes incorporate the heavy atoms into the metabolically active 87 biomass (typically a small subset of the total). The result is an enrichment of heavy atoms in 88 biomarker molecules [i.e. phospholipid fatty acids (PLFAs), DNA or RNA) that can be extracted 89 and separated from the light (non-labeled) biomass. Biomarkers are then analyzed to identify the 90 organisms responsible for substrate metabolism (29–31). Previous, SIP investigations of the 91 metabolism of BTEX compounds have largely focused on anaerobic benzene and toluene 92 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom degradation, using PLFA and rRNA-based finger printing techniques to identity the active 93 populations (32–35). DNA-, RNA-based benzene SIP investigations have been conducted in 94 coal-tar contaminated sediment, gasoline-contaminated water, and soil (31, 36, 37). However, 95 the high diversity of both habitat types and geochemical conditions where contaminants occur 96 warrant additional SIP investigations. 97 Here we present a systematic approach using SIP, cultivation, and molecular techniques 98 to understand the microbial populations responsible for degrading benzene in a naturally99 occurring groundwater microbial community. The contaminated site is one with a history of 100 coal-tar contamination (38–40), that features benzene within a complex mixture of organic 101 compounds. We used SIP-microcosm incubations to identity the bacterial community members 102 responsible for benzene degradation in site waters. The SIP investigation was followed by 103 cultivating representatives of the community and verifying their ability to degrade benzene. We 104 also used qPCR to establish expression of a putative dioxygenase during benzene degradation in 105 a Variovorax sp. isolate that showed high 16S rRNA similarities to the populations identified in 106 the SIP study. Finally, we found enhanced signals for Variovorax and its dioxygenase catabolic 107 gene in site waters dosed with benzene. This study demonstrates how the combination of 108 cultivation-independent and -dependent methodologies can be used to gain insight into the 109 populations and their catabolic genes involved in contaminant biodegradation. 110 111 MATERIALS and METHODS 112 Site description and sample collection. The study site is a coal-tar contaminated aquifer located 113 in South Glens Falls, New York, the site’s microbiology and geochemistry is well documented 114 (13, 41, 42). Well water samples were taken from two monitoring wells (Well 12 and Well 36) as 115 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom described in Bakermans and Madsen 2002 (42). Previously collected site sediment was stored at 116 4°C. Sterile sediment was prepared by air drying site samples, dispensing to 50-ml screw117 cap glass tubes, and applying gamma-irradiation (2.5 Mrad from the 60Co source in Ward 118 Nuclear Reactor, Cornell University); sterility was verified by the absence of colonies after 119 plating a 10-1 dilution on LB media and incubating at room temperature for 10 days. 120 Benzene biodegradation assays. Laboratory biodegradation assays for groundwater were set up 121 within 24 hours of sample collection. Eighty ml of groundwater was dispensed to 160-ml serum 122 bottles and sealed with teflon-faced butylrubber septa. Four treatments were established for 123 bottles that received both unlabeled (C) and labeled (C) benzene: (i) site water only 124 (unamended); (ii) amended with 10% v/v sterile-sediment; (iii) amended with 10 mM (NH4)3PO4 125 ; and (iv) an abiotic control that received 10N HCl to achieve a pH <2. Liquid benzene was added 126 (neat) through to septa (or immediately prior to sealing the serum bottles) with a 2μl syringe to 127 final concentrations ranging from 1.2 to 20 ppm. Bottles were incubated at 10 °C (ambient for 128 the study site) in the dark without shaking. At various times over a 360-hour period, headspace 129 gases were sampled using a 250μl gastight syringe (Hamilton) and analyzed for benzene by 130 GC/MS, as described below. Benzene metabolism by isolated bacterial cultures was