Sludge and sediment samples collected from bioreactors and black-odorous urban rivers were employed as inoculum for culture setup (Table 1). These samples were acquired directly by filling sterile 50 ml plastic Falcon tubes that were capped and transported to the laboratory at an ambient temperature. To control exposure of the samples to oxygen, Falcon tubes were sealed with Parafilm, and microcosm setup was performed in anaerobic chamber soon after their arrivals. For granular sludge, it was smashed into floc-form sludge before inoculation. Defined anaerobic mineral medium in 160 ml serum bottles for microbial cultivation was prepared as described (He et al., 2003; Wang and He, 2013a), which contains salts, trace elements and vitamins. L-cysteine and Na₂S⋅9H₂O (0.2 mM each) were added to the medium to achieved reduced conditions. The bottles were sealed with black butyl rubber septa and secured with aluminum crimp caps. The organohalide-fed cultures were transferred in 100 ml medium supplemented with 10 mM lactate, 10 mM 2-bromoethanesulphonate (BES, to inhibit methanogen growth), and 1 mM PCE or 10 ppm chloromethane. The control cultures without organohalide-amendment were transferred in the same mineral medium. Unless stated otherwise, cultures were incubated at 30°C in the dark without shaking. All the experiments were set up in duplicates.

Headspace samples of chloroethenes (i.e., PCE, TCE, cis-DCE, trans-DCE, VC and ethane)and chloromethane were injected manually with a glass, gastight, luer lock syringe (Hamilton, Reno, NV, United States) into a gas chromatography (GC) 7890N equipped with a flame ionization detector (Agilent, Wilmington, DE, United States) and a GS-GasPro column (30 m × 0.32 mm; Agilent, Wilmington, DE, United States) as described (Wang and He, 2013b). The standards compounds (with analytical pure or above) were purchased from Sigma–Aldrich.

The FISH experiment was performed according to protocols described previously (Amann et al., 1995). Granular sludge samples were fixed in a 4% paraformaldehyde solution for 8 h at 4°C, and embedded in Optimal Cutting Temperature (O.C.T.) compound (Fisher Healthcare, Houston, TX, United States). Then the freezing granules were cut into 15 μm-thick sections with CM3050S cryostat (Leica, Germany). Hybridization was performed at 46°C for 4 h with oligonucleotide probes Dhe1259 (Yang and Zeyer, 2003), EUBmix and ARCH915 (Amann et al., 1995) targeting Dehalococcoides, bacteria and archaea, respectively. Dhe1259 and EUBmix/ARCH915 for dual-staining FISH were labeled with Cyanine 3 (Cy3) and Cy5, respectively. FISH-stained images were captured CLSM (Leica TCS-SP2, Germany).

Community gDNA was extracted using the FastDNA Spin Kit for Soil (MP Biomedicals, Carlsbad, CA, United States) according to the manufacturer’s instructions. The 16S rRNA gene was amplified with the U515F forward primer and U909R reverse primer as described (Narihiro et al., 2015). Illumina Miseq sequencing (Illumina, San Diego, CA, United States) service was provided by BGI (Shenzhen, China). The provided pair-end (2 × 300 nd) demultiplexed sequences were assembled and filtered using Mothur v.1.33 (Schloss et al., 2009). Quantitative Insights Into Microbial Ecology (QIIME, v1.8.0) was employed for the subsequent processing and downstream analysis (Caporaso et al., 2010).

Raw Illumina Miseq sequencing reads were deposited into NCBI Sequence Read Archive (SRA) with accession no. SRP112682.

Dehalococcoides as an obligate dehalogenating bacterial group can only utilize organohalides as electron acceptors to support their growth (Löffler et al., 2013). In this study, sediment and sludge samples from black-odorous urban rivers and anaerobic bioreactors, respectively, were selected to screen the presence of Dehalococcoides (Table 1). PCR amplification with Dehalococcoides genus-specific primers, FpDHC1/RpDHC1377 (Hendrickson et al., 2002), showed the positive detection of Dehalococcoides in all urban river sediment samples, as well as in a granular sludge sample collected from a full-scale mesophilic UASB reactor treating petrochemical wastewater (Table 1). And the petrochemical wastewater contains organic compounds generated from terephthalic-acid industry, e.g., terephthalic-acid, benzoic acid, toluic acid, acetic acid and other intermediate compounds and byproducts (Lykidis et al., 2011).

