|Home | About | Journals | Submit | Contact Us | Français|
Polybrominated diphenyl ethers (PBDEs) have attracted attention recently due to their proven adverse effects on animals and their increasing concentrations in various environmental media and biota. To gain insight into the fate of PBDEs, microcosms established with soils and sediments from 28 locations were investigated to determine their debromination potential with an octa-brominated diphenyl ether (octa-BDE) mixture consisting of hexa- to nona-BDEs. Debromination occurred in microcosms containing samples from 20 of the 28 locations when they were spiked with octa-BDE dissolved in the solvent trichloroethene (TCE), which is a potential cosubstrate for stimulating PBDE debromination, and in microcosms containing samples from 11 of the 28 locations when they were spiked with octa-BDE dissolved in nonane. Debromination products ranging from hexa- to mono-BDEs were generated within 2 months. Notably, the toxic tetra-BDEs accounted for 50% of the total product. In sediment-free culture C-N-7* amended with the octa-BDE mixture and nonane (containing 45 nM nona-BDE, 181 nM octa-BDEs, 294 nM hepta-BDE, and 19 nM hexa-BDE) there was extensive debromination of the parent compounds, which produced hexa-BDE (56 nM), penta-BDEs (124 nM), and tetra-BDEs (150 nM) within 42 days, possibly by a metabolic process. A 16S rRNA gene-based analysis revealed that Dehalococcoides species were present in 11 of 14 active microcosms. However, unknown debrominating species in some of the microcosms debrominated the octa-BDE mixture in the absence of other added halogenated electron acceptors (such as TCE). These findings provide information that is useful for assessing microbial reductive debromination of higher brominated PBDEs to less-brominated congeners, a possible source of the more toxic congeners (e.g., penta- and tetra-BDEs) detected in the environment.
Since they were first developed in the 1960s, polybrominated diphenyl ethers (PBDEs) have been used as flame retardant additives in an array of common household and industrial appliances. As a result of their widespread use, PBDEs have become ubiquitous environmental contaminants, and increasing levels have been detected in the air, soil, and water (5, 12). In a recent study, Leung et al. reported the highest PBDE concentrations in soil samples (2.7 to ~4.3 ppb) and combusted residues (33.0 to ~97.4 ppb) that were collected in Guiyu, Guangdong Province, China (18). More worrisome is the fact that increasing concentrations of PBDEs have also been detected in marine mammals, birds, fishes, and human tissues (3, 14, 20, 30), and 63 ppm of PBDEs in bird eggs is the highest level ever found in biota (23). The PBDE concentrations in both environmental samples and biota have been increasing exponentially, with a doubling time of 4 to 6 years (5, 12). Although the PBDEs comprise 209 different congeners designated 1 to 209, the PBDE congeners most often detected in biota (e.g., human tissues) include tetra-brominated diphenyl ether (tetra-BDE) (congener 47), penta-BDEs (congeners 99 and 100), and hexa-BDEs (congeners 153 and 154), which may have originated directly from a commercially available penta-BDE technical mixture or indirectly via breakdown of an octa- or deca-BDE technical mixture (10, 12). PBDEs began to receive worldwide scientific and public attention when a temporal study performed from 1972 to 1997 revealed increasing concentrations of PBDEs in Swedish human breast milk (19). Toxicological studies of rodents using a commercial penta-BDE mixture (including tetra-, penta- and hexa-BDEs) and congeners in a commercial octa-BDE mixture (such as hepta-BDE [congener 183] and octa-BDE [congener 203]) revealed developmental neurotoxicity, reproductive toxicity, liver toxicity, and disruption of thyroid hormone levels (24, 26, 29).
To date, studies of PBDEs have focused mainly on detection of these compounds in the environment and their potential adverse health effects; only a few studies have reported microbial debromination of PBDEs (7, 10, 22, 25). Recently, He et al. demonstrated debromination of a technical octa-BDE mixture by pure isolates of Dehalococcoides species, which generated hepta- to di-BDEs after 6 months of incubation (10). Additionally, microbes belonging to the genera Dehalobacter and Desulfitobacterium were also found to be able to debrominate individual PBDE congeners present in commercial octa-BDE mixtures (10, 22). However, the debromination of PBDEs in both studies required the presence of a primary electron acceptor (e.g., chloroethenes or chlorophenols); in other words, debromination occurred cometabolically.
