Response of Soil-Associated Microbial Communities to Intrusion of

Article pubs.acs.org/est

Response of Soil-Associated Microbial Communities to Intrusion of Coal Mine-Derived Acid Mine Drainage Justin S. Brantner†,‡ and John M. Senko*,†,‡,§ †

Department of Biology, ‡Integrated Biosciences Program, and §Department of Geosciences, The University of Akron, Akron, Ohio 44325, United States S Supporting Information *

ABSTRACT: A system has been identified in which coal mine-derived acid mine drainage (AMD) flows as a 0.5-cm-deep sheet over the terrestrial surface. This flow regime enhances the activities of Fe(II) oxidizing bacteria, which catalyze the oxidative precipitation of Fe from AMD. These activities give rise to Fe(III) (hydr)oxide-rich deposits (referred to as an iron mound) overlying formerly pristine soil. This iron mound has developed with no human intervention, indicating that microbiological activities associated with iron mounds may be exploited as an inexpensive and sustainable approach to remove Fe(II) from AMD. To evaluate the changes in microbial activities and communities that occur when AMD infiltrates initially pristine soil, we incubated AMD-unimpacted soil with site AMD. Continuous exposure of soil to AMD induced progressively greater rates of Fe(II) biooxidation. The development of Fe(II) oxidizing activities was enhanced by inoculation of soil with microorganisms associated with mature iron mound sediment. Evaluation of pyrosequencing-derived 16S rRNA gene sequences recovered from incubations revealed the development of microbial community characteristics that were similar to those of the mature iron mound sediment. Our results indicate that upon mixing of AMD with pristine soil, microbial communities develop that mediate rapid oxidative precipitation of Fe from AMD.

■

INTRODUCTION Coal mine-derived acid mine drainage (AMD) is produced when oxygenated water induces the oxidation of coal seam-associated FeS phases that are exposed during mining activities, yielding fluids with low pH (typically 3.0−4.5) and high Fe(II) concentrations.1,2 AMD remains one of the greatest threats to surface water quality in Appalachian coal mining regions of the United States, with over 10 000 km of AMD-impacted streams.3 Given that AMD may emerge from abandoned mines for hundreds of years, the anticipated costs of AMD treatment are prohibitive, and currently employed treatment technologies often require levels of investment (i.e., money, labor, energy) that preclude widespread treatment efforts.4,5 The major goal of coal mine-derived AMD treatment is the removal of dissolved Fe(II), which is frequently accomplished using limestone-based systems.6,7 Dissolution of limestone-CaCO3 increases pH of the fluids, enhancing the kinetics of Fe(II) oxidation (R1) and subsequent hydrolysis and precipitation of Fe(III) (hydr)oxides (R2). 4Fe2 + + O2 + 4H+ → 4Fe3 + + 2H 2O

(R1)

Fe3 + + 3H 2O → Fe(OH)3 + 3H+

(R2)

Given the unpredictable performance and economic limitations of limestone-based and other AMD treatment strategies,9 it has been proposed that AMD “sheet flow” systems may be exploited to oxidatively precipitate Fe from AMD.10−13 In such systems, AMD flows as a 0.5−1-cm sheet over the terrestrial surface, enhancing aeration of AMD and the activities of Fe(II) oxidizing bacteria (FeOB), which oxidatively precipitate Fe from AMD (R1 and R2). Such activities give rise to massive Fe(III) (hydr)oxide deposits which are referred to as “iron mounds” (alternatively referred to as “terraced iron formations”).10−13 FeOB activities in iron mounds may lead to removal of over 90% of dissolved Fe(II) in AMD.10−13 Several iron mounds have been identified in which robust oxidative precipitation of Fe from AMD is occurring with no human intervention to stimulate these activities, so these systems are attractive models for new AMD treatment strategies.10−13 Implicit in the observation that iron mounds have developed with no human intervention is the fact that upon initial mixing of AMD with formerly pristine soil, microbial communities were altered, and over time, culminated in communities capable of robust Fe(II) oxidizing activities. Therefore, to evaluate the dynamics of the initial stages of iron mound development, we challenged pristine soil with coal minederived AMD in laboratory incubations and assessed the dynamics of responses of microbial activities and communities.

However, such systems require routine maintenance (e.g., flushing or limestone replacement) to remove Fe(III) phases that accumulate on limestone surfaces (referred to as “armoring”), thus limiting further CaCO3 dissolution and neutralization capacity.8 © XXXX American Chemical Society

Received: May 7, 2014 Revised: June 26, 2014 Accepted: June 27, 2014

A

dx.doi.org/10.1021/es502261u | Environ. Sci. Technol. XXXX, XXX, XXX−XXX

Environmental Science & Technology

Article

oxidizing activities (with results shown in Figure 1), AMD (either filter-sterilized or nonsterile) was provided as the fluid for slurry

Companion experiments were also conducted to determine if inoculation of pristine soil with mature iron mound microbial communities might enhance the development of Fe(II) oxidizing activities.