monitored in 131 serum bottles prepared as above, but incubated at 21 °C and shaken at 120 rpm. 132 133 Stable isotope probing. After ~70% of the benzene was consumed in the above microcosms, 134 both Cand Cbenzene treatments were sacrificed for nucleic acid extraction. The DNA was 135 extracted using Fast DNASPIN kit as previously described in DeRito et al. 2005 (33). The 136 triplicate samples from each treatment were pooled and processed by CsCl ultracentrifugation to 137 create an isotopic DNA gradient, separating the C and C-DNA (30). The location of labeled 138 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom (heavy) DNA in the gradient was determined by including a tube containing C-DNA (from 139 Pseudomonas putida grown on C-glucose) in the ultracentrifuge rotor. After pulling this heavy 140 fraction from both the Cand Cbenzene treatments, dilutions were prepared, and 16S PCR 141 reactions as per DeRito et al. 2005 (30) were performed. The pool of 16S rRNA PCR amplicons 142 representing the active benzene-degrading populations was operationally defined as the dilution 143 of DNA from the C treatment that successfully amplified, while the corresponding dilution 144 from the C treatment failed to amplify (30). This C-benzene-derived set of amplicons was 145 cloned and sent for sequencing (Applied Biosystems model 3700) at the Cornell University 146 Biotechnological Facility (30). 147 Cultivation of isolates on solid media. A site-specific medium consisting of filter sterilized (0.2 148 micron, Corning) well water and noble agar (Difco, BD) was prepared. At the conclusion of the 149 Well 36 incubation, two separate dilution series (10, 10, 10 fold) were spread plated and 150 incubated at 10°C in the presence and absence of benzene vapor in the headspace. After 15 days 151 of observation, small translucent colonies appeared on both the control and benzene-exposed 152 plates. At day 22, sterile colony transfer pads (RepliPlate, FMC) were used to replica plate the 153 cultures onto Stanier’s Basal Salt Minimal Media (BSM) agar plates as per Burlage 1998 (43). 154 After 18 more days of incubation in benzene vapor the minimal media plates yielded colonies ~2 155 mm in diameter. Nine colonies with distinct morphologies that grew only in the presence of 156 benzene were selected and streaked for single isolation. Isolates were restreaked and grown three 157 successive times in the presence of benzene and were picked and grown in R2A medium. A 158 portion of the R2A culture was preserved by adding 50% glycerol and freezing at -80°C. Isolates 159 were revived and streaked on BSM media and displayed growth only in the presence of benzene. 160 161 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom Dioxygenase expression in minimal media. Isolate MAK3 was grown in BSM containing 0.5 162 g/l glucose, washed 3x in BSM and added to 80 mL BSM liquid media in 1-l serum bottles for 163 final concentration of approximately 1 x 10 cfu/ml. The benzene (7 ppm) degradation 164 experiment included three treatments: (i) Variovorax strain MAK3 only (ii) Variovorax strain 165 MAK3 cells killed with 5M hydrochloric acid (iii) without inoculum. Cultures were incubated at 166 21°C, not shaken. Concurrently, a glucose (0.5 g/l) control treatment for quantification of 167 dioxygenase gene expression was prepared in BSM. Subsamples of each culture were taken 168 periodically (at 0, 21, 27, 36 hours) and immediately frozen in liquid N2 for RNA extraction. 169 170 Dioxygenase expression in nutrient-amended well water. In the final degradation experiment 171 80 mL of Well 36 water amended with filter-sterilized (NH4)3PO4 (final concentration of 10 172 mM) was added to serum bottles. Variovorax Strain MAK3 cells, grown as above were added at 173 an initial concentration 2 x 10 cfu/mL. Liquid benzene was added to each treatment at 1.2 ppm. 174 Cultures were incubated at 21°C and not shaken. Subsamples of the Variovorax amended and 175 unamended treatments were taken at 3 time points corresponding with 15, 75 and 100 percent 176 benzene loss and immediately frozen in liquid N2 for RNA extraction. 