Microbial Populations Identified by Fluorescence In Situ Hybridization ...

1 downloads 0 Views 271KB Size Report
Jul 6, 2006 - Microorganisms are an integral part of the biogeochemical pro- cesses in wetlands, yet microbial communities in sediments within constructed ...
Published online July 6, 2006

Microbial Populations Identified by Fluorescence In Situ Hybridization in a Constructed Wetland Treating Acid Coal Mine Drainage Duongruitai Nicomrat, Warren A. Dick,* and Olli H. Tuovinen isms. Acidity is also generated by abiotic reactions when ferric Fe hydrolyzes and precipitates as schwertmannite [Fe8O8(OH)6SO4], jarosite [KFe3(SO4)2(OH)6], and goethite (a-FeOOH) in AMD (Gagliano et al., 2004). Minor and trace metals in gangue minerals may also solubilize in the course of action and precipitate downstream from the source. A passive treatment system that routes AMD through a constructed wetland is one remediation approach for improving water quality that has been evaluated in many mining regions. Previous studies of constructed AMD wetlands have described experimental systems that have ranged from laboratory-scale columns to pilotscale wetland cells in the field (Morales et al., 2005; Song et al., 2001; Whitehead and Prior, 2005). The success of wetland systems for AMD treatment, however, has been variable (Johnson and Hallberg, 2005; Kleinmann et al., 1998; Morales et al., 2005; Vile and Wieder, 1993). In general, the understanding of the biogeochemical processes underlying their performance has been limited. Constructed wetlands usually have multiple cells to increase the contact and hydraulic residence times, thereby also extending the reaction time, equilibration, and sedimentation times. These wetland cells are designed to promote aerobic oxidation processes in the water column and oxic surface sediments. The reactions include the oxidation of soluble S compounds to sulfate and ferrous Fe [Fe(II)] to ferric Fe, leading to precipitates that comprise ferric hydroxysulfates and oxides. Wetland design parameters often also include addition of organic matter to create anaerobic bottom sediments. Microbial decomposition of organic substances in these anaerobic zones helps to maintain low redox potential and produces low-molecular weight organic compounds that serve as electron donors for the bacterial reduction of sulfate. Biogenic sulfide formation then leads to the precipitation of Fe sulfides. Constructed wetland cells are claylined for containment and some contain limestone in the bottom layer for neutralization. Sources of organic matter range from straw to animal manure and municipal sewage sludge. The present study is concerned with microbiological analysis of oxic surface sediments in a field-scale constructed wetland system that was treating acid drainage from an abandoned coal mine in southeastern Ohio, USA. This region has numerous closed and abandoned coal mine sites that produce copious amounts of AMD

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

ABSTRACT Microorganisms are an integral part of the biogeochemical processes in wetlands, yet microbial communities in sediments within constructed wetlands receiving acid mine drainage (AMD) are only poorly understood. The purpose of this study was to characterize the microbial diversity and abundance in a wetland receiving AMD using fluorescence in situ hybridization (FISH) analysis. Seasonal samples of oxic surface sediments, comprised of Fe(III) precipitates, were collected from two treatment cells of the constructed wetland system. The pH of the bulk samples ranged between pH 2.1 and 3.9. Viable counts of acidophilic Fe and S oxidizers and heterotrophs were determined with a most probable number (MPN) method. The MPN counts were only a fraction of the corresponding FISH counts. The sediment samples contained microorganisms in the Bacteria (including the subgroups of acidophilic Fe- and S-oxidizing bacteria and Acidiphilium spp.) and Eukarya domains. Archaea were present in the sediment surface samples at ,0.01% of the total microbial community. The most numerous bacterial species in this wetland system was Acidithiobacillus ferrooxidans, comprising up to 37% of the bacterial population. Acidithiobacillus thiooxidans was also abundant. Heterotrophs in the Acidiphilium genus totaled 20% of the bacterial population. Leptospirillum ferrooxidans was below the level of detection in the bacterial community. The results from the FISH technique from this field study are consistent with results from other experiments involving enumeration by most probable number, dot-blot hybridization, and denaturing gradient gel electrophoresis analyses and with the geochemistry of the site.

A

(AMD) is a serious environmental pollution problem, detrimental to aquatic life as it adversely impacts water and sediment quality in receiving waterways. Acid mine drainage is a long-term problem and has few short-term treatment options at the source or downstream. It is characterized by low pH values and high concentrations of Fe and sulfate (Evangelou, 1995). In abandoned coal mines, the primary source of acidity is Fe pyrite (FeS2), which is oxidized to ferric Fe [Fe(III)] and sulfate. The oxidation yields net acidity and is accelerated by several orders of magnitude by acidophilic Fe- and S-oxidizing microorganCID MINE DRAINAGE

D. Nicomrat, Environmental Science Graduate Program, The Ohio State University, Columbus, OH 43210. Present address: National Science and Technology Development Agency, 111 Thailand Science Park, Paholyothin Road, Klong 1, Klong Luang, Pathumthani 12120, Thailand. W.A. Dick, Environmental Science Graduate Program and School of Environment and Natural Resources-Soil Science, The Ohio State University, 1680 Madison Avenue, Wooster, OH 44691. O.H. Tuovinen, Environmental Science Graduate Program and Department of Microbiology, The Ohio State University, Columbus, OH 43210. Received 24 Aug. 2005. *Corresponding author ([email protected]).

Abbreviations: AMD, acid mine drainage; DAPI, 49,6-diamidino-2phenylindole; DGGE, denaturing gradient gel electrophoresis; dUTP, 29-deoxyuridine-59-triphosphate; EDTA, ethylenediaminetetraacetic acid; FISH, fluorescence in situ hybridization; MPN, most probable number; PBS, phosphate buffered saline; PCR, polymerase chain reaction; SDS, sodium dodecyl sulfate; SSC, sodium chloride–sodium citrate; TE, trishydroxymethylaminomethane–EDTA.