■

MATERIALS AND METHODS Material Collections from Field Site. All samples were collected near an AMD-impacted site referred to as the Mushroom Farm, located in North Lima, OH.13−15 Pristine (AMD-unimpacted) soil from the property was collected by excavating the top 3 cm of soil and vegetation, and the underlying soil was placed in sterile containers. Iron mound sediment was collected from the top 2 cm of deposits. Emergent AMD was collected as needed for soil/sediment incubations from a pool upstream of the iron mound and placed in sterile bottles. Materials were placed on ice (for soil/sediment incubations) or dry ice (for nucleic acid-based microbial community analysis) and transported to The University of Akron. Soil, iron mound sediment, and AMD collected for microcosm incubations were stored at 4 °C before use (no more than 1 week). Soil and iron mound sediment for nucleic acid based analysis were stored at −80 °C until further processing. In addition, 1 L of AMD intended for nucleic acid based microbial community analysis was passed through a Supor-200 0.2-μm membrane filter (Pall Corporation, Port Washington, NY) and the membrane was stored at −80 °C before further processing. Soil/Sediment Incubations. Soil/sediment incubation to evaluate Fe(II) oxidizing activities associated with pristine soil and AMD included 4 g of soil in 125-mL Erlenmeyer flasks capped with aluminum foil. For soil/sediment incubations to evaluate shifts in microbial community structure in response to AMD intrusion, 5 g of pristine soil, 5 g of iron mound sediment, or 4 g of soil seeded with 1 g of iron mound sediment was placed in 125-mL Erlenmeyer flasks and capped with aluminum foil. As needed, 50 mL of AMD (filter-sterilized (0.2 μm) or nonsterile) or filter-sterilized synthetic acid mine drainage (SAMD)6 was added to soil/sediment. Fluid samples were periodically obtained from incubations, and pH and dissolved Fe(II) were measured. Soil/sediment incubation slurry pH was measured using an Orion 370 PerpHect pH meter (ThermoFisher Scientific, Waltham, MA). To measure dissolved Fe(II), solids were separated from fluid by centrifugation, and Fe(II) in the supernatant was determined colorimetrically by ferrozine assay16 using a Helios Zeta UV−vis spectrophotometer (ThermoFisher Scientific, Waltham, MA). Incubations were conducted in a semicontinuous format,17 whereby 50% total volume of the fluid layer in each incubation was removed and replaced with fresh AMD or SAMD. Fluid replacement was generally conducted upon removal of ≥95% of dissolved Fe(II) from solution. Incubations that contained nonsterile soil/ sediment were conducted in triplicate. Sterilized control incubations were conducted using autoclaved soil or formaldehyde-treated (1% by volume) iron mound sediment suspended in filter-sterilized AMD or SAMD, and conducted in duplicate. First-order rate constants (k) of Fe(II) oxidation were determined for each microcosm environment after each fluid replacement by least-squares linear regression analysis of Fe(II) concentrations plotted against time18 using the following equation: ln[Fe(II)t ] = − kt + ln[Fe(II)initial ]

Figure 1. (●) Fe(II) concentrations and (○) pH in semicontinuous incubations containing pristine soil and AMD (A), pristine soil and filter-sterilized AMD (B), heat-deactivated soil and AMD (C), and heatdeactivated soil and filter-sterilized AMD (D). The numbers next to plotted Fe(II) concentrations represent first-order rate constants (k) of Fe(II) oxidation (d−1) after each AMD replacement event. Error bars represent one standard deviation.

preparation. Subsequent fluid exchanges were made using filtersterilized SAMD. For experiments to evaluate the changes in microbial communities associated with mixing of AMD with pristine soil (with results shown in Figures 2−5), AMD was initially provided to incubations, and replaced with filtersterilized AMD every 2 days, and freshly collected AMD every 6 days. Filter-sterilized AMD was used for some of the fluid replacement events to avoid adding AMD-associated microbial communities that were altered due to storage. In cases where fluid was replaced with nonsterile AMD, it was added to incubations ≤2 h after collection. After 0, 6, 12, 18, and 24 d of incubation, samples were obtained for culture-dependent enumeration of FeOB and nucleic acid-based analysis of microbial communities (described below). Samples intended for nucleic acid based analysis were stored at −80 °C until further processing. Microbial Enumerations. Culture-dependent enumerations of aerobic acidophilic FeOB were carried out by plate counts using FeTSB medium, which contains Fe(II) as an electron donor.19 Plates were monitored for 3 weeks, and colony forming units (CFU) were counted positive based on the appearance of orange/red colonies. Nucleic Acid-Based Microbial Community Analysis. Prior to DNA extraction, iron mound sediment and samples collected from soil/sediment incubations were washed with 0.3 M ammonium oxalate (pH adjusted to 3 using oxalic acid) to remove Fe(III) prior to DNA extraction.10,20 DNA was extracted

(1)

For experiments to evaluate the relative contributions of soiland AMD-associated microorganisms to the development Fe(II) B



Article

Fe(II) oxidation in incubations that contained nonsterile soil or AMD was microbiologically mediated. Initial rates of Fe(II) oxidation were greatest in incubations that included nonsterile soil and AMD (Figure 1A). Incubations that contained soil with filter-sterilized AMD and autoclaved soil with AMD exhibited relatively low rates of Fe(II) oxidation upon initial mixing of materials (Figure 1B and C). With continued exposure to AMD, rates of Fe(II) oxidation in all incubations increased upon each fluid replacement event, and reached similarly high levels after four fluid replacement events (Figure 1). These results indicate that components of both soil- and AMD-associated microbial communities contribute to the development of rapid Fe(II) oxidizing activities. To evaluate the changes in soil- and AMD-associated microbial communities and activities in response to AMD intrusion, we incubated pristine soil with AMD, and compared the biogeochemical processes associated with these incubations to processes associated with mature iron mound sediment in parallel incubations. As such, processes associated with incubations containing mature iron mound sediment were viewed as a final “steady state” that soil-associated microbial communities might approach with sustained exposure to AMD. Additionally, soil was mixed with a small portion of iron mound sediment (referred to as “IM-seeded soil”) to determine if inoculating soil with mature iron mound sediment might enhance the rate of development of robust Fe(II) oxidizing activities. In incubations that contained iron mound sediment, Fe(II) oxidation occurred concomitantly with a decrease in pH (Figure 2A and D), indicating oxidative precipitation of Fe(II) as Fe(III) (hydr)oxides (R1 and R2). With the first five fluid replacement events (in the first 10 d of incubation), first-order rate constants (k) of Fe(II) oxidation increased, before stabilizing at a k of approximately 0.17 d−1 (Figure 2G). Abundances of culturable FeOB remained at approximately 104 CFU mL−1 throughout the incubations (Figure 2G), indicating that a robust Fe(II) oxidizing microbial community had developed in the iron mound sediment. In incubations that contained pristine soil, Fe(II) oxidation with concomitant pH decrease was observed (Figure 2B and E). First-order rate constants of Fe(II) oxidation increased with each of the first six fluid replacement events, before stabilizing at a k of approximately 0.17 d−1 (Figure 2H). Increased rates of Fe(II) oxidation were accompanied by an increase in the abundances of FeOB in the incubations (Figure 2H). Rates of Fe(II) oxidation increased after fewer fluid replacement events in incubations that contained pristine soil than in incubations that contained iron mound sediment (Figure 2G−I). However, Fe(II) was lost from solution in formaldehydedeactivated control incubations due to abiotic oxidation of Fe(II) and adsorption on solid phases (Figure 2B and C). As such, it is likely that the measured Fe(II) oxidation rates in pristine soil incubations were influenced by the abiotic oxidation of Fe(II), particularly in the first 10 days of the incubations. Incubations that included IM-seeded soil exhibited patterns of Fe(II) oxidation similar to those of incubations that included pristine soil only (Figure 2B and C). Rates of Fe(II) oxidation increased more rapidly than incubations that contained only pristine soil, and exceeded those observed in incubations that contained iron mound sediment. FeOB abundances were similar to those observed in the iron mound sediment-containing incubations (Figure 2G−I), indicating that the development of efficient Fe(II) oxidizing microbial communities can be enhanced by the addition of mature iron mound sediment to soil.