177 178 Phylogenetic analysis of benzene degrading isolates. Nine isolates were identified using 16S 179 rRNA gene sequencing. 16S rRNA PCR was performed on individual colonies using 27F/1492R 180 primers (Table S1). The 25 μL 16S rRNA reactions included: 5μl 5x MyTaq buffer (15 mM 181 MgCl2, 5mM dNTPs, (Bioline), 0.5 μL 27F/1492R primers (20μM), 0.1 μL MyTaq DNA 182 Polymerase, (5U/μL, Bioline) in thermocycler (MJ Research PTC-200) conditions: initial 183 denaturation [95°C, 5min], 32 cycles of denaturation [94°C, 1 min], annealing [55°C, 1:30 min] 184 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom and extension [72°C, 1:30 min], and a final extension [72°C, 10 min]. The 16S rRNA PCR 185 amplicons were gel purified using QIAquick gel extraction kit (Qiagen), and ligated into a 186 pCR2.1 plasmid vector (TOPO TA Cloning, Invitrogen) as per manufacturer’s protocol. Plasmid 187 were transformed into chemically component E. coli (One Shot TOP10, Invitrogen) cells and 188 grown for blue/white screening. White colonies were picked and verified by PCR with M13F/R 189 primers (Invitrogen). Clones containing the insert were grown over night in Luria-broth with 190 kanamycin (50 μg/μL) and the plasmids were extracted for sequencing using Zyppy Plasmid 191 Miniprep (Zymo). Sequencing was performed by Cornell University Life Sciences Core 192 Laboratories Center, with an Applied Biosystems Automated 3730 DNA Analyzer using Big 193 Dye Terminator chemistry and AmpliTaq-FS DNA Polymerase. Consensus 16S rRNA 194 sequences were built from 4 independently sequenced reactions using primer sets: M13F/M13R, 195 27F/1492R, 530F/519R, 1114F/1100R using ebiox (Version 1.5.1) (Table S1). The isolate 196 sequences compared with GenBank nucleotide database library with BLASTn and Ribosomal 197 Database Project for taxonomic identification. Phylogenetic trees were constructed using MEGA 198 5.05 (44). The isolates, SIP-clones and reference sequences were aligned with ClustalW and 199 assembled using a neighbor-joining algorithm, with 1,000 bootstrap replications. 200 201 Variovorax RHD primer design. Primers were designed to target the aromatic-ring202 hydroxylating dioxygenase beta subunit (Vapar_5383) of Variovorax paradoxus S110 203 (NC_012792.1) using NCBI Primer-BLAST tool. The primers VarRHDF/VarRHDR amplify an 204 182bp region of the gene (Table S1). 25 μL PCR VarRHD reaction conditions included: 0.1μL 205 Taq (5 U/μL) DNA Polymerase (Bioline), 5μL 5X MyTaq Buffer (15 mM MgCl2, 5mM dNTPs, 206 Bioline), 0.5μL 20μM VarioRHD F/R, 1 μL DNA Template. Thermocycler conditions: Initial 207 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom denaturation [95°C, 10 min] then 40 cycles of denaturation [95°C, 1 min], annealing [60°C, 1 208 min] and extension [72°C, 30 sec], and a final extension [72°C, 10 min]. To validate the 209 specificity of the primers the amplicon was gel purified with QIAquick gel extraction kit 210 (Qiagen) cloned, sequenced and compared to the NCBI reference using Blastn. 211 212 Cell extraction, RNA, RT-PCR and qPCR. Cells from the degradation experiments were 213 subsampled from the microcosm experiments concentrated by centrifugation (7000g), 214 immediately frozen in liquid nitrogen and stored at -80C until processing. RNA extractions were 215 performed with Quick-RNA kits (Zymo). The extraction buffer was applied directly to the frozen 216 pellet and the procedure was followed as per manufacturer’s protocol. Total nucleic acid 217 extractions were treated with Dnase (Invitrogen) and converted to cDNA using SuperScript III 218 (Invitrogen) First Strand for reverse transcription. The cDNA was quantified using qPCR with 219 dioxygenase-specific primers VarioRHD F/R and Variovorax-specific 16S rRNA primers 220 VarF/R (45) (Table S1). Quantitative PCR was performed on an Applied Biosystems 7300 Real 221 Time PCR System. Standards were made by serial dilution of gel purified VarF/R and 222 VarioRHDF/R PCR amplicons and comparing them with known quantities of lambda DNA 223 using Quant-iT (PicoGreen dsDNA reagent, Invitrogen P7581). The 16S rRNA and RHD 224 reactions were run under PCR conditions including 12.5 μL Master Mix (SYBR Select, Applied 225 Biosystems), 0.3 μL RHD/16S (20uM) 240nM, 0.3 μL VarR (20uM) 240nM, 10.9 H2O, 1.0 μL 226 template cDNA. Quantitative PCR thermocycler conditions included 2 min at 50°C, 15min at 227 95°C, 40 Cycles (15 sec at 95°C, 30 sec at 58°C), and a disassociation curve was included to 228 assess amplification specificity. 