The Pennsylvania State University the Graduate School

The Pennsylvania State University

The Graduate School

Department of Crop and Soil Sciences

DISTRIBUTION AND DIVERSITY OF SULFUR-REDUCING PROKARYOTES

IN SULFUR-RICH PEAT SOILS

A Thesis in

Soil Science

Carolina Elvira María Yáñez Prieto

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

December 2006

The thesis of Carolina Elvira María Yáñez Prieto was reviewed and approved* by the following:

Carmen Enid Martínez Assistant Professor of Soil Environmental Chemistry Thesis Advisor Chair of Committee

Mary Ann Bruns Associate Professor of Soil Science and Microbial Ecology

Peter J. Landschoot Professor of Turfgrass Science

Katherine H. Freeman Professor of Geosciences and Associate Head of Graduate Programs and Research, Department of Geosciences

John M. Regan Assistant Professor of Environmental Engineering

David M. Sylvia Professor of Soil Microbiology Head of the Department of Crop and Soil Sciences

*Signatures are on file in the Graduate School

iii ABSTRACT

Geochemical transfers between the Lockport Dolomite Formation and overlying wetlands near the town of Manning in Western New York have resulted in peat soils that contain high levels of sulfur, zinc, and cadmium. These peat soils are artificially drained for agricultural purposes and they have undergone drastic seasonal changes in redox for the last 60 years. Sulfur cycle reactions driven by prokaryotes potentially influence the availability, mobility and the biogeochemical control of Zn and Cd retention in the peats.

The objectives of this research were to determine the presence of sulfate-reducing prokaryotes (SRP), to detect enzymatic pathways used for sulfur respiration, to evaluate the influence of temporal and spatial changes in sulfate-reducing populations and to determine the oxidation states of S present in peat soils from the Manning region.

Surface peat samples were collected from the Manning peatland region of

Western New York in July 2002 (dry season). The surface soils were collected from the edge of a field, along a Zn phytotoxicity gradient starting in an area where there was no plant growth and moving to a zone of typical plant growth. Additionally, intact soil vertical profiles (soil cores) were collected during dry (July 2002) and wet (March 2004) seasons. The soil cores were collected along a lateral transect where the center of the field had high enough levels of Zn (phytotoxic levels) to prevent plant growth and moving to areas of stunted, and then typical plant growth.

The diversity of prokaryotes involved in sulfur reduction was studied in peat soils containing high natural levels of sulfur (4,200 to 40,500 mg kg-1), zinc (137.5 to 71175

mg kg-1) and cadmium (<1 to 310.7 mg kg-1). Diversity in microbial populations was iv analyzed using classical and molecular approaches and the microbiological data were

coupled to the spectroscopic characterization (S-XANES) of the oxidation states of sulfur in the peats under investigation. Sulfur-XANES analyses reveal that sulfide/thiol groups

(reduced forms of S) and sulfonate groups (oxidized forms of S) are the two dominant S species in surface peats. While the percentage of the total S present in oxidized forms is relatively constant (~10%) in vertical profiles, about 40% and 60% of the total S exist as sulfide/thiol groups (reduced forms of S) in surface and deep soils, respectively. The use of ribosomal intergenic spacer analyses (RISA) for the characterization of bacterial communities revealed the abundance and diversity of microorganisms. Dominant bacterial populations inhabiting surface soils were very similar, while those located in deep soils were dynamic, responding to changes induced by depth and season.

Enumerations of sulfate-reducing prokaryotes (SRPs) in surface soils indicated that SRPs using acetate or lactate as electron donors represent only a small fraction (~103 g-1 dry

soil) of total cell densities determined by microscopy (~109 g-1 dry soil). Enrichment for

SRPs showed that typical sulfate reducing bacteria (delta subgroup of Proteobacteria,

among others) do not seem to be present in surface soils although 40% of the total S is in

reduced (thiol/sulfide) forms. Instead, sulfur cycling in these metalliferous surface soils is

driven by Gram-positive bacteria related to Clostridium spp. and may involve the use of

sulfonates as electron acceptors. Results of PCR-amplification of enzymes involved in

the sulfur respiration pathway [adenosine-5’-phosphosulfate reductase (apsA) and

dissimilatory sulfite reductase (dsrAB)] in DNA from surface soils supported these

observations. No dsrAB genes were detected and phylogenetic association of cloned

sequences showed that apsA sequences recovered from these soils had diverged from v apsA of known SRPs. Alternative pathways for sulfur dissimilation are proposed for these surface soils. The enzyme anaerobic sulfite reductase (asrA) was detected in the peats and phylogenetic analysis of asrA sequences revealed a wide distribution and diversity of this gene.