from washed samples, pristine soil, and AMD (immobilized on membrane filter) using MoBio PowerBiofilm DNA isolation kits (MoBio Laboratories, Inc., Carlesbad, CA). Partial 16S rRNA gene sequencing using bacterial tag-encoded FLX amplicon pyrosequencing (bTEFAP) was conducted by Molecular Research Laboratories, LP (Shallowater, TX).21 Universal primers designed from the 515 and 806 positions of the Escherichia coli 16S rRNA gene were used in a 30-cycle, singlestep PCR reaction using HotStarTaq Plus Master Mix Kit (Qiagen, Valencia, CA) using parameters that included initial denaturation at 94 °C for 3 min, 28 cycles of 94 °C for 30 s, 53 °C for 40 s, and 72 °C for 1 min, and a final elongation step at 72 °C for 5 min. Amplified PCR products were mixed in equal concentrations and purified with Agencourt Ampure beads (Agencourt Bioscience Corporation, MA). Sequencing of PCR amplicons was completed with a Roche (Roche Diagnostics Corp., Indianapolis, IN) 454 FLX Titanium instrument and reagents using manufacturer protocols. Sequences were processed initially by removing all barcodes and primers. All sequences that included less than 200 bp, chimeras, ambiguous base calls, and all homopolymers >6 bp were discarded.22 Sequence libraries for this project have been submitted to the Sequence Read Archive (SRA) under accession numbers in Table S1 of the Supporting Information (SI). Sequences were analyzed in the MacQIIME (http://www. wernerlab.org/software/macqiime) environment, using default parameters of QIIME scripts.23 Shannon diversity indices were determined in the QIIME environment for rarefied sequence libraries (6240 sequences) described above. While in the QIIME environment, operational taxonomic units were assigned based on 97% sequence similarity (OTU0.03), and taxonomic assignments were made using the RDP-II classifier function.24 The PyNAST algorithm25 was used against the Greengenes core set26 to align OTU sequences, and a single phylogenetic tree was constructed using FastTree 2.27 Distance matrices were developed using the weighted UniFrac metric28,29 through iterative rarefaction to 6573 sequences by means of Jack-knife sampling using OTU0.03 tables generated from each sample. Temporal shifts in microbial community structure for each longterm microcosm environment were visualized using principle coordinate analysis (PCoA) based on weighted UniFrac distance matrix.28,29 Selected OTU0.03 that comprised large fractions of phyla prominently represented in sequence libraries were compared to sequences contained in the National Center for Biotechnology Information (NCBI) database using the Basic Local Alignment Search Tool (BLASTN).30

■

RESULTS AND DISCUSSION Microbial Activities Associated with Soil and Iron Mound Sediment Incubations. To evaluate the development of Fe(II) oxidizing microbial activities upon intrusion of AMD into pristine soil, incubations were constructed containing pristine soil and emergent AMD from the Mushroom Farm site. These incubations were operated in a “semi-continuous” format,17 and SAMD was used for fluid replacement events after initial incubation construction. With continuous exposure to the acidic and Fe(II) rich conditions of AMD, rates of microbiological Fe(II) oxidation increased, and pH decreased concomitantly with removal of Fe(II) from solution, suggestive of the oxidative precipitation of Fe(II) (R1 and R2; Figure 1). Less oxidative precipitation of Fe(II) was observed in incubations that contained autoclaved soil and filter-sterilized AMD than in nonsterile incubations (Figure 1D), suggesting that C



Article

Figure 2. Fe(II) concentrations (panels A−C), pH (panels D−F), (⧫) first-order rate constants of Fe(II) oxidation (k) after each AMD replacement event and (◊) culturable FeOB abundances (panels G−I), and Shannon Indices (panels J−L) in incubations containing AMD mixed with iron mound sediment (first column), pristine soil (second column), and IM-seeded soil (third column). In panels A−F, open and closed shapes represent data from nonsterile and formaldehydedeactivated incubations, respectively. Dashed lines in panels G−I represent first-order rate constants for Fe(II) oxidation in formaldehyde-deactivated controls. Solid lines in panels J−L represent Shannon Indices of microbial communities associated with iron mound sediment (red lines), AMD (green lines), and pristine soil (blue lines). Error bars in panels A−I represent one standard deviation.

Figure 3. Relative abundances of 16S rRNA gene OTU0.03 detected in libraries from pristine soil, iron mound sediment, AMD, and incubations containing AMD mixed with iron mound sediment, pristine soil, and IM-seeded soil after incubation for 0, 6, 12, 18, and 24 days. Relative OTU0.03 abundances are depicted at phylum-level and class-level (in the cases of the Proteobacteria) taxonomic resolution. Profiles of starting material (i.e., pristine soil, iron mound sediment, and AMD) are provided for reference in the left panels.