In addition to debromination of PBDEs by pure cultures, a previous study demonstrated that in anaerobic sludge 5% of added deca-BDE (congener 209) was debrominated to nona- and octa-BDEs (total amount of product, 0.5 nmol) after 238 days of incubation (7). Moreover, another study showed that deca-BDE was debrominated to products ranging from nona-BDEs to hexa-BDEs in 3.5 years with anaerobic sediments as the inocula (25). These findings suggest that microbial reductive debromination of highly brominated congeners, such as deca-, nona-, octa-, and hepta-BDEs, may contribute to formation of less-brominated PBDEs in the environment, which are potentially more toxic (e.g., tetra- and penta-BDEs). Additionally, debromination of less-brominated PBDE congeners, such as di-BDE, to mono-BDE and diphenyl ether was demonstrated in a fixed-film plug flow biological reactor (21). Besides microbial debromination, highly brominated PBDEs were also found to be transformed to lower congeners via photodegradation or in vivo metabolism in aquatic and terrestrial animals (1, 16).
This study was initiated to obtain information about the distribution of microorganisms capable of debrominating highly brominated PBDE congeners to more toxic daughter products or the final product diphenyl ether by assessing microcosm samples collected from various locations. Debromination of an octa-BDE mixture was evaluated in the presence of the potential energy-generating cosubstrate trichloroethene (TCE) (PBDEs dissolved in TCE) or in the presence of the relatively inert solvent nonane (PBDEs dissolved in nonane). The latter experiment provided, for the first time, information about the possible microbes living on the energy generated from the debromination of an octa-BDE mixture in the absence of any cosubstrate, such as TCE or another primer compound. Initial insights into the key debrominating microbes were obtained by using genus-specific 16S rRNA gene-based techniques.
A commercial octa-BDE mixture (consisting of nona-BDE [congener 207], octa-BDEs [congeners 203, 196, and 197], hepta-BDE [congener 183], and hexa-BDE [congener 153]) was purchased from Sigma-Aldrich (St. Louis, MO). Isooctane (2,2,4-trimethylpentane; analysis quality) was obtained from Tedia Company Inc. (Fairfield, OH). Deca-bromobiphenyl (DBB) was purchased from AccuStandard (New Haven, CT). Trichloroethene (TCE), n-nonane, and diphenyl ether with a minimum purity of 99.5% were obtained from Sigma-Aldrich (St. Louis, MO).
Samples were collected in three regions, East Asia (Wuhan in Hubei and Guiyu in Guangdong, China), Southeast Asia (Singapore), and North America (San Francisco in California). The characteristics of the sampling sites are shown in Table Table1.1. The soil and sediment samples were acquired by filling sterile 50-ml plastic Falcon tubes that were capped and transported to the laboratory at the ambient temperature. Within 1 week, microcosms were established aseptically in a Bactron anaerobic chamber (Sheldon Manufacturing, Inc., Cornelius, OR), in which ~5-g portions of collected samples were distributed into 60-ml serum bottles containing ~30 ml of bicarbonate-buffered defined mineral salts medium (9). Each sample from the 28 locations was spiked with pyruvate (10 mM), lactate (10 mM), or acetate (10 mM) in duplicate bottles for each condition. The media were reduced by adding l-cysteine and Na2S·9H2O (0.2 mM each) (9). The vitamin solution of Wolin et al. (27) and vitamin B12 (25 mg/liter) (8) were added to all samples. Unless stated otherwise, H2 was added to acetate-containing microcosms at a partial pressure of 3.4 × 104 Pa (10 ml). Prior to addition of H2, the same volume of headspace was removed to ensure that there was minimal change in the pressure in the bottles. All the bottles were crimp sealed with butyl rubber septa (Geo-Microbial Technologies, Inc., Ochelata, OK). Either TCE (1.5 μl) or nonane (3 μl) carrying dissolved PBDEs (0.05 g of the octa-BDE mixture dissolved in 10 ml TCE or nonane) was added to the samples and controls. The amount of the PBDE-TCE preparation added to the bottles was about one-half the amount of the PBDE-nonane preparation added due to concerns about the inhibitory effects of TCE or the potential synergistic combined toxic effects of the TCE and PBDEs on the microbes. All the sample bottles were incubated in an upright position (to minimize sorption to the butyl rubber septa) at 30°C without agitation in the dark.
