the Role of Reactive Oxygen Species - ACS Publications

Subscriber access provided by Queen Mary, University of London

Article

Responses of Microbial Community Structure in Fe(II)-bearing Sediments to Oxygenation: the Role of Reactive Oxygen Species Sicong Ma, Man Tong, Songhu Yuan, and Hui Liu ACS Earth Space Chem., Just Accepted Manuscript • DOI: 10.1021/ acsearthspacechem.8b00189 • Publication Date (Web): 04 Apr 2019 Downloaded from http://pubs.acs.org on April 5, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Earth and Space Chemistry

Responses of Microbial Community Structure in Fe(II)-bearing Sediments to Oxygenation: the Role of Reactive Oxygen Species Sicong Ma†, Man Tong*,†, Songhu Yuan†, Hui Liu†,‡ †State

Key Laboratory of Biogeology and Environmental Geology, China University

of Geosciences, 388 Lumo Road, Wuhan, 430074, P. R. China ‡School

of Environmental Studies, China University of Geosciences, 388 Lumo Road,

Wuhan, 430074, P. R. China

RECEIVED DATE (to be automatically inserted after your manuscript is accepted if required according to the journal that you are submitting your paper to)

* To whom correspondence should be addressed. E-mail: [email protected] (M. Tong), Phone: +86-27-67848629, Fax: +86-27-67883456.

ACS Paragon Plus Environment

ACS Earth and Space Chemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ABSTRACT Responses of microbial community structure in subsurface to oxygen (O2) have been previously attributed to fundamental changes in microbial metabolic lifestyles. Here we show that reactive oxygen species (ROS) produced from oxygenation of Fe(II)-bearing sediments play an important role in the variation of microbial community structure. Sediments sampled along a vertical redox gradient were exposed to laboratory air. After a 10-h oxygenation, the living bacteria counts in sediments at the depth of 100 cm decreased from 6.1 to 5.7 orders of magnitude, 4.39 mg/g Fe(II) was oxidized and 27.6 μM hydroxyl radicals ( • OH) was produced. Bacteria inactivation correlated well with • OH produced upon Fe(II) oxygenation. Control experiments with the addition of ROS quencher and hydrogen peroxide further proved that the bacteria inactivation was induced by ROS rather than O2. 16S rRNA analysis indicated that the relative abundance of Anaerolineaceae increased from 15.4% to 29.9% but of Nistrospiraeceae and Geobacteraceae decreased, pointing to the suppression of nitrification, denitrification and iron respiration but promotion of organic matter degradation. These findings reveal an important but overlooked mechanism for the influence of O2 on microbial community structure.

KEYWORDS: Microbial community structure, Fe redox cycling, Reactive oxygen species, Subsurface sediments, Antimicrobial effect

INTRODUCTION Microbes in subsurface play an crucial role in a number of biogeochemical


Page 2 of 37

Page 3 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


processes including the cycling of nutrients (i.e., C, N, P, S) and the transformation of organic and inorganic matters.1 Microbes are sensitive to environmental changes (i.e., O2,2,3 temperature,4 salinity,5 alkalinity,6 dissolved organic matters7 and nutrients8), among which O2 is a strong selective force for microbes as a preferred electron acceptor. It has been well documented that redox conditions regulate the microbial abundance, diversity, and community structure in subsurface.9,10 Distinctive microbial community structures has been found along the vertical profiles of sediments as the redox conditions shift from the oxic surface zones to the anoxic deeper zones.9‒14 Aerobic microbes dominating in oxic zones require different enzymatic and metabolic capabilities from the anaerobic microbes in anoxic zones.11,14 However, in many natural and artificial processes (e.g., surface water and groundwater interactions and riverbank filtration treatment), O2 may intrude into anoxic subsurface,15‒18 which likely induces the changes of microbial communities and subsequently affects the associated biogeochemical processes. In order to gain more insights into the biogeochemical processes involved in these redox fluctuating zones, it is crucial to know how O2 perturbations influence subsurface microbial communities. The perturbation of O2 changes the availability of electron acceptors in subsurface, which could lead to fundamental changes in microbial metabolic lifestyles.9 The responses of microbial community structures in sediments to O2 perturbation have been largely documented. Bryant et al. observed a pronounced increase in the activity of metal-reducing communities upon sediments oxygenation.2 Danczak et al. found that the abundance and activity of chemolithoautotrophs and heterotrophs were



positively correlated with O2 concentration.19 Wang et al. also observed an enrichment of heterotrophic microorganisms with elevated O2 concentration, whereas they observed a pronounced decrease in the abundance of nitrate-reducing bacteria when the concentration of O2 increased.20 Overall, microbial communities in sediments are dynamic assemblages of populations bearing different tolerances toward O2.9,21 In the face of O2 perturbation, the obligate anaerobes may be inactivated, while the aerobes and facultative microorganisms as well as the microbes with tolerance mechanisms to redox stress might be enriched.22‒24 The combined outcome results in changing community structure. In addition to these direct effects on microbes, O2 may cause indirect oxidative stress on microbes by producing reactive oxygen species (ROS) through the interaction with reducing components in subsurface.25 Fe(II) is an abundant component in subsurface environment under reducing conditions.26 Production of ROS (i.e., •O2-, H2O2, •OH) from the interaction of O2 with Fe(II)-bearing sediments27 and various Fe(II)-bearing minerals (i.e., nontronite,28 mackinawite,29 pyrite30 and magnetite31) has been confirmed in recent years. ROS, especially •OH (standard reduction potential: 2.8V), has strong oxidizing ability and could induce serious oxidative stress to living organisms.32 Studies have been reported on the antimicrobial effect of ROS derived from Fe(II) oxidation against bacteria (i.e., Escherichia coli,33,34 Staphylococcus aureus,35 Shewanella oneidensis MR-1,25 cyanobacteria36), virus (i.e. MS2, ФX174)37 and fungus (i.e. Aspergillus versicolor)38. Generally, the toxicity of ROS comes from their ability to oxidize a large number of cellular constituents such as DNA disruption,39,40 oxidation of proteins and amino


Page 4 of 37

Page 5 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


acids,41 and lipid peroxidation of membrane fatty acids.42 Although the production of ROS upon oxygenation of Fe(II)-bearing sediments and the biotoxicity of ROS against microbes have been largely reported, little is known about whether the ROS produced upon oxygenation of Fe(II)-bearing sediments impact the microbial diversity, abundance and community structure in subsurface environment. In this study, we took four sediment samples along a depth gradient representing different redox conditions and Fe(II) contents. The variations of living bacteria counts, the oxidation of Fe(II) and the production of •OH were measured during the sediments oxygenation. The influence of O2 partial pressure and H2O2 concentration was studied. The changes of community structures of indigenous microorganisms upon oxygenation were tested by 16S rRNA analysis. Our main objective is to unravel the response of microbial community to the oxygenation of Fe(II)-bearing sediments and the potential role of ROS in this process.