Published in J. Environ. Qual. 35:1329–1337 (2006). Technical Reports: Wetlands and Aquatic Processes doi:10.2134/jeq2005.0325 ª ASA, CSSA, SSSA 677 S. Segoe Rd., Madison, WI 53711 USA

1329

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

1330

J. ENVIRON. QUAL., VOL. 35, JULY–AUGUST 2006

that pollute several thousand kilometers of local waterways, ultimately discharging to the Ohio River. Fullscale treatment systems for abating AMD pollution are piloted at some Ohio sites. The mine examined in this study was abandoned in the 1920s and the site was modified in the early 1990s to treat acid drainage by channeling it through a full-scale constructed wetland system. While the geochemistry and water quality have been monitored at many constructed wetland sites receiving acid coal mine drainage, the underlying transformations involve microbiological reactions that are not well elucidated in these environments. In a previous paper (Nicomrat et al., 2006), we characterized the microbial populations at the wetland site by amplification of 16S rRNA genes followed by denaturing gradient gel electrophoresis (DGGE) and sequence analysis. The DGGE technique demonstrated that Fe- and S-oxidizing acidithiobacilli were present at this site, but these estimates did not lend themselves to estimating actual cell numbers. In the present study, microorganisms at the wetland site were enumerated and fractionated to domain and target species populations by a fluorescent in situ hybridization technique (FISH). The FISH approach alleviates cultivation-based enumeration of microorganisms as well as DNA extraction and amplification. Analysis with FISH has been extensively used to identify acidophiles in bioleaching operations or AMD in metal mines (Espejo and Romero, 1997; Norris et al., 1996; Pizarro et al., 1996). Several 16S rDNA oligonucleotide probes have been designed for FISH analysis of acidophilic bacteria such as Acidithiobacillus ferrooxidans, Acidithiobacillus thiooxidans, Acidithiobacillus caldus, Leptospirillum ferrooxidans, and Acidiphilium spp. Probes have also been designed for several acidophilic archaea such as Ferroplasma acidarmanus and Sulfolobus spp. (Bond et al., 2000a, 2000b; Edwards et al., 1999, 2000; Peccia et al., 2000; Schrenk et al., 1998). A. ferrooxidans and A. thiooxidans share high homology (.98.5%) of 16S rRNA sequences and some of the FISH probes designed for these species show cross-species hybridization (Goebel and Stackebrandt, 1994). A. ferrooxidans is often found to be abundant by cultural methods in acid coal mine drainage systems. L. ferrooxidans has been reported to be dominant in some bioleaching processes and acid drainage from sulfide mines (Boon et al., 1995; Rawlings et al., 1999; Schrenk et al., 1998). While FISH data have greatly contributed to the understanding of the diversity of microbial populations in metal mines and bioleaching operations, this tool has not been used to profile microorganisms in coal mines and in constructed wetland systems treating acid coal mine drainage. The current knowledge of microbial communities in coal mines and AMD passive treatment systems is largely based on traditional, cultivation-dependent techniques and recent polymerase chain reaction (PCR)– based molecular approaches. Additional information on microbial community analysis in this study was sought by comparison between FISH and most probable number (MPN)–based enumeration of the major bacterial groups in the constructed wetland system. Thus this study also

provides insight into culturability of acidophiles from a constructed wetland habitat. The information in this paper is important in developing biological monitoring methods for wetland systems that are dependent on specific microbial physiological functions, such as Fe oxidation, for treatment efficiency. MATERIALS AND METHODS Wetland Site and Sample Collection The coal mine at the study site is located near Carbondale, Athens County, southeastern Ohio. The mine was abandoned in 1923. The wetland was constructed by the Ohio Department of Natural Resources in 1991 to treat the acid drainage from two deep coal mine shafts. The main seep entering the wetland is treated sequentially in six 1020-m2 cells. Also entering the wetland is a minor seep that is treated separately in three small cells before the two flows join in Cell 7. Each cell is divided into thirds by retaining boards to avoid flow channeling. The effluent from the wetland discharges into Hewett Fork, which is a tributary to Raccoon Creek. The watershed drains to the Ohio River. The cells were clay lined and layered with 30 cm of limestone and 38 cm of organic substrate (manure compost or spent mushroom compost depending on the cell). Cells 1 and 4 sampled in this study were layered with mushroom compost. The banks of the wetland cells were planted with cattails (Typha spp.). The influent pH of this wetland has ranged between 2.0 and 3.9 on an annual basis. The average conductivity has been 1.45 dS m21, Fe(II) concentration 110 mg L21, and sulfate concentration 1400 mg L21 (Shimala, 2000). The oxidation of Fe(II) to Fe(III) and the complete oxidation of S compounds to sulfate were the main biogeochemical reactions in the oxic surface layer. The oxic sediments were composed of Fe(III) precipitates containing goethite, schwertmannite, and minor amounts of jarosite (Gagliano et al., 2004). Figure 1 shows a flowchart for the study. Samples were analyzed by the FISH technique, 49,6-diamidino-2-phenylindole (DAPI) counts, and most probable number (MPN) enumeration with selective media. Grab samples of the uppermost oxic sediment layer (approximately 0- to 15-cm depth) were collected from Cells 1 and 4 in December 1998 and in March, Oxic surface samples from wetland cells 1 and 4

Sample fixation with formaldehyde and dehydration with ethanol series

Hybridization with fluorescein-labeled oligonucleotide probes Total counts Domains Target species

Total DAPI based fluorescent cell counts

Relative composition of microbial community in oxic surface sediments

Fig. 1. Experimental flowchart of this study.