To minimize sorption effects on quantification of PBDEs, 20 actively debrominating microcosms of the 28 microcosms containing octa-BDEs and TCE (octa-BDE/TCE microcosms) and 11 actively debrominating microcosms of the 28 microcosms containing octa-BDEs and nonane (octa-BDE/nonane microcosms) were transferred to fresh medium to perform a quantitative assay for PBDEs. It should be noted that only one microcosm that exhibited the most rapid debromination activity and had a distinctly different debromination profile was transferred for samples from one location. Before transfer, the microcosm bottles were vigorously shaken and left for 30 s so that the suspended solids could settle, and then 1-ml portions of supernatant (containing minute amounts of solids) were transferred into duplicate bottles containing 30 ml of fresh basal salts medium spiked with the same carbon source and substrate (PBDEs) as the parental bottles. The active transferred microcosms were then analyzed by gas chromatography-mass spectrometry (GC-MS) as described below to obtain the results shown in Table Table1.1. During the experiments, control microcosms (two microcosms for each type of soil or sediment) were prepared in the same way as the samples, except that the inoculated medium was autoclaved prior to addition of PBDEs. Subsequent transfers were performed as described above.
Biweekly, 1-ml portions of supernatants (from original microcosms) or liquid samples (from transferred cultures) were removed from the bottles with a disposable plastic syringe (Becton, Dickinson and Company, Franklin Lakes, NJ) after vigorous shaking. Each 1-ml sample was placed in a 4-ml amber glass vial, which was then spiked with the same volume of isooctane. The mixture was vortexed for 5 min and then shaken for 16 h in the dark to minimize exposure to light. The sample was then centrifuged at 31,550 × g for 5 min. The upper isooctane layer containing PBDEs was transferred to a 2-ml amber glass vial and concentrated to 400 μl with N2.
After liquid-liquid extraction, PBDEs were analyzed by GC-MS with a model GC 6890/MSD 5975 apparatus (Agilent Technologies, Inc., Santa Clara, CA) that was equipped with a Restek Rxi-5ms column (15 m; inside diameter, 0.25 mm; film thickness, 0.25 μm; Restek Corporation, Bellefonte, PA). Helium was used as the carrier gas at a flow rate of 1.2 ml/min. Based on experiments conducted using different injector temperatures (250 to 320°C) for a fixed analyte concentration, the optimum injector temperature (300°C) was selected, at which the highest peak response occurred. This observation correlates well with the report of Björklund et al. that a relatively high injector temperature facilitates detection of higher brominated congeners (2). Therefore, the injection mode selected was pulsed splitless with an injector temperature of 300°C. The initial oven temperature used for GC was 110°C, after which the temperature was increased to 310°C at a rate of 15°C/min and kept at 310°C for 5 min; the total program time was 25.7 min. The MSD transfer line, ion source, and quadrupole temperatures used were 300°C, 280°C, and 150°C, respectively. The analysis was done by using selected ion monitoring in the electron impact ionization mode.
The substrate (PBDEs) was quantified by constructing five-point calibration curves for the congeners in an octa-BDE mixture consisting of nona-BDE (congener 207), octa-BDEs (congeners 203, 196, and 197), hepta-BDE (congener 183), and hexa-BDE (congener 153). The concentrations of the congeners in the mixture were computed by using ratios obtained from a previous study performed by La Guardia et al. (15). For quantification of product peaks, five-point calibration curves were constructed by using the average peak areas of congeners in the same homolog group in an analytical standard (98% pure, dissolved in n-nonane) which contained 39 congeners from mono- to hepta-BDEs (Cambridge Isotope Laboratories, Inc., Andover, MA). The peak responses for the same homolog group had an acceptable variation of 12% for hepta- to tetra-BDEs, while the variation was up to 28% for tri- to mono-BDEs. Currently, average calibration curves are the most feasible way to quantify unknown debromination products. A total of nine calibration curves (from mono-BDE to nona-BDE) were constructed. Deca-bromobiphenyl (DBB) was used as an internal standard and was added to samples prior to PBDE extraction, which accounted for losses during the extraction process. The extraction efficiency for the samples was determined by normalizing the amount of DBB in PBDE samples with the known initial amount of DBB added to the samples. The extraction efficiencies for DBB ranged from 71% to 84%. The concentrations of PBDEs were computed by dividing the concentrations obtained from the calibration curves by the extraction efficiencies for the corresponding samples. The instrument detection limits for PBDE congeners ranged from 0.070 to 0.224 nM. Chloroethenes were analyzed as described previously (11).
