Environ. Sci. Technol. 2006, 40, 310-317
Stimulation and Molecular Characterization of Bacterial Perchlorate Degradation by Plant-Produced Electron Donors JOSHUA D. SHROUT,* GARRETT C. STRUCKHOFF, GENE F. PARKIN, AND JERALD L. SCHNOOR Department of Civil and Environmental Engineering, The University of Iowa, Iowa City, Iowa 52242
Root homogenate from poplar trees (Populus deltoides × nigra DN34, Imperial Carolina) stimulated perchlorate degradation in microcosms of soil and water samples collected at a perchlorate contaminated site, the Longhorn Army Ammunition Plant (LHAAP), located outside Karnack, Texas. Direct use of root products by perchloratedegrading bacteria was shown for the first time as six pureculture bacteria isolated from LHAAP perchloratedegrading microcosms degraded perchlorate when given root products as the sole exogenous source of carbon and electron donor. Nonenriched environmental consortia were able to utilize root products for perchlorate degradation, regardless of prior exposure to perchlorate. Microcosms that contained perchlorate-contaminated groundwater (MW-3) or uncontaminated surface water (Harrison Bayou) as inoculum degraded approximately 240 and 160 mg L-1 perchlorate, respectively, using root products (approximately 440 mg L-1 as COD) over 38 days. The predominant bacterial species in these aqueous microcosms, identified by DGGE, depended only upon the source inoculum as similar sequences were obtained whether root products or lactate was the electron donor. Sequences from DGGE bands that matched species within Dechloromonas, a genus consisting of many perchlorate degraders, were identified in all perchlorate-degrading microcosms. This study demonstrates the ability of root products to drive perchlorate respiration by bacteria and the potential for successful achievement of perchlorate rhizodegradation using in situ phytoremediation.
Introduction Perchlorate contamination is a significant problem in several areas of the United States, and this is of concern because perchlorate has been linked to abnormal thyroid function and other human health problems (1-6). While the future of federal regulation of perchlorate in drinking water is unclear, a recent National Academy of Science NRC committee recommended an oral reference dose (0.7 µg kg-1 d-1) that would set a drinking water standard of 24.5 µg L-1 perchlorate (7). As methods for effective remediation of * Corresponding author present address: Department of Microbiology, The University of Iowa, 540 EMRB, Iowa City, IA 52242; phone: (319)335-8240; fax (319)335-7949; e-mail: joshua-shrout@ uiowa.edu. 310
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perchlorate are sought, the most promising techniques appear to involve use of bacteria that respire and degrade perchlorate to chloride and water (8-13). Additionally, if perchlorate can be treated at the point(s) of contamination this is likely to lessen health risk and reduce costs. One remedy for highly contaminated soils and groundwater might be to treat perchlorate by engineering an in situ bioremediation zone where perchlorate-degrading bacteria become stimulated to degrade perchlorate. As interest in perchlorate bioremediation has increased, so have reports of isolation of perchlorate-degrading bacteria (9, 14-16). It is known that perchlorate degraders require an electron donor source to drive the respiration of perchlorate. Bacterial perchlorate reduction can be quite rapid and laboratory studies with both pure-strain and mixed-culture perchlorate-degrading bacteria have shown perchlorate degradation half-lives on the order of hours (9, 13, 15-17). However, few studies have addressed the capabilities of these organisms to respire perchlorate under conditions likely to be found at perchlorate bioremediation sites. Optimal conditions are rarely observed in situ. Specifically, an electron donor is needed for reduction of perchlorate (9, 18-20) but is often in short supply in the subsurface. Therefore, an exogenous electron donor must often be added in significant quantities (at significant cost). The challenge of in situ perchlorate remediation is to provide an effective electron donor source for bacteria that respire perchlorate. Recent reports in the literature have suggested the potential use of root products as bacterial substrates for bioremediation (21-25). Additionally, Leigh et al. (21) showed that the potential turnover of roots in the soil is significant; 58% of fine roots produced by mulberry saplings were sloughed from the plant at the end of a six-month growing season. Several studies have reported the use of compounds in root exudate as an electron donor and carbon source for bacterial degradation of pollutants such as PCBs and atrazine (21, 23). Research performed with poplar trees has shown a stimulation of bacterial populations in the rhizosphere capable of transforming pollutants (22). The potential for removal of perchlorate using plants has also been established as perchlorate degradation in simulated and field-scale rhizosphere zones was found to be significant (26-30). It is known that many simple carbon compounds are contained in plant roots that should be suitable for bacterial growth (31). Therefore, there is great potential for success in perchlorate degradation using root products, but using root products as electron donors for bacteria has not been directly investigated. The