MPN enumeration of acidophilic ironoxidizers, sulfuroxidizers, and heterotrophs

1331

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

NICOMRAT ET AL.: MICROORGANISMS IN A WETLAND TREATING ACID MINE DRAINAGE

August, and October 1999 using a sterile container. The oxic zone was determined by Gagliano et al. (2004) and Shimala (2000) based on color and chemical properties. The pH values of the samples ranged between pH 2.1 and 3.9 at the time of the sampling. The samples collected in December and March were initially stored at 48C for 2 mo followed by 2 mo at 2208C before fixation with paraformaldehyde and FISH analysis. The August and October samples were immediately fixed with paraformaldehyde and dehydrated with ethanol series and kept at 2208C until analyzed. Samples for MPN enumeration and other cultivation-based work were stored at 48C.

Microorganisms, Media, and Most Probable Number Enumeration Several reference microorganisms were used as positive and negative controls and as a source of DNA for the FISH and dot-blot analyses. The reference microorganisms were Acidithiobacillus ferrooxidans DSM 9465 (obtained from W. Sand, University of Hamburg), Acidithiobacillus thiooxidans FG01 (O. Carcia Jr., Sa˜o Paulo State University), Escherichia coli DH5a (this laboratory), Ferroplasma acidarmanus (P.L. Bond, University of East Anglia and J.F. Banfield, University of California at Berkeley), and Archaeoglobus fulgidus (C.J. Daniels, The Ohio State University). F. acidarmanus was received as paraformaldehyde-fixed cells along with a genomic DNA sample, and A. fulgidus was received as a cell pellet that required no further culture work. A. ferrooxidans and A. thiooxidans were grown in inorganic mineral salts media with ferrous sulfate (Silverman and Lundgren, 1959) and elemental S (Garcia, 1991) as the respective sources of energy. Bacteria in sediment samples were enumerated with an MPN technique. Samples of Fe precipitates were vortexed and diluted in sterile acidified media with Fe(II), elemental S, or glucose as the substrate. The MPN media with Fe(II) and elemental S were formulated as for A. ferrooxidans and A. thiooxidans. The MPN media for acidophilic heterotrophs with glucose (10 g L21) as the substrate followed the formulation adopted for Acidiphilium spp. (Peccia et al., 2000). Five replicates were prepared for each decimal dilution tube. The tubes were incubated at 22 6 28C for at least 4 wk. Growth was scored based on the color change from light green to red-

brown for Fe oxidizers and by turbidity for acidophilic heterotrophs. A decrease from pH 4.6 to ,pH 2 was used to score positive tubes of S oxidizers. Elemental S, initially hydrophobic, settled as small particles in positive MPN tubes. Negative controls without inocula were used in all incubations.

FISH Analysis Microorganisms were enumerated with a FISH protocol (Alfreider et al., 1996; Glo¨ckner et al., 1996; Manz et al., 1992) that was slightly modified for this study. Samples (1 g) of Fe precipitates and cells from MPN cultures were fixed with three volumes of freshly prepared 4% paraformaldehyde-phosphate buffered saline (PBS; 130 mM NaCl, 10 mM Na-phosphate buffer, pH 7.2) for 1 to 3 h at 48C. The samples were centrifuged at 10 000 3 g for 3 min and washed in PBS and then resuspended in a PBS and 95% ethanol solution (1:1 v/v). The sample solution was stored at 48C. Paraformaldehyde-fixed cells (10 mL) in PBS and ethanol (1:1 v/v) were spread on 10-well Teflon coated glass slides (CelLine, Portsmouth, NH), which were coated with 0.1% (w/v) gelatin and 0.01% (w/v) KCr(SO4)2 (Amann et al., 1995; Sandaa et al., 1999). The samples were dried at 228C for 2 to 24 h. The slides were cooled to 48C followed by the addition of 40 mL of ice-cold lysozyme solution (250 mg lysozyme mL21; Sigma, St. Louis, MO) in 100 mM Tris-HCl (pH 7.5) to the cell smear. After 5 to 20 min of digestion, the slides were washed for 3 min at 48C in TE (10 mM Tris-HCl, 0.1 mM ethylenediaminetetraacetic acid [EDTA], pH 7.5), and in 1% (v/v) Nonidet P-40 for 10 min, followed by a brief wash in PBS buffer. The samples were subsequently dehydrated in 50, 80, and 100% ethanol for 3 min each and air-dried at 228C at least 1 h. Samples were kept dry at 48C in a desiccator or a sealed bag until used for hybridization. The oligonucleotides used in this study were rDNA-targeted probes specific for species, groups, and domains (Table 1). The probes were 39-end labeled with fluorescein-11-dUTP (29-deoxyuridine-59-triphosphate) (Molecular Probes, Eugene, OR) by treating with terminal transferase enzymes (Gibco BRL, Rockville, MD) per manufacturer’s instructions. The formamide concentrations (Table 1) varied depending on the melting temperatures of the oligonucleotide sequences. After hybridization, the slides were briefly rinsed with washing

Table 1. The oligonucleotide probes used in fluorescence in situ hybridization (FISH) analysis and primers used for polymerase chain reaction (PCR) amplification.

Designation

16S rRNA target position†

Nucleotide sequence‡

Specificity

Formamide/NaCl§

Reference

%/mM Probes for FISH analysis Acdp821 Arch 915 EUB338 Euk 502 LF581 TF539 Thio820

821–842 915–934 338–354 502–517 581–598 539–556 830–851

Univ-1392

1392–1378

Acidiphilium spp. most archaea most bacteria most eukaryotes L. ferrooxidans A. ferrooxidans A. ferrooxidans and A. thiooxidans 59-ACG GGC GGT GTG TRC-39 all species 16S rDNA primers for PCR amplification and detection of archaea

59-ACC AAA CAT CTA GTA TTC ATC G-39 59-GTG CTC CCC CGC CAA TTC CT-39 59-CCT ACG GGA GGC AGC AG-39 59-ACC AGA CTT GCC CTC C-39 59-CGG CCT TTC ACC AAA GAC-39 59-CAG ACC TAA CGT ACC GCC-39 59-ACC AAA CAT CTA GTA TTC ATC G-39