Cells (1 ml from original microcosm samples and transferred samples) used for DNA extraction were centrifuged at 14,000 rpm for 15 min at 4°C. After the supernatant was removed, genomic DNA was extracted from cell pellets by using an UltraClean soil DNA isolation kit (MO BIO Laboratories, Inc., Carlsbad, CA) according to the manufacturer's instructions. PCR was carried out with an Eppendorf Master Cycler ep gradient S thermocycler (Eppendorf AG, Hamburg, Germany) with universal bacterial primers (primers 8F and 1541R), group-specific primers for Chloroflexi (primers 348F and 884R), and genus-specific primers for Dehalococcoides (primers 730F and 1350R), Dehalobacter (primers 179F and 1007R), and Desulfitobacterium (primers 205F and 1020R) (4, 6, 13, 17, 28). Nested PCR was performed by using direct PCR (primers 8F and 1541R) products as templates for subsequent amplification with specific primers, such as primers specific for Chloroflexi (primers 348F and 884R) or Dehalococcoides (primers 730F and 1350R). The PCR products (3 μl) were resolved on 1% agarose gels and stained in an aqueous ethidium bromide solution (0.5 mg/ml) for 20 min. The bands were visualized by UV excitation, and images were obtained with a digital camera (Bio-Rad, Hercules, CA).
To prepare for sequencing, PCR products of the partial Dehalococcoides 16S rRNA gene were excised from an agarose gel and purified by using a QIAquick PCR purification kit (Qiagen GmbH, Germany). The purified PCR products were sent to 1st BASE Pte. Ltd. (Singapore) for sequencing with primers 730F and 1350R specific for Dehalococcoides (4).
In this study, the samples were designated based on the locations from which they were collected and the solvents used to dissolve PBDEs. The abbreviations used for locations were as follows: C, Wuhan; GY, Guiyu; S, Singapore; and U, San Francisco. For solvents the abbreviations used were T (TCE) and N (nonane). For example, C-T-7 indicates a sample from Wuhan with the PBDEs dissolved in TCE.
The assembled nucleotide sequences have been deposited in the GenBank database under accession numbers FJ594420 to FJ594427.
Soil and sediment samples collected from 28 locations were used as inocula in microcosm studies to determine their abilities to debrominate congeners in a technical octa-BDE mixture. After an initial incubation for 1 month, debromination products appeared in microcosms (amended with lactate, pyruvate, or acetate) containing samples from 20 locations (octa-BDE/TCE microcosms), and they were measured qualitatively based on the appearance of new product peaks compared to control samples. In order to minimize sorption effects on the quantification of PBDEs, the 20 active octa-BDE/TCE debrominating microcosms (one microcosm from each location) and autoclaved controls were transferred once to fresh medium. The initial (and 2-month) PBDE concentrations in the control bottles were 30 ± 9 nM (27 ± 6 nM) for nona-BDE, 123 ± 14 nM (115 ± 19 nM) for octa-BDEs, 178 ± 32 nM (166 ± 27 nM) for hepta-BDE, and 12 ± 3 nM (11 ± 2 nM) for hexa-BDE, indicating that PBDE sorption to the solids in the transferred microcosms did not play a significant role. Table Table11 shows the concentrations of debromination products in the 20 transferred active microcosms on day 60. The penta- and tetra-BDE debromination products were detected in most samples, while less-brominated congeners, such as tri- and di-BDEs, were formed in 60% and 20% of the 20 active microcosms, respectively. Overall, tetra-BDEs accounted for 50% of the total debromination products in all active microcosms. Debromination products were not observed in autoclaved controls throughout the experiment. Table Table11 also shows the dechlorination endpoints for the carrier solvent TCE.
It is noteworthy that of all the microcosms, microcosm GY-T-2 exhibited the most extensive debromination of the octa-BDE mixture, generating hexa-BDE (18.3 nM), penta-BDEs (10 nM), tetra-BDEs (26.3 nM), tri-BDE (10.3 nM), di-BDE (11.9 nM), and mono-BDE (3.6 nM) after incubation for 2 months. Interestingly, diphenyl ether was also observed in microcosm GY-T-2 (peak not quantified).