EXPERIMENTAL METHODS Chemicals. Sodium benzoic (99.5%), p-hydroxybenzoic acid (p-HBA, 99%), ethanol (99.7%), hydrogen peroxide (H2O2, 30%), 2,2’-bipyridine (BPY, 99.5%) and 1, 10-o-phenanthroline were purchased from Sinopharm Chemical Reagent Co., Ltd., China. BacLight LIVE-DEAD kit (SYTO 9 and PI) was obtained from Invitrogen (USA).N,N-diethyl-p-phenylenediamine (DPD,98%) was acquired from Aike Reagent Co., China. Deionized (DI) water (18.2 MΩ·cm) produced from a Heal Force NW ultrapure water system was used in all of the experiments. All of other chemicals used



in this experiment were of above analytical grade. Sediment Sampling. A total of 4 sediment samples were taken from a river-groundwater interaction zone which located in Jianghan Plain, central China (114.42890°E, 30.53312°N). This sampling site was chosen because our previous study observed a high production of • OH upon oxygenation of sediments here.27 Sediments were taken by a 45-mm gouge auger at the depth of 20, 40, 60 and 100 cm, reflecting the profile of redox conditions and Fe(II) contents. The sediments at the depth of 20 and 40 cm were within oxic zone, the 60-cm sediments were within sub-oxic zone, and the 100-cm sediments were within anoxic zone. The groundwater table was 50 cm below the ground when sampling. Sediments were wrapped successively by a aluminum foil and sealed in vacuum bags immediately, and subsequently they were transported to the lab and stored at 4 ℃. Analysis of Fe(II), total Fe, •OH production and microorganisms were done within 2 weeks. Batch Experiments. Batch experiments were conducted in serum bottles which wrapped in aluminum foil to avoid light interaction, and all of the experiments were carried out in duplicate. Five grams of sediments were mixed with 50 mL of deoxygenated DI water in an anaerobic glove box (COY, U.S.A.) filled with 92% N2 and 8% H2. The unsealed serum bottles were then moved out from the glove box and exposed to laboratory air for oxygenation in a shaking incubator (25 ± 2℃, 220 rpm). At different time intervals (0, 2, 4, 8, 10 h), about 3 mL of suspensions were taken out for the analysis of living bacteria counts and Fe(II) contents. To determine the cumulative concentrations of • OH produced during sediments oxygenation, 10 mM


Page 6 of 37

Page 7 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


BA was added into the sediment suspensions to capture the produced •OH. At different time intervals (0, 2, 4, 8, 10 h), about 2 mL of suspension was taken out and filtered through a 0.22-μm membrane, and 1 mL of filtrate was immediately mixed with 1 mL of methanol to quench the further produced •OH. The cumulative concentration of •OH was probed by the transformation of BA to p-HBA. Control experiments in the absence of O2 were carried out in the anaerobic glove chamber, and another control experiment was performed with the addition of 100 mM ethanol to quench •OH. To explore the influence of O2 partial pressure on bacteria inactivation, batch experiments were carried out in sealed serum bottles in a sacrifice mode to avoid the interference of laboratory air during sampling. One gram of sediments at the depth of 100 cm was mixed with 10 mL of deoxygenated water in 25-mL serum bottles in an anaerobic glove chamber. Then the bottles were sealed with thick butyl rubber stoppers and an aluminum cap. A 0.8-mL and 1.2-mL of air was injected into the container by a syringe to obtain an initial O2 partial pressure of 0.0168 atm and 0.0112 atm, respectively. At different time intervals (0, 1, 2, 4, 8, 10 h), two bottles were sacrificed for the analysis of living bacteria counts and Fe(II) contents, another two bottles were sacrificed for the analysis of cumulative concentrations of •OH. To test the effect of H2O2 on bacteria inactivation, batch experiments were also carried out in sealed serum bottles in a sacrifice mode. 1, 10, 50 and 100 mM of H2O2 was respectively added into the sediment suspension which containing 1 gram of sediment and 10 mL of deoxygenated water in an anaerobic glove chamber. The



containers were sealed with thick butyl rubber stoppers and an aluminum cap to avoid laboratory air. After a 2-h reaction, samples were taken out for analysis of the living bacteria number and residual concentration of H2O2. Analysis. Living bacteria were quantified by a microscopic quantitative assay using the BacLight LIVE-DEAD viability kit. A 2-mL sediment suspension was mixed with 0.9 mL of 5% ethanol and 0.1 mL of 27% NaCl solution. After settling for 10 min, 1 mL of the supernatant was taken out and mixed with 1 μL of SYTO 9 and 1 μL of propidium iodide (PI). After a 15-min reaction, the cells were filtered out by a 0.2 μm nuclepore filter (Whatman, polycarbonate) and observed under a fluorescent microscope (DM2500, Leica). The red-flurescent nucleic acid stain PI penetrate only dead cells with disrupted membranes, while the green-fluorescent nucleic acid stain SYTO9 enters all bacterial cells (live and dead). Since PI exhibits a stronger affinity for nucleic acids than SYTO9, SYTO9 will be displaced by PI and the cells will fluoresce in red when both dyes are present.43 Therefore, cells with a compromised membrane that are considered to be dead or dying stained red, whereas cells with an intact membrane stained green, we counted the number of green cells as living bacteria. Fe(II) and total Fe in sediments were measured after digestion by 6 M HCl for 24 hours. Fe(II) was measured by the 1, 10-o-phenanthroline analytical method at 510 nm. Total Fe concentrations were assayed through the reduction of Fe3+ to Fe2+ by hydroxylamine-HCl. The cumulative concentration of •OH was measured using the transformation of