use of plants to provide needed electrons (and carbon) is particularly appealing due to the simplicity, cost, and aesthetics associated with their use. Additionally, as studies have shown the ability of plants to take up perchlorate directly, the use of plants with bacteria could lead to a synergistic perchlorate rhizodegradation scheme (26, 32-34). If 17% of photosynthate from young growing trees is diverted to root turnover, typical carbon supply rates would be approximately 100 g carbon (m2-day)-1 (i.e., 17% × 600 g carbon (m2-day)-1 primary production rate) (35). Research has shown that bacteria exist that can readily degrade perchlorate under the right conditions and suggests that perchlorate-degrading bacteria are ubiquitous in nature (9, 19). We hypothesized that perchlorate degradation could be stimulated using plant root products as the sole added source of electron donor for perchlorate-degrading bacteria. Here we report on perchlorate degradation by bacteria from soil and water samples collected from a perchloratecontaminated site that used root products as electron donors. 10.1021/es051130g CCC: $33.50
2006 American Chemical Society Published on Web 11/30/2005
TABLE 1. Inoculum and Enrichment Conditions Used for Isolation of Six Perchlorate Degrading Bacteria isolate
enrichment source microcosm
electron donor for enrichmenta
organism doubling time (h)b
Isolate JDS1 Isolate JDS2 Isolate JDS3 Isolate JDS4 Dechloromonas sp. JDS5c Dechloromonas sp. JDS6c
INF sediment + lactate INF sediment + lactate INF sediment + lactate Building 25C soil + IBJ-004 water + lactate Building 25C soil + IBJ-004 water + hydrogen Building 25C soil + IBJ-004 water + hydrogen
10 mM lactate 10 mM lactate 10 mM lactate 10 mM acetate 40% H2 headspace 40% H2 headspace
6.9 5.3 5.4 5.6 4.5 4.8
a All organisms isolated using 5 mM perchlorate in Freshwater medium. lactate. c See reference 52 for more detail.
b
The specific objectives of this work were (1) to assess the ability of plant root products to directly serve as an electron donor for bacterial perchlorate reduction; (2) to characterize pure-culture bacteria from a perchlorate-contaminated site that were capable of utilizing root products; and (3) to determine the diversity of bacteria cultivated in mixed-species perchlorate-degrading enrichments fed root products.
Electron Donor Utilization Experiments. Testing of the contribution of root products to perchlorate degradation under defined conditions was performed using batch experiments. Serum bottles containing Freshwater medium (supplied initially at 2× strength to account for dilution) and filter-sterile root homogenate containing 100 mg L-1 perchlorate to a volume of 50 mL in 60-mL a serum bottle with a headspace of N2/CO2 (80/20, v/v) were inoculated with Strain JDS4 and monitored for perchlorate reduction. Additional treatments containing root homogenate alone were also monitored. All treatments were performed in triplicate and were incubated at 20 °C in the dark on a shaker table at 160 rpm. The ability of six pure-culture bacteria isolated from LHAAP microcosms, isolate JDS1, isolate JDS2, isolate JDS3, isolate JDS4, Dechloromonas sp. JDS5, and Dechloromonas sp. JDS6 (JDS1-JDS6) to utilize lactate, acetate, hydrogen, succinate, propionate, butyrate, formate, casamino acids, glucose, ethanol, benzoate, benzene, or ferrous iron as electron donors for growth on 5 mM perchlorate was investigated. Additionally, the ability of these strains to grow using lactate, acetate, hydrogen, glucose, ethanol, benzoate, or benzene with 5 mM nitrate was investigated. Cultures were inoculated into 10 mL of sterile Freshwater medium under N2/CO2 gas (80/20, v/v) in 25-mL anaerobic pressure tubes. The medium contained 5 mM perchlorate or 5 mM nitrate (sodium nitrate) as an electron acceptor and one source of electron donor (in the form of dissolved sodium salt or acid, or 40% headspace in the case of hydrogen). Similarly, all isolates were investigated for the potential to grow with 10 mM acetate using the electron acceptors chlorate, nitrate, oxygen, fumarate, sulfate, sulfite, selenate, ferric iron, tetrachloroethene (PCE), and trichloroethene (TCE). The ability of isolates to grow on some electron acceptors with 10 mM lactate or 40% hydrogen headspace was also investigated. All tubes containing PCE, TCE, or benzene were stoppered with Teflon coated rubber septa to limit sorption. Positive growth was determined by ∼10-fold increase in OD600. Sterile techniques were used to inoculate each of the tubes with bacteria and they were incubated in the dark at 30 °C. Additional batch microcosms that contained LHAAP natural waters as inoculum were examined. A perchloratecontaining groundwater sample from Burning Ground #3 monitoring well MW-3 and a surface water sample containing no perchlorate from Harrison Bayou adjacent to the LHAAP were investigated. Freshwater medium was inoculated with 45 mL of site water and sterile root homogenate or lactate to a volume of 57 mL in 60-mL serum bottles with a headspace of N2/CO2 (80/20, v/v). Perchlorate (60 mg L-1) was added from an aqueous concentrated stock solution to Harrison Bayou microcosms. No initial perchlorate addition was required for MW-3 microcosms (initial concentration ) 240 mg L-1). All bottles were incubated at 20 °C in the dark on a shaker table at 160 rpm. Duplicate microcosms were investigated for each treatment.