229 230 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom Gas chromatography/mass spectrometry analysis of benzene. A Hewlett-Packard HP6890 231 gas chromatograph (Wilmington, DE) linked with a HP 5973 mass selective detector was used to 232 quantify benzene. The gas chromatograph (GC) was fitted with a Hewlett-Packard HP-5 233 phenylmethyl-siloxane capillary column (Model no. HP 19091J-433, Capillary 30.0m x 250 um 234 x 0.25 um), carrying high-purity helium gas supplied by Airgas (Elmira, NY). The GC 235 parameters included, a split injection (20:1), inlet temperature at 150°C, initial GC oven temp 236 was at 50°C and ramped 15°C /min to a final temperature of 100°C. The mass spectrometer 237 detector was operating at 2000eV and a vacuum of 2 x 10 tor, in scan mode from m/z 50 to 500 238 (benzene m/z = 78). Benzene was measured by taking 100 μL headspace samples with a 250μL 239 gas tight syringe (Hamilton, NV). Calibration curves were constructed using external standards 240 with known amounts of benzene (EMD, Germany). Benzene concentrations were averaged and 241 compared by standard deviation from triplicate chambers. 242 243 Accession number(s). The sequences from this study have been deposited in GenBank under 244 accession numbers KX665551-KX665559 and KX670396-KX670409 for isolates and clones 245 respectively. 246 247 RESULTS 248 Degradation of benzene by well-water communities 249 Microcosm incubations were conducted to demonstrate the ability of microbial 250 communities in Wells 12 and 36 to degrade benzene. Water samples from both wells were dosed 251 with 3 ppm benzene and treated four ways: (i) site water only (unamended), (ii) water amended 252 with 10% v/v sterile-sediment, (iii) water amended with 10 mM (NH4)3 PO4; and (iv) an abiotic 253 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom control poisoned with HCl. Benzene persisted in the abiotic treatments for Wells 36 and 12 254 (Figure 1 Panels A and B, respectively) during the >300-h incubation. The community in the 255 sedimentand nutrient-amended treatments from Well 36 degraded the benzene completely in 256 300 and 360 hours, respectively (Figure 1A). Similarly, the community from Well 12 degraded 257 benzene in 200 and 250 hours for nutrient and sediment amended microcosms, respectively 258 (Figure 1B). Surprisingly, benzene persisted in the well water unamended with nutrients or 259 sterile sediment for >350 h. The lack of benzene biodegradation by microorganisms in the 260 unamended incubations was likely a result of nutrient limitation. Clearly benzene biodegradation 261 was equivalent in the nutrientand sediment-amended incubations. We conclude that nitrogen 262 and phosphorus sources associated with the sterile aquifer solids were capable of supporting 263 benzene metabolism by the native community. These results demonstrate that the microbial 264 communities in site waters were capable of degrading benzene, but that nitrogen and/or 265 phosphorus limitation can occur if site waters are incubated in the absence of site aquifer solids. 266 267 Stable isotope probing to identify active benzene-degrading populations 268 To identify the members of the aquifer microbial community that were active in benzene 269 metabolism, a stable isotope probing (SIP) experiment was performed in parallel with the 270 degradation assays. Treatments were prepared identically as in the above degradation assay 271 except C-labelled liquid benzene was added to a set of microcosms. After 70% of the C272 labelled benzene was degraded the microcosms were sacrificed. Parallel, (unlabeled) C273 benzene treatments were also processed. After DNA extraction, CsCl ultracentrifugation allowed 274 the heavy (C) fraction to be collected from both Cand C-treatments (30). The location of 275 the heavy DNA in the CsCl gradient corresponded to the band where DNA from C-grown 276 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom control cells appeared. Dilutions were prepared. The first dilution that failed to yield 16S rRNA 277 gene amplicons in the C treatment, but continued to yield amplicons in the C treatments was 278 operationally defined as enriched in benzene-derived C DNA (30). 