Page 8 of 37

Page 9 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


benzoic acid to p-HBA as a probe reaction.44 The concentration of p-HBA was measured in an LC-15C HPLC (Shimadzu) equipped with a UV detector and an Inter Sustain C18 column (4.6 × 250 mm). The cumulative •OH concentration was estimated by p-HBA concentration with a conversion factor of 5.87. The detection limit was 0.1 μM for p-HBA, which corresponds to 0.59 μM for •OH.44 H2O2 was analyzed by a modified N,N-diethyl-p-phenylenediamine (DPD) method at 551 nm using a UV-vis spectrophotometer.45 During the measurement, BPY and Na2EDTA were added to complex Fe2+ and Fe3+, respectively. DNA Extraction. Genomic DNA extraction and further analysis were conducted by Shanghai Personal Biotechnology Co., Ltd., China. DNA was extracted from sediments by Soil DNA Kit (Omega, D5625-01) following the manufacturer's protocol. The extracted genomic DNA was quantified and checked for purity at A260/280 nm by UV spectrophotometer (Eppendorf, Bio Photometer). PCR Amplification of 16S rRNA Gene and Sequencing. The polymerase chain reaction (PCR) amplification was performed with the bacterial V4-16S rRNA gene specific primers 520F (5’-GCACCTAAYTGGGYDTAAAGNG-3’) with a unique barcode and 802R (5’-TACNVGGGTATCTAATCC-3’).46 PCR was conducted using Q5 high-fidelity DNA polymerase (NEB, M0491L) at 25 μL final volume with 20 ng template DNA, and the other PCR components are shown in Table S1. PCR reactions were carried out with initial denaturation at 98 ℃ for 5 min followed by 25-27 cycles of 98 ℃ for 10 s, 50 ℃ for 30 s, 72 ℃ for 30 s, and ended with a final extension at 72 ℃ for 5min. Amplicon sizes were confirmed and selected by 2% agarose gel



electrophoresis in 1.0X TAE buffer, then the amplicons were purified using the AxyPrep DNA Gel Extraction Kit (Axygen, AP-GX-250). After purification, the amplicons were quantified by a Microplate reader (BioTek, Flx800) with Quant-iT PicoGreen dsDNA Assay Kit (Invitrogen, P7589). In the final mixture, samples were normalized in equimolar amounts with a final volume of 50 μL at 2 ng/μL amplicon. The sequencing of amplification was accomplished with MiSeq after the library was prepared following the manufacturer’s instruction (Illumina). Before sequencing, linearization enzyme was used to make sure the library spliced into single strands, then dyed by four kinds of nucleotides (ddATP, ddGTP, ddCTP and ddTTP) which contain disparate fluorescent dye. The template was complemented at a time by a removable blocking group. Data Analysis. Based on the phred algorithm or contained ambiguous base calls, the raw data was filtered by the truncation of reads which did not have an average quality of 20 over 5-bp sliding window. Filtered paired-end reads were merged by FLASH (v1.2.6), which is an accurate and fast tool to merge paired-end reads. The minimum required overlap length is 10-bp, requiring 100% consensus between the overlapping regions of the forward and reverse paired-end. By using the Quantitative Insights into Microbial Ecology (QIIME) software,47 the sequences were trimmed and assigned to samples based on the barcodes. According to the seed-based UCLUST algorithm,48 similar sequences were clustered at an identity threshold of 97%, which is a generally acceptable definition for operational taxonomic unit (OTU).49 The longest sequences in each OTU were


Page 10 of 37

Page 11 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


selected for further analyses. The bacterial 16S rRNA gene is widely used to measure the diversity of microbial community. To calculate the richness of microbial community, the ACE and Shannon estimators were used for the calculation of diversity of microbial community.50,51 Taxonomy of microbial communities was evaluated at the levels of phylum to genus using the QIIME, and the potential metabolic function was also forecasted.52

RESULTS AND DISCUSSION Inactivation of bacteria upon sediment oxygenation. Mixed suspensions of 5 g sediments and 50 mL DI water were exposed to laboratory air for a 10-h oxygenation. As shown in Figure 1a, no significant inactivation of bacteria was observed after oxygenation of sediments at the depth of 20, 40 and 60 cm (P> 0.05), but living bacteria counts in sediment at a depth of 100 cm decreased from 6.1 to 5.7 orders of magnitude after oxygenation, which is statistically significant (P< 0.05). Figure 1b demonstrated that the Fe(II) contents in sediments at the depth of 100 cm was much higher (6.87 mg/g) than that of 20, 40 and 60 cm (< 1 mg/g), which indicates bacteria inactivation only occurred upon oxygenation of Fe(II)-rich sediments. After a 10-h oxygenation, about 64% of Fe(II) (4.39 mg/g) was oxidized in the sediment at the depth of 100 cm, along with a production of 27.6 μM •OH (Figure 1c). But for the sediments at the depth of 20, 40 and 60 cm, the cumulative concentration of •OH was negligible (Figure 1c). In control experiments conducted under anoxic conditions (Figure S1), no observable changes can be seen for bacteria, Fe(II) and •OH, which



indicates O2 is prerequisite for Fe(II) oxygenation, • OH production and bacteria inactivation. As can be seen in Figure 2, the variation tendency of living bacteria counts, Fe(II) contents and •OH production during oxygenation of 100-cm sediment correlated very well with each other, implying that the •OH was produced from Fe(II) oxygenation, which might induce bacteria inactivation. The production of •OH from oxidation of subsurface sediments has been proved in our previous study, Fe(II)-bearing minerals are the predominant contributor to •OH production.27 In order to identify the role of ROS in bacteria inactivation, 100 mM ethanol was added to quench ROS produced during sediment oxygenation. It is worth noting that the minimal inhibitory concentration of ethanol for microorganisms inactivation was reported to be 5%,53 which is an order of magnitude higher than the concentration in this study. So the antimicrobial effect of 100 mM ethanol should be negligible, which is in agreement with our observation that the addition of 100 mM ethanol caused negligible changes in bacteria numbers during a 10-h reaction (Figure 3). As depicted in Figure 3, the bacteria inactivation was suppressed by ethanol significantly (P< 0.01), indicating that ROS is the dominant factor for bacteria inactivation. Overall, the above findings demonstrate that bacteria could be killed upon oxygenation of Fe(II)-bearing sediments, ROS rather than O2 is the key species for bacteria inactivation.


Page 12 of 37

Page 13 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 1. Variations of (a) living bacteria number, (b) total Fe(II) concentration and (c) •OH produced upon oxygenation of sediments at different depths. The reaction conditions were based on pH 7.0-7.4, 5 g sediments and 50 mL DI water exposed to air for 10 hours. The groundwater table was about 50 cm below the ground at sampling time. T-tests were used to compare the differences of living bacteria number between sediments before and after oxygenation, P< 0.05 indicates the difference is statistically significant.



Figure 2. Time profiles of variations in (a) living bacteria number, (b) total Fe(II) concentration and (c) production of •OH upon oxygenation of sediments at a depth of 100 cm.

Figure 3. Variations of living bacteria number during oxygenation of sediments at the depth of 100 cm. The reaction conditions were based on pH 7.0-7.4, 5 g sediments and 50 mL DI water exposed to air. T-tests were used to compare the differences of living bacteria number between systems with and without ethanol, P< 0.05 indicates the difference is statistically significant.