Experimental Section Perchlorate-Contaminated Site Description. The Longhorn Army Ammunition Plant (LHAAP) located outside Karnack, Texas, is a site with confirmed subsurface perchlorate contamination (36). Perchlorate-containing Pershing missile rockets were constructed at the site from the 1950s through the 1980s. Additionally, missile rockets were dismantled at the site in the late 1980s and 1990s as part of disarmament stipulated in the INF Treaty of 1987 (5, 37). These activities resulted in perchlorate contamination. Concentrations of perchlorate have been detected as high as 250 mg L-1 in groundwater samples collected from the site. The source inocula for bacterial enrichments and isolations described below made use of soil and water samples collected from perchlorate-contaminated areas (Burning Ground #3, INF Holding Pond, and Building 25-C) as well as areas without perchlorate contamination (still on LHAAP property). Isolation of Perchlorate-Reducing Bacteria. Bacteria were isolated from LHAAP soil microcosms fed lactate or hydrogen that showed repeated removal of perchlorate (38). Standard anaerobic techniques were used for all isolations (39). “Freshwater” medium (40) was heated near boiling under N2/CO2 gas (80/20, v/v), and 10 mL was dispensed into 25-mL glass anaerobic pressure tubes, capped with thick butyl-rubber stoppers to maintain an anaerobic headspace (N2/CO2, 80/20, v/v), and sterilized by autoclaving. Media contained 5 mM perchlorate (as sodium perchlorate) as an electron acceptor and 10 mM lactate (as lactic acid), 10 mM acetate (as acetic acid), or 40% (headspace) H2 as an electron donor. Aliquots of LHAAP INF-Pond and LHAAP Building 25-C soil microcosms (0.1-0.5 mL) were inoculated into 10 mL of sterile medium and incubated at 30 °C in the dark (Table 1). Positive enrichments were identified by an increased optical density and were transferred (1 in 10, v/v) into fresh medium. After two transfers, isolated colonies (including isolates JDS1-JDS6) were obtained by growth of these enrichments in 1.2% Noble agar roll tubes containing 5 mM perchlorate and 10 mM lactate, 10 mM acetate, or 40% (headspace) H2 (39). Preparation of Root Products. Root homogenate from hybrid poplar tree cuttings (Populus deltoides × nigra DN34, Imperial Carolina) grown hydroponically in 25% Hoagland solution (41) was prepared by blending harvested plant roots into deionized water using a Biospec Products (Bartlesville, OK) hand-held homogenizer. This slurry was then filtersterilized (0.2 µm) to form the root homogenate used for these studies.
