Molecular Characterization and In Situ Quantification of Anoxic Arsenite Oxidizing Denitrifying Enrichment Cultures

Abstract

To explore bacteria involved in the oxidation of arsenite (As(III)) under denitrifying conditions, three enrichment cultures (ECs) and one mixed culture (MC) were characterized that originated from anaerobic environmental samples. The oxidation of As(III) (0.5 mM) was dependent on NO₃⁻ addition and N₂-formation was dependent on As(III) addition. The ratio of N₂-N formed to As(III) fed approximated the expected stoichiometry of 2.5. A 16S rRNA gene clone library analysis revealed three predominant phylotypes. The first, related to the genus Azoarcus from the division β-Proteobacteria, was found in the three ECs. The other two predominant phylotypes were closely related to the genera Acidovorax and Diaphorobacter within the Comamonadaceae family of β-Proteobacteria and one of these was present in all of the cultures examined. Fluorescent in situ hybridization (FISH) confirmed that Azoarcus accounted for a large fraction of bacteria present in the ECs. The Azoarcus clones had 96% sequence homology with Azoarcus sp. strain DAO1, an isolate previously reported to oxidize As(III) with nitrate. FISH analysis also confirmed that Comamonadaceae were present in all cultures. Pure cultures of Azoarcus and Diaphorobacter were isolated and shown to be responsible for nitrate-dependent As(III) oxidation. These results taken as a whole suggest that bacteria within the genus Azoarcus and the family Comamonadaceae are involved in the observed anoxic oxidation of As(III).

Introduction

Arsenic (As) contaminated drinking water poses a risk to millions of people around the world (Smedley & Kinniburgh, 2002). Numerous studies have provided compelling evidence linking As in drinking water with cancer and other medical disorders (ATSDR, 2007). Arsenic contamination in groundwater generally results from naturally occurring As-bearing geological material rather than specific anthropogenic sources (Welch, et al., 2000, Smedley & Kinniburgh, 2002). Arsenic usually occurs as either arsenite (As(III), H₃AsO₃) or arsenate (As(V), H₂AsO₄⁻ and HAsO₄²⁻) in circumneutral aqueous environments. The mobility of As in the environment is highly influenced by microbial transformations, which affect As speciation (Oremland & Stolz, 2003). A large diversity of anaerobic microorganisms have been discovered that reduce As(V) to As(III) utilizing two biochemical systems. One system involves a cytoplasmic As(V) reductase (arsC) as part of a specific As efflux mechanism in resistant strains (Silver & Phung, 2005, Lloyd & Oremland, 2006). The other system involves a periplasmic dissimilatory As(V) reductase (arrA) whereby microorganisms gain energy for growth utilizing As(V) as a terminal electron acceptor (Malasarn, et al., 2004, Stolz, et al., 2006). The formation of As(III) from the microbial reduction of As(V) increases the public health risk, since As(III) is generally considered to be the more mobile and toxic form of As (Smedley & Kinniburgh, 2002, Sierra-Alvarez, et al., 2004).

The oxidation of As(III) to As(V) has the potential to immobilize soluble As and lower the risk of As contamination in groundwater due to the improved adsorption of As(V) on some metal (hydr)oxides such as those of aluminum (Lin & Wu, 2001, Hering & Dixit, 2005). Microorganisms from physiologically diverse groups, including both heterotrophs and autotrophs, can oxidize As(III) to As(V) in the presence of elemental oxygen (O₂) in various environments (Stolz, et al., 2006, Inskeep, et al., 2007). The heterotrophic oxidation of As(III) is catalyzed by periplasmic enzymes, most likely as a detoxification mechanism because energy for growth is not derived from the reaction (Silver & Phung, 2005). However, some chemolithotrophic As(III)-oxidizing bacteria can grow using the energy gained from the oxidation of As(III) (Santini, et al., 2000, Rhine, et al., 2007). As(III) oxidases are heterodimeric structures with two subunits containing molybdopterin and Rieske iron-sulfur domains (Santini & vanden Hoven, 2004, Stolz, et al., 2006, Rhine, et al., 2007). The full diversity and distribution of As(III) oxidase genes in environmentally relevant bacteria is not presently known (Silver & Phung, 2005). Inskeep et al. (Inskeep, et al., 2007) and Rhine et al. (Rhine, et al., 2007) recently designed primer sets to examine As(III) oxidase-like genes (aroA, asoA and aoxB) in environmental samples. The results suggest that a variety of aerobic As(III) oxidizers are widespread in As-contaminated environments.

Compared with aerobic As(III)-oxidizing microorganisms, little is known about anoxic As(III)-oxidizers. Although oxidation with O₂ is more favorable based on biochemical energetic considerations, alternative oxidants with lower reduction potentials are feasible for the oxidation of As(III). The data of Senn and Hemond (Senn & Hemond, 2002) were one of the first indications that anoxic As(III) oxidation occurs in the environment, based on the observation that seasonal changes in arsenic speciation were related to nitrate in anoxic lake water. Recently it has come to light that nitrate can be utilized by anaerobic microorganisms to gain energy from As(III) (Oremland, et al., 2002). A facultative chemolithoautotrophic arsenite oxidizing bacterium, Alkalilimnicola ehrlichi sp strain MLHE-1, capable of oxidizing As(III) to As(V) linked to partial denitrification of nitrate to nitrite, was isolated from an arsenic-containing soda lake in California (Mono Lake, CA) (Oremland, et al., 2002, Hoeft, et al., 2007). Subsequently, two anaerobic chemolithoautotrophic As(III)-oxidizing denitrifying bacteria were isolated from arsenic-contaminated soil and shown to oxidize As(III) with a stoichiometry consistent with the complete denitrification of nitrate to dinitrogen (N₂) gas (Rhine, et al., 2006) as shown in equation 1. The occurrence of the nitrous oxide gene, nosZ, was also demonstrated in the isolates (Rhine, et al., 2006). The two novel strains, DAO1 and DAO10, are phylogenetically similar to Azoarcus and Sinorhizobium on the basis of 16S rRNA sequences, respectively.