The antimicrobial effect of ROS derived from oxygenation of Fe(II)-bearing


Page 14 of 37

Page 15 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


minerals has been reported.33,35,54 Generally, the toxicity of ROS comes from their ability to oxidize a large number of cellular constituents such as DNA,39,40 proteins and amino acids,41 and phosphatide42. The antimicrobial process could take place in vivo or in vitro. Williams et al. reported that Fe(II)-containing clays killed antibiotic-resistant human pathogens by promoting Fe2+ solubility. Soluble Fe2+ diffused into the cells and was oxidized with a production of •OH, which resulted in microbes inactivation.33 Dong group identified the role of clay structural Fe(II) in antibacterial process by incubating Escherichia coli with reduced nontronite (rNAu-2) under air-saturated conditions.54 They demonstrated that the • OH generated from structural Fe(II) oxygenation damaged the cardiolipin, which triggered the influx of Fe2+ into the cell and ultimately leaded to the bacteria inactivation. In this study, the bacteria inactivation was suppressed by adding ethanol which only scavenged extracellular • OH, so the extracellular process was identified to be the main mechanism for bacteria inactivation.

Effect of initial O2 partial pressure on bacteria inactivation Since the amount of O2 intruding into the subsurface is usually limited, we further evaluated the bacteria inactivation upon sediment oxygenation under lower O2 partial pressure. The two O2 partial pressure conditions (0.0112 atm and 0.0168 atm) represent the theoretical oxygen demand for 50% and 75% of oxidation of reactive Fe(II) in 100-cm sediments. As can be seen in Figure 4a, the living bacteria number decreased by 0.38 orders of magnitude under air conditions (O2 partial pressure: 0.21



atm). When initial O2 partial pressure decreased to 0.0168 atm and 0.0112 atm, bacteria inactivation was still observed, which is 0.19 and 0.15 orders of magnitude, respectively. Correspondingly, the extent of Fe(II) oxygenation decreased with a decreasing in initial O2 partial pressure (Figure 4b). Figure 4c showed that 27.5 μM • OH was produced under air conditions. When the initial O2 partial pressure decreased to 0.0168 atm and 0.0112 atm, the cumulative concentration of • OH decreased to 12.7 and 9.5 μM, respectively. These results proved that bacteria could be inactivated upon sediment oxygenation with a very limited amount of O2, and O2 could influence the inactivation of bacteria by controlling the amount of • OH produced upon Fe(II) oxygenation.


Page 16 of 37

Page 17 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 4. Effect of initial O2 partial pressure on the variations of (a) living bacteria number, (b) total Fe(II) concentration and (c) •OH produced upon oxygenation of sediments at the depth of 100 cm. The reaction conditions were based on pH 7.0-7.4, 1 g sediments and 10 mL DI water. Bacteria inactivation induced by H2O2. H2O2 is a more persistent and stable ROS than • OH, which has been detected in a wide range of surface waters and groundwater.55 H2O2 is also an intermediate generated during the transformation of O2 to •OH and a product of bacterial aerobic respiration. Moreover, H2O2 is widely used as an oxidant during in-situ chemical oxidation (ISCO) for the remediation of



contaminated groundwater and soil, so high concentrations of H2O2 could be introduced into the subsurface. To simulate the long-term cumulative effect of H2O2 on microbes in natural environment as well as the influence of H2O2 on microbes in subsurface environment which perturbed by anthropogenic activities, 1–100 mM H2O2 was added into the sediment suspensions under anoxic conditions. After a 2-h reaction (Figure 5), no significant bacteria inactivation was observed in all sediments when 1 mM H2O2 was added (P> 0.05). The residual H2O2 after reaction was determined to be 1 μM. This negligible change of bacteria could be attributed to the competitive consumption of H2O2 by reducing substances in sediments. The concentration of total organic carbon (TOC) in sediment suspension is about 1 g/L, which is 30 times higher than that of H2O2 (0.034 g/L). S(-II) and other reducing components (i.e. Mn(-II)) were not detected in all of the sediments. Therefore, the natural organic matters (NOM) might be the main competitive species consuming ROS. When the initial concentration of H2O2 was elevated to 10 mM, the bacteria in 60 and 100-cm sediments was inactivated by 0.3 and 0.6 orders of magnitude, respectively. T-test analysis demonstrated the differences were statistically significant (P< 0.05). But the inactivation was still insignificant for 20 and 40-cm sediments (P> 0.05). When initial H2O2 concentration was further elevated to 50 mM, bacteria in sediments at the depth of 60 and 100 cm were inactivated completely (P< 0.01), and the living bacteria number in sediments at the depth of 20 and 40 cm significantly decreased by 0.7 and 0.6 orders of magnitude (P< 0.05), respectively. With an addition of 100 mM H2O2, complete inactivation of bacteria was observed in all of the


Page 18 of 37

Page 19 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


samples (P< 0.01). These findings demonstrated that H2O2 exhibited an antimicrobial effect against bacteria in Fe(II)-bearing sediments, more H2O2 leaded to more bacteria inactivation, the threshold concentration for inactivation varies with each sediment.

Figure 5. Variations of living bacteria number in sediments at different depths after reacting with H2O2 for 2 h. The reaction conditions were based on pH 7-7.4, 1 g sediments, 10 mL DI water, anoxic conditions. T-tests were used to compare the differences of living bacteria number between systems without H2O2 and with different concentrations of H2O2, P> 0.05 indicates the difference is statistically insignificant, P< 0.05 indicates the difference is statistically significant, P< 0.01 indicates the difference is markedly significant. The antimicrobial effects of H2O2 against bacteria including Escherichia coli, Staphylococcus aureus, Xanthobacter flavus and Salmonella sp. have been largely studied.39,40,56–61 On the one hand, H2O2 could kill bacteria directly by influencing its metabolism process.61 On the other hand, H2O2 may produce •OH by reacting with



Fe(II) (Fenton reaction), which inducing bacteria inactivation indirectly.39,40,62,63 Imlay group observed a significant death of Escherichia coli because of DNA damage through an in vivo Fenton reaction. They found H2O2 itself couldn’t directly oxidize DNA, but it reacted rapidly with Fe2+ to form •OH or Fe(IV) which induced DNA damage, the intracellular free Fe2+ was identified to catalyze this in vivo Fenton reaction.39,40 Similarly, Kim et al. observed serious damages to cell membrane integrity and respiratory activity after exposing Escherichia coli cells to Fe2+ under deaerated conditions, the generation of intracellular ROS during the reaction of Fe2+ with intracellular H2O2 was proved to be responsible for bactericidal activities.62,63 In this study, the bacterial inactivation upon oxygenation of 100-cm sediments was more significant than that of other sediments when H2O2 concentration was lower than 50 mM. Considering the Fe(II) contents in 100-cm sediments are much higher than other sediments, the enhancement of antibacterial effect was probably because •OH produced from Fenton reaction increased the bio-toxicity. Moreover, the shallower sediments are disturbed by O2 more often than the deeper sediments, so the bacteria in shallower sediments could adjust allocation of resources and life strategy to acclimate to the oxidative stress. In addition to H2O2 and •OH, O2•– could also be generated as an intermediate ROS through a one electron transfer pathway during the interaction of O2 with Fe(II). It has been documented that O2•– itself couldn't directly inactivate bacterial, but it could induce indirect DNA damage by reacting with Fe(II) to produce H2O2 and •OH.64 Moreover, O2•– has the ability to leach the iron from iron-sulfur clusters in cells,