Doubling time of bacteria grown using 5 mM perchlorate and 10 mM
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Molecular Techniques and DGGE Analysis. Cells from 10-mL cultures of JDS1-JDS6 were harvested by centrifugation, resuspended in 1 mL of sterile water, and lysed by adding 20 µL of chloroform and incubating for 10 min at 90 °C. Primers specific to bacterial DNA encoding 16S rRNA were used (primer 8F [5′-AGAGTTTGATCCTGGCTCAG-3′] and primer 1525R [5′-AGGAGGTGATCCAGCC-3′]) to amplify DNA by PCR (9). DNA was further purified using standard molecular techniques for Sanger-based sequencing performed by the DNA Facility at the University of Iowa. DNA of cells from MW-3 and Harrison Bayou microcosms (2 mL) were purified using a Qiagen (Valencia, CA) DNeasy Tissue kit. Primers specific to bacterial DNA encoding 16S rRNA were used (primer 341F [5′-CCTACGGGAGGCAGCAG3′] and primer 907R [5′-CCGTCAAATCMTTGAGTTT-3′]) (42, 43). An additional 40 base-pair GC-rich sequence ([5′CGCCCGCCGCGCCCCGCGCCCGTCCCGCCGCCCCCGCCCG-3′]) was attached to the 5′-end of primer 341F as a DNA clamp for DGGE separation. A 5-µL aliquot of purified DNA extract was added in 50 µL of Qiagen HotStar Master Mix with primers 341F (1 mM) and 907R (1 mM). DNA was amplified by PCR with an initial period of 15 min at 95 °C; 30 cycles of 1 min at 95 °C, 1 min at 55 °C, and 2 min at 72 °C; and an end cycle of 10 min at 72 °C. Products (10-15 µL) from these incubations were separated using denaturing gel gradient electrophoresis (DGGE) by a 30-80% denaturant gradient at 60 °C using a fixed voltage of 100 V for 20 h (42). DNA was recovered from washed excised bands by incubation with sterile water (50 µL) overnight at 4 °C. Recovered DNA was amplified by repeating the PCR protocol described above, checked for purity by DGGE as described above, purified using a Qiagen (Valencia, CA) QIAquick Extraction Kit, and sequenced by the DNA Facility at the University of Iowa. Recovered DNA sequences were compared against known sequences in the GenBank database. A phylogenetic tree was generated to compare the recovered sequences from this study against (1) similar sequences in GenBank and (2) other relevant environmental bacteria. Sequences for the tree were organized with a multiple sequence global alignment using ClustalW (44). The aligned sequences were then edited using BioEdit (http://jwbrown.mbio.ncsu.edu/BioEdit/bioedit. html) to retain only the commonly shared region (427 bp) across all 41 species. This edited alignment was then utilized by PAUP* 4.0 to perform a maximum parsimony analysis to generate a phylogenetic tree (45). A bootstrap analysis with 100 replicates was performed using a heuristic search strategy to provide the probability for each indicated branch-point. The bootstrap support values are indicated at the branch nodes of a maximum parsimony tree of the phylogenetic relationships. Analyses. Quantitation for perchlorate was performed using a Dionex (Sunnyvale, CA) DX-500 ion chromatograph equipped with a Dionex ASRS suppressor operating in external water mode with a regenerant of 10 mM sulfuric acid. Separation was achieved with a Dionex AS11 column by a sodium hydroxide eluent. A Dionex CD20 conductivity detector performed detection and peak areas were integrated by Dionex PeakNet software. Injection volumes depended upon the initial concentration of perchlorate. The detection limit for perchlorate was approximately 0.0001 mM (0.01 mg L-1). Chemical oxygen demand (COD), used as a bulk measure of electron donor of root products, was determined by closed reflux method using Hach (Loveland, CO) COD vials (46).
Results and Discussion Stimulation of Perchlorate Degradation. Perchlorate degradation in enrichments containing soil collected from a perchlorate-contaminated site required the addition of exogenous electron donor. Microcosms containing LHAAP 312
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FIGURE 1. Stimulation of perchlorate degradation in LHAAP soils with addition of electron donor (lactate). Inoculum was from the INF Pond or Building 25C of the LHAAP. Lactate was added on Day 0, Day 21, and Day 83. soil showed an ability to degrade all added perchlorate (250 mg L-1) when fed lactate (200 mg L-1 COD, cumulative) (Figure 1). While some degradation (e40%) of the initial perchlorate dose was observed in unamended microcosms, degradation of subsequent perchlorate additions was only achieved in microcosms amended with lactate (data not shown). The ability to develop a perchlorate-degrading enrichment from LHAAP soil is in agreement with other studies that have suggested the general ubiquity of perchlorate-degrading bacteria in nature (9, 19) including perchlorate-contaminated sites such as the LHAAP (47, 48). Therefore it was apparent that the potential for bacteria to degrade perchlorate was present; however, low supply of electron donor limits bacterial activity when concentrations of perchlorate are high (g50 mg L-1), which is relevant to many perchlorate-contaminated sites (49, 50). Perchlorate Degradation by Pure Culture Bacteria. Some of the specific perchlorate-degrading bacteria present in these