By analysis of dsrAB genes in deep peat soils, we identified sulfate-reducing prokaryotes in soil profile samples deeper than 45 cm. Furthermore, the sequences of dsrAB genes present in peat soil samples from both dry and wet seasons have not been previously described in the literature. Thus, our results suggest that novel sulfur-reducing prokaryotes are present in these peat soils and that they might be specialized colonists in these metalliferous soil environments. vi TABLE OF CONTENTS

LIST OF FIGURES ...... viii

LIST OF TABLES...... xi

ACKNOWLEDGEMENTS...... xii

Chapter 1 Introduction ...... 1

SULFUR CYCLE...... 2 METALLIFEROUS SOILS OF THE MANNING REGION...... 6 OBJECTIVES...... 8 REFERENCES ...... 10

Chapter 2 Sulfur-Reducing Prokaryotes in Surface Peats Subjected to Variable Redox Conditions ...... 12

ABSTRACT ...... 13 INTRODUCTION ...... 15 MATERIALS AND METHODS ...... 17 RESULTS...... 23 DISCUSSION...... 40 CONCLUSIONS ...... 44 REFERENCE ...... 46

Chapter 3 Novel Functional Genes and Enzymatic Pathways Involved in Dissimilatory Reduction of Sulfur Compounds in Agriculturally Drained Peats...... 57

ABSTRACT ...... 58 INTRODUCTION ...... 60 MATERIALS AND METHODS ...... 63 RESULTS AND DISCUSSION...... 66 CONCLUSIONS ...... 78 REFERENCE ...... 79

Chapter 4 Sulfate-Reducing Bacteria Populations in Sulfur-Rich Peat Soils as a Function of Soil Depth and Moisture Content...... 87

ABSTRACT ...... 88 INTRODUCTION ...... 90 MATERIALS AND METHODS ...... 92 RESULTS...... 97 DISCUSSION...... 109 vii CONCLUSIONS ...... 113 REFERENCES ...... 114

Chapter 5 General Conclusions ...... 121

viii LIST OF FIGURES

Figure 1-1: Biological sulfur cycle...... 4

Figure 2-1: Sulfur K-edge (2472 eV) XANES spectral analysis for the soils labeled Peat 2 (a) and Peat 8 (b). The spectra show normalized intensity versus relative energy (relative to the elemental S K-edge spectrum). The solid line is the experimental spectrum, the fine broken lines are the Gaussian curves (six), the thick broken lines are the arctangents (two), and the solid line with circles represents the fit to the experimental spectrum...... 27

Figure 2-2: Phylogenetic affiliation of isolates from MPN cultures based on comparative analyses of 16S rRNA gene sequences. The tree was created after comparison of partial (1300-nucleotides) 16S rRNA sequences using the neighbor-joining method. Bootstrap values obtained after 100 resamplings. Bar corresponds to 10 nucleotide substitutions per 100 sequence positions...... 33

Figure 2-3: RISA from surface peat soils obtained with primers specific for Bacteria. The intergenic spacer region was PCR amplified and resolved on a non-denaturating 4% polyacrylamide gel. (a) RISA in DNA extracted directly from eight surface soils. Lane 1, peat 1; Lane 2, peat 2; Lane 3, peat 3; Lane 4, peat 4; Lane 5, peat 5; Lane 6, peat 6; Lane 7, peat 7; Lane 8, peat 8; M, molecular weight marker. RISA bands present in all fingerprintings (30 in total) are represented schematically. (b) RISA of peats 1, 2 and 8. The bands selected for sequencing are marked with a number. Lane 1, soil 1; Lane 2, soil2; Lane 3, soil 8; M, molecular weight marker...... 36

Figure 2-4: RISA from SRB-specific enrichments. The intergenic spacer region was PCR amplified using bacterial primers and resolved on a non- denaturating 4% polyacrylamide gel. Three soils were studied: Peat 1 (lanes 1, 2, and 3), Peat 2 (lanes 4, 5, and 6) and Peat 8 (lanes 7 and 8). Acetate enrichments are shown in lanes 2, and 5; lactate enrichments are shown in lanes 3, 6, and 8. Unenriched soils are shown in lanes 1, 4, and 7. M, molecular weight marker. Bands selected for sequencing are marked with a number...... 38