Page 20 of 37

Page 21 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


which increases the amount of free Fe2+ that is available to catalyze •OH production.64 Changes in microbial community structure upon sediment oxygenation. The variations of species diversity index of microorganisms upon sediment oxygenation are shown in Table S2. After a 10-hour oxygenation, the diversity indices of microorganisms changes insignificantly. Figure 6a demonstrated the variations of taxonomy and distribution of microorganisms at phylum level. Proteobacteria, Chloroflexi, Acidobacteria and Nitrospirae constituted about 80% of the sequences of sediments (Fig. 6a, Table S3), and the structure was similar in sediments at different depths. After a 10-h oxygenation, different variations of microorganisms at phylum level were observed. The relative abundance of Proteobacteria in sediments at the depth of 20 cm barely changed, but that in 60 and 100-cm sediments decreased by about 5%. A significant increase of Chloroflexi from 26.18% to 42.64% was observed in 100-cm sediments, which was not observed in 20 and 60-cm sediments. The relative abundance of Nistrospirae in 100-cm sediments decreased from 12.61% to 7.60% after oxygenation, but no obvious change was found in 20 and 60-cm sediments. The phylum of Acidobacteria didn’t change significantly in all of the sediments. Figure 6b showed the variations of the microorganism distribution at class level. The microbial community structure of 100-cm sediments was different from the 20 and 60-cm sediments. The relative abundance of Deltaproteobacteria and Anaerolineae in 100-cm sediments was about 2 times higher than that in 20 and 60-cm

sediments.

But

the

relative

abundance


of

Betaproteobacteria,


Alphaproteobacteria, Subgroup_6, Subgroup_17 and KD4-96 in 100-cm sediments was less than that in 20 and 60-cm sediments. After a 10-h oxygenation, the relative abundance of Anaerolineae in 100-cm sediments increased from 15.43% to 29.90%, which was recognized as the domain class contributing the variation in the phylum of Chloroflexi.

Figure 6. Taxonomy and distribution of microorganisms at (a) phylum and (b) class levels.


Page 22 of 37

Page 23 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


The variations of the microorganism distribution at order level and family level are shown in Table 1. Anaeroloneales was the most abundant order in 100-cm sediments, and its relative abundance increased from 15.43% to 29.90% after sediment oxygenation (Table 1a). The family of Anaerolineaceae was recognized as the only family contributing the variation in the order of Anaeroloneales, which was also the domain kind of microorganisms at family level in sediments at a depth of 100 cm (Table 1b). Anaerolineaceae has been identified to be an organic matter degrader,65 which is a keystone microbe playing an important role in shaping soil microbial communities.66–67 The increase of Anaerolineaceae may promote the degradation of organic matters and the transformation of heavy metals (e.g., Cd) in soil and sediments.66–69 Table 1. Taxonomy and distribution of microorganisms at order and family levels (a) Order level Abundance

0h

10 h

Taxon

20 cm

60 cm

100 cm

20 cm

60 cm

100 cm

Anaerolineales

6.16%

7.65%

15.43%

8.64%

8.15%

29.90%

Nitrospirales

12.84%

7.64%

12.61%

10.43%

9.66%

7.60%

Rhizobiales

6.95%

14.16%

4.23%

7.61%

9.77%

3.52%

Gemmatimonadales

6.61%

5.34%

0.71%

7.67%

7.07%

0.36%

Nitrosomonadales

4.49%

4.56%

0.09%

4.87%

3.92%

0.03%

Sva0485

0.07%

0.90%

10.18%

0.36%

1.02%

8.99%

Myxococcales

4.18%

3.73%

1.31%

3.82%

4.11%

0.59%

(b) Family level Abundance

0h


10 h


Page 24 of 37

Taxon

20 cm

60 cm

100 cm

20 cm

60 cm

100 cm

Anaerolineaceae

6.16%

7.65%

15.43%

8.64%

8.15%

29.90%

Xanthobacteraceae

4.88%

10.12%

2.00%

5.95%

7.60%

1.89%

Gemmatimonadaceae

6.61%

5.34%

0.71%

7.67%

7.07%

0.36%

Nitrospiraceae

2.99%

2.93%

8.86%

3.08%

3.77%

4.85%

0319-6A21

8.40%

3.48%

0.19%

6.31%

4.13%

0.07%

Since the variation of Anaerolineaceae has the ability to shape microbial communities in sediments, we investigated other changes in microbial function including nitrification, denitrification and iron respiration upon sediments oxygenation by QIIME. As shown in Figure 7, the family of Geobacteraceae, which plays a dominant role in iron respiration,70,71 decreased after sediment oxygenation. No obvious variation of the function of aerobic ammonia oxidation was observed after sediment oxygenation, but decreases of aerobic nitrite oxidation and nitrate reduction function were observed. The relative abundance of function of aerobic nitrite oxidation decreased from 12.37% to 7.58% in 100-cm sediments, but that in 20 and 60-cm sediments only decreased by 2%. The relative abundance of family of Nitrospiraceae decreased from 8.86% to 4.85% upon oxygenation of 100-cm sediments, but no changes were observed in 20 and 60-cm sediments (Table 1). Overall, these finding demonstrated that oxygenation of Fe(II)-bearing sediments induced a changing in microbial community structure. The variations of microbial community structure could be induced by O2 or ROS. As Geobacteraceae was reported to be strictly anaerobic, the decreasing abundance of Geobacteraceae may be mainly due to the fundamental changes in metabolic lifestyle induced by O2. The abundance of Nistrospiraeceae which recognized as aerobes also decreased upon


Page 25 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


sediment oxygenation, which could be attributed to the DNA damage induced by ROS generated from Fe(II) oxidation. Surprisingly, the abundance of anaerobic Anaerolineaceae increased upon sediments oxygenation, the mechanism was unknown as the genus of this Anaeroloneaceae was not identified, which needs further investigation. Overall, the relative abundance of Anaerolineaceae increased but of Nistrospiraeceae and Geobacteraceae decreased, pointing to the suppression of nitrification, denitrification and iron respiration but promotion of organic matter degradation.