enrichments were identified. Six perchlorate-degrading bacteria, isolate JDS1, isolate JDS2, isolate JDS3, isolate JDS4, Dechloromonas sp. JDS5, and Dechloromonas sp. JDS6 (JDS1JDS6), were enriched and isolated from the perchloratedegrading microcosm described above (Figure 1) or similar enrichments that were shown to consistently degrade perchlorate when hydrogen or lactate was added for over one year (Table 1). All isolates were Gram-negative rods approximately 1 µm long that displayed motility in wet cell mounts; however, isolate JDS4 was observed as having significantly faster motility than the other five isolates. Doubling times for these bacteria were between 4.5 h (JDS5) and 6.9 h (JDS1) (Table 1). Comparatively, Dechloromonas agitata strain CKB and Dechlorosoma suillum strain PS, two previously well-characterized perchlorate degrading bacteria (51), were grown alongside the six isolates and the doubling times were 4.1 h and 6.9 h, respectively. Analyses for chloride indicated nearly 100% recovery from added perchlorate, and no chlorate or chlorite was detected. Analysis of the 16S rRNA gene sequences recovered from the six pure-culture bacteria JDS1-JDS6 places them all within the Proteobacteria class (Figure 2). From phylogenetic analysis of partial 16S rDNA sequence, bacteria JDS1, JDS2, and JDS3 appear most closely related to known dechlorinating organisms (i.e., dehalorespiration of chlorinated aliphatics) in the -Proteobacteria. Strain JDS4 was the most unique genotype identified of all six isolates. This strain showed the least phylogenetic relation to any described perchloratedegrading bacterium. The phenotypic characteristics of JDS4 were also unique as this strain showed the greatest versatility to utilize alternate electron donors and acceptors as compared to the other five isolates (e.g., only isolate capable of growth on ethanol and nitrate). Strains Dechloromonas sp.
FIGURE 2. 16S rDNA phylogeny of LHAAP perchlorate-degrading isolates (bold*) and bacterial sequences recovered using DGGE from perchlorate-degrading microcosms (bold). The maxiumum parsimony tree generated using PAUP 4.0* includes the closest relatives of sequences identified using Blastn as well as relevant environmental bacteria. The bootstrap support values from a majority-rule consensus tree are indicated at the branch nodes. Numbers in parentheses indicate the GenBank accession numbers. JDS5 and Dechloromonas sp. JDS6 are most similar to other perchlorate-degrading bacteria within the β-Proteobacteria (52). These newly isolated bacteria were tested for their ability to use tree root homogenates as a source of carbon and electrons to drive perchlorate respiration. This was examined because one potential perchlorate bioremediation treatment scheme involves the planting of trees to create an active rhizodegradation community of bacteria. Perchlorate was readily degraded by isolate JDS4 over time with homogenate from poplar tree roots (Populus deltoides × nigra DN34, Imperial Carolina) as the sole electron donor and carbon source (Figure 3). Perchlorate was degraded by JDS4 inoculated into sterile medium (2 mL of growing cells, OD600 ≈ 0.5, in 50 mL) containing 100 mg L-1 perchlorate and root homogenate (160 mg L-1 COD). Lack of 100% perchlorate removal was attributed to an inability to prepare completely anaerobic root homogenate and subsequent use of available electrons for aerobic respiration. (While all serum bottle preparation and transfers were performed in an anaerobic glovebox, root homogenate was prepared aerobically and the sole mechanism to remove oxygen was diffusion in an anaerobic headspace; other methods were not employed due to concerns of maintaining a sterile homogenate solution.)
FIGURE 3. Degradation of perchlorate by isolate JDS4 fed root homogenate. Error bars represent standard deviation of three replicates. The presence of oxygen (potentially inhibitory to perchlorate reduction) is of less concern for larger, mixed-culture, openatmosphere systems, as perchlorate has been shown to be removed in the presence of oxygen in separate studies (26, 38). No appreciable degradation of perchlorate was observed in treatments containing root homogenate without bacterial VOL. 40, NO. 1, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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TABLE 2. Electron Donors Tested for Growth with 5 mM Perchlorate at 30 °Ca
TABLE 3. Electron Acceptors Tested for Growth with 10 mM Acetate at 30 °Ca
electron donor
JDS1
JDS2
JDS3
JDS4
JDS5b
JDS6b
electron acceptor
lactate (10mM) acetate (10mM) hydrogen (40% headspace) succinate (5mM) propionate (5mM) butyrate (5mM) formate (5mM) casamino acids (1 g L-1) glucose (5mM) ethanol (5mM) benzoate (5mM) benzene (0.1mM) Fe (II) (5mM)
+ + -
+ + -
+ + -
+ + -
+ + +
+ + +
+ + *
+ + *
+ + *
+ + + *
+ + * *
+ + * *
-
-
-
+ -
-
-
perchlorate (5mM) chlorate (5mM) nitrate (5mM) oxygen (20%- -atm) sulfate (5mM) fumarate (5mM) sulfite (5mM) selenate (5mM) Fe (III) (5mM) TCE (0.1mM) PCE (0.1mM)
a -, No Growth observed. +, Growth observed, ∼10-fold increase in OD600. *, Only poor growth observed, ∼2-fold increase in OD600.b See ref 52.