ARC/EUK1373R 1373–1355

59-AGG GGG CAG GGA CGT ATT C-39

FER3–20 FER656

59-TCC GGT TGA TCC TGC CGG-39 59-CGT TTA ACC TGA CCC GAT C-39

3–20 656–674

† E. coli numbering (Brosius et al., 1981). ‡ Nucleotide abbreviations; R 5 A or G. § Percent formamide and mM NaCl in the in situ hybridization buffer.

most archaea and eukaryotes most archaea F. acidarmanus

20/220 25/150 30/100 20/220 25/150 25/150 20/220

Peccia et al. (2000) Schrenk et al. (1998) Manz et al. (1992) Schrenk et al. (1998) Schrenk et al. (1998) Schrenk et al. (1998) Peccia et al. (2000)

20/220

Zheng et al. (1996) Hugenholtz et al. (1998b)

25/150

Hugenholtz et al. (1998a) Edwards et al. (2000)

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

1332

J. ENVIRON. QUAL., VOL. 35, JULY–AUGUST 2006

buffer at 258C and then washed for 20 min at 488C with preheated washing buffer that contained 20 mM Tris-HCl (pH 7.4), a probe-dependent concentration of NaCl, 7 mM sodium dodecyl sulfate (SDS), and 5.3 mM EDTA. The NaCl concentrations in the washing buffer were 220, 150, and 100 mM for 20, 25, and 30% formamide, respectively. The slides were rinsed in sterile, filtered distilled water and air-dried. Hybridized cells were counter-stained with 49,69-diamidino2-phenylindole (DAPI) (1 mg per mL) for 10 min and washed in PBS and distilled water (Amann et al., 1995). The slides were covered with VectaShield mounting medium (antifading agent; Vector Laboratories, Burlingame, CA) and examined under epifluorescence microscopy with a high-pressure mercury bulb (50 W) (Axioplan; Carl Zeiss, Thornwood, NY) using the filter sets BP365, FT395, and LP397 for DAPI and BP485, FT510, and LP515–565 for fluorescein. Hybridized and DAPI-stained cells were enumerated with a 100 3 1.3 numerical aperture oil immersion lens and from digitized video images captured by IPLab Spectrum (Signal Analytics, Bothell, VA). More than 1000 DAPI-stained cells were counted for each sample in the well. Twenty fields of DAPI-stained cells and cells hybridizing with the probe were randomly selected for counting. Each field has an area of 0.01 mm2. Means and standard deviations were calculated from the 20 randomly chosen fields from each filter. Phase contrast microscopy was used to examine some samples to observe morphological diversity and extent of cell aggregation. To confirm the low levels of Archaea detected using FISH, primer pairs specific for Archaea were used to amplify 16S rDNAusing DNA extracted from wetland samples. Primer pairs were FERS-20 and ARC/EUK1373R and FER656 and ARC/ EUK1373R (Table 1). The amplification procedure was as described by Hugenholtz et al. (1998b) and Edwards et al. (2000).

Dot-Blot Hybridization and DNA Purification and Amplification Dot-blot hybridization was performed to complement the relative abundance data of species as revealed by FISH analysis. Hybridization was performed with 16S rDNA gene sequences amplified from reference cultures used as the target DNA. Unamplified DNA from wetland samples containing a fluorescent label was used as the probe. Target DNA from reference cultures was amplified using the universal primers 8f (Amann and Stahl, 1992) and 1492r (Lane, 1991). The PCR amplification reaction (total volume of 50 mL) contained 10 to 50 ng DNA, 5 mL 103 reaction buffer (Gibco BRL), 2.5 mM MgCl2, 200 mM of dNTP mixture, 250 nM of each primer, and 2.5 U of Taq DNA polymerase (Gibco BRL). The thermocycling program used for the amplification was 948C for 3 min, 948C for 45 s followed by 30 cycles of 508C for 30 s, and 728C for 1.5 min and finally holding at 728C for 10 min. The amplified target DNA was diluted in buffer (10 mM Tris-HCl, 1 mM EDTA; pH 8.0), denatured for 5 min at 958C, and chilled immediately on ice. The DNA was dot-blotted onto a positively charged nylon membrane (Boehringer-Mannheim, Indianapolis, IN) under vacuum and fixed for 5 min by UV cross-linking. For preparation of the probe DNA from wetland sediment samples, a 1-g aliquot of the sample was pretreated with 0.3 M ammonium oxalate as described by Nicomrat et al. (2006). Cell pellets from both reference cultures and wetland samples were washed with buffer solution (200 mM Tris [pH 8.0]; 20 mM EDTA; 200 mM NaCl) and suspended in 1.0 mL of buffer solution that contained 2% SDS. Cells were broken with a bead beating protocol combined with freezing (2808C) and thawing (708C) for six to eight repeat cycles (Nicomrat et al.,

2006). The protocol used 0.3 g of acid-washed zirconium-silica beads (0.1-mm diameter) and one volume of phenol solution. After centrifugation, protein and cell debris in the supernatant were removed with an equal volume of phenol–chloroform– isoamyl alcohol (25:24:1), and left-over phenol was extracted with an equal volume of chloroform–isoamyl alcohol. Nucleic acids were precipitated from the supernatant by adding approximately 2 volumes of isopropanol. After 1 h at 2808C, the solution was centrifuged and traces of isopropanol were completely removed by pipetting. The DNA-containing pellets were resuspended in 10 mL of TE solution (10 mM Tris and 1 mM EDTA; pH 8.0) and kept at 48C for immediate use or stored at 2208C. Aliquots of this DNA were then randomly labeled with fluorescein-11-dUTP, using a primer labeling and detection kit, according to the manufacturer’s instructions (Amersham-Pharmacia, Piscataway, NJ). After hybridization of target and probe DNA for 4 h at 558C, the membranes were washed twice in buffer with low stringency condition (5 3 SSC and 0.1% SDS; 1 3 SSC is 0.15 M NaCl plus 0.015 M sodium citrate and SDS is sodium dodecyl sulfate) at 228C for 10 min, followed by three 15-min washes with a high stringency buffer (0.1 3 SSC and 0.1% SDS) at 568C. The membranes were incubated in the antibodyconjugated alkaline phosphatase solution, washed, incubated in substrate solution, and exposed to X-ray film as described by the manufacturer (Amersham). Negative controls were DNA from pCR2.1 vector and 16s rDNA from Escherichia coli amplified using universal primers (8f and 1492r; see above). Positive control included unamplified DNA from wetland samples collected in December. No signal was detected for negative controls and the positive control yielded hybridization signal (data not shown).