Figure 7. Variations of relative abundance of functional microbial upon oxygenation. Environmental implications. We identify here that ROS produced from oxygenation of Fe(II)-bearing sediments play an important role in the variation of microbial community structure. This new finding could be significant because Fe(II) is abundant component in subsurface environment under reducing conditions, and the interaction of O2 with sediments is common in many natural and artificial processes.15‒18 The biogeochemical cycling of many nutrients (i.e., C, N, P, S, Fe,



Mn) and trace elements (i.e., As, Cr, U) are directly mediated by functional microbes.1 In addition, the biogeochemical behavior of other substances may be indirectly coupled to microbially-mediated transformations of natural organic matter and mineral phases.1 Based on our finding, ROS produced from Fe(II) oxygenation would potentially inactivate the functional microbes relating to the cycling of elements, thus reshaping the microbial community and influencing the biogeochemical cycling of elements and substances.

CONCLUSIONS This study investigated the responses of microbial community to the oxygenation of Fe(II)-bearing sediments and the potential role of ROS in this process. After a 10-h oxygenation, the living bacteria counts in sediment at the depth of 100 cm decreased from 6.1 to 5.7 orders of magnitude, along with an oxidation of Fe(II) and production of • OH. Bacteria inactivation correlated well with • OH produced upon Fe(II) oxygenation. Control experiments with the addition of ROS quencher and hydrogen peroxide further proved that the bacteria inactivation was induced by ROS rather than O2. 16S rRNA analysis indicated that the microbial community structure changed upon sediment oxygenation, the relative abundance of Anaerolineaceae increased from 15.4% to 29.9%, while of Nistrospiraeceae and Geobacteraceae decreased, pointing to the suppression of nitrification, denitrification and iron respiration but promotion of organic matter degradation. These findings reveal an important but overlooked mechanism for the influence of O2 on microbial community structure.


Page 26 of 37

Page 27 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Supporting Information Available Additional information: Figures S1; Tables S1–S3. This material is available free of charge on the ACS Publications website.

Data Deposition The raw sequence data have been deposited in the NCBI Sequence Read Archive (BioProject accession no. SRP182765).

ACKNOWLEDGEMENTS This work was supported by the National Natural Science Foundation of China (NSFC, No. 41672353; 41703113; 41521001), China Scholarship Council (201606415076), China Postdoctoral Science Foundation (2016M590727) and the Natural Science Foundation of Hubei Province, China (2018CFA028).

REFERENCES (1) Ehrlich, H. Z.; Newman, D.K., Ed.; Geomicrobiology; CRC press: Boca Riton, 2009. (2) Bryant, L. D.; Little, J. C.; Bürgmann, H. Response of sediment microbial community structure in a freshwater reservior to manipulations in oxygen availability. Microbiol. Ecol. 2012, 80 (1), 248‒263. (3) Meyerhof, M. S.; Wilson, J. M.; Dawson, M. N.; Beman, J. M. Microbial



community diversity, structure and assembly across oxygen gradients in meromictic marine lakes, Palau. Environ. Microbiol. 2016, 18 (12), 4907‒4919. (4) Koretsky, C. M.; Moore, C. M.; Lowe, K. L.; Meile, C.; DiChristina, T. J.; van Cappellen, P. Seasonal oscillation of microbial iron and sulfate reduction in saltmarsh sediments (Sapelo Island, GA, USA). Biogeochemistry. 2003, 64 (2), 179‒203. (5) Evangelisti, M.; D’Amelia, D.; Di Lallo, G.; Thaller, M.; Migliore, L. The relationship between salinity and bacterioplankton in three relic coastal ponds (Macchiatonda Wetland, Italy). J. Water Resour. Prot. 2013, 5 (9), 859‒866. (6) Bier, R. L.; Voss, K. A.; Bernhardt, E. S. Bacterial community responses to a gradient of alkaline mountaintop mine drainage in Central Appalachian streams. ISME J. 2015, 9 (6), 1378‒1390. (7) Crump, B. C.; Kling, G. W.; Bahr, M.; Hobbie, J. E. Bacterioplankton community shifts in an arctic lake correlate with seasonal changes in organic matter source. Appl. Environ. Microbiol. 2003, 69 (4), 2253‒2268. (8) Hu, A.; Wang, H.; Li, J.; Liu, J.; Chen, N.; Yu, C. P. Archaeal community in a human-disturbed watershed in southeast China: diversity, distribution, and responses to environmental changes. Appl. Microbiol. Biotechnol. 2016, 100 (10), 4685‒4698. (9) DeAngelis, K. M.; Silver, W. L.; Thompson, A. W.; Firestone, M. K. Microbial communities acclimate to recurring changes in soil redox potential status. Environ. Microbiol. 2010, 12 (12), 3137‒3149. (10) Chen, J.; Hanke, A.; Tegetmeyer, H. E.; Kattelmann, I.; Sharma, R.; Hamann, E.; Hargesheimer, T.; Kraft, B.; Lenk, S.; Geelhoed, J. S.; Hettich, R. L. Impacts of


Page 28 of 37

Page 29 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


chemical gradients on microbial community structure. ISME J. 2017, 11 (4), 920‒931. (11) Lüdemann, H., Arth, I.; Liesack, W. Spatial changes in the bacterial community structure along a vertical oxygen gradient in flooded paddy soil cores. Appl. Environ. Microbiol. 2000, 66 (2), 754‒762. (12) Gao, D.W.; Fu, Y.; Tao, Y.; Li, X. X.; Xing, M.; Gao, X. H.; Ren, N. Q. Linking microbial community structure to membrane biofouling associated with varying dissolved oxygen concentrations. Bioresour. Technol. 2011, 102 (10), 5626‒5633. (13) Spietz, R.L., Williams, C.M., Rocap, G. and Horner‒Devine, M.C. A dissolved oxygen threshold for shifts in bacterial community structure in a seasonally hypoxic estuary. PloS one. 2015, 10 (8), e0135731. (14) Lipson, D.A., Raab, T.K., Parker, M., Kelley, S.T., Brislawn, C.J. and Jansson, J. Changes in microbial communities along redox gradients in polygonized Arctic wet tundra soils. Environ. Microbiol. Rep. 2015, 7 (4), 649‒657. (15) Kumar, A. R.; Riyazuddin, P. Seasonal variation of redox species and redox potentials in shallow groundwater: A comparison of measured and calculated redox potentials. J. Hydrol. 2012, 444, 187‒198. (16) Beck, M.; Dellwig, O.; Schnetger, B.; Brumsack, H. J. Cycling of trace metals (Mn, Fe, Mo, U, V, Cr) in deep pore waters of intertidal flat sediments. Geochim. Cosmochim. Acta 2008, 72 (12), 2822‒2840. (17) Hester,E. T.; Gooseff, M. N. Moving beyond the banks: Hyporheic restoration is fundamental to restoring ecological services and functions of streams. Environ. Sci. Technol. 2010, 44 (5), 1521‒1525.