$$5{\mathrm{H}}_3{\text{AsO}}_3+2{{\text{NO}}_3}^{-}\to 5{{\text{HAsO}}_4}^{2-}+{\mathrm{N}}_2++8{\mathrm{H}}^{+}+{\mathrm{H}}_2\mathrm{O}$$ [eq 1]

In this study, we report on bacteria involved in the anoxic oxidation of As(III) linked to denitrification in three enrichment cultures (ECs) and one mixed culture (MC) that were established from sludges and sediments with no prior exposure to As (Sun, et al., 2008) as well as from a biofilm of an anoxic reactor exposed to nitrate and As(III). Complete denitrification in the ECs and the MC was confirmed by direct measurement of N₂ formation coupled with As(III) oxidation. The cultures were characterized by 16S rDNA gene clone library and fluorescent in situ hydridization (FISH) analysis (Amann, et al., 1995, Sanz & Kochling, 2007). The clone libraries were used to identify the predominant phylotypes in the cultures. FISH probes were then utilized to quantify the predominant phylotypes as fractions of the total cells. The results provide insights into the types of bacteria responsible for the widespread occurrence of anoxic As(III)-oxidizing denitrifying bacteria in the environment.

Material and Methods

Enrichment cultures

The three ECs differed based on the source of the original inoculum. The inoculum for enrichment culture 1 (EC1) was anaerobic effluent containing suspended biofilm from a laboratory-scale bioreactor (Department of Chemical and Environment Engineering, University of Arizona, Tucson, AZ) which coupled the anoxic oxidation of As(III) with denitrification. The bioreactor was fed with 3.75 mM As(III) and 6.4 mM NO₃⁻ using a 1 d hydraulic retention time. The original inoculum of the bioreactor was granular sludge obtained form a sulfoxidizing-denitrification reactor. The effluent sample was collected after 498 days of operation. The inoculum for enrichment culture 2 (EC2) was a methanogenic granular sludge obtained from a UASB treating alcohol distillery wastewater (Nedalco, Bergen op Zoom, The Netherlands). The inoculum for enrichment culture 3 (EC3) and for the mixed culture (MC) was derived from duck pond sediment obtained at the Agua Caliente Park (Tucson, Arizona). The culture volumes during the initial feeding were different from the routine volumes used for the ECs. The initial feed was performed in 500 mL serum bottles, with 250 mL media. EC1 received 10 mL of effluent (with resuspended biofilm), EC2 and EC3 received 5 g of wet sludge and sediment, respectively. The MC can be considered a subset of EC3 because it was inoculated with a colony cultured from a plate inoculated from the 5th transfer of EC3 (the colony used for inoculation was a single colony that upon later examination contained a mixed culture). The plate contained the same medium used for the ECs with 25 g L⁻¹ sterile noble agar (Difco Laboratories, Detroit, MI) and was incubated anaerobically for approximately 4 weeks under an N₂ atmosphere. Azoarcus sp strain EC3-pb1 and Diaphorobacter sp. strain MC-pb1 were isolated by dilution to extinction of EC3 and MC from. The highest dilution still providing a positive anoxic oxidation of As(III) with nitrate (10⁻⁷ in both cases) was used to isolate the pure cultures.

The three ECs, the MC and the pure cultures were maintained in the absence of O₂ in a basal mineral medium amended with 0.5 mM As(III) as electron donor, 10 mM NO₃⁻ as electron acceptor, and 8 mM HCO₃⁻ as the sole carbon source. The cultures were incubated in 160 mL serum flasks with a liquid volume of 100 mL that were sealed with butyl rubber stoppers. The medium and headspace was purged for 20 min with 80:20% N₂:CO₂. Cultures were serially transferred (5% vol:vol) to fresh medium after incubations of 2–3 weeks at 30°C and confirmation of As(III) oxidation (measured by As(V) formation). The enrichment process was continued for 25 transfers.

Medium composition

The standard As(III)/NO₃⁻-containing basal medium (A/N-BM) (pH 7.0–7.2) was prepared using ultra pure water (Milli-Q system; Millipore) and contained (mg/L): K₂HPO₄ (638); KH₂PO₄·2H₂O (1700); NH₄Cl (593); MgCl₂·6H₂O (166); MgSO₄·7H₂O (22); CaCl₂ (22), and 2 mL/L of a trace element solution containing (mg/L): FeC1₃·4 H₂O (2000); CoCl₂·6 H₂O (2000); MnCl₂·4 H₂O (500); AlCl₃·6 H₂O (90); CuCl₂·2H₂O (30); ZnCl₂ (50); H₃BO₃ (50); (NH₄)₆Mo₇O₂₄·4 H₂O (50); Na₂SeO₃·5 H₂O (100); NiCl₂·6 H₂0 (50); EDTA (1000); resazurin (200); HCl 36% (1 mL). The basal medium was amended with 8 mM HCO₃⁻ (NaHCO₃), 0.5 mM As(III) (NaAsO₂) and 10 mM NO₃⁻ (KNO₃). The basal medium with NO₃⁻ was sterilized by autoclave, while NaHCO₃ and As(III) were sterilized using membrane filters (0.2 μm).

Experimental incubations

Once the cultures were established, they were used to evaluate the time course of As(III) oxidation under denitrifying conditions using the A/N-BM medium. Samples were periodically removed for measurement of As(III), As(V) and NO₃⁻. Various controls were utilized based on the requirements of each experiment. Abiotic controls were prepared without adding enrichment culture inoculum. Heat killed controls were prepared by autoclaving the flasks added with inoculum for 20 min at 121°C, on three consecutive days. Controls lacking As(III) were included to measure endogenous consumption of NO₃⁻. All assays were conducted in triplicate. To avoid the contamination of carry-over NO₃⁻ from old cultures to fresh medium, the cultures (5% volume of previous culture) were centrifuged in sterilized eppendorf tubes at 10,000 rpm for 10 min, the pellets were collected and washed into sterilized MiliQ water for two cycles. The pellets were resuspended into same volume of sterilized MiliQ water and transferred to the experiments.

In one experiment the impact of low concentrations of organic carbon supplements were evaluated. In this case the A/N-BM medium was supplemented with yeast extract (1.0 mg l⁻¹) or pyruvate (1.7 mg l⁻¹). The experiment was inoculated with EC1 at 1% (v/v).