RESULTS AND DISCUSSION Microscopic Observations of Microorganisms in Oxic Sediment Samples Iron precipitate samples from the Carbondale wetland contained microorganisms that varied in size and shape, ranging from ,0.5 to .10 mm in length (Fig. 2A). Under phase contrast microscopy, some cells were aggregated with Fe precipitates and some were planktonic. Eukaryotic organisms, mostly unicellular cells and filamentous fungi, were also associated with Fe precipitates. Eukaryotic cells varied from ,2 to .10 mm in size (Fig. 2B). Large rods, 5 to 6 mm long, were abundant in some samples. Fragments of algae and fungal hyphae were observed, especially in the October samples. Bacteria varied from ,0.5 to 3 mm in length and were generally rod shaped or spherical and many were planktonic (Fig. 2C). Bacterial cells hybridized with probe TF539 were rod shaped, 1 to 3 mm in length, consistent with the morphological characteristics of A. ferrooxidans (Fig. 3A). Cells hybridized with probe Acdp821 for Acidiphilium spp. were mostly round and rod shaped (Fig. 3B). Very few archaea could be detected in these samples. Signals for archaeal cells were sporadic and the cells were small rod, oval, or round shaped, and did not adhere to Fe precipitates.

Cells Counts with DAPI and Universal FISH Probes In general, FISH-hybridized cells showed lower fluorescence intensity than DAPI-stained cells. The cell

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

NICOMRAT ET AL.: MICROORGANISMS IN A WETLAND TREATING ACID MINE DRAINAGE

1333

Fig. 2. Photomicrographs of iron precipitate samples. (A) Stained with 49,6-diamidino-2-phenylindole (DAPI) (top) and hybridized with universal probe Univ-1392 (bottom). (B) stained with DAPI (top) and hybridized with eukaryotic universal probe Euk 502 (bottom). (C) Stained with DAPI (top) and hybridized with bacterial universal probe EUB338 (bottom). Scale bar marker: 10 mm.

count determined with probe Univ-1392 was 2.4 3 108 cells mL21 Fe precipitate. The FISH counts in the August and October samples were approximately 40% of the DAPI-stained cell counts (5.7 3 108 cells mL21 Fe precipitate). The DAPI-stained cells included some fungal and bacterial cells that did not hybridize against Univ-1392. The FISH counts may be low due to probe specificity that leads to omission of some cells by the

FISH technique, low rDNA content in dormant and/or inactive microorganisms, and thick cell walls that resist the probe’s ability to penetrate through the cell wall (Bidnenko et al., 1998). Thus some cells in the environment may give a weak or no signal in the FISH assay because they are in dormancy or have a very low number of rRNA genes. It is also possible that some cells may have lysed during the FISH labeling treatment.

Fig. 3. Photomicrographs of cells in Fe precipitate samples. (A) Stained with 49,6-diamidino-2-phenylindole (DAPI) (top) and hybridized with probe TF539 for A. ferrooxidans (bottom). (B) Stained with DAPI (top) and hybridized with probe Acdp821 for Acidiphilium spp. (bottom). Scale bar marker: 5 mm.

7

Thio820

2.10 3 10 7 4.00 3 10 7 6.75 3 10 6 2.19 3 10 7 9.49 3 10 7 2.10 3 10 6 2.24 3 10 6 2.45 3 10 7 3.14 6 3.42 3 10 14 34

4.18 7.90 3.18 1.36 1.43 3.51 1.96 3.17 1.85

7

Acdp821

Given these constraints, it was not possible to optimize conditions for the detection of strong and weak fluorescent signals from microorganisms. For comparative purposes, the counts of FISHhybridized cells using various probes were normalized against the microorganisms hybridized with the Univ1392 probe. Thus the relative abundance of 100% refers to Univ-1392 counts of cells. This was based on the assumption that the Univ-1392 probe included both cells that were counted by other specific probes plus cells in samples that were not counted by these probes.

3 10 7 3 10 6 3 10 6 3 10 7 3 10 6 3 10 6 3 10 6 3 10 7 6 2.80 3 10 8 20

J. ENVIRON. QUAL., VOL. 35, JULY–AUGUST 2006

g sediment 4 1.39 3 10 4 1.90 3 10 3 2.02 3 10 3 1.61 3 10 4 1.07 3 10 3 4.12 3 10 3 1.12 3 10 3 3.13 3 10 3 6.95 6 6.73 3 10 0.003 NA

7

TF539

† Not applicable.

Mean of each probe % of Univ-1392 counts % of EUB338 counts

October

August

March

December

Cell 1 Cell 4 Cell 1 Cell 4 Cell 1 Cell 4 Cell 1 Cell 4 Cells 1 and 4

1.06 2.20 1.82 7.63 2.91 1.66 7.46 1.88 2.28

8

3 10 8 3 10 8 3 10 6 3 10 8 3 10 8 3 10 8 3 10 8 3 10 8 6 2.21 3 10 100 NA†

7

1.42 3 10 7 9.80 3 10 6 8.35 3 10 6 1.97 3 10 7 7.33 3 10 7 9.40 3 10 8 7.20 3 10 8 1.60 3 10 8 1.46 6 2.38 3 10 64 NA

7

9.99 3 10 8 1.43 3 10 8 1.50 3 10 6 5.61 3 10 8 2.79 3 10 7 5.13 3 10 7 1.02 3 10 6 7.20 3 10 7 9.32 6 9.56 3 10 41 100