inoculum; this verified that root products provided an electron donor for bacterial processes but did not reduce perchlorate directly. Additional investigation showed that all six bacterial isolates could degrade perchlorate using root homogenate or root exudate (Table S1 in the Supporting Information). To our knowledge, this is the first report of perchlorate degradation by pure culture bacteria that utilized root products as a sole carbon and energy source. These bacterial isolates were further characterized for their ability to use a variety of substrates for comparison with other known perchlorate-degrading bacteria. All six isolates were found to utilize lactate, acetate, and propionate as electron donors when using perchlorate as an electron acceptor (Table 2). Growth was characterized by an observable increase in optical density (OD600) (e.g., at least 1 order of magnitude) within a 2-week period or less. One difference in the trends observed for these isolates was the ability of JDS4 to grow with 5 mM ethanol and 5 mM nitrate but not with 5 mM ethanol and 5 mM perchlorate. Dechloromonas sp. JDS5 and Dechloromonas sp. JDS6 are the first autotrophic perchlorate-degrading bacteria isolated from a perchloratecontaminated site (52). No isolates were able to grow on or degrade 0.1 mM benzene under the conditions investigated (with 5 mM perchlorate or 5 mM nitrate). No isolates were able to ferment lactate or acetate. The six isolates were also examined for their ability to utilize alternate electron acceptors to perchlorate. All isolates could use 5 mM chlorate for growth with 10 mM acetate (Table 3). Bacteria JDS1, JDS2, and JDS3 were unable to grow using nitrate with any electron donor investigated (lactate, acetate, hydrogen, glucose, ethanol, benzoate, or benzene). Only isolate JDS4 was able to grow aerobically. Aerobic growth of JDS1, JDS2, and JDS3 was observed initially but could not be repeated from anaerobically maintained inocula when tested several months after the initial observation. Because chlorinated aliphatics such as TCE are cocontaminants with perchlorate in some areas of the LHAAP (36), the isolates were also tested for the ability to degrade such compounds. No growth using PCE or TCE or degradation of PCE or TCE was observed. The investigation of TCE or PCE degradation by isolate JDS1, isolate JDS2, and isolate JDS3 was of particular relevance considering the phylogenetic similarity (results below) of these bacteria with other dehalogenating species. Despite their genetic relatedness, these three isolates did not show dechlorinating activity. Nonetheless, the phylogeny of isolates JDS1, JDS2, and JDS3 are extremely interesting given their close relationship to a 314
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JDS1 JDS2 JDS3 JDS4 JDS5b JDS6b + + -
+ + -
+ + -
+ + + + + -
+ + + + -
+ + + + -
a -, No Growth observed. +, Growth observed, ∼10-fold increase in OD600. b See ref 52.
dechlorinating bacteria (bacterium DCE10), providing some evidence that bacteria capable of degrading both perchlorate and TCE might exist in nature. Degradation by a Characterized Microbial Community. Since LHAAP soils were shown to contain perchloratedegrading bacteria that were starved for an electron donor and that pure culture perchlorate-degrading bacteria were capable of using root products to drive perchlorate respiration, we tested the ability to directly promote perchlorate degradation in contaminated LHAAP materials. Perchloratecontaminated groundwater was used as inoculum in microcosm batch experiments. Uncontaminated surface water from Harrision Bayou, a surface waterway bordering the LHAAP, was also tested. Perchlorate degradation was stimulated in root-homogenate amended microcosms started with both perchlorate-contaminated and previously uncontaminated site water samples. Microcosms containing perchlorate-containing groundwater from LHAAP monitoring well MW-3 showed degradation of perchlorate, and complete degradation of perchlorate was achieved in one of two duplicates amended with root homogenate (initial COD ) 185 mg L-1) (Figure 4A). Most perchlorate degradation occurred after more root exudate (260 mg L-1 COD) was added on Day 34. Microcosms amended with lactate (190 mg L-1 COD and 260 mg L-1 COD, initially and on Day 34, respectively) degraded perchlorate similarly to the homogenate-fed microcosm that degraded all perchlorate. (Lactate was used as a positive control; based upon initial results and those of other studies (9, 38, 48)). Perchlorate was also degraded with root homogenate in microcosms containing previously uncontaminated surface water from Harrison Bayou. Apparent degradation of perchlorate in microcosms started from LHAAP soil samples with no known perchlorate exposure further suggested the possibility for using root homogenate to stimulate perchlorate degradation (Table S2 in the Supporting Information). Previous acclimation to perchlorate, therefore, was