Figure 3-1: PCR amplification of the genes for adenosine-5’-reductase subunit A (apsA). Lane 1, peat 1; Lane 2, peat 2; Lane 3, peat 3; Lane 4, peat 4; Lane 5, peat 5; Lane 6, peat 6; Lane 7, peat 7; Lane 8, peat 8; Lane 9, blank (negative control, no DNA added); Lane 10, Desulfovibrio vulgaris. M, molecular marker...... 68

Figure 3-2: Neighbor-joining phylogenetic tree showing the associations for apsA genes. Cloned sequences were denoted indicating soil origin (S, 1 to 8) and clone analyzed (CL). Tree was rooted with Archeoglobus fulgidus. The ix numbers on the branches are the results of 100 bootstrap replicates and indicate the frequency with which the sequences grouped together in the way shown...... 69

Figure 3-3: DNA sequence alignment used for the design of primers for asrA genes. Sequences were aligned using Clustal X and screened by visual inspection. Conserved region (black boxes) served as template for design of degenerated primers...... 75

Figure 3-4: PCR amplification of the genes for anaerobic sulfite reductase subunit A. asrA in enrichments (A): Lane 1 and 2, enrichment for Clostridium peat 1; Lane 3 and 4, enrichment for Desulfovibrio peat 1; Lane 5 and 6, enrichment for Clostridium peat 2; Lane 7 and 8, enrichment for Desulfovibrio peat 2; Lane 9 and 10, enrichment for Desulfotomaculum peat 2; Lane 11 and 12, enrichment for Clostridium peat 8; Lane 13 and 14, enrichment for Desulfovibrio peat 8; Lane 15 and 16, enrichment for Desulfotomaculum peat 8; Lane 17, blank (negative control, no DNA added); Lane 18, Clostridium subterminale ATCC 25774: M, molecular marker. asrA in isolates (B): Lane 1, CYP1; Lane 2, CYP2; Lane 3, CYP3; Lane 4, CYP4; Lane 5, CYP5; Lane 6, CYP6; Lane 7, CYP7; Lane 8, CYP8; Lane 9, CYP9; Lane 10, CYP10; Lane 11, CYP11; Lane 12, CYP12; Lane 13, negative control; Lane 14, Clostridium subterminale ATCC 25774...... 76

Figure 3-5: Neighbor-joining phylogenetic tree of asrA gene sequences (ca. 450 nt) from clones obtained from Clostridium (C) and SRB-enrichments (DV for Desulfovibrio and DM for Desulfotomaculum). Peat origin is indicated by S1 (peat 1), S2 (peat 2) or S8 (peat 8). Thermoplasma acidophilum sequence served as outgroup for rooting the tree. The numbers on the branches are the results of 100 bootstrap replicates...... 77

Figure 4-1: Variations in Zn and Cd concentrations and pH values in soil cores (high, medium and low zinc) collected during dry (top) and wet (bottom) seasons...... 99

Figure 4-2: Percentage of total sulfur at estimated electronic oxidation state. Most reduced (sulfides and thiols), +5 and +6 oxidation states are represented...... 103

Figure 4-3: Polymerase chain reaction analysis in soil DNA extracted from soil cores from dry and wet seasons. Samples are separated per season and per soil cores (high, medium and low zinc). Soil section analyzed are indicated as surface (S), deep (D) and bottom (B). A) RISA from peat soils obtained with primers specific for Bacteria. B) Detection of dsrAB genes. Expected size for PCR product is indicated. Products for dsrAB genes selected for clone libraries are marked (*)...... 105 x Figure 4-4: Phylogenetic tree showing the affiliation of dsrAB clones from peat samples W-8 and D-14. Tree was created after comparisons of dsrAB nucleotide sequences (sequences longer than 1700 bp with exception of AY015594 and AY015596; accession numbers in parenthesis) using neighbor-joining analysis. The dsrAB tree was rooted with Thermodesulfovibrio islandicus and bootstrap values were calculated after 100 resamplings. Bar corresponds to 10 nucleotide substitutions per 100 sequences positions...... 107