16S rRNA genes in the 279 DNA fraction were amplified and cloned using universal bacterial 16S rRNA primers. From a 280 library of 156 clones, fourteen displaying unique RFLP profiles were chosen for sequencing, ten 281 from Well 36 and four from Well 12. A phylogenetic analysis of the fourteen SIP-generated 282 (Figure 2) sequences showed that γand β-Proteobacteria, specifically the genera 283 Herminiimonas, Acidovorax, Variovorax and Pseudomonas, represented the predominant active 284 benzene degrading microorganisms. Results from the clone library revealed that Well 36 was 285 highly enriched with two sequences of Variovorax which together comprised 21% of the clone 286 library and one Acidovorax sequence represented another 25% of the library. 287 288 16S rRNA gene sequencing of benzene-degrading isolates 289 Incubations were established to isolate benzene-degrading organisms from the well water 290 degradation experiment. Nine isolates exhibiting distinct morphologies were successfully 291 cultivated and yielded robust colonies in the presence but not in the absence of benzene vapor 292 (Figure S1). The nine isolates were then transferred to BSM broth at room temperature with 293 benzene as the sole carbon source. Five of the nine isolates successfully degraded benzene in 294 liquid culture over the incubation period of 70 hours (Figure S2). The remaining four isolates, 295 including the control, did not show significant loss of benzene over the same period. We 296 speculate that the four isolates may not have degraded benzene in liquid culture because of the 297 toxicity of dissolved benzene at 16 ppm or slow growth rates. Alternatively, they may have been 298 able to grow on agar plates (used during isolation) because of carbon scavenging from the agar. 299 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom The 16S rRNA gene of the isolates were sequenced and compared with those derived 300 from the C-labelled DNA fraction of the SIP experiment (Figure 2). The results indicated that 301 several of the isolates shared >97% identity to SIP clones. Most notably isolate MAK5 had 97% 302 identity with a SIP clone (36B1 clone3), which comprised 25% of clone library and MAK3, 303 shared 98% with two SIP sequences (36B1 clone88 and 36B1 clone89) that comprised 21% of 304 the clone library. The largest group of sequenced isolates (MAK 1, 2, 4, 10) clustered with genus 305 Rhodococcus, which was not identified in the SIP experiment. The reason for the absence of 306 Rhocococci from the SIP library is uncertain; it may have been due to the limited size of the 307 clone library or to resistance of Gram-positive Rhodococci to cell lysis. 308 309 Dioxygenase gene expression by strain MAK3 during benzene degradation 310 Isolate MAK3 was chosen for further investigation because of its high sequence similarity and 311 representation in the SIP clone library. qPCR primers were designed based on a putative 312 dioxygenase gene annotated in the Variovorax paradoxus S110 genome. The specificity of the 313 primers was verified using conventional PCR on isolate MAK3 and by sequencing the 182 bp 314 amplicon (data not shown). The results confirmed 96% identity to the annotated dioxygenase 315 gene in Variovorax paradoxus S110. 