Most probable number (MPN)

MPN assays were performed in A/N-BM. Culture samples (1 mL) were taken after 23 transfers, were homogenized by vortexing, and were serially diluted in 10-fold increments to 10⁻⁹ in sterile medium under anaerobic conditions. Each dilution was set up with five replicate MPN tubes. The tubes were incubated in an orbital shaker (110 rpm) at 30°C under anaerobic conditions with N₂/CO₂ (80:20) in the headspace. After 3–4 weeks incubation, samples from each tube were removed and analyzed for As(V). Conversion of >80% of the As(III) to As(V) was considered as a positive tube. Finally an MPN table for five tubes (APHA, 1999) was utilized to enumerate the number of bacteria with As(III)-oxidizing capability.

Batch assays to determine the terminal product of denitrification

End products of denitrification were measured by monitoring gaseous nitrogen species in the headspace of bioassays flushed with He/CO₂ (80:20, v:v) in lieu of N₂/CO₂ as described above. Gaseous nitrogen measured in As(III) spiked samples (3.0 mM) was compared to endogenous controls. Microbial reduction of NO₃⁻ coupled to As(III) oxidation was assessed in shaken batch bioassays inoculated with EC1 and EC3. The NO₃⁻ concentration utilized was 2.5 mM for the experiments. Samples were supplied 50 g L⁻¹ of activated aluminum (AA-400G, Alcan Bauxite and Alumina) adsorbents to lower the effective aqueous concentration of As(III) so as to minimize its toxicity. As(III) was supplied at 3.0 mM; however by using this procedure, the aqueous equilibrium concentration of As(III) was 0.59 mM. The high concentration of As(III) supplied was required to have enough electron equivalents to measure N₂ production properly. Headspace samples were analyzed periodically for N₂ and N₂O with a pressure lock gas tight syringe (1710RN, 100 μl (22s/2″/2), Hamilton Reno, Nevada USA) to confirm denitrification. Liquid samples were analyzed for NO₃⁻ and NO₂⁻. Flushed headspace controls incubated with just water were monitored to measure background levels of N₂. The background level was low.

Analytical methods

Samples (1 mL) were taken from sealed anaerobic serum flasks by piercing the stoppers using sterile 1.0 mL syringes with 16-gauge needles. All liquid samples were centrifuged (10 min, 10,000 rpm) or membrane filtered (0.45 μm) immediately after sampling and stored in polypropylene vials. Samples for As analysis were stored at −20°C. As(III) and As(V) were analyzed by ion chromatography–inductively coupled plasma–mass spectroscopy (IC-ICP-MS). The system consists of an IC (Agilent 1100) and an ICP-MS (Agilent 7500a) with a Babington nebulizer as the detector. The operating parameters were as follows: Rf power 1500 watts, plasma gas flow 15 L/min, carrier flow 1.2 L/min, arsenic was measured at 75 m/z and terbium (IS) measured at m/z 159. The injection volume was 10 μL. The detection limit for the various arsenic species was 0.1 μg/L.

Nitrate and nitrite were analyzed by suppressed conductivity ion chromatography using a Dionex 500 system (Sunnyvale, CA, USA) fitted with a Dionex IonPac AS11 analytical column (4 × 250 mm) a AG16 guard column (4 mm × 40 mm). During each injection the eluent (KOH) used was 20 mM for 20 min. The same procedure also was suitable as an alternative analysis of As(V).

N₂ and N₂O were analyzed using a Hewlett Packard 5890 Series II gas chromatograph fitted with a Carboxen^™ 1010 Plot column (30 m × 0.32 mm) and a thermal conductivity detector. The temperature of the column, the injector port and the detector were 220, 110 and 100 °C, respectively. Helium was used as the carrier gas and the injection volume was 100 μL.

Other analytical determinations (e.g., pH, TSS, VSS, etc.) were conducted according to Standard Methods (APHA, 1999).

16S rRNA gene clone libraries

Community genomic DNA was extracted from 5 mL samples taken from each EC and the MC by a modification of the Current Protocols extraction method for Genomic DNA from bacteria (Ausubel, et al., 1995). Briefly, cells were pelleted and resuspended in TE buffer (10 mM Tris Cl, 1 mM EDTA, pH 8.0) and subjected to three freeze-thaw cycles in liquid nitrogen and boiling water. Cells were then lysed and the DNA purified according to the protocol. Before the extraction all sterile tubes, caps, and solutions used in the protocol were exposed to UV light for 10 min in a UVC-508 Ultraviolet Crosslinker (Ultra-Lum, Claremont, CA) to remove any potential DNA contamination. The extracted DNA was stored at −20°C. Extraction blanks were processed in parallel throughout the full procedure as negative controls to evaluate potential DNA contamination from reagents. The DNA was quantified with TBS-380 Mini-Fluorometer (Tuner BioSystems, Sunnyvale, CA) Using Molecular Probes’ PicoGreen dsDNA Quantitation Reagent (Molecular Probes, Inc., Eugene, Oregon).

The 16S rRNA gene was PCR amplified from community DNA extracts using universal primers 27F and 1492R (Lane, 1991). Each 25-μL reaction contained 0.5 mM of each primer, 0.2 mM of each dNTP, 1x buffer consisting of 10 mM Tris-HCl, 50 mM KCl, 2.0 mM MgCl₂ (pH 8.3), 5% dimethyl sulfoxide (DMSO), 0.5 U of Taq DNA polymerase (Roche, Indianapolis, IN), and 2.5 μL of DNA extract. The DNA was amplified with an initial denaturation step of 95°C for 5 min followed by 30 cycles of: 94°C for 1 min, 60°C for 1 min and 72°C for 1.15 min, with a final extension for 10 min at 72°C in a Perkin Elmer GeneAmp PCR System 2400 (PerkinElmer Inc., Boston, MA). The PCR products were visualized on a 1% GenePure LE agarose gel (Intermountain Scientific Corp., Kaysville, UT) and then purified using a Quick PCR purification kit (Qiagen, Chatsworth, CA).