Characterization of Microbial Communities Associated with Pristine Soil, Iron Mound Sediment, and AMD. We used 454 pyrosequencing-derived partial 16S rRNA gene sequences to evaluate the microbial community composition of iron mound sediment, AMD, and pristine soil that were used as “starting materials” for soil and iron mound sediment incubations. Information regarding numbers of sequences in each sequence library is provided in Table S1 of the SI. Shannon Indices and rarefaction analysis of sequence libraries indicated that the microbial diversity of iron mound sediment and AMD was lower than that of pristine soil (Figure 2J; Figure S1 of the SI), which is consistent with previous observations of lowdiversity microbial communities associated with AMD-impacted systems.1,31 The soil-derived sequence library contained a wide distribution of phylotypes attributable to several phyla (Figure 3). Whereas the phylum-level distributions of OTU0.03 were more uniform than those observed in other soil surveys,32−34 the most prominent phyla (e.g., Proteobacteria, Planctomycetes, Chloroflexi, Acidobacteria, and Actinobacteria) were similar to those surveys. The majority of 16S rRNA gene sequences detected in AMD (67%) were attributable to Betaproteobacteria and Euryarchaeota, and 63% of sequences detected in the iron mound sediment were attributable to Gammaproteobacteria (Figure 3). One OTU0.03 (Figure 4A) comprised 66% of Euryarchaeotal sequences, and while similar phylotypes to this have been detected in AMD-impacted systems, it could not be reliably assigned using BLASTN to an organism represented in culture. However, using the RDP-II classifier function, this

OTU0.03 could be assigned to the Parvarchaea, which have been detected in AMD-impacted environments.56 Two OTU0.03 (Figure 4B and C) attributable to the neutrophilic FeOB Leptothrix ochracea36 and Sideroxydans lithotrophicus37 comprised 59% of total Betaproteobacterial phylotypes detected in AMD. Alterations to Microbial Communities Associated with Soil and Iron Mound Sediment Incubations. We also used a 454 pyrosequencing-based survey of 16S rRNA gene sequences to evaluate changes in microbial communities that accompanied the development of greater rates of Fe(II) oxidizing activity in the soil/sediment incubations. The diversity (as indicated by Shannon Indices) of microbial communities associated with incubations containing iron mound sediment remained low throughout the incubations, and community diversity decreased over the course of incubations that contained soil (including incubations that contained IM-seeded soil). These results suggest that the relatively harsh geochemical conditions associated with the AMD induced alterations in the soilassociated microbial communities (Figure 2J−L). The alterations in community structure were visualized upon comparison of microbial communities in the starting materials with the incubations that contained those components by principal coordinates analysis (PCoA) using the UniFrac metric (Figure 5). Little deviation from the iron mound was observed in communities associated with iron mound sediment incubations, but microbial communities associated with soil-containing D



Article

Figure 4. Relative abundances of selected abundant OTU0.03 within sequences attributable to the Euryarchaeota, Betaproteobacteria, Gammaproteobacteria, and Firmicutes that were detected in incubations containing AMD mixed with (blue) pristine soil, (red) iron mound sediment, or (pink) IM-seeded pristine soil. OTU0.03 were selected based on their numerically abundant representation in phyla that comprised large fractions of pristine soil, mature iron mound sediment, or AMD. Red, green, and blue dashed lines represent relative abundances of the OTU0.03 in iron mound sediment, AMD, and pristine soil, respectively. The underlying table provides information on the similarity of the respective OTU0.03 to sequences of organisms represented in culture and organisms detected in culture-independent environmental surveys. GenBank accession numbers of sequences used for comparison are provided in Table S2 of the SI.

(Figure 5), indicating that with sustained mixing of AMD with soil, a microbial community structure developed that was more similar to that of the iron mound sediment. The development of a microbial community more similar to the iron mound sediment was enhanced by inoculation of soil with iron mound-associated microorganisms (Figure 5). Similarly, the number of OTU0.03 shared between pristine soil and incubations that contained AMD mixed with soil decreased with longer exposure to AMD (Figure S2A of the SI). The number of OTU0.03 shared between iron mound sediment and incubations that contained AMD mixed with soil increased throughout the incubations (Figure 2B of the SI). This increase in OTU0.03 shared between iron mound sediment and incubations was enhanced by seeding soil with iron mound sediment (Figure S2 of the SI). These results indicate that continuous mixing of pristine soil with AMD leads to the development of more “iron mound-like” microbial communities. A taxonomic evaluation of 16S rRNA gene sequences recovered from incubations that contained iron mound sediment indicated little change in the structure of the microbial communities with sustained exposure to AMD (Figure 3). Throughout the incubations, phylotypes attributable to the Gammaproteobacteria remained numerically dominant in sequence libraries (Figure 3). Approximately 90% of these Gammaproteobacteria-affiliated phylotypes were attributable to Metallibacterium sp. X11, which, along with closely related M.

Figure 5. PCoA of microbial communities associated with iron mound sediment, pristine soil, and AMD (red, blue, and green +, respectively) and incubations containing AMD mixed with iron mound sediment (red shapes), pristine soil soil (blue shapes), and IM-seeded soil mixed (pink shapes) after (●) 0, (■) 6, (⧫) 12, (▲) 18, and (▼) 24 days using weighted UniFrac.28,29 Red circle is to aid in visualizing clustering of iron mound-associated microbial communities. Arrow is to aid in visualizing the separation of soil-associated microbial communities from pristine soil with continuing exposure to AMD.

incubations separated from the pristine soil and toward the iron mound sediment along PCo1 as the incubations progressed E