Figure 4-5: Confocal laser scanning microscopy of undisturbed peat particles. A) Tridimensional (stacked x-y planes) view of peat particles labeled with LIVE/DEAD® BacLight™ stain. Dead bacteria appear in red and alive bacteria in green. ZnS autofluorescence is indicated in blue. Purple zones result from the concurrence of ZnS and bacteria (white box). B) Z axis view of stained particles. C) DIC Tridimensional (stacked x-y planes) view of peat particles...... 108

Figure 5-1: Schematic representation of the dissimilatory sulfate reduction and the enzymatic pathways found in soils from the Manning peatland region of Western New York...... 123

xi LIST OF TABLES

Table 1-1: Important forms of sulfur and their oxidation states...... 3

Table 2-1: Elemental concentrations (dry weight basis), soil organic matter (SOM), water content, exchangeable Zn, and pH of the peats (peats 1 to 8)...... 25

Table 2-2: Percentage of the total soil sulfur at estimated electronic oxidation state for surface peats (peat 1 to 8)...... 26

Table 2-3: Total cell number estimation and number of culturable sulfate-reducing bacteria in selected peats (peat 1, 2 and 8)...... 29

Table 2-4: 16S rRNA analysis of isolates obtained from enrichments with acetate or lactate in selected peats (peats 1, 2, and 8)...... 32

Table 2-5: Sequence analysis of bands excised from RISA fingerprinting derived from bacterial 16S rRNA extracted from surface peat soils (peat 1, 2 and 8)...... 37

Table 2-6: Sequence analysis of bands excised from RISA fingerprinting derived from bacterial 16S rRNA extracted from enrichments...... 39

Table 3-1: List of bacterial strains used in this study...... 71

Table 4-1: Elemental concentrations (dry weight basis), soil organic matter (SOM), water content, and pH of peat samples collected during dry season (July 2002)...... 100

Table 4-2: Elemental concentrations (dry weight basis), soil organic matter (SOM), water content, and pH of peat samples collected during wet season (March 2004)...... 101

xii ACKNOWLEDGEMENTS

Now that my journey at State College is about to come to an end, I would like to take a moment to express my gratitude to all the persons that have contributed to the

accomplishment of this chapter of my life.

My first acknowledgements are for my husband Alexander. Without your infinite

support and encouragement I wouldn’t be here. The separation seemed sometimes

unbearable but at the end, it helped us to become the solid couple that we are now. Ты -

радость всей моей жизни. Спасибо тебе за поддержку, любовь и терпение на

протяжении всего пути. Я тебя люблю.

My gratitude goes to my adviser Dr. Carmen Enid Martínez. Many thanks for

taking the chance of choosing me as your first Ph.D. student. All your support, guidance,

patience, work and enthusiasm for research in soils were very inspiring for my future

career. It was a wonderful experience working with you and I will try to do my best to

follow all your advices.

Thanks to my co-adviser Dr. Mary Ann Bruns. I appreciate your guidance,

support and help in the discovery of the microbial world hidden in soils. I would also like

to thank my remaining committee members Dr. Peter J. Landschoot, Dr. Katherine H.

Freeman, and Dr. John M. Regan for their advice and input into this project.

Most of all I would like to thank my parents, Hugo Yáñez Piña and Graciela

Prieto Fonseca, for teaching me the value of a good education. Thanks for encouraging

me to discover the world and accept my decisions even though that signified to be far

away from our home in Chile. Los quiero mucho. Thanks to my brothers Hugo and xiii Andrés (and their respective beautiful families) for their encouragements and unconditional love and support.

I also thank my friends at State College for their support and company, in best and worst times. Special thanks to my friend Beatriz, for her friendship and her unlimited

“psychological” assistance.

And at last but not least, many thanks to my friends René and Israel. Your friendship has been one of the greatest gift I could ask. Thank you for making me feel like home and for all the great memories! xiv

“Judge your success by what you had to give up in order to get it” Dalai Lama.

To Alexander

Chapter 1

Introduction 2

SULFUR CYCLE

Sulfur is the 10th most abundant element in the universe (about 5th by weight). It

o occurs naturally as gypsum (CaSO4•H2O), as pyrite (FeS2), as elemental S and as part of

soil and marine sediments. Sulfur is an important element for life. It stabilizes proteins

and participates in the transfer of hydrogen mediated by enzymes in redox metabolism

(4).