The octa-BDE mixture dissolved in the solvent nonane (octa-BDE/nonane) was added to the microcosms in order to investigate the possibility of debromination without addition of any external electron acceptor (e.g., TCE) other than PBDEs. Similar to the octa-BDE/TCE microcosms, microcosms were prepared with samples from 28 locations, and the 11 active microcosms (one microcosm from each location where there was activity) containing the octa-BDE mixture and nonane were transferred to fresh medium amended with lactate (10 mM), pyruvate (10 mM), or acetate (10 mM)-H2 (3.4 × 104 Pa or 10 ml) for quantification studies. The initial (and 2-month) PBDE concentrations in the control bottles were 58 ± 7 nM (56 ± 11 nM) for nona-BDE, 231 ± 24 nM (216 ± 34 nM) for octa-BDEs, 332 ± 47 nM (342 ± 51 nM) for hepta-BDE, and 25 ± 10 nM (23 ± 7 nM) for hexa-BDE. The observed debromination products in the 11 transferred microcosms are shown in Table Table1.1. The microcosms which exhibited high debromination rates with octa-BDE/TCE (e.g., microcosms C-T-9, C-T-10, GY-T-1, GY-T-2, GY-T-10, GY-T-11, and S-T-2) did not exhibit debromination with octa-BDE/nonane (data not shown). Similarly, bacteria in microcosms showing rapid debromination with octa-BDE/nonane (e.g., microcosms GY-N-9, GY-N-12, and GY-N-16) did not debrominate the octa-BDE mixture dissolved in TCE (data not shown due to the absence of microbial activity).
Debromination was generally less extensive in octa-BDE/nonane microcosms than in octa-BDE/TCE microcosms. After 60 days, hexa-BDEs were detected in all active samples, while penta- and tetra-BDEs appeared only in 5 of the 11 samples (Table (Table1).1). When nonane was the carrier solvent for the octa-BDE mixture, debromination of certain substrate congeners (e.g., hepta-BDE [congener 183] and octa-BDE [congener 203]) was more evident than it was in octa-BDE/TCE-amended microcosms, as observed in the GC-MS chromatograms (data not shown). Another observation was that microcosms amended with acetate-H2 and with octa-BDE/nonane (e.g., microcosms C-N-7ace, U-N-1ace, and U-N-2ace) were able to debrominate a wider range of substrate congeners (nona-, octa-, and hepta-BDEs) than the other microcosms (only hepta-BDE and hexa-BDE debromination was observed) listed in Table Table11 (data not shown).
Microcosm C-N-7ace was transferred to fresh medium amended with acetate-H2, resulting in a sediment-free culture (designated C-N-7*). After 42 days of incubation, the concentrations of all congeners in the octa-BDE mixture (such as nona-BDE [congener 207], octa-BDEs [congeners 196, 197, and 203], and hepta-BDE [congener 183]) decreased, and less-brominated congeners formed, including hexa-BDE (56 nM), three unidentified penta-BDE congeners (124 nM), and two unidentified tetra-BDE congeners (150 nM) (Fig. (Fig.1).1). With the generation of debromination products, significant amounts of the substrate congeners were removed, as reflected by reductions in the concentrations of nona-BDE (46% removal of the initial 45 nM), octa-BDEs (57% removal of the initial 181 nM), hepta-BDE (75% removal of the initial 294 nM), and hexa-BDE (55% removal of the initial 19 nM) (Fig. (Fig.1B).1B). No debromination beyond tetra-BDEs was detected even after an extended incubation period (8 months) and amendment with additional H2 (3.4 × 104 Pa or 10 ml). The concentrations of total PBDEs in the control bottles are shown in Fig. Fig.1B1B (inset). Many factors, such as losses during sampling, sorption effects, and analysis errors, contributed to the discrepancies in the data for the control bottles. Quantification of PBDEs in environmental samples has been shown to be a significant challenge.
In addition to culture C-N-7*, sediment-free culture U-N-1* exhibited the most complete conversion of substrate PBDE congeners to hexa-BDEs when it was amended with 10 mM acetate and 3.4 × 104 Pa H2 and with octa-BDE/nonane (Fig. (Fig.2).2). On day 105, the concentrations of hexa-BDE congeners 153 and 154 were 202.67 nM and 91.67 nM, and this was concomitant with the debromination of 84% of nona-BDE, 79% of octa-BDEs, and 68% of hepta-BDE. Although large amounts of the substrates were removed, debromination did not proceed beyond hexa-BDEs even after extended incubation for 1 year and amendment with additional acetate and H2.