Article

schef f leri, A4F5, WJ2, and YE-D1-10-CH, constitute a group of acidophilic microorganisms capable of aerobic Fe(II) oxidation and organo- or lithotrophic Fe(III) reduction (Figure 4D).39,42 Iron mound-associated microbial communities appeared to be quite resistant to influence by AMD-associated microorganisms, likely since these communities were well adapted to continuous intrusion of AMD. Indeed, phylotypes attributable to neutrophilic FeOB that were detected in AMD diminished in abundance over the course of incubations that contained iron mound sediment (Figure 4B and C). Taxonomic evaluation of 16S rRNA gene sequences recovered from incubations that contained pristine soil and AMD revealed an increase in the relative abundances of phylotypes attributable to Gammaproteobacteria, Firmicutes, and Betaproteobacteria with sustained exposure of previously pristine soil to AMD (Figure 3). Although OTU0.03 assignable to Metallibacterium spp. (Figure 4D) increased in relative abundance within Gammaproteobacterial phylotypes, OTU0.03 assignable to the closely related acidophilic FeOB A4F5 (Figure 4E) comprised a progressively larger fraction of the Gammaproteobacteria as soil was incubated with AMD. Similarly, an OTU0.03 (Figure 4F) attributable to the acidophilic FeOB Ferrovum myxofaciens44 increased in relative abundance within the Betaproteobacteria, reaching a relative abundance comparable to that of the mature iron mound sediment. An additional OTU0.03 (Figure 4G) attributable to the acidophilic FeOB C4C642 increased in relative abundance with sustained incubation of soil with AMD, but this phylotype did not comprise a large fraction of Betaproteobacterial sequences in mature iron mound sediment. Additionally, OTU0.03 attributable to Herbispirillum spp.46,47 (Figure 4H) comprised a fraction of the Betaproteobacterial sequences in pristine soil-containing incubations similar to that of iron mound-containing incubations at the conclusion of the incubations. The role of these organisms in the context of acidophilic Fe(II) metabolism is unclear, since these organisms are organotrophic and generally encountered in circumneutral settings.46,47 The increase in relative abundance of Firmicutesaffiliated phylotypes during incubation of pristine soil with AMD (Figure 3) is likely attributable to the ability of these organisms to sporulate, since most OTU0.03 were attributable to Bacillus spp., whose relative abundances within the Firmicutes did not change throughout the incubations (Figure 4I). Taxonomic distributions of 16S rRNA gene sequences recovered from incubations that contained IM-seeded soil and AMD exhibited patterns similar to those observed in soilcontaining incubations when continuously exposed to AMD (Figure 3). We observed increased relative abundances of Gammaproteobacteria-, Firmicutes-, and Betaproteobacteriaaffiliated phylotypes, which were mostly attributable to acidophilic FeOB (in the cases of Gamma- and Beta-proteobacterial phylotypes) and Bacillus spp. (Figure 4). Although these patterns of microbial community response to AMD intrusion were similar to those observed in soil incubations that were not IM-seeded, the increase in relative abundances of Gammaproteobacterial phylotypes proceeded more rapidly in the IM-seeded incubations (Figure 3). These results indicate that the addition of mature iron mound sediment-associated microoganisms enhanced the development of microbial communities that were capable of efficient Fe(II) oxidation. Sources of FeOB. The results of experiments that included incubations of pristine soil with AMD, pristine soil with filtersterilized AMD, and autoclaved soil with AMD indicated that the development of high rates of Fe(II) oxidizing activity was most

rapid in the presence of active microorganisms in both soil and AMD (Figure 1). Evaluation of the microbial communities upon mixing of soil and AMD led to a similar conclusion, as the acidophilic FeOB-attributable Gamma- and Beta-proteobacterial phylotypes that were abundant in libraries from iron mound incubations were detected in both soil and AMD (albeit at relatively low abundances). These phylotypes increased in relative abundance with sustained incubation of soil with AMD (Figure 4D−G). The acidophilic FeOB detected in pristine soil, which increased in relative abundance upon exposure to AMD, may represent components that are common to many soils and are part of the “rare biosphere” of soils.32,60,61 Whereas OTU0.03 shared between soil and soil-containing incubations declined (Figure S2A of the SI), OTU0.03 shared between iron mound sediment and soil-containing incubations increased during incubations (Figure S2B of the SI). An initial increase in OTU0.03 shared between AMD and soil-containing incubations after 6 d incubation was followed by a decrease in shared OTU0.03 (Figure S2C of the SI). Euryarchaeota-attributable phylotypes, which were relatively abundant in the AMD-derived sequence library, were not prominent components of microbial communities in incubations (Figure 3). Phylotypes attributable to neutrophilic Betaproteobacterial FeOB L. ochracea36 and S. lithotrophicus37 initially increased in incubations, but subsequently decreased over time (Figure 4B and C). The pH of emergent AMD at the MF is approximately 4.5,13 which may have been tolerable for these organisms, but sustained oxidative precipitation of Fe and consequent pH decrease (Figure 1) favored the development of microbial communities in which acidophilic Metallibacterium and Ferrovum spp. were the most prominent FeOB (Figure 4D−G). These results indicate that the initial development of FeOB communities may be largely attributable to inoculation of soil by AMD, but with sustained intrusion of AMD into soil, FeOB that are initially less abundant comprise a larger fraction of the FeOB community. Environmental Implications. Our results indicate that the intrusion of AMD into soil that was not previously impacted by acidic, Fe(II)-rich AMD fluids leads to rapid changes in soil- and AMD-associated microbial communities, culminating in communities that are capable of efficient oxidative precipitation of Fe from coal mine-derived AMD, such as that observed at the Mushroom Farm and other iron mounds.10−12,62 These iron mound systems may serve as models upon which engineered systems mimicking the hydrodynamics of mature iron mounds may be exploited for inexpensive and sustainable AMD treatment.10−12,62 We have demonstrated the dynamics of this process in the absence of human intervention to stimulate Fe(II) oxidation, but our results also indicate that the development of robust Fe(II) oxidizing activities may be enhanced by the addition of material (and associated microbial communities) from mature iron mounds.