To profile microbial communities of those Dehalococcoides-containing environmental samples, Miseq 16S rRNA gene sequencing was performed, showed the very different microbial community structure in samples between Dehalococcoides-containing granular sludge and urban river sediments (Figure 1). In granular sludge collected from the UASB reactor, acidogenic populations, Syntrophorhabdus (of Syntrophorhabdaceae) and Syntrophus, formed syntrophic interactions with methanogenic Methanosaeta and Methanosarcinaceae (Figure 1). Surprisingly, the obligate organohalide-respiring Dehalococcoides presented abundant in the full-scale UASB reactor, accounting for 1.83% of the total microbial community, comparable with the relative abundance of Dehalococcoides in enrichment cultures dechlorinating PCBs (Wang and He, 2013a) and PCE (Lee et al., 2015). The presence of abundant obligate organohalide-respiring Dehalococcoides implied that the TA-wastewater contained uncharacterized organohalide compound(s). In the UASB reactor, acetate and H₂ generated from degradation of aromatic compounds in petrochemical wastewater by Syntrophorhabdus, Syntrophus and other syntrophs, together with low redox potential and the uncharacterized organohalide compounds, provide ideal growth niches for the fastidious Dehalococcoides. No other obligate dechlorinating bacteria, e.g., Dehalogenimonas and Dehalobacter, were found in the granular sludge sample. In a control sample collected from a lab-scale anaerobic sludge digester without organohalide amendment, no known dechlorinating bacteria can be detected (Figure 1). The highly similar microbial community structures of the three black-odorous river sediments, distinguish themselves from the community compositions of the granular sludge, especially the predominant lineages of Chloroflexi (i.e., Longilinea, GCA004, WCHB1-05 and Anaerolinaceae) and Proteobacteria (i.e., Syntrophobacter and Dechloromonas) (Şimsir et al., 2017) (Figure 1). Dehalococcoides were shown the appearance in the microbial community, on which indicate the potential of organohalides’ contamination.

To further evaluate the dechlorination activities, perchloroethene (PCE) was spiked into microcosms established with those Dehalococcoides-containing sediment and sludge samples. After around 2 months’ incubation, PCE dechlorination activities were observed in all three microcosms with the river sediment inocula (data not shown). Subsequent consecutive culture transfers of the three microcosms generated three active cultures which reductively dechlorinate PCE into vinyl chloride (VC) or ethene (Figure 2). No dechlorination activity was observed in the control microcosm established with digester sludge (Figure 2D).

In contrast to PCE dechlorination in sediments of the three black-odorous urban rivers, microcosms inoculated with the Dehalococcoides-containing granular sludge showed negative PCE-dechlorination activity. To identify potential organohalides to support the growth of Dehalococcoides in the granular sludge, organohalide pollution in the petrochemical wastewater as influent of the UASB reactor was evaluated. The petrochemical wastewater was generated from a AMOCO process that oxidize para-xylene to terephthalic-acid, using a homogeneous catalyst of cobalt and manganese together with bromide as a promoter, in which bromomethane was generated as a byproduct (Tomás et al., 2013). Due to difficulties in obtaining bromomethane, dehalogenation activity test was performed with chloromethane as a homolog alternative to bromomethane. In chloromethane-fed culture, over 70% chloromethane was dechlorinated within 8 days (Figure 3). No obvious dechlorination activity was observed in abiotic control.

The partial 16S rRNA gene sequences (∼400 bp) generated from Miseq sequencing of V4–V5 hypervariable regions were unable to differentiate Dehalococcoides between Cornell and Victoria subgroups. Therefore, Dehalococcoides genus-specific primers (i.e., FpDHC1/RpDHC1377) were utilized to generate longer 16S rRNA gene sequences (∼1300 bp) to identify the Dehalococcoides in the anaerobic granular sludge. Phylogenetic analysis showed the close clustering of Dehalococcoides in TA-degrading granules with D. mccartyi 195 in Cornell subgroup (Figure 4A), sharing 99% 16S rRNA gene sequence similarity (2 bp difference over 1311 bp) with that of strain 195.

To provide insight into the spatial distribution of Dehalococcoides in the granular sludge, FISH was conducted with Dehalococcoides-specific, bacterial and archaeal oligonucleotide probes (Amann et al., 1995; Yang and Zeyer, 2003). FISH analysis showed the scattered distribution of Dehalococcoides inside granules, closely colonized with other bacteria (Figure 4B) but separated from archaea (Figure 4C). Degradation of aromatic compounds by fermentative bacteria is thermodynamically restricted and will become endergonic (ΔG > 0) as metabolic byproducts (e.g., acetate and H₂) accumulate in the biosystem. Similar with methanogenic archaea, Dehalococcoides might form syntrophic interactions with aromatic compound degrading acidogens in the granular sludge: the degradation of aromatic compounds by Syntrophorhabdus and other syntrophs provide acetate as carbon source and H₂ as electron donor for the halorespiration of Dehalococcoides; correspondingly, Dehalococcoides help maintain acetate and H₂ at low concentration in the biosystem and ‘pull’ degradation of aromatic compounds toward completion through consuming metabolic byproducts generated by acidogenic bacteria. The close colonization of Dehalococcoides with syntrophic bacteria could facilitate the interspecies transfer of H₂ (Mao et al., 2015).