Table Table11 shows that microcosms amended with different carbon sources or carrier solvents had different debromination profiles. For further study, microcosm C-T-7 or C-N-7 was transferred into four different growth media, including medium amended with pyruvate and octa-BDE/nonane, medium amended with acetate-H2 and octa-BDE/nonane, medium amended with acetate-H2 and octa-BDE/TCE, and medium amended with pyruvate and octa-BDE/TCE. After 4 weeks of incubation, no debromination was observed in the culture spiked with pyruvate and octa-BDE/nonane. In contrast, debromination occurred with generation of tetra-, tri-, and di-BDEs when the culture was amended with pyruvate and octa-BDE/TCE (Fig. (Fig.3).3). Simultaneously, TCE was dechlorinated to dichloroethenes (DCEs) either in the presence or in the absence of the octa-BDE mixture, indicating that the TCE dechlorinator(s) debrominated the octa-BDE mixture in a cometabolic manner. In the acetate-H2-amended medium spiked with octa-BDE/nonane, substantial amounts of hexa-, penta-, and tetra-BDEs were produced, while significant amounts of tetra-BDEs appeared in the culture spiked with octa-BDE/TCE (Fig. (Fig.33).
To summarize, as shown in Fig. Fig.3,3, in the presence of TCE, pyruvate was a more suitable carbon source or electron donor for dehalogenators to generate less-brominated congeners, such as tri- and di-BDEs.
To acquire information about the possible dehalogenating microbes present in the microcosms, Dehalococcoides, Dehalobacter, and Desulfitobacterium genus-specific primer pairs were utilized to target the 16S rRNA genes of genomic DNA extracted from 14 distinctly different microcosm samples (different in terms of debromination endpoint and product concentration) (Fig. (Fig.4).4). The genera mentioned above were chosen due to previous reports on their abilities to debrominate PBDEs (10, 22). Because of the structural similarity between PBDEs and polychlorinated biphenyls (PCBs), Chloroflexi primers targeting the 16S rRNA genes of both Dehalococcoides species and o-17/DF-1-like microorganisms (PCB-dechlorinating microorganisms) were also tested with the DNA extracted from the microcosms in this study (6).
PCR-amplified 16S rRNA gene products confirmed that there was successful extraction of DNA from the 14 microcosms when the genomic DNA was targeted with universal primers (primers 8F and 1541R). When the genomic DNA of the microcosms was targeted with Dehalococcoides primers (primers 737F and 1350R), 8 of 14 samples produced amplicons that were the expected size (620 bp) (Fig. (Fig.4A,4A, lanes 2 to 9). Notably, all eight samples were spiked with the octa-BDE mixture dissolved in TCE. A nested PCR using the same Dehalococcoides primers (primers 737F and 1350R) was performed for the 16S rRNA gene for the six microcosm samples that were negative in the direct PCR test described above. Three of these samples (samples from microcosms GY-T-2, GY-N-16, and U-N-2) produced positive Dehalococcoides signals in the nested PCR, while the GY-N-5, U-N-1*, and C-N-7* samples did not produce any bands (Fig. (Fig.4A,4A, lanes 11 to 16). Bands exhibiting positive signals were sequenced and were found to be affiliated with the Pinellas or Cornell group of the Dehalococcoides cluster with a sequence similarity of 98 to 100% for 500 bp (GenBank accession no. FJ594420 to FJ594427), suggesting that Dehalococcoides may be responsible for PBDE debromination. Meanwhile, no Dehalobacter and Desulfitobacterium amplicons were detected with the direct PCR approach, while nested PCR detected the presence of Dehalobacter in samples from microcosms GY-T-2, C-T-10, and C-T-14 and Desulfitobacterium in a sample from microcosm GY-N-5. These results indicate that Dehalobacter and Desulfitobacterium species may play minor roles in debromination of PBDEs in the microcosms used in this study.