The purified PCR products were cloned into plasmid vector pCR^®2.1-TOPO^® using the TOPO TA cloning system (Invitrogen, Carlsbad, CA) according to the protocol described by the manufacturer. The control TOPO^® Cloning reaction was performed using the reagent included in the kit in parallel with real samples. The plasmid DNA was purified using PureLink^™ Quick Plasmid Miniprep Kit (Invitrogen, Carlsbad, CA) according to the protocol described by the manufacturer. Purified plasmid DNA was sequenced using four primers including the vector primers T7 (5′-TAATACGACTCACTATAGGG-3′) and M13R (5′-AGGAAACAGCTATGACCATG-3′) and universal internal primers 518F and 1070R (Lane, 1991). Nearly full-length 16S rRNA gene sequences were obtained using an Applied Biosystems 3730xl DNA Analyzer (Applied Biosystems, Foster City, CA) (University of Arizona Research Laboratory Genomic Analysis and Technology Core, Tucson, AZ).

The number of clones analyzed for each culture was determined using a rarefaction curve to estimate the diversity (Analytic Rarefaction 1.3, UGA Stratigraphy Lab, University of Georgia, Atlanta, GA). An exponential model, y = a × [1−exp(−b × x)], was used based on the formulation of Tipper (Tipper, 1979) to fit the clone distribution data. Sequences were aligned by using FAKII Fragment Assembly Kernel (FAKtory, Biotechnology Computing Facility, University of Arizona, Tucson, AZ) and compared to known sequences through the Basic Local Alignment Search Tool (BLAST) (http://www.ncbi.nlm.nih.gov/BLAST/). The clones were clustered into phylotypes on the basis of having sequence similarities of >99. 5%. ARB program package (Ludwig, et al., 2004) was used for the phylogenetic analysis. Sequence data were aligned and a tree, including 16S rRNA gene sequences from reference bacterial strains (GenBank) and unique phylotypes recovered from each of the three ECs and the MC, was constructed using the Maximum Likelihood algorithm (Felsenstein, 1981) with a bacterial filter of the fastDNAml program (Olsen, et al., 1994) in the ARB program package. Tree confidence was tested using the Hasegawa method (Hasegawa, et al., 1985). The sequences obtained in this work (of selected clones representing each phylotype obtained in each culture) have been deposited in the GenBank data base with accession numbers: EC1-1 (EU708507); EC2-2 (EU741793); MC-1 (EU741795); EC3-10 (EU708501); EC1-12 (EU708506); EC1-7 (EU708505); EC3-1 (EU708500); EC2-7 (EU741794); EC1-17 (EU708503); EC1-10 (EU708508); EC2-2 (EU741793); EC1-33 (EU708504); MC-9 (EU741796); and EC3-11 (EU708502).

DNA extraction of pure cultures and sequencing of their 16S rRNA genes

To identify the pure culture isolates, the DNA was extracted from washed pellets in a microcentrifuge tube resuspended in 40 μl of sterile miliQ water. The resuspended pellet was subjected to 3 cycles of boiling (10 sec) and freezing in liquid N₂ followed by 15 minutes of boiling which was halted by placing the microcentrifuge tubes on ice. The PCR and sequencing were conducted as described above for the gene clone libraries. The GenBank accession numbers for 16S rRNA genes of the pure cultures were FJ514096 and FJ514095 for Azoarcus sp strain EC3-pb1 and Diaphorobacter sp. strain MC-pb1, respectively.

FISH analysis

In order to quantify the bacterial composition of enrichment cultures, 10 mL of each culture were fixed with formaldehyde (4% in phosphate saline buffer, (PBS)) for 4 h. After that, the samples were washed with PBS and stored in a mixture of PBS:ethanol 50:50 v:v at −20°C until they were hybridized. 10 μl of each sample were placed on to multi-well slides. All samples were further dehydrated by immersion in 50, 80, and 100% ethanol solutions for 3 min each time. FISH was applied according to protocols described by Amann et al (1990, 1995). Samples were hybridized with the probes (Table S1) Azo644, specific for Azoarcus (Hess, et al., 1997); DEN220, which hybridizes with the Comamonadaceae family (Ginige, et al., 2005) (to target members of the genera Alicycliphilus and Diaphorobacter within the Acidovorax cluster); and the universal probe for Bacteria domain Eub338 (Amann, et al., 1990). The probes were purchased from Biomers (Ulm, Germany). The formamide concentration used for hybridization (2 h at 46°C) was 20% (Azo644), 40% (DEN220) and 35% (Eub338) and the NaCl for washing (15 min at 48°C) was 215 mM (Azo644), 46 mM (DEN220) and 70 mM (Eub338). After hybridization the samples were stained with 4′, 6′-diamin phenylindol (DAPI, 1 mg/mL) in order to determine the total cells present. Twenty randomly selected fields (approximately 1000 DAPI-stained cells) were analyzed using a Zeiss Axiovert 200 fluorescence inverted microscope.

Staining of polyhydroxbutryates (PHB)

Evidence for the presence PHB and related polyhydroxyalkanate polymers was done according to the Nile Red method. A stock solution of Nile Red was prepared (1 mg ml⁻¹) in 100% DMSO (Degelau, et al., 1995, Spiekermann, et al., 1999). Before staining, a bacterial cell pellet from 30 ml ECs were collected by centrifuge 10 min (12,000 rpm), and resuspended in saline solution (0.9% sodium chloride). Heat-fixed smears of the bacterial cells were stained with the Nile Red stock solution at 55°C for 30 min in the dark (Ostle & Holt, 1982, Degelau, et al., 1995). After being stained, the slides were washed with MiliQ water to remove excess stain for 1 min. The stained smear was blotted dry with bibulous paper, remoistened with MiliQ water, and covered with a No. 1 glass cover slip. The slides were examined by using a Leitz Diaplan microscope fluorescence phase contrast system (Leitz Wetzlar, Germany) with 100 × 1.32 oil-immersion ICT PL Fluotar OEL objective.