316 We hypothesized that Variovorax MAK3’s ring hydroxylating dioxygenase (RHD) gene 317 would be upregulated (highly expressed) by the culture during benzene degradation. To test this 318 we prepared microcosm incubations in the presence and absence of benzene and two negative319 controls: one with no added carbon and the other with glucose as the growth substrate. Over the 320 course of the experiment (Figure 3), biomass was subsampled at times corresponding with 321 measurements of benzene concentration. qRT-PCR quantification of the putative dioxygenase 322 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom gene transcripts showed that Variovorax MAK3 cells were most active between 21 and 27 hours 323 (14% to 57% benzene depletion); this amounts to a ten-fold increase in RHD transcript 324 abundance (Figure 3). The elevated transcript levels persisted to the final time point when 98% 325 of the benzene had been consumed. Absolute transcript copies of the MAK3 RHD gene 326 increased from approximately 1.0 x 10 copies/ng at 14% benzene depletion to 1.3 x 10 327 copies/ng at 57% benzene depletion and 1.0 x 10 copies/ng at 98% depletion. Over the same 328 time periods the RHD transcript levels in the glucose did not increase and the levels were similar 329 to those in the no-benzene control. 330 Additionally, Variovorax-specific 16S rRNA primers (45) were used in qPCR assays to 331 monitor the abundance of Variovorax cells during the experiment. The results indicate that 16S 332 rRNA copies increased 150% in the benzene treatment, while the treatment without benzene 333 decreased by 62% during the experiment. The glucose treatment demonstrated the largest 334 increase, 16S rRNA copies rose 410% (Figure S4). 335 The above results show that an increase in the dioxygenase transcript abundance 336 corresponded to the rapid degradation of benzene and an increase in 16S rRNA copy number, 337 associated with cell growth. The comparison of the dioxygenase results between the benzene and 338 glucose treatments strongly suggest that the putative dioxygenase gene is specifically involved in 339 the catabolism of benzene and not in general growth or metabolism. Furthermore, the difference 340 in 16S rRNA gene abundance found in the presence versus the absence of benzene confirms that 341 Variovorax MAK3 can derive its carbon and energy solely from benzene. 342 343 Dioxygenase gene expression by the native populations during benzene degradation 344 345 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom The final series of experiments were performed to test the hypothesis that the 346 Variovorax-related, naturally-occurring populations are active in the degradation of benzene in 347 the microbial community native to the site’s groundwater. The 16S rRNA and RHD gene qPCR 348 assays described above were applied to nitrogenand phosphorousamended Well 36 349 incubations with and without added Variovorax MAK3 cells (with and without added benzene). 350 Samples were sacrificed for biomass extraction at times in the experiment when approximately 351 0%, 25% and 90% of the initial benzene concentration was consumed. The addition of 352 Variovorax MAK3 accelerated benzene degradation substantially (3-fold) compared to the native 353 community alone (Figure 4A). The inoculated treatment consumed ~100% of the benzene in 90 354 hours, while the native community required 300 h to degrade the benzene. In the inoculated 355 treatment, absolute transcript abundance for the MAK3 RHD gene increased over time 356 corresponding well with the accelerated loss of benzene 357 during the first 100 hours of the experiment (Figure 4B); RHD transcripts were not detected 358 initially but increased to 246 copies/ng when 25% of benzene had been consumed and 362 359 copies/ng when 90% had been consumed. These patterns in RDH gene expression are consistent 360 with pure culture experiment (Figure 3) and indicate that benzene loss is mechanistically tied to 361 RDH gene activity. We hypothesized that Variovorax populations related to strain MAK3 were 362 present and active in the uninoculated well-water microcosms shown in Figures 4A and 4B. 363 qPCR assays of Variovorax-specific 16S rRNA and RHD transcripts showed that at 200 hours 364 (when the native community was most active; Figure 4B), added benzene caused a 6-fold 365 enhancement in the ratio of expressed RDH genes to 16S rRNA copies (Figure 4C). Clearly, 366 Variovorax MAK3 and related well-water populations are adapted to degrading benzene in site 367 waters. 