When Chloroflexi-affiliated microbes in the microcosms were screened, PCR amplicons of the expected size (536 bp) were observed for 9 of 14 samples (Fig. (Fig.4B).4B). The five samples which did not produce any bands in this analysis produced bands (Fig. (Fig.4B)4B) when nested PCR (with Chloroflexi-specific primers) was performed using the universal primer pair-amplified PCR products as the DNA templates, indicating that microorganisms other than Dehalococcoides (perhaps members of the Chloroflexi) were involved in PBDE debromination in the non-Dehalococcoides-containing microcosms (e.g., microcosms GY-N-5, U-N-1*, and C-N-7*).
PBDEs in the environment can end up in various forms, ranging from mono- to nona-BDEs and diphenyl ether, due to environmental microbial activity, a conclusion which is supported by the results of this and previous studies (7, 10, 21, 22, 25). This investigation also provided preliminary evidence that debrominating microbes are widespread in various regions and different climatic zones. More importantly, the highly toxic penta- and tetra-BDEs constituted the major homolog group of debromination products in the microcosms studied. Our findings demonstrate, for the first time, that debromination of an octa-BDE mixture occurs in the absence of a more readily utilized electron acceptor (e.g., TCE or primer compounds), suggesting that microbe-mediated debromination may be easier in the natural environment than previously thought.
Culture C-N-7* is the first known culture capable of significant removal (46 to 75%) of congeners present in a commercial octa-BDE mixture and conversion to penta- and tetra-BDEs (Fig. (Fig.1)1) within 42 days, which is much faster and more complete than that obtained with other known defined or undefined cultures (which generally require 0.3 to 3.5 years to remove ~10% of the added PBDEs and require the presence of the cosubstrate TCE or other primer compounds) (10, 22, 25). Since cultures U-N-1* and U-N-2* also significantly debrominated the octa-BDE mixture in nonane to hexa-BDEs, our results correlate well with previous observations that hexa-, penta-, and tetra-BDEs are the PBDEs most often detected in environmental samples (5, 12). Further studies are required to confirm that cultures C-N-7*, U-N-1*, and U-N-2* respire solely with PBDEs. It is also noteworthy that less-brominated products, such as di- and mono-BDEs, were observed in microcosms containing samples from Guiyu (e.g., microcosms GY-T-2, GY-T-1, and GY-T-11) after incubation for 2 months (Table (Table1).1). These daughter compounds were not detected in the original soil or sediment samples, confirming that they were indeed debromination products and not historical PBDE congeners. The generation of the less-brominated congeners (mono- and di-BDEs) may have been due to debromination of historical PBDE congeners in the sediments, such as penta-, tetra-, or tri-BDE congeners in microcosm GY-T-2 (Table (Table1).1). However, we cannot rule out the possibility that microcosm GY-T-2 is capable of debrominating congeners in the octa-BDE mixture to mono-BDE, since microbes from sites at Guiyu had been exposed to high concentrations of PBDEs for 12 years (18).
During the investigation of octa-BDE mixture debromination, we observed that the carrier solvent correlated with the distribution of Dehalococcoides species. The different debromination profiles or endpoints could be attributed to the presence of diverse Dehalococcoides species, as corroborated by various 16S rRNA gene sequences, or to a microbe(s) other than a Dehalococcoides species. In this study, TCE promoted the growth of Dehalococcoides species (Fig. (Fig.4A,4A, lanes 2 to 9) cultivated with octa-BDE/TCE in the microcosms, indicating the importance of this genus in the dehalogenation process in the environment. Interestingly, the debromination of octa- and nona-BDEs was more efficient in Dehalococcoides-free cultures C-N-7* and U-N-1* than in Dehalococcoides-containing microcosms (e.g., microcosms C-T-2, C-T-3, C-T-7, C-T-10, C-T-14, GY-T-11, S-T-1, and U-T-2). The distinctly different debromination abilities of Dehalococcoides-free microcosms and cultures (e.g., GY-N-5*, C-N-7*, and U-N-1*) warrant further investigations of the identities of the microorganisms present and their roles in the debromination process.
Together, the results of this study provide valuable insights into the role of environmental microbes in the fate of PBDEs in the natural environment, and they complement previous studies. The high debromination rates for higher brominated PBDEs shown here, coupled with the use of these compounds as flame retardants in many consumer products, increase the threat that more-toxic less-brominated congeners will be released following microbial degradation. It is our hope that this assessment serves as a platform for further, in-depth studies to address these environmental threats.
This research was supported by the Singapore Agency for Science, Technology and Research (A*STAR) of the Science and Engineering Research Council under project 062 101 0028.
Published ahead of print on 11 December 2009.