(18) Diem, S.; Cirpka, O. A.; Schirmer, M. Modeling the dynamics of oxygen consumption upon riverbank filtration by a stochastic‒convective approach. J. Hydrol. 2013, 505, 352‒363. (19) Danczak, R. E.; Yabusaki, S. B.; Williams, K. H.; Fang, Y.; Hobson, C.; Wilkins, M. J. Snowmelt induced hydrologic perturbations drive dynamic microbiological and geochemical behaviors across a shallow riparian aquifer. Front. Earth Sci. 2016, 4, 57. (20) Wang, X., Zhang, Y., Zhang, T.; Zhou, J. Effect of dissolved oxygen on elemental sulfur generation in sulfide and nitrate removal process: characterization, pathway, and microbial community analysis. Appl. Microbiol. Biotechnol. 2016, 100 (6), 2895‒2905. (21) Schimel, J.; Balser, T. C.; Wallenstein, M. Mocrobial stress-response physiology and its implications for ecosystem function . Ecol. Soc. Am. 2007, 88 (6), 1386‒1394. (22) Pett‒Ridge, J., Silver, W. L.; Firestone, M. K. Redox fluctuations frame microbial community impacts on N-cycling rates in a humid tropical forest soil. Biogeochemistry. 2006, 81 (1), 95‒110. (23) Carucci, A.; Lindrea, K.; Majone, M.; Ramadori, R. Different mechanisms for the anaerobic storage of organic substrates and their effect on enhanced biological phosphate removal (EBPR). Water. Sci. Tech. 1999, 39 (6), 21–28. (24) Niviere, V.; Fontecave, M. Discovery of superoxide reductase: an historical perspective. J. Biol. Inorg. Chem. 2004, 9 (2), 119–123. (25) Chen, R., Liu, H., Tong, M., Zhao, L., Zhang, P., Liu, D.; Yuan, S. H. Impact of


Page 30 of 37

Page 31 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Fe (II) oxidation in the presence of iron-reducing bacteria on subsequent Fe (III) bio‒reduction. Sci. Total Environ. 2018, 639, 1007‒1014. (26) Ilbert, M.; Bonnefoy, V. Insight into the evolution of the iron oxidation pathways. Biochim. Biophys. Acta. 2013, 1827 (2), 161‒175. (27) Tong, M.;

Yuan, S.;

Ma, S.;

Jin, M.;

Liu, D.;

Cheng, D.;

Liu, X.;

Gan, Y.; Wang, Y. Production of abundant hydroxyl radicals from oxygenation of subsurface sediments. Environ. Sci. Technol. 2015, 50 (1), 214‒221. (28) Liu, X.;

Yuan, S.;

Tong, M.; Liu, D. Oxidation of trichloroethylene by the

hydroxyl radicals produced from oxygenation of reduced nontronite. Water

Res.

2017, 113, 72‒79. (29) Cheng, D.;

Yuan, S.;

Liao, P.; Zhang, P. Oxidizing impact induced by

mackinawite (FeS) nanoparticles at oxic conditions due to production of hydroxyl radicals. Environ. Sci. Technol. 2016, 50 (21), 11646‒11653. (30) Zhang, P.; Yuan, S. Production of hydroxyl radicals from abiotic oxidation of pyrite

by

oxygen

under

circumneutral

conditions

in

the

presence

of

low-molecular-weight organic acids. Geochim. Cosmochim. Acta 2017, 218, 153‒166. (31) Ardo, S. G.;

Nélieu, S.;

Ona‒Nguema, G.;

Delarue, G.;

Brest, J.;

Pironin, E.; Morin, G. Oxidative degradation of nalidixic acid by nano-magnetite via Fe2+/O2-mediated reactions. Environ. Sci. Technol. 2015, 49 (7), 4506‒4514. (32) Dixon, S. J. and Stockwell, B. R. The role of iron and reactive oxygen species in cell death. Nat. Chem. Biol. 2013, 10 (1), 9–17. (33) Williams, L. B.; Metge, D. W.; Eberl, D. D.; Harvey, R. W. Turner, A. G.; Prapaipong, P.; Poret-Peterson, A. T. What makes a natural clay antibacterial? Environ. Sci. Technol. 2011, 45, 3768−3773. (34) Delarie, C.; van Genuchten, C. M.; Nelson, K. L.; Amrose, S. E.; Gadgil, A. J. Escherichia coli attenuation by Fe electrocoagulation in synthetic Bengal



Page 32 of 37

groundwater: effect of pH and natural organic matter. Environ. Sci. Technol. 2015, 49 (16), 9945–9953. (35) Tran, N.; Mir, A.; Mallik, D.; Sinha, A.; Nayar, S.; Webster, T. J. Bactericidal effect of iron oxide nanoparticles on Staphylococcus aureus. Int. J. Nanomed. 2010, 5, 277–283. (36) Swanner, E. D.;

Mloszewska, A. M.;

Cirpka, O. A.;

Schoenberg, R.;

Konhauser, K. O.; Kappler, A. Modulation of oxygen production in Archaean oceans by episodes of Fe (II) toxicity. Nat. Geosci. 2015, 8 (2), 126–130. (37) You, Y. W.; Han, J.; Chiu, P.C.; Jin, Y. Removal and inactivation of waterborne viruses using zerovalent iron. Environ. Sci. Technol. 2005, 39 (23), 9263–9269. (38) Diao, M. H. and Yao, M.S. Use of zero-valent iron nanoparticles in inactivating microbes. Water Res. 2009, 43 (20), 5243–5251. (39) Imlay, J. A.; Linn, S. Toxic DNA damage by hydrogen peroxide through the Fenton reaction in vivo and in vitro. Science 1988, 240 (4852), 640–642. (40) Park, S.; Imlay, J. A. High levels of intracellular cysteine promote oxidative DNA damage by driving the Fenton reaction. J. Bacteriol. 2003, 185 (6), 1942–1950. (41) Valko, M.; Morris, H.; Cronin, M. T. D. Metals, toxicity and oxidative stress. Curr. Med. Chem. 2005, 12 (10), 1161–1208. (42) Girotti, A. W. Lipid hydroperoxide generation, turnover, and effector action in biological systems. J. Lipid. Res. 1998, 39 (8), 1529–1542. (43) Stocks, S. M. Mechanism and use of the commercially available viability stain, BacLight. Cytometry A. 2004, 61 (2), 189‒195.