■

ASSOCIATED CONTENT

S Supporting Information *

Two tables providing information on pyrosequencing reads (with SRA accession numbers) and accession numbers of GenBank sequences in Figure 4. Two figures show rarefaction curves derived from sequence libraries and shared OTU0.03 between systems. This material is available free of charge via the Internet at http://pubs.acs.org/. F



■

Article

(17) Roden, E. E.; Urrutia, M. M. Ferrous iron removal promotes microbial reduction of crystalline iron(III) oxides. Environ. Sci. Technol. 1999, 33, 1847−1853. (18) Senko, J. M.; Bertel, D.; Quick, T. J.; Burgos, W. D. The influence of phototrophic biomass on Fe and S redox cycling in an acid mine drainage-impacted system. Mine Water Environ. 2011, 30, 38−46. (19) Johnson, D. B. Selective solid media for isolating and enumerating acidophilic bacteria. J. Microbiol. Methods 1995, 23, 205−218. (20) Nicomrat, D.; Dick, W. A.; Tuovinen, O. H. Assessment of the microbial community in a constructed wetland that receives acid coal mine drainage. Microb. Ecol. 2006, 51, 83−89. (21) Dowd, S. E.; Callaway, T. R.; Wolcott, R. D.; Sun, Y.; McKeehan, T.; Hagevoort, R. G.; Edrington, T. S. Evaluation of the bacterial diversity in the feces of cattle using 16S rDNA bacterial tag-encoded FLX amplicon pyrosequencing (bTEFAP). BMC Microbiol. 2008, 8, 125. (22) Gontcharova, V.; Youn, E.; Wolcott, R. D.; Hollister, E. B.; Gentry, T. J.; Dowd, S. E. Black Box Chimera Check (B2C2): a Windows-based software for batch depletion of chimeras from bacterial 16S rRNA gene datasets. Open Microbiol. J. 2010, 4, 47−52. (23) Caporaso, J. G.; Kuczynski, J.; Stombaugh, J.; Bittinger, K.; Bushman, F. D.; Costello, E. K.; Fierer, N.; Pena, A. G.; Goodrich, J. K.; Gordon, J. I.; Huttley, G. A.; Kelley, S. T.; Knights, D.; Koenig, J. E.; Ley, R. E.; Lozupone, C. A.; McDonald, D.; Muegge, B. D.; Pirrung, M.; Reeder, J.; Sevinsky, J. R.; Tumbaugh, P. J.; Walters, W. A.; Widmann, J.; Yatsunenko, T.; Zaneveld, J.; Knight, R. QIIME allows analysis of highthroughput community sequencing data. Nat. Methods 2010, 7, 335− 336. (24) Wang, Q.; Garrity, G. M.; Tiedje, J. M.; Cole, J. R. Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 2007, 73, 5261−5267. (25) Caporaso, J. G.; Bittinger, K.; Bushman, F. D.; DeSantis, T. Z.; Andersen, G. L.; Knight, R. PyNAST: A flexible tool for aligning sequences to a template alignment. Bioinformatics 2010, 26, 266−267. (26) DeSantis, T. Z.; Hugenholtz, P.; Larsen, N.; Rojas, M.; Brodie, E. L.; Keller, K.; Huber, T.; Dalevi, D.; Hu, P.; Andersen, G. L. Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Appl. Environ. Microbiol. 2006, 72, 5069−5072. (27) Price, M. N.; Dehal, P. S.; Arkin, A. P. FastTree 2 − Approximately maximum likelihood trees for large alignments. PLoS One 2010, 5, 3. (28) Lozupone, C.; Knight, R. UniFrac: A new phylogenetic method for comparing microbial communities. Appl. Environ. Microbiol. 2005, 71, 8228−8235. (29) Lozupone, C.; Hamady, M.; Kelly, S. T.; Knight, R. Quantitative and qualitative β diversity measures lead to different insights into factors that structure microbial communities. Appl. Environ. Microbiol. 2007, 73, 1576−1585. (30) Altschul, S. F.; Madden, T. L.; Schagger, A. A.; Zhang, J.; Zhang, Z.; Miller, W.; Lipman, D. J. Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res. 1997, 25, 3389−3402. (31) Hallberg, K. B.; Coupland, K.; Kimura, S.; Johnson, D. B. Macroscopic streamer growths in acidic, metal-rich mine waters in North Wales consist of novel and remarkably simple bacterial communities. Appl. Environ. Microbiol. 2006, 72, 2022−2030. (32) Elshahed, M. S.; Youssef, N. H.; Spain, A. M.; Sheik, C.; Najar, F. Z.; Sukharnikov, L. O.; Roe, B. A.; Davis, J. P.; Schloss, P. D.; Bailey, V. L.; Krumholz, L. R. Novelty and uniqueness patterns of rare members of the soil biosphere. Appl. Environ. Microbiol. 2008, 74, 5422−5428. (33) Sheik, C. S.; Beasley, W. H.; Elshahed, M. S.; Zhou, X.; Luo, Y.; Krumholz, L. R. Effect of warming and drought on grassland microbial communities. ISME J. 2011, 5, 1692−1700. (34) Lauber, C. L.; Ramirez, K. S.; Aanderud, Z.; Lennon, J.; Fierer, N. Temporal variability in soil microbial communities across land-use types. ISME J. 2013, 7, 1641−1650. (35) Amaral-Zettler, L.; Zettler, E. R.; Theroux, S. M.; Palacios, C.; Aguilera, A.; Amils, R. Microbial community structure across the tree of life in the extreme Rió Tinto. ISME J. 2011, 5, 42−50.

AUTHOR INFORMATION

Corresponding Author

*Phone: +1 330-972-8047; e-mail: [email protected]; mail: Department of Geosciences, The University of Akron, 126 Crouse Hall, Akron, OH 44325, USA. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This work was funded by The Ohio Water Resources Research Center and National Science Foundation EAR Geobiology and Low Temperature Geochemistry award 0851847. We thank Cheryl Socotch from the Ohio Department of Natural Resources for valuable background information on the MF. We thank John Wilson, the Mushroom Farm land manager, for allowing access to the site. We thank Amy Milsted (UA Department of Biology) for her contributions to the work.