not required for root exudates to stimulate perchlorate degradation. All added perchlorate (70 mg L-1) was degraded in less than 10 days for Harrison Bayou water incubated with root homogenate (initial ) 185 mg L-1) (Figure 4C). While the rate of root homogenate utilization compared to lactate utilization was not determined, it is noteworthy that all Harrison Bayou microcosms degraded perchlorate within 10 days of the first known exposure to perchlorate. This demonstrates the potential for bacterial rhizodegradation of perchlorate at a site like the LHAAP. More perchlorate (90 mg L-1) and electron donor (260 mg L-1 COD) were added on Day 34 and most perchlorate was degraded within 4 days (Day 38). These microcosms were sacrificed after 38 days to perform molecular community analyses to assess differences in the communities that developed in these perchlorate-
FIGURE 4. Perchlorate removal and community profile of root homogenate-fed LHAAP microcosms. (A and C) Perchlorate removal over 38 days. Water was from (A) a perchlorate-contaminated monitoring well (MW-3) or (C) a stream adjacent to a perchlorate-contaminated site (Harrison Bayou). Roots or lactate were added as electron donors on Day 0 and Day 34. More perchlorate was added on Day 34 to Harrison Bayou microcosms. (B and D) DGGE separation of DNA encoding 16S rRNA amplified from microcosm samples fed homogenate or lactate from (B) monitoring well MW-3 or (D) Harrison Bayou. DNA was extracted after 34 days or 38 days incubation for duplicate microcosms that were fed lactate (Lac) or root homogenate (Root) as an electron donor. DNA extract from Dechloromonas agitata CKB was used as a standard for comparison in DGGE analysis. contaminated versus previously unexposed perchloratedegrading enrichments. Genomic DNA encoding 16S rRNA from all MW-3 and Harrison Bayou microcosms was recovered and amplified and then separated by denaturing gel gradient electrophoresis (DGGE). Few differences were observed between root product-amended and lactate-amended microcosms as the bands observed after DGGE analysis were nearly the same independent of the added electron donor and depended only upon the source inoculum (Figure 4). The observable DNA bands identified were different in perchlorate-contaminated MW-3 microcosms compared to those microcosms started from uncontaminated Harrison Bayou. Additionally, MW-3 and Harrison Bayou sequences showed no match to any of the six LHAAP bacteria JDS1-JDS6. Species LH1 was the only band observed in both duplicates for lactate-amended microcosms that was not observed in any root productamended microcosms. Some bands in Harrison Bayou microcosms were observed in only one of four microcosms. All bands present on DGGE gels were further investigated to determine the DNA sequences; however, some bands could not be sequenced. An analysis of sequences obtained from DGGE bands (i.e., those labeled “LH” in Figure 3) indicated a mixed bacterial population had been enriched in MW-3 and Harrison Bayou microcosms (Figure 2). Phylogenetic analysis suggested some LH sequences may represent bacteria within new genera; however, this could not be determined from the partial 16S sequences amplified from these experiments. The sequences of DGGE bands from the same vertical position for multiple lanes (utilized as qualitative controls) were highly similar except for position LH4 where two distinct sequences were recovered. Sequences from each vertical position with the greatest number of
nucleotide bases were utilized for phylogenetic analysis after editing the sequences to remove beginning and end regions with poor chromatographic response. A comparison of recovered sequences from DGGE bands with many perchlorate-degrading bacteria (including JDS1JDS6) and other well-described environmental bacteria showed highest similarity to several different bacterial genera (Figure 2). Both MW-3 and Harrison Bayou samples indicated the presence of at least one species that most likely is within the genus Dechloromonas. Species LH4B from MW-3 microcosms and species LH10 and LH11 from Harrison Bayou microcosms appear to be Dechloromonas species. This is significant as many of the perchlorate degrading bacteria isolated are from this genus (9, 16, 51, 53) and these bacteria likely contributed to the perchlorate degradation in these microcosms. For example, no perchlorate degradation was observed in the absence of band LH4 in MW-3 microcosms (cf., Figure 1 and Figure 4A), which supports the idea that degradation of perchlorate in MW-3 microcosms was due to bacteria identified by band LH4. Furthermore, the lack of perchlorate degradation observed by one root-amended microcosm could be explained by insufficient numbers of LH4 bacteria within 38 days. The presence of band LH4 recovered after 38 days suggests that perchlorate would have been degraded over time with the addition of more root homogenate. While the bacterial species performing