21

Arch 915

Cell counts

EUB338 Euk 502 Univ-1392 Location Sample

Iron precipitate samples collected in December and March were not immediately fixed with paraformaldehyde before storage at 48C. Storage may lower cellular RNA content, and cell decay may also contribute to loss in signal and cell counts. The FISH (Table 2) show that counts generally varied by one order of magnitude or less between wetland Cells 1 and 4 at all sampling dates. Furthermore, counts among the four sampling dates within each wetland cell also varied by an order of magnitude or less except for very few exceptions. These data indicated no cellular decay or loss of counts in the first two sets of samples when samples were not immediately fixed for FISH analysis. Therefore, the data for the first two sampling dates were included in averages of cell counts for the various probes or cell types. For the counts averaged for each probe (Table 2), Bacteria (EUB338) accounted for about 41% of the cells hybridized with Univ-1392 probe. The averaged FISH counts for Eukarya (Euk 502) were 64% of the Univ1392 counts. Numerous eukaryotic microorganisms have been shown to live in AMD-impacted habitats (Baker et al., 2004; Gross and Robbins, 2000). Algae were also prominent among eukaryotes. Eukaryotes in these samples may originate from the wetland banks and sustain themselves with runoff nutrients and organic compounds from the manure layer and decaying plant material. By comparison with Eukarya and Bacteria, the FISH counts for the Archaea were five orders of magnitude lower. The low level of Archaea in the oxic sediments was confirmed by PCR amplification of 16S rRNA genes with archaeal specific primers and by dot-blot hybridization. No PCR product was detected on amplification of 16S rDNA with the Archaea-specific primers FER3–20 and ARC/EUK1373. No amplified 16S rRNA genes of F. acidarmanus were found after amplification with the primers FER656 and ARC/EUK1373. Hybridization with fluorescein-labeled probes of genomic DNA from Fe precipitate samples gave no signal for the target 16S rDNA of Ferroplasma acidarmanus and the reference archaeon Archaeoglobus fulgidus (Fig. 4). Thus these results demonstrated that very few Archaea were present in the oxic wetland sediments samples, and their function and identity would be speculative at best. Leptospirillum ferrooxidans (LF581) was not detected by the FISH technique and thus their counts were at least six order orders of magnitude lower than the Univ1392 counts.

4.80 3 10 7 3.58 3 10 7 6.30 3 10 6 1.91 3 10 7 9.21 3 10 7 2.77 3 10 6 6.22 3 10 6 2.88 3 10 7 3.47 6 3.21 3 10 15 37

Analysis of Microorganisms with FISH

Table 2. Fluorescence in situ hybridization (FISH) counts for wetland sediment samples.

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

1334

1335

NICOMRAT ET AL.: MICROORGANISMS IN A WETLAND TREATING ACID MINE DRAINAGE

100

Percent of FISH Counts

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

38%

Fig. 4. Dot-blot hybridization of DNA extracted from December samples from Cell 1 (A and B) and Cell 4 (C and D). Amplified 16S rDNA from A. ferrooxidans (A1 and C1), A. thiooxidans (A2 and C2), Archaeoglobus fulgidus (B1 and D1), and Ferroplasma acidarmanus (B2 and D2) was hybridized by using fluoresceinlabeled rDNA from Fe precipitates.

The FISH results for probes TF539 (A. ferrooxidans), Thio820 (A. ferrooxidans and A. thiooxidans), and Acdp821 (Acidiphilium spp.) accounted for 37, 34, and 20% of the total bacterial (EUB338) counts (Table 2). As a percentage of total counts measured using the universal probe Univ-1392, the values for the TF539, Thio820, and Acdp821 were 15, 14, and 8%, respectively. These data are consistent with the abundance of A. ferrooxidans, A. thiooxidans, and Acidiphilium spp. in Fe precipitate samples. A. ferrooxidans and A. thiooxidans were also readily detected by dot-blot hybridization (Fig. 4). The Thio820 counts also include some TF539 counts since A. thiooxidans and A. ferrooxidans are closely related and share similar features and hybridize against this probe (Peccia et al., 2000). Probe TF539 is reportedly specific to A. ferrooxidans (Peccia et al., 2000). Although Thio820 does not differentiate between these two species, it is clear from the dot-blot hybridization data that A. thiooxidans was also present in these environmental samples. Cumulatively, these three probes accounted for 91% of the bacterial (EUB338) cell counts (Table 2). In any case, the remaining cell counts are a much smaller fraction than the counts of cells that hybridized and were identified using the three species-specific FISH probes. While the data are, of course, subject to variability in cell counts due to the inherent nature of fieldwork, they indicate that most of the bacterial community could be resolved with only three FISH probes. With the August sampling, bacteria were also enumerated with an MPN protocol using media that contained Fe(II) for iron oxidizers, elemental S for S oxidizers, and glucose for acidophilic heterotrophs. The MPN counts of these culturable bacterial groups were compared to the counts obtained with the FISH technique using TF539, Thio820, and Acdp821 as the probes for the same samples. The results indicated that most of the acidophilic bacteria in the wetland system enumerated with FISH

10 3.1%

1

0.24%

0.1

MPN Fe 2

MPN S0

MPN Glucose

Fig. 5. Culturability of acidophilic bacteria in oxic wetland sediments. The most probable number (MPN) counts of acidophilic Fe and S oxidizers, and glucose-oxidizing heterotrophs were compared to fluorescence in situ hybridization (FISH) counts obtained with probes TF539 (A. ferrooxidans), Thio820 (A. thiooxidans), and Acdp821 (Acidiphilium spp.) that were taken to be 100%. The corresponding FISH counts are listed in Table 2.

were not detected using the commonly used culture methods described in this paper (Fig. 5). The MPN counts with ferrous sulfate media were 3% of the corresponding FISH counts with TF539 as the probe. For elemental S media, the MPN counts were 38% of the corresponding Thio820 counts. The MPN counts of acidophilic heterotrophs were 0.2% of the Acdp821 (Acidiphilium spp.). Thus these data showed that a varying fraction of the target acidophiles could be recovered with a viable counting protocol. Based on the MPN counts, S oxidizers were 10 and 105 times more numerous than the Fe oxidizers and acidophilic heterotrophs, respectively, which contrasts with results from the probe counts that include both culturable and unculturable microorganisms.