The major reservoirs of sulfur in earth are the lithosphere (24.3 x1018 kg), the

hydrosphere (oceans 1.3 x1018 kg), and soils (2.6 x1014 kg) (11). Sulfur can be classified

into two broad groups: organic and inorganic forms. Organic sulfur in soil organic matter

constitutes more than 90% of the total sulfur present in surface soils (5). Organic sulfur is

found in two broad categories: organic sulfates (R-O-S) and carbon-bonded sulfur (C-S).

Organic sulfates include compounds such as sulfate esters (C-O-S), sulfamates (C-N-S)

and sulfate thioglycosides (N-O-S). Carbon-bonded sulfur is found in amino acids,

sulfinates, sulfones, sulfonates, and sulfoxides (5, 12).

Inorganic sulfur generally accounts for less than 25% of total sulfur in agricultural

soils (5). Sulfur exists in a wide range of oxidation states (Table 1-1). Sulfide, elemental

sulfur, thiosulfate, tetrathionate, and sulfate are the most common inorganic forms found

in soils (4, 11). In well-drained soils, most of the inorganic sulfur typically occurs as

sulfate, whereas in poorly drained or waterlogged soils the main form is sulfide and often

S0 (12).

Table 1-1: Important forms of sulfur and their oxidation states. Compound Formula Oxidation state(s) Sulfide S2- -2 2- Polysulfide Sn -2,0 Sulfura S8 0 2- Hyposulfide dithionite) S2O4 +3 2- Sulfite SO3 +4 b 2- Thiosulfate S2O3 -1, +5 2- Dithionate S2O6 +4 2- Trithionate S3O6 -2, +6 2- Tetrathionate S4O6 -2, +6 2- Pentathionate S5O6 -2, +6 2- Sulfate SO 4 +6

a Occurs in an octagonal ring in crystalline form. b Outer sulfur has an oxidation state of -1; the inner sulfur has an oxidation state of +5. Modified from (4).

Figure 1-1: Biological sulfur cycle.

5 The microbial metabolism of sulfur compounds comprises a major portion of the global sulfur cycle (Figure 1-1). Microorganisms can participate either by assimilation or dissimilation of sulfur compounds and by reducing or oxidizing them. The dissimilatory reduction of sulfate or sulfur is carried out by specialized groups of bacteria and archaea and the energy is obtained by oxidative phosphorylation under anaerobiosis. Large amounts of sulfide are produced under these conditions; sulfide can be oxidized aerobically by chemolithotrophic bacteria to generate energy.

Various forms of oxidized sulfur can serve as terminal electron acceptors in the respiration of some prokaryotes under anaerobic conditions. The sulfur compounds include sulfate, thiosulfate, and sulfur. The utilization of sulfur compounds for energy- yielding processes seems to have developed very early during prokaryotic evolution (8).

Extensive research has been done on prokaryotic sulfate reduction. Sulfate- reducing prokaryotes are a heterogeneous group of microorganisms comprising both bacteria and archaea that have as a common feature the ability to use sulfate as a terminal electron acceptor. Five phylogenetic groups of sulfate-reducing prokaryotes (SRP) have been defined based on rRNA sequence analysis (3, 9): low G+C Gram positive spore forming bacteria (Desulfotomaculum spp.), Gram-negative mesophilic bacteria of δ- proteobacteria (Desulfovibrio spp.), thermophilic bacteria of the Thermodesulfobacterium division), bacterial members of the Nitrospira division (Thermodesulfovibrio spp.), and thermophilic archaeal (Archaeoglobus spp.).

6 METALLIFEROUS SOILS OF THE MANNING REGION

Natural peatlands retain nutrients and heavy metals. These characteristics of peatlands contribute to the improvement of streams and watershed water quality.

Elevated natural concentrations of Cd and Zn have often been associated with soils high in organic matter and S in various parts of the world (1). These soils typically arise from parent materials enriched in these metals. Well-known examples of high-Cd soils are those derived from marine black shales (6), but other parent materials, such as Silurian- age dolomites, can also be enriched in certain potentially toxic metals (2).

Draining metalliferous peatlands for agricultural or other land uses, reduces downstream water quality through the release of sulfide-bound cadmium (Cd) and zinc

(Zn) by oxidation. These metals can be accumulated by plants, causing phytotoxicity

(toxicity to plants). This situation appears to have arisen in peatland regions in Western

New York State, referred to as the Manning peatlands. A comprehensive study in metalliferous peat bogs in Western New York was conducted by Cannon in 1955 (2), who noted inclusions of sphalerite (ZnS), galena (PbS) and gypsum (CaSO4) in outcroppings of Lockport Dolomite, the Silurian-age bedrock near the town of Manning.