Results

Anoxic As(III) conversion by enrichment cultures

The three ECs and the MC were incubated under anoxic conditions with As(III) either in the presence or absence of NO₃⁻ (Fig. 1). In treatments incubated with NO₃⁻, As(V) formation started after approximately 1 d and the reaction came to completion after 4–6 d. The maximum reaction rate, calculated from the formation of As(V), was 221 ± 9, 221 ± 8, 175 ± 12, and 138 ± 1 μMd⁻¹ for EC1, EC2, EC3, and MC, respectively. The reactions were dependent on the presence of NO₃⁻ as evidenced by the lack of any significant conversion in incubations without added NO₃⁻ (Fig. 1). The reactions did not occur in bottles receiving the medium with As(III) and NO₃⁻ but lacking inoculum (Fig. 1) or with heat-killed inoculum (data not shown).

Fig. 1.

Formation of As(V) by cultures under denitrifying condition. Incubated with 0.5 mM As(III) and 10 mM NO₃⁻: EC1 (▲), EC2 (■), EC3 (●), and MC (◆) Control with only 0.5 mM As(III) and no NO₃⁻: EC1 (△), EC2 (□), EC3 (○), and MC (◇) Abiotic control (X).

MPN analysis of the As(III)-oxidizing denitrifying populations performed after the 23^rd transfer revealed that 4.9×10⁷, 2.2×10⁸, 1.7×10⁷, and 2.3×10⁷ cells were present per mL of EC1, EC2, EC3 and MC, respectively; corresponding to 3.3 × 10¹² to 4.4 × 10¹⁴ cells produced per mol of As(V) formed.

EC1 was tested for its ability to utilize low concentrations of organic carbon sources for growth under denitrifying conditions. The treatments amended with yeast extract and pyruvate (1.0 and 1.7 mg l⁻¹, respectively) showed no significant difference in the rate or extent of As(III) oxidation compared with the autotrophic oxidation conditions without any organic carbon added (data not shown).

Experiments performed to determine whether NO₃⁻ consumption and N₂ formation are linked to As(III) oxidation are shown in Fig. 2. In EC cultures amended with both As(III) and NO₃⁻, N₂ was formed as an end product (Fig. 2A), NO₃⁻ was consumed (Fig. 2B), and NO₂⁻ was transiently observed with its appearance and disappearance correlating closely with the consumption of NO₃⁻ and formation of N₂ respectively (Fig. 2C). In the absence of As(III) and in the abiotic controls, no consumption of NO₃⁻ or appearance of either NO₂⁻ or N₂ was observed. The molar ratios of As(III) consumption to the NO₃⁻ consumption and N₂-N formation (corrected for background endogenous denitrification) are provided in Table 1. The molar ratios of As(III) fed to NO₃⁻ consumption and N₂-N formation from the adsorbent amended EC1 and EC3 enrichment cultures were close to the theoretical ratio of 2.5 expected for As(III) oxidation linked to complete denitrification to N₂ (equation 1).

Fig. 2.

The influence of As(III) on the formation of N₂-N (Panel A), consumption of NO₃⁻ (Panel B) and accumulation of NO₂⁻ (Panel C) by EC1 (squares) and EC3 (triangles) cultures under denitrifying conditions. Legend: full treatment with 3 mM As(III) and 2.5 mM NO₃⁻ (■ and ▲), endogenous control with only 2.5 mM NO₃⁻ (□ and △), and abiotic control (●). All assays contained activated alumina (50 g L⁻¹).

Table 1.

Summary of denitrification linked to As(III) oxidation when adsorbed onto AA (50 g l⁻¹)

Parameters	EC1		EC3
	As(III)+NO3−	NO3− only	As(III)+NO3−	NO3− only
As(III) fed (mM)	3.10±0.02	—	3.10±0.02	—
NO3− consumed (mM)	1.41±0.07	0.18±0.01	1.56±0.23	0.12±0.07
Corrected† NO3− consumed (mM)	1.23±0.07	—	1.44±0.18	—
N2-N formed (mmol/Lliquid)	1.24±0.06	0.02±0.01	1.18±0.03	0.01±0.02
Corrected N2-N formed (mmol/Lliquid)	1.21±0.06	—	1.17±0.02	—
Corrected N2-N formed/corrected NO3− consumed	0.99±0.06	—	1.12±0.13	—
Mol As(III) fed/corrected mol NO3− consumed	2.53±0.17	—	2.17±0.24	—
Mol As(III) fed/corrected mol N2-N formed	2.56±0.14	—	2.64±0.03	—

^†Corrected for the endogenous nitrate consumption or N₂-N formation determined in the nitrate treatment lacking added As(III)

Community composition of enrichment cultures

The composition and structure of the microbial community in the ECs and MC were analyzed by preparing a 16S rRNA gene clone library for each. Rarefaction analysis for each clone library (Figure S1) suggested that 46, 18, 21 and 20 clones were sufficient for capturing the community composition in the clone libraries corresponding to EC1, EC2, EC3, and MC, respectively. A total of eight unique phylotypes were obtained from the four cultures. Of the eight unique phylotypes recovered, 6, 3, 3, and 2, were found in EC1, EC2, EC3, and MC respectively (Fig. 3). Note that the 2 phylotypes found in the MC were also recovered from EC3 which was expected since the colony used to inoculate the MC was from the EC3 (Fig. 4).

Fig. 3.

Phylogenetic distributions in the four cultures. Diagrams show the relative abundance of 16S rRNA phylotypes of clones from each culture with a total of 8 unique phylotyptes found in the study. Out of these 8 unique phylotyptes, 6, 3, 3, and 2 were found in EC1, EC2, EC3, and the MC respectively.

Fig. 4.

Phylogenetic relationships of the EC and MC clones recovered in this study. Maximum likelihood analysis was used to generate the tree from 16S rRNA gene sequences of bacterial strains and unique phylotypes represented by clones recovered from the four cultures labeled with EC#-# (the first # refers to the enrichment culture, the second # refers to the clone number), or MC-# (where the # refers to the clone number). Genbank accession numbers are indicated in parentheses. Confidence values for tree topology (1,000 replicates) are given for nodes with ≥ 50% support.