Page 33 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(44) Mopper, K.; Zhou, X. Hydroxyl radical photoproduction in the sea and its potential impact on marine processes. Science 1990, 250 (4981), 661–664. (45) Katsoyiannis, I. A.; Ruettimann, T.; Hug, S. J. pH dependence of Fenton reagent generation and As (III) oxidation and removal by corrosion of zero valent iron in aerated water. Environ. sci. technol. 2008, 42 (19), 7424–7430. (46) Ewing, B.; Hillier, L.; Wendl, M. C.; Green, P. Base-calling of automated sequencer traces usingPhred. I. Accuracy assessment. Genome Res. 1998, 8 (3), 175– 185. (47) 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. QIIME allows analysis of high‒throughput community sequencing data. Nat. Methods 2010, 7 (5), 335. (48) Edgar, R. C. Search and clustering orders of magnitude faster than BLAST. Bioinformatics 2010, 26 (19), 2460–2461. (49) Lauber, C. L.; Hamady, M.; Knight, R.; Fierer, N. Pyrosequencing-based assessment of soil pH as a predictor of soil bacterial community structure at the continental scale. Appl. Environ. Microbiol. 2009, 75 (15), 5111–5120. (50) Pitta, D. W.; Pinchak, W. E.; Dowd, S. E.; Osterstock, J.; Gontcharova, V.; Youn, E.; Dorton, K.; Yoon, I.; Min, B. R.; Fulford, J. Rumen bacterial diversity dynamics associated with changing from bermudagrass hay to grazed winter wheat diets. Microb. Ecol. 2010, 59 (3), 511–522. (51) Shannon, C. E. A mathematical theory of communication. ACM SIGMOBILE mobile computing and communications review 2001, 5 (1), 3–55. (52) White, J. R.; Nagarajan, N.; Pop, M. Statistical methods for detecting



differentially abundant features in clinical metagenomic samples. PLoS Comput. Biol. 2009, 5 (4), e1000352. (53) Oh, D. H., Marshall, D. L. Antimicrobial activity of ethanol glycerol monolaurate or lactic acid against Listeria monocytogenes, Int. J. Food Microbiol. 1993, 20 (4), 239‒246. (54) Wang, X.; Dong, H.; Zeng, Q.; Xia, Q.; Zhang, L.; Zhou, Z. Reduced iron-containing clay minerals as antibacterial agents. Environ. Sci. Technol. 2017, 51 (13), 7639–7647. (55) Yuan, X.; Nico, P. S.; Huang, X.; Liu, T.; Ulrich, C.; Williams, K. H.; Davis, J. A. Production of hydrogen peroxide in groundwater at Rifle, Colorado. Environ. Sci. Technol. 2017, 51 (14), 7881–7891. (56) Kikuchi, Y.; Sunada, K.; Iyoda, T.; Hashimoto, K.; Fujishima, A. Photocatalytic bactericidal effect of TiO2 thin films: dynamic view of the active oxygen species responsible for the effect. J. Photoch. Photobio. A 1997, 106 (1‒3), 51–56. (57) Cho, M.; Chung, H.; Choi, W.; Yoon, J. Linear correlation between inactivation of E. coli and OH radical concentration in TiO2 photocatalytic disinfection. Water Res. 2004, 38 (4), 1069–1077. (58) Scully, N. M.; Cooper, W. J.; Tranvik, L. J. Photochemical effects on microbial activity in natural waters: the interaction of reactive oxygen species and dissolved organic matter. FEMS Microbiol. Ecol. 2003, 46 (3), 353–357. (59) Imlay, J. A. Cellular defenses against superoxide and hydrogen peroxide. Annu. Rev. Biochem. 2008, 77, 755–776.


Page 34 of 37

Page 35 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(60) Brandi, G.; Cattabeni, F.; Albano, A.; Cantoni, O. Role of Hydroxyl Radicals in Escherzchza Colz Killing Induced by Hydrogen Peroxide. Free Radical Res. Commun. 1989, 6 (1), 47–55. (61) Imlay, J. A.; Linn, S. Bimodal pattern of killing of DNA-repair-defective or anoxically grown Escherichia coli by hydrogen peroxide. J. Bacteriol. 1986, 166 (2), 519–527. (62) Kim, J. Y.; Park, H.; Lee, C.; Nelson, K. L, Sedlak, D. L.; Yoon, J. Inactivation of Escherichia coli by nanoparticulate zero valent iron and ferrous ion. App. Environ. Microbiol. 2010, 76, 7668–7670. (63) Kim, J. Y.; Park, H.; Lee, C.; Nelson, K. L, Sedlak, D. L.; Yoon, J. Inactivation of Escherichia coli by nanoparticulate zero valent iron and ferrous ion. App. Environ. Microbiol. 2010, 76, 7668–7670. (64) Keyer, K., Gort, A. S. Superoxide and the production of oxidative DNA damage, J. Bacteriol. 1995, 177 (23), 6782‒6790. (65) Ansola, G.; Arroyo, P.; de Miera, L. E. S. Characterisation of the soil bacterial community structure and composition of natural and constructed wetlands. Sci. Total Environ. 2014, 473, 63–71. (66) Sherry, A.; Gray, N.; Ditchfield, A.; Aitken, C.; Jones, D.; Röling, W.; Hallmann, C.; Larter, S.; Bowler, B.; Head, I. Anaerobic biodegradation of crude oil under sulphate-reducing conditions leads to only modest enrichment of recognized sulphate-reducing taxa. Int. Biodeterior. Biodegrad. 2013, 81, 105–113. (67) Savage, K. N.; Krumholz, L. R.; Gieg, L. M.; Parisi, V. A.; Suflita, J. M.; Allen,



Page 36 of 37

J.; Philp, R. P.; Elshahed, M. S. Biodegradation of low-molecular-weight alkanes under

mesophilic,

sulfate-reducing

conditions:

metabolic

intermediates

and

community patterns. FEMS Microbiol Ecol 2010, 72 (3), 485–95. (68) Sutton, N. B.; Maphosa, F.; Morillo, J. A.; Al‒Soud, W. A.; Langenhoff, A. A.; Grotenhuis, T.; Rijnaarts, H. H.; Smidt, H. Impact of long term diesel contamination on soil microbial community structure. Appl. Environ. Microbiol. 2012, AEM. 02747–12. (69) Meng, D.; Li, J.; Liu, T.; Liu, Y.; Yan, M.; Hu, J.; Li, X.; Liu, X.; Liang, Y.; Liu, H. Effects of redox potential on soil cadmium solubility: Insight into microbial community. J. Environ. Sci. 2019, 75, 224–232. (70) Coupland, K.; Johnson, D. B. Evidence that the potential for dissimilatory ferric iron reduction is widespread among acidophilic heterotrophic bacteria. FEMS Microbiol. Lett. 2008, 279 (1), 30–35. (71) Lin, B.; Braster, M.; van Breukelen, B. M.; van Verseveld, H. W.; Westerhoff, H. V.; Röling, W. F. Geobacteraceae community composition is related to hydrochemistry and biodegradation in an iron-reducing aquifer polluted by a neighboring landfill. Appl. Environ. Microbiol. 2005, 71 (10), 5983–5991.


Page 37 of 37 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


for TOC only


the Role of Reactive Oxygen Species - ACS Publications

Recommend Documents