CONCLUSIONS Analysis with FISH revealed numerous Bacteria and Eukarya in Fe-precipitates collected from the oxic surface sediment layers of the constructed wetland. In contrast, Archaea constituted only a small fraction of abundance. Cells were aggregated with Fe precipitates and were also freely suspended. The bacterial communities at this wetland site were dominated by A. ferrooxidans, A. thiooxidans, and acidophilic heterotrophs in the Acidiphilium genus. Combined they represented a total of 91% of total bacterial counts in these samples. L. ferrooxidans was not detected by FISH analysis. The MPN counts and the parallel comparative FISH analysis indicated that cultivation recovered only a minor fraction of Fe oxidizers and S oxidizers as well as acidophilic heterotrophs of the Acidiphilium type. There were no

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

1336

J. ENVIRON. QUAL., VOL. 35, JULY–AUGUST 2006

discernible differences in cell counts of microbial populations between Cells 1 and 4 or among sampling dates in the constructed wetland. Microbial diversity as determined by FISH, enumeration by most probable number, and dot-blot hybridization were consistent, that is, the bacterial population was dominated by A. ferrooxidans and A. thiooxidans, and Acidiphilium spp. were also a major faction of the community. This agrees with previous results based on DGGE analyses of amplified 16S rDNA sequences (Nicomrat et al., 2006). Thus the oxic surface layers of field-scale wetlands receiving acid mine drainage are dominated by relatively few types of microorganisms, information that is important for wetland management practices. ACKNOWLEDGMENTS We thank J.F. Banfield, P.L. Bond, O. Garcia Jr., and W. Sand for reference cultures. D. Nicomrat gratefully acknowledges a fellowship from the Royal Thai Government in partial support of this study. Salary and research support were provided to W.A. Dick by state and federal funds appropriated to the Ohio Agricultural Research and Development Center.

REFERENCES Alfreider, A., J. Pernthaler, R. Amann, B. Sattler, F.O. Glo¨ckner, A. Wille, and R. Psenner. 1996. Community analysis of the bacterial assemblages in the winter cover and pelagic layers of a high mountain lake by in situ hybridization. Appl. Environ. Microbiol. 62: 2138–2144. Amann, R.I., W. Ludwig, and K.-H. Schleifer. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59:143–169. Amann, R.I., and D.J. Stahl. 1992. Dual staining of natural bacterioplankton with 49,6-diamidino-2-phenylindole and fluorescent oligonucleotide probes targeting kingdom-level 16S rRNA sequences. Appl. Environ. Microbiol. 58:2158–2163. Baker, B.J., M.A. Lutz, S.C. Dawson, P.L. Bond, and J.F. Banfield. 2004. Metabolically active eukaryotic communities in extremely acidic mine drainage. Appl. Environ. Microbiol. 70:6264–6271. Bidnenko, E., C. Mercier, J. Tremblay, P. Taillier, and S. Kulakauskas. 1998. Estimation of the state of the bacterial cell wall by fluorescent in situ hybridization. Appl. Environ. Microbiol. 64:3059–3062. Bond, P.L., G.K. Druschel, and J.F. Banfield. 2000a. Comparison of acid mine drainage microbial communities in physically and geochemically distinct ecosystems. Appl. Environ. Microbiol. 66: 4962–4971. Bond, P.L., S.P. Smriga, and J.F. Banfield. 2000b. Phylogeny of microorganisms populating a thick, subaerial, predominantly lithotrophic biofilm at an extreme acid mine drainage site. Appl. Environ. Microbiol. 66:3842–3849. Boon, M., G.S. Hansford, and J.J. Heijnen. 1995. The role of bacterial ferrous iron oxidation in the bio-oxidation of pyrite. p. 153–181. In T. Vargas et al. (ed.) Biohydrometallurgical processing. Vol. I. Microbiology and dissolution mechanisms in bioleaching. Bioleaching processes for gold, copper and non-sulfide ores. Univ. of Chile, Santiago. Brosius, J., T.L. Dull, D.D. Sleeter, and H.F. Noller. 1981. Gene organization and primary structure of a ribosomal RNA operon from Escherichia coli. J. Mol. Biol. 148:107–127. Edwards, K.J., P.L. Bond, T.M. Gihring, and J.F. Banfield. 2000. An archaeal iron-oxidizing extreme acidophile important in acid mine drainage. Science 287:1796–1799. Edwards, K.J., B.M. Goebel, T.M. Rodgers, M.O. Schrenk, T.M. Gihring, M.M. Cardona, B. Hu, M.M. McGuire, R.J. Hamers, N.R. Pace, and J.F. Banfield. 1999. Geomicrobiology of pyrite (FeS2) dissolution: Case study at Iron Mountain, California. Geomicrobiol. J. 16:165–179.