■

REFERENCES

(1) Baker, B. J.; Banfield, J. F. Microbial communities in acid mine drainage. FEMS Microbiol. Ecol. 2003, 44, 139−152. (2) Cravotta, C. A., III Dissolved metals and associated constituents in abandoned coal-mine discharges, Pennsylvania, USA. Part 1: Constituent quantities and correlations. Appl. Geochem. 2008, 23, 166−202. (3) Herlihy, A. T.; Kaufman, P. R.; Mitch, M. E. Regional estimates of acid mine drainage impact on streams in the Mid-Atlantic and Southeastern United States. Water, Air, Soil Pollut. 1990, 50, 5854− 5861. (4) Younger, P. L. The longevity of minewater pollution: A basis for decision-making. Sci. Total Environ. 1997, 194/195, 457−466. (5) United States Geologic Survey (USGS). Coal Mine Drainage Projects in Pennsylvania; USGS. http://pa.water.usgs.gov/projects/ energy/amd (March 31, 2014). (6) Cravotta, C. A. III; Trahan, M. K. Limestone drains to increase pH and remove dissolved metals from acidic mine drainage. Appl. Geochem. 1999, 14, 581−606. (7) Johnson, D. B.; Hallberg, K. B. Acid mine drainage remediation options: A review. Sci. Total Environ. 2005, 338, 3−14. (8) Ziemkiewicz, P. F.; Skousen, J. G.; Simmons, J. Acid mine drainage treatment with armored limestone in open limestone channels. Mine Water Environ. 2003, 22, 118−129. (9) Johnson, D. B.; Hallberg, K. B. Pitfalls of passive mine water treatment. Re/Views Environ. Sci. Bio/Technol. 2002, 1, 335−343. (10) Senko, J. M.; Wanjugi, P.; Lucas, M.; Bruns, M. A.; Burgos, W. D. Characterization of Fe(II) oxidizing bacterial activities and communities at two acidic Appalachian coalmine drainage-impacted sites. ISME J. 2008, 2, 1134−1145. (11) DeSa, T. C.; Brown, J. F.; Burgos, W. D. Laboratory and field-scale evaluation of low-pH oxidation at Hughes Borehole, Portage, Pennsylvania. Mine Water Environ. 2010, 29, 239−247. (12) Brown, J. F.; Jones, D. S.; Mills, D. B.; Macalady, J. L.; Burgos, W.D. Application of a depositional facies model to an acid mine drainage site. Appl. Environ. Microbiol. 2012, 77, 545−554. (13) Gouin, M.; Saracusa, E.; Clemons, C. B.; Senko, J.; Kreider, K. L.; Young, G. W. A mathematical model of a passive scheme for acid mine drainage remediation. GEM − Int. J. Geomath. 2013, 4, 27−53. (14) Cutright, T. J.; Senko, J.; Sivaram, S.; York, M. Evaluation of the phytoextraction potential at an acid mine drainage-impacted site. Soil Sed. Contam. Int. J. 2012, 21, 970−984. (15) Brantner, J. S.; Haake, Z. J.; Burwick, J. E.; Menge, C. M.; Hotchkiss, S. T.; Senko, J. M. Depth-dependent geochemical and microbiological gradients in Fe(III) deposits resulting from coal minederived acid mine drainage. Front. Microbiol. 2014, submitted. (16) Stookey, L. L. Ferrozine: A new spectrophotometric reagent for iron. Anal. Chem. 1970, 42, 779−781. G