perchlorate reduction in these MW-3 and Harrison Bayou microcosms could not be specifically identified, these organisms (LH4B, LH10, and LH11) within Dechloromonas likely contributed to the reduction of perchlorate. The presence of Dechloromonas bacteria at the LHAAP is also confirmed by the isolation of Dechloromonas sp. JDS5 and Dechloromonas sp. JDS6 (described above and in 52). VOL. 40, NO. 1, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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Those sequences of DGGE bands not closely related to Dechloromonas were most closely associated with other known soil bacteria. Species LH2 is likely within the genus Rhizobiumsthese bacteria are most commonly associated with the roots of legumes (54). Species LH9 was most closely related to Shewanella putrefaciens, a facultative, ironreducing bacterium (54). Species LH1, LH3, LH4A, LH5, LH6, LH7, and LH8 did not show particular similarity to known culturable bacteria utilized for phylogenetic analysis; however, a “Blastn” search suggested highest similarity to the genera Dechloromonas (LH1, 91% similarity, and LH4B, 96% similarity), Pseudomonas (LH8, 91% similarity), Ralstonia (LH3, 83% similarity) and Green Non-Sulfur (LH5, 94% similarity) for some of these sequences. Environmental Significance. This is the first report of bacterial perchlorate degradation using direct addition of root products as an electron donor. Root products were shown capable of promoting perchlorate degradation for pure culture bacteria and uncharacterized water samples. Perchlorate degradation occurred when using inocula with and without prior exposure to perchlorate. This is the first report of the bacterial phylogeny developed in a perchlorate-degrading enrichment from perchloratecontaminated site materials. Sequences from DGGE bands that showed high phylogenetic similarity to Dechloromonas species, the predominant genera of perchlorate degrading bacteria, were identified in all perchlorate-degrading microcosms. While this result was not unexpected, it is interesting that no sequences were associated with other known perchlorate-degrading bacteria outside the genus Dechloromonas. Overall, the predominant bacterial species identified in water microcosms by DGGE were essentially the same independent of the added electron donor and depended only upon the source inoculum. The results of this research support the hypothesis that plants can be used to foster a diverse, perchlorate-degrading, rhizosphere bacterial community. The ability to stimulate rhizodegradation of perchlorate has also been field tested at the LHAAP. A 0.28-hectare (0.7-acre) planting area containing 425 poplar trees removed an estimated 45% (0.1135 kg ClO4d-1) of perchlorate applied by irrigation (0.2528 kg ClO4d-1) over a 2-year period (Table S3 in the Supporting Information). These direct demonstrations of bacterial perchlorate degradation using root products and the removal of perchlorate within a field-scale phytoremediation/rhizodegradation zone are evidence for the potential of successful full-scale development of a passive perchlorate rhizodegradation system. Clearly, further investigation of plants as an electron donor delivery system for perchlorate-degrading bacteria is necessary to develop this potential engineering strategy to control migration of contaminant plumes and treat perchlorate contamination.
Acknowledgments This work was funded by the U.S. Army Operations Command (Grant Program DAAA09-00-C-0016 and DAAA09-02-C-0071) and the University of Iowa-National Science Foundation Research Training Grant (DBI-96-02247). Logistical support of the W. M. Keck Phytotechnologies Laboratory at the University of Iowa is appreciated. Romy Chakraborty and John D. Coates of the University of CaliforniasBerkeley provided instruction for the isolation of perchlorate bacteria. Assistance with DNA sequence work was provided by Andrew Hawkins, Caroline S. Harwood, Christopher Weber, Benoit van Aken, and Jeremy Rentz. Some root products were prepared by Maija Seppanen and Jeremy Rentz. Craig Just and Collin Just provided assistance with analytical measurements. 316
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Supporting Information Available Results from experiments showing degradation of perchlorate by perchlorate-degrading bacteria using Poplar tree root homogenate and exudate (Table S1), degradation of perchlorate by LHAAP soil microcosms fed root homogenate (Table S2), and summary perchlorate mass balance calculations for the LHAAP field-scale phytoremediation demonstration control volume between April 2003 and March 2004 (Table S3). This material is available free of charge via the Internet at http://pubs.acs.org.
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Received for review June 15, 2005. Revised manuscript received October 21, 2005. Accepted October 24, 2005. ES051130G
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