Espejo, R.T., and J. Romero. 1997. Bacterial communities in copper sulfide ores inoculated and leached with solution from a commercial-scale copper leaching plant. Appl. Environ. Microbiol. 63: 1344–1348. Evangelou, V.P. 1995. Pyrite oxidation and its control: Solution chemistry, surface chemistry, acid mine drainage (AMD), molecular oxidation mechanisms, microbial role, kinetics, control, ameliorates and limitations, microencapsulation. CRC Press, Boca Raton, FL. Gagliano, W.B., M.R. Brill, J.M. Bigham, F.S. Jones, and S.J. Traina. 2004. Chemistry and mineralogy of sediments in a constructed mine drainage wetland. Geochim. Cosmochim. Acta 68:2119–2128. Garcia, O., Jr. 1991. Isolation and purification of Thiobacillus ferrooxidans and Thiobacillus thiooxidans from some coal and uranium mines of Brazil. Rev. Microbiol. (Sa˜o Paulo) 22:1–6. Glo¨ckner, F.O., R. Amann, A. Alfreider, J. Pernthaler, R. Psenner, K. Trebesius, and K.-H. Schleifer. 1996. An optimized in situ hybridization protocol for planktonic bacteria. Syst. Appl. Microbiol. 19:403–406. Goebel, B.M., and E. Stackebrandt. 1994. Cultural and phylogenetic analysis of mixed microbial populations found in natural and commercial bioleaching environments. Appl. Environ. Microbiol. 60:1614–1621. Gross, S., and E.I. Robbins. 2000. Acidophilic and acid-tolerant fungi and yeasts. Hydrobiologia 433:91–109. Hugenholtz, P., B.M. Goebel, and N.R. Pace. 1998a. Impact of cultureindependent studies on the emerging phylogenetic view of bacterial diversity. J. Bacteriol. 180:4765–4774. Hugenholtz, P., C. Pitulle, K.L. Herschberger, and N.R. Pace. 1998b. Novel division level bacterial diversity in a Yellowstone hot spring. J. Bacteriol. 180:366–376. Johnson, D.B., and K.B. Hallberg. 2005. Acid mine drainage remediation options: A review. Sci. Total Environ. 338:3–14. Kleinmann, R.L.P., R.S. Hedin, and R.W. Nairns. 1998. Treatment of mine drainages by anoxic limestone drains and constructed wetlands. p. 303–319. In A. Geller et al. (ed.) Acidic mining lakes: Acid mine drainage, limnology and reclamation. Springer Verlag, Berlin. Lane, D.J. 1991. 16S/23S rRNA sequencing. p. 115–175. In E. Stackebrandt and M. Goodfellow (ed.) Nucleic acid techniques in bacterial systematics. John Wiley & Sons, New York. Manz, W., R. Amann, W. Ludwig, M. Wagner, and K.-H. Schleifer. 1992. Phylogenetic oligonucleotide probe for the major subclasses of Proteobacteria: Problems and solutions. Syst. Appl. Microbiol. 15:593–600. Morales, T.A., M. Dopson, R. Athar, and R.B. Herbert, Jr. 2005. Analysis of bacterial diversity in acidic pond water and compost after treatment of artificial acid mine drainage for metal removal. Biotechnol. Bioeng. 90:543–551. Nicomrat, D., W.A. Dick, and O.H. Tuovinen. 2006. Assessment of the microbial community in a constructed wetland that receives acid coal mine drainage. Microb. Ecol. 51:83–89. Norris, P.R., D.A. Clark, J.P. Owen, and S. Waterhouse. 1996. Characteristics of Sulfobacillus acidophilus sp. nov. and other moderately thermophilic mineral-sulphide-oxidizing bacteria. Microbiol. 142: 775–783. Peccia, J., E.A. Marchand, J. Silverstein, and M. Hernandez. 2000. Development and application of small-subunit rRNA probes for assessment of selected Thiobacillus species and members of the genus Acidiphilium. Appl. Environ. Microbiol. 66:3065–3072. Pizarro, J., E. Jedlicki, O. Orellana, J. Romero, and R.T. Espejo. 1996. Bacterial populations in samples of bioleached copper ore as revealed by analysis of DNA obtained before and after cultivation. Appl. Environ. Microbiol. 62:1323–1328. Rawlings, D.E., H. Tributsch, and G.S. Hansford. 1999. Reasons why ‘Leptospirillum’-like species rather than Thiobacillus ferrooxidans are the dominant iron-oxidizing bacteria in many commercial processes for the biooxidation of pyrite and related ores. Microbiol. 145:5–13. Sandaa, R.-A., Ø. Enger, and V. Torsvik. 1999. Abundance and diversity of Archaea in heavy-metal-contaminated soils. Appl. Environ. Microbiol. 65:3293–3297. Schrenk, M.O., K.J. Edwards, R.M. Goodman, R.J. Hamers, and J.F. Banfield. 1998. Distribution of Thiobacillus ferrooxidans and Leptospirillum ferrooxidans: Implications for generation of acid mine drainage. Science 279:1519–1522.

Reproduced from Journal of Environmental Quality. Published by ASA, CSSA, and SSSA. All copyrights reserved.

NICOMRAT ET AL.: MICROORGANISMS IN A WETLAND TREATING ACID MINE DRAINAGE

Shimala, J.R. 2000. Hydrogeochemical characterization of the Carbondale wetland, Athens county, Ohio: Evaluation of acid mine drainage remediation alternatives. M.S. thesis. Ohio Univ., Athens, OH. Silverman, M.P., and D.G. Lundgren. 1959. Studies on the chemoautotrophic iron bacterium Ferrobacillus ferrooxidans. I. An improved medium and a harvesting procedure for securing high cell yields. J. Bacteriol. 77:642–647. Song, Y., M. Fitch, J. Burken, L. Nass, S. Chilukiri, N. Gale, and C. Ross. 2001. Lead and zinc removal by laboratory-scale constructed wetlands. Water Environ. Res. 73:37–44.

1337

Vile, M.A., and R.K. Wieder. 1993. Alkalinity generation by Fe(III) reduction versus sulfate reduction in wetlands constructed for acid mine drainage treatment. Water Air Soil Pollut. 69:425–441. Whitehead, P.G., and H. Prior. 2005. Bioremediation of acid mine drainage: An introduction to the Wheal Jane Wetlands Project. Sci. Total Environ. 338:15–21. Zheng, D., E.W. Alm, D.A. Stahl, and L. Raskin. 1996. Characterization of universal small-subunit rRNA hybridization probes for quantitative molecular microbial ecology studies. Appl. Environ. Microbiol. 62:4504–4513.