368 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom 369 Discussion 370 Our aim was to use groundwater microcosms as a model system to investigate benzene 371 degradation by microbial communities in coal-tar contaminated groundwater. Although benzene 372 is a minor constituent of coal-tar, the continued presence of other BTEX compounds on site and 373 the ubiquity of benzene as an environmental pollutant makes it an important compound to study 374 (46). In these experiments, we attempted to create relevant naturally occurring field conditions in 375 our incubations; we fully acknowledge, however, that the behavior and composition of microbial 376 communities in our laboratory model incubations are not necessarily indicative of the function of 377 microbial communities in situ (3). 378 We demonstrated in samples from Well 12 and Well 36,that separating the groundwater 379 from the sediment matrix altered the biodegradation potential of the community (Figure 1). A 380 nutrient amendment was required to support benzene biodegradation activity. Previous 381 investigations have demonstrated that nutrient additions do not necessarily lead to drastic 382 alteration of community function (47, 48). Moreover, the sequences recovered from the SIP 383 experiments represented γ-and β-Proteobacteria, which is a finding consistent with field 384 investigations of benzene-degrading communities (12). 385 Convergent lines of cultivation-dependent and cultivation-independent inquiry confirmed 386 that the members of γ-and β-proteobacteria within the groundwater community were responsible 387 for benzene biodegradation in the microcosms. We coupled SIP with cultivation to directly 388 address potential weaknesses in SIP methodological procedures (4, 49). The isolation and pure389 culture degradation experiments confirmed that the organisms native to the groundwater, 390 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom specifically Variovorax MAK3 and Acidovorax MAK5 were capable of benzene degradation 391 (Figure S2 and S3). 392 Cultivation approaches that supply BTEX compounds in vapor phase have been used in 393 prior investigations (12, 22). Hendrickx et al (2006) reported the cultivation of Acidovorax and 394 Variovorax spp. from soil samples using BTEX mixtures. The results of SIP and cultivation 395 experiments from this study reinforce a growing body of evidence that Acidovorax and 396 Variovorax are active benzene degrading organisms in many BTEX contaminated sites (50–52). 397 Variovorax spp. in particular have been associated with soil habitats, and characterized as a 398 metabolically diverse group of organisms with promising bioremediation and biotechnological 399 applications (53–55). 400 To our knowledge this is the first investigation demonstrating benzene degradation by 401 Variovorax spp. in pure culture, while simultaneously quantifying expression of the oxygenase 402 gene involved in benzene attack (Fig 3). The RT qPCR data that suggested the putative 403 dioxygenase (Vapar_5383) plays a role in benzene biodegradation by Variovorax MAK3. 404 However, correlation does not prove the putative RHD is responsible for benzene degradation 405 and quantitative PCR has inherent limitations particularly concerning the primer specificity and 406 SYBR green assays. To improve the confidence that the primers were amplifying the target gene 407 the amplicon was sequenced and the qPCR reaction was optimized to achieve a single peak in 408 the disassociation curve. The transcript abundances from the pure-culture experiment were 409 consistent with the findings of Kong and Nakastu (56), who investigated RNA-extraction 410 methods for quantification of aromatic oxygenase genes (56). We feel it is reasonable to presume 411 that putative dioxygenase (homologous to Vapar_5383) enables benzene attack by Variovorax 412 on N ovem er 9, 2017 by gest ht://aem .sm .rg/ D ow nladed fom MAK3, though acknowledging that only gene knockout experiments would provide definitive 413