The 105 clones analyzed in this study fell into four phylogenetic divisions, α-Proteobacteria β-Proteobacteria, γ-Proteobacteria, and Flavobacteria accounting for 1.9%, 84.8%, 9.5%, and 3.8% of all the clones from the four cultures, respectively. The phylogenetic relationships among the phylotypes recovered are shown in Figure 4. The β-Proteobacteria accounted for a large majority of all the clones and four of the eight phylotypes recovered were most closely related to members of this group (Azoarcus, Alicycliphilus, Diaphorobacter and unclassified Burkholderiales). One phylotype belonged to the α-Proteobacteria and was most closely related to Rhizobiaceae. Two phylotypes belonged to the γ-Proteobacteria and were most closely related to Stenotrophomonas and Dokdonella, and the remaining phylotype was affiliated with Flavobacteria and was most closely related to Kaistella.

Three out of the eight phylotypes identified in the ECs and MC dominated in terms of clones analyzed. One of these phylotypes (including clones EC1-7, EC3-1, EC2-7) was 100% likely to belong to the genus Azoarcus (according to the RDP Classifier (Cole, et al., 2005)) and was found in the three ECs (Fig. 3 and 4). The Azoarcus phylotype accounted for 22 to 62% of the clones in the clone libraries of the ECs. The remaining two dominant phylotypes were related to the genera Alicycliphilus (clones EC1-1, EC2-2; 98%; RDP Classifier) or Diaphorobacter (clones MC-1, 99%; EC3-10, 96%; RDP classifier) which both closely related to Acidovorax belonging to the Comamonadaceae. All of the ECs and the MC contained one of the two phylotypes related to the Comamonadaceae (Fig. 3 and 4). The ECs originating from biofilm and sludge (EC1 and EC2) contained the phylotype related to the Alicycliphilus genus of this cluster, which accounted for 41 to 61% of the clones. The cultures originating from pond sediment contained the phylotype most closely related to Diaphorobacter, which accounted for 29 and 90% of the clones in EC3 and MC, respectively.

The remaining five phylotypes were present as small fractions of the clones examined. The phylotype most closely related to Dokdonella (clone EC1-10; 100%; RDP Classifier) accounted for 11 to 17% of the clones in the the ECs originating from biofilms and sludge (EC1 and EC2). The phylotype most closely related to Kaistella (clones EC3-11, MC-9; 100%; RDP Classifier) accounted for about 10% of the pond sediment-derived cultures (EC3 and MC). The phylotypes most closely related to Stenotrophomonas (clone EC1-17; 100%; RDP Classifier), Rhizobiaceae (clone EC1-33; 100%; RDP Classifier), and unclassified Burkholderiales (clone EC1-12; 94%; RDP Classifier), were from the EC started with the biofilm incolum (EC1) and each accounted for about 4% of all the clones in that EC.

FISH analysis showed that the cells detected by the eubacteria probe, Eub338, ranged from 75 to 80% of the total DAPI-stained cells in the different ECs and MC (Fig. 5). The Azoarus-specific probe, Azo644, hybridized with 38 to 72% of DAPI-stained cells in EC1, EC2 and EC3. As expected, there was no hybridization of Azo644 in the MC which did not contain Azoarcus-like clones. The Comamonadaceae-specific DEN220 probe (used to target Alicycliphilus and Diaphorobacter) detected a smaller proportion of the total cells ranging from 4 to 20% of total DAPI-stained cells in the four cultures.

Fig. 5.

FISH analysis of the EC and MC communities using universal (Eub338) and group-specific (Azo644 and DEN220) probes. Results are presented as percentages relative to DAPI counts (set at 100%)

It should be noted that the FISH results and clone library results are not identical. For example in EC1, the clone library results indicate that the Azoarcus and Comamonadaceae clones are approximately equal in number. In contrast, the FISH results indicate that the Azoarcus is strongly dominant. It should be recognized that both of these techniques have associated biases which likely explain this discrepancy (Hongoh, et al., 2003, Alonso-Saez, et al., 2007). However, these techniques, used in combination are in agreement as to the presence or absence of Azoarcus and Comamonadaceae in the ECs and MC. Further, these techniques provide strong evidence that in addition to Azoarcus members of the Comamonadaceae are capable of carrying out As(III) oxidation under anoxic conditions.

Anoxic oxidation of As(III) by isolated pure cultures

Bacteria representing the dominant clones from the clone library were isolated to confirm their ability to oxidize As(III) in the absence of O₂. The two dominant phylotypes from EC3 were isolated by dilution to extinction with the MPN method. Azoarcus sp. strain EC3-pb1 (99.5% homology with clone EC3-1) was isolated from EC3. Diaphorobacter sp. strain MC-pb1 (99.7% homology with clone MC-1) was isolated as a reperesentative of the Comamonadaceae phylotype from MC (a subset of EC3). The isolated pure bacterial cultures were maintained for 6 serial transfers of 5% (v/v) with an incubation period of approximately 2 weeks with a medium containing 0.5 mM As(III) and 10 mM NO₃⁻. Before each transfer, the oxidation of As(III) to As(V) was confirmed. In the final transfer a triplicate experiment was set up with controls to confirm the dependence of the oxidation on the presence of nitrate and live cells. Treatments with live cells of Azoarcus sp. strain EC3-pb1 and Diaphorobacter sp. strain MC-pb1 converted As(III) to a measured concentration of 0.52±0.08 and 0.51±0.06 mM As(V) after 10 d, respectively. Controls lacking cells or with heat killed cells of the isolates converted less than 0.025 mM of the As(III). Likewise there was less than 0.025 mM conversion in controls with live cells but lacking nitrate addition.

Staining for PHB

Since Azoarcus is known for the production of the food storage polymer, PHB (Reinhold-Hurek & Hurek, 2006), the three ECs containing Azoarcus were tested for the presence of PHB by testing for fluorescence after staining with Nile Red dye. The ECs harvested on day 10 were dominated by fluorescing cells accounting for approximately 80 to 90% of the cells observed. Starved cell controls (ECs incubated 2 weeks at 30° C followed by 4 months storage at 4° C and subsequently reincubated for 2 weeks at 25°C), which presumably would have consumed PHB, had only 5 to 10% of the cells fluorescing after Nile Red staining. Controls without cells that were stained showed no fluorescence.