Article

(36) Fleming, E. J.; Langdon, A. E.; Martinez-Garcia, M.; Stepanauskas, R.; Poulton, N. J.; Masland, E. D. P.; Emerson, D. What’s new is old: Resolving the identity of Leptothrix ochracea using single cell genomics, pyrosequencing and FISH. PLoS One 2011, 6, 17769. (37) Weiss, J. V.; Rentz, J. A.; Plaia, T.; Neubauer, S. C.; Merrill-Floyd, M.; Lilburn, T.; Bradburne, C.; Megonigal, J. P.; Emerson, D. Characterization of neutrophilic Fe(II)-oxidizing bacteria isolated from the rhizosphere of wetland plants and description of Ferritrophicum radicicola gen. nov., and Sideroxydans paludicola sp. nov. Geomicrobiol. J. 2007, 24, 559−570. (38) Roden, E. E.; McBeth, J. M.; Blöthe, M.; Percak-Dennett, E. M.; Fleming, E. J.; Hollyoke, R. R.; Luther, G. W., III; Emerson, D.; Schieber, J. The microbial ferrous wheel in a neutral pH groundwater seep. Front. Microbiol. 2012, 3, 172. (39) Delavat, F.; Lett, M.-C.; Lièvremont, D. Novel and unexpected bacterial diversity in an arsenic-rich ecosystem revealed by culture dependent approaches. Biol. Direct 2012, 7, 28. (40) Guo, X.; Yin, H.; Cong, J.; Dai, Z.; Liang, Y.; Liu, X. RubisCO gene clusters found in a metagenome microarray from acid mine drainage. Appl. Environ. Microbiol. 2013, 79, 2019−2026. (41) Falteisek, L.; Cepička, I. Microbiology of diverse acidic and nonacidic microhabitats within a sulfidic ore mine. Extremophiles 2012, 16, 911−922. (42) Wu, X.; Wong, Z. L.; Sten, P.; Engblom, S.; Osterholm, P.; Dopson, M. Microbial community potentially responsible for acid and metal release from an Ostrobothnian acid sulfate soil. FEMS Microbiol. Ecol. 2013, 84, 555−563. (43) Santofimia, E.; González-Toril, E.; López-Pamo, E.; Gomariz, M.; Amils, R.; Aguilera, A. Microbial diversity and its relationship to physicochemical characteristics of the water in two extreme acidic pit lakes from the Iberian Pyrite Belt (SW Spain). PLoS One 2013, 8, 66746. (44) Johnson, D. B.; Hallberg, K. B.; Hedrich, S. Uncovering a microbial enigma: Isolation and characterization of the streamergenerating, iron-oxidizing, acidophilic bacterium Ferrovum myxofaciens. Appl. Environ. Microbiol. 2014, 80, 672−680. (45) Bohorquez, L. C.; Delgado-Serrano, L.; López, G.; Osorio-Forero, C.; Klepac-Ceraj, V.; Junca, H.; Zanbrano, M. M. In-depth characterization via complementing culture-independent approaches of the microbial community in an acidic hot spring of the Colombian Andes. Microb. Ecol. 2012, 63, 103−115. (46) Hoque, M. S.; Broadhurst, L. M.; Thrall, P. H. Genetic characterization of root-nodule bacteria associated with Acacia salicina and A. stenopylla (Mimosaceae) across south-eastern Autralia. Int. J. Syst. Evol. Microbiol. 2011, 61, 299−309. (47) Dobritsa, A. P.; Reddy, M. C. S.; Samadpour, M. Reclassification of Herbaspirillum putei as a later heterotypic synonym of Herbispirillum huttiense, with the description of H. huttiense subsp. huttiense subsp. nov. and H. huttiense subsp. putei subsp. nov., comb. nov., and description of Herbaspirillum aquaticum sp. nov. Int. J. Syst. Evol. Microbiol. 2010, 60, 1418−1426. (48) Cretoiu, M. S.; Korthals, G. W.; Visser, J. H. M.; van Elsas, J. D. Chitin amendment increases soil suppressiveness toward plant pathogens and modulates the Actinobacterial and Oxalobacterial communities in an experimental agricultural field. Appl. Environ. Microbiol. 2013, 79, 5291−5301. (49) Hoyles, L.; Honda, H.; Logan, N. A.; Halket, G.; La Ragione, R. M.; McCartney, A. L. Recognition of greater diversity of Bacillus species and related bacteria in human faeces. Res. Microbiol. 2012, 163, 3−13. (50) Agarwal, S.; Gupta, M.; Choudhury, B. Bioprocess development for nicotinic acid hydroxamate synthesis by acyltransferase activity of Bacillus smithii strain IITR6b2. J. Ind. Microbiol. Biotechnol. 2013, 40, 937−946. (51) Lepleux, C.; Uroz, S.; Collignon, C.; Churin, J.-L.; Turpault, M.P.; Frey-Klett, P. A short term mineral amendment impacts the mineral weathering bacterial communities in an acidic forest soil. Res. Microbiol. 2013, 164, 729−739. (52) Marasco, R.; Rolli, E.; Fusi, M.; Cherif, A.; Abou-Hadid, A.; ElBahairy, U.; Borin, S.; Sorlini, C.; Daffonchio, D. Plant growth promotion potential is equally represented in diverse grapevine root-

associated bacterial communities from different biopedoclimatic environments. BioMed. Res. Int. 2013, 2013, 491091. (53) Dunbar, J.; Eichorst, S. A.; Gallegos-Graves, L. V.; Silva, S.; Xie, G.; Hengartner, N. W.; Evans, R. D.; Hungate, B. A.; Jackson, R. B.; Megonigal, J. P.; Schadt, C. W.; Vilgalys, R.; Zak, D. R.; Kuske, C. R. Common bacterial responses in six ecosystems exposed to 10 years of elevated atmospheric carbon dioxide. Environ. Microbiol. 2012, 14, 1145−1158. (54) Kumar, N.; Omoregie, E. O.; Rose, J.; Masion, A.; Lloyd, J. R.; Diels, L.; Bastiaens, L. Inhibition of sulfate reducing bacteria in aquifer sediment by iron nanoparticles. Water Res. 2014, 51, 64−72. (55) Partanen, P.; Hultman, J.; Paulin, L.; Auvinen, P.; Romantschuk, M. Bacterial diversity at different stages of the composting process. BMC Microbiol. 2010, 10, 94. (56) Baker, B. J.; Comolli, L. R.; Dick, G. J.; Hauser, L. J.; Hyatt, D.; Dill, B. D.; Land, M. L.; VanBerkmoes, N. C.; Hettich, R. L.; Banfield, J. F. Enigmatic, ultrasmall, uncultivated Archaea. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 8806−8811. (57) Coupland, K.; Johnson, D. B. Evidence for the potential for dissimilatory iron reduction is widespread among acidophilic heterotrophic bacteria. FEMS Microbiol. Lett. 2008, 279, 30−35. (58) Lu, S.; Gischkat, S.; Reiche, M.; Akob, D. M.; Hallberg, K. B.; Küsel, K. Ecophysiology of Fe-cycling bacteria in acidic sediments. Appl. Environ. Microbiol. 2010, 76, 4272−4281. (59) Ziegler, S.; Waidner, B.; Itoh, T.; Schumann, P.; Spring, S.; Gescher, J. Metallibacterium schlef f leri gen. nov., sp. nov., an alkalinizing gammaproteobacterium isolated from an acidic biofilm. Int. J. Syst. Evol. Microbiol. 2013, 63, 1499−1504. (60) Sogin, M. L.; Morrison, H. G.; Huber, J. A.; Welch, D. M.; Huse, S. M.; Neal, P. R.; Arrieta, J. M.; Herndl, G. J. Microbial diversity in the deep sea and the underexplored rare biosphere. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 12115−12120. (61) Galand, P. E.; Casamyor, E. O.; Kirchman, D. L.; Lovejoy, C. Ecology of the rare microbial biosphere of the Arctic Ocean. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 22427−22432. (62) Larson, L. N.; Fitzgerald, M.; Singha, K.; Gooseff, M. N.; Macalady, J. L.; Burgos, W. Hydrogeochemical niches associated with hyporheic exchange beneath an acid mine drainage-contaminated system. J. Hydrol. 2013, 501, 163−174.

H


Response of Soil-Associated Microbial Communities to Intrusion of

Recommend Documents