Discussion

Evidence of As(III) oxidation linked to denitrification

Multiple lines of evidence demonstrate that the bacterial communities in the ECs and MC evaluated in this study linked the anoxic oxidation of As(III) to denitrification. First, the formation of As(V) from As(III) was dependent on the presence of NO₃⁻ and inoculum. No oxidation occurred in controls lacking either inoculum or NO₃⁻. Second, the production of N₂ was linked to the addition of As(III) to the cultures and the yield of N₂ corresponded to the expected stoichiometry from the electron equivalents in As(III). Third, bacteria appeared to benefit from the process as evidenced by the large number of transfers (25×) performed in this study. Likewise, the MPN assay indicated high cell densities of microorganisms were produced with up to 4 × 10¹⁴ cells mol⁻¹ As(V) formed. The enrichment cultures were sustained without addition of organic C, and small additions of organic C (yeast extract or pyruvate) did not stimulate growth. These findings indicate the cultures are likely chemolithoautotrophic, obtaining energy from As(III) and most likely obtaining carbon from added HCO₃⁻/CO₂.

The As(III) oxidizing bacterial community – Azoarcus and Comamonadaceae

One phylotype belonging to the β-Proteobacteria and related to the genus Azoarcus constituted a large fraction of the clones analyzed in all the ECs studied. FISH analysis supported the dominance of this genus, 38 to 72% of the cells in these cultures were detected with the Azoarcus probe. The Azoarcus phylotype was closely related to Azoarcus strains PbN1 (98%) and EbN1 (97%) (Rabus & Widdel, 1995) (Figure 4). These strains are known for their ability to degrade propyl- and ethylbenzene (Szaleniec, et al., 2007). In general, the Azoarcus genus is known for the ability to degrade aromatic substrates under anaerobic denitrifying conditions (Reinhold-Hurek & Hurek, 2006). Azoarcus strain EbN1 and a number of closely related aromatic-degrading strains within the Azoarcus/Thauera cluster are being proposed as a new genus, “Aromatoleum”. In fact, EbN1 is known as Aromatoleum aromaticum, a metabolically versatile representative of the group capable of degrading a wide variety of aromatic and non-aromatic compounds under both anoxic and aerobic conditions (Szaleniec, et al., 2007, Wohlbrand, et al., 2007).

The Azoarcus clones from this study had 96% similarity to Azoarcus strain DAO1, which is an isolate from an arsenic-contaminated site and is known to link As(III) oxidation to denitrification (Rhine, et al., 2006). Therefore, there is a precedent within the genus Azoarcus for the anoxic oxidation of As(III). Azoarcus sp. strain EC3-pb1, isolated from EC3 of this study, was also shown to be responsible for anoxic As(III) oxidation. Azoarcus strain DAO1 does not contain a gene that hybridizes with degenerate primers designed from known sequences of arsenite oxidases (Rhine, et al., 2007). Nonetheless, it is interesting to note that the key enzyme produced by “Aromatoleum aromaticum” strain EbN1, ethylbenzene dehydrogenase (Johnson, et al., 2001, Kloer, et al., 2006, Szaleniec, et al., 2007) is related to arsenite oxidases (Inskeep, et al., 2007, Rhine, et al., 2007) since both contain molybdate cofactors and are members of the DMSO family.

The genome of “Aromatoleum aromaticum” strain EbN1 (97% similarity with EC1-7 and EC3-1) has been sequenced (Rabus, 2005, Rabus, et al., 2005). The genome contains putative genes for arsenic resistance in the ars operon family (arsC, arsD, arsR, and arsA) (Rabus, et al., 2005). The ars operon could potentially explain the ability of the Azoarcus strains in the ECs to be viable in the presence of 0.5 mM arsenic (in some experiments 1.0 mM). The genome also contains a complete set of denitrification genes (e.g. NarG, NirS, NorC, and NosZ) (Rabus, 2005) which could account for the observed production of N₂ from NO₃⁻.

All of the cultures examined in this study (ECs and MC) contained clones representing one of two phylotypes related to a cluster of genera (Acidovorax, Alicycliphilus and Diaphorobacter) within the family Comamonadaceae which also belongs to the β-Proteobacteria (Fig. 4). The fraction of clones belonging to this group ranged from 29 to 90% depending on the culture. FISH analysis with the DEN220 probe confirmed the presence of Comamonadaceae but suggested they constitute a smaller fraction of the community (Fig. 5). One Comamonadaceae cluster phylotype recovered from sludge-derived ECs (EC1 and EC2) was related to the genus Alicycliphilus. This clones have 99% similarity to an uncultured clone from activated sludge, Alicycliphilus strain R-24611. These clones also have 99% similarity to several other Acidovorax isolates. An example is an H₂-consuming nitrate reducing isolate Acidovorax strain Ic31 from a hydrogenotrophic denitrification batch reactor (Vasiliadou, et al., 2006), indicating a precedent for autotrophic denitrification in closely related species. Another example is Acidovorax avenae strain C1 which was isolated from a low O₂ phenol-degrading nitrate-reducing culture (Baek, et al., 2003).

The second Comamonadaceae cluster phylotype was obtained from the pond sediment-derived cultures (EC3 and MC) and was related to the genus Diaphorobacter. A number of published isolates belonging to species Diaphorobacter nitroreducens have 99% similarity with clones EC3-10 and MC-1 recovered in this study. These are known for their ability to degrade polyhydroxyalkanoic acid bacterial energy storage polymer, poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV), under denitrifying conditions (Khan & Hiraishi, 2001, Khan, et al., 2002, Khan & Hiraishi, 2002 ). Interestingly, the genus Azoarcus is known for the production of poly-β-hydroxybutyrate (PHB) as energy storage polymers, which accumulate in cells as granules (Reinhold-Hurek & Hurek, 2006, Serafim, et al., 2006). In this study, Nile Red dye staining confirmed the presence of PHB in As(III)-grown cells of the ECs. Thus, the PHB produced by Azoarcus could potentially benefit the Comamonadaceae members of the community related to Diaphorobacter by serving as a source of carbon and energy suggesting one possible interaction between two important members of the EC communities.

Acidovorax sp. JS42, which is a well known aerobic nitroaromatic degrading bacterium (Lessner, et al., 2003), also has 99% similarity with clones EC3-10 and MC-1. The genome of Acidovorax sp. JS42 has been sequenced and the genome contains ars operon genes (arsA, arsR and arsC) as well as a gene for an arsenical-resistance protein which could explain the ability of these microorganisms to tolerate arsenic. The genome also contains genes involved in denitrification (nitrate reductase, nitrite reductase, nitric oxide reductase large subunit, and nitrous oxide reductase) as well as one gene for an esterase for PHB depolymerization.

There is no precedent for anoxic arsenite oxidation in the Comamonadaceae cluster. In the MC, Diaphorobacter was dominant which indicates that it is likely involved in using As(III) as an energy source. Diaphorobacter sp. strain MC-pb1, isolated from the MC was shown here to be responsible for nitrate-dependent As(III) oxidation in the absence of O₂, confirming this suspected role. Recently, an aerobic As(III) oxidizing strain (GW2) was isolated in Chinese sediments that belongs to Acidovorax sp (Fan, et al., 2008) and it was shown to possess an arsenic oxidase gene. The Diaphorobacter-related 16S RNA gene clones MC-1 and EC4-10 of this study had 95% homology with Acidovorax sp. GW2. Strains in the Comamonadaceae cluster are also known for their ability to link Fe(II) oxidation to dentrification (Straub, et al., 2004).

The As(III) oxidizing bacterial community – less dominant members

A phylotype belonging to the γ-Proteobacteria was recovered from sludge-derived ECs (EC1 and EC2). This phylotype has 97% similarity to Dokdonella fugitive, an isolate obtained from potting soil (Cunha, et al., 2006) (Figure 4). A second phylotype belonging to the γ-Proteobacteria was recovered only from EC1. This phylotype (clone EC1-17) has 99% similarity to Stenotrophomonas sp. YC-1 (Figure 4), isolated from an organophosphorous pesticide manufacturing wastewater treatment plant (Yang, et al., 2006). Two additional isolates which have 99% similarity to EC1-17 include Stenotrophomonas acidaminiphila sp. nov. which is a strictly aerobic bacterium isolated from an upflow anaerobic sludge blanket reactor (Assih, et al., 2002) and Stenotrophomonas sp. BO, which was isolated from a microaerophilic denitrification reactor fed methane (Costa, et al., 2000). The latter isolate can carry out denitrification with acetate to predominantly N₂O and to a lesser extent, N₂. Stenotrophomonas nitritireducens L2 which has a 98% similarity to EC1-17 was isolated from a biofilter treating ammonia (Finkmann, et al., 2000) and the strain was able to reduce NO₂⁻ to N₂O but could not reduce NO₃⁻.

The phylotype belonging to the Flavobacteria was recovered only from the pond sediment-derived cultures (EC3 and MC). This phylotype contains 2 clones (EC3-11 and MC-9), that have 99% similarity with Kaistella koreensis strain Chj707 which was isolated from a fresh water stream in Korea and is known to reduce nitrate (Kim, et al., 2004).

A phylotype belonging to the α-Proteobacteria was recovered from EC1. This phylotype (EC1-33) has 99% similarity to Amorphomonas oryzae strain B26 in the family Rhizobiaceae (Fig. 4). Amorphomonas oryzae is a free-living nitrogen fixing bacterium isolated from rice roots. There are also closely related Rhizobium-Sinorhizobium strains that are known for their ability for aerobic chemolithoautotrophic oxidation of arsenite such as strains NT-2, NT-3 and NT-4 which have 96% similarity with EC1-33 (Santini, et al., 2002). Other bacteria that are related included aerobic As(III)-oxidizing strains Rhizobium NT-26 (Santini, et al., 2000) Sinorhizobium sp. strain GW3 (EF550173) and Agrobacterium strain GW4 (EF550174). The latter two strains were isolated from sediments naturally enriched with arsenic (Fan, et al., 2008). There is also a precedent in the “Rhizobium” cluster for anoxic oxidation of As(III). Sinorhizobium strain DAO10 (99.8% similarity with EC1-33) is reported to be As(III)-oxidizing denitrifying bacterium (Rhine, et al., 2006). DAO10 was shown to contain an arsenite oxidase like gene with 73.6% amino acid similarity with arsenite oxidase (aroA) from NT-26 (Rhine, et al., 2007).

The last phylotype recovered belongs to the β-Proteobacteria and was recovered from EC1. The clone EC1-12 has 97 to 98% similarity to uncultured bacterial clones (AY548933 and AY548944) from the order Burkholderiales (Fig. 4) that were obtained from anaerobic ammonium oxidizing (Anammox) microbial communities. The family Oxalobacteraceae within the Burkholderiales contains at least one species, Herminiimonas arsenicoxydans strain ULPAs1^T (AY728038) which is known for aerobic As(III) oxidation and contains arsenite oxidase genes (Muller, et al., 2006).

Conclusion

Clone library and FISH analysis suggest that phylotypes representing Azoarcus and Comamonadaceae are the main As(III) oxidizers in the microbial communities of the three ECs studied. For Azoarcus, this conclusion is supported by the fact that a related isolate from this genus (DAO1) can link As(III) oxidation to denitrification (Rhine, et al., 2006) and by the fact that Azoarcus sp. strain EC3-pb1 isolated in this study could also carry out the reaction. Furthermore, the closely related isolate EbN1 contains arsenic resistance genes as well as a full set of denitrification genes. This study is the first report of anoxic As(III) oxidation by members of the Comamonadaceae. The fact that As(III) was oxidized in the MC, which contained no Azoarcus but contained an enrichment of Diaphorobacter (Comamonadaceae family) implies that the phylotype related to Diaphorobacter is responsible for anoxic As(III) oxidation in the MC. This was confirmed by isolating Diaphorobacter sp. strain MC-pb1 which oxidized As(III) with nitrate. The occurrence of aerobic As(III) oxidizers in the Comamonadaceae cluster (Fan, et al., 2008), together with the occurrence of arsenic resistance- and denitrification genes in the genome of the closely related Acidovorax sp. strain JS42, support this potential role.