High average concentrations of Zn and Pb were measured not only in the dolomitic rock and the surrounding peats, but also in fossilized Silurian reef fauna (2). Soluble Zn is presumed to have entered these wetlands by means of drainage and groundwater emanating from the weathering of dolomitic rock and fossilized reef fauna containing unusually high concentrations of Zn (2). Cannon (2) observed that zinc (among other

7 metals) “has been concentrated by residual enrichment through decay of plant remains in the peat and by precipitation from the ground water”.

Phytotoxicity in vegetable crops was noted in the early 1940’s, not long after the bogs were drained (10). Very high concentrations of zinc (exceeding 500 mg/kg, dry weight basis) were measured in plant shoots (10). Although cadmium in the soils and crops was not measured at that time, subsequent studies have indicated that cadmium and several other heavy metals were present at high concentrations in these peats. It appeared, then, that processes in the soil surface were enhancing zinc solubility and phytoavailability shortly after the peats had been drained. Martínez et al (7) demonstrated that very high total concentrations of zinc and cadmium and severe phytotoxicity have persisted for decades since the last investigation in 1955 (2, 7). A high percentage of the total S in organic-rich soils was found in reduced forms. The authors suggested that sulfur biogeochemical cycling may play an important role in the retention of Zn and Cd in these peat soils (7).

OBJECTIVES

The present work involves investigations on the microbial communities and processes in peat soils overlying the Lockport Dolomite formation of Western New York.

In particular, the interest is focused on bacteria of the sulfur cycle and their potential influence in the biogeochemical control of Zn and Cd retention by sulfur. The broad aim of this research is to help us understand the impact of microorganisms on biogeochemical processes in soils.

Hypotheses: Bacterial populations involved in S cycling are present and active in

naturally metalliferous peat soils. The distribution, abundance and diversity of these

populations vary with soil depth and seasonal changes. These populations may be in close

relation with sulfide-metal minerals.

The aim of this research is to characterize the presence and activity of SRPs in

these organic soil environments and to assess their potential effect on the formation and

dissolution of metal sulfide minerals. The objectives of this work are (analytical approach in parenthesis):

1. Compare classical and molecular methods in characterizing microbial

populations and groups of sulfate reducers in surface soils from the

Manning peatland region. (Microscopic counts, Most Probable Number

[MPN] enumeration, Ribosomal Intergenic Spacer Analysis [RISA]).

2. Detect the genes encoding for enzymes involved in sulfur respiration.

(PCR amplification of dsrAB, apsA and asrA genes)

9 3. Evaluate changes in sulfate-reducing populations as a function of temporal

and spatial variability. (RISA, PCR amplification of dsrAB)

4. Determine the oxidation states of S present in organic-matter rich aerobic

soils. (X-ray absorption near edge spectroscopy [XANES])

This research advances basic knowledge of biogeochemical processes for environmental management. The relationship of microbial processes and populations with Zn, Cd, and S solubility in these peatlands will help to understand the importance of bacteria involved in the mobilization of metals in soils.

REFERENCES

1. Alloway, B. J. 1995. Heavy metals in soils, 2nd ed. Blackie Academic &

Professional, London.

2. Cannon, H. L. 1955. Geochemical relations of zinc-bearing peat to the Lockport

dolomite, Orleans County, New York., p. 119-185, Geological survey bulletin

1000-d. United States Goverment Printing Office, Washington, D.C.

3. Castro, H. F., N. H. Williams, and A. Ogram. 2000. Phylogeny of sulfate-

reducing bacteria. FEMS Microbiol. Ecol. 31:1-9.

4. Ehrlich, H. L. 2002. Geomicrobiology of sulfur, p. 549-620, Geomicrobiology,

4th ed. Marcel Dekker, New York.

5. Germida, J. J., and S. D. Siciliano. 2002. Sulfur cycle in soils, p. 3104-3116. In

G. Bitton (ed.), Encyclopedia of environmental microbiology. Wiley, New York.

6. Kim, K.-W., and I. Thornton. 1993. Influence of uraniferous black shales on

cadmium, molybdenum and selenium in soils and crop plants in the Deog-Pyoung

area of Korea. Environ. Geochem. Health 15:119-133.

7. Martínez, C. E., M. B. McBride, M. T. Kandianis, J. M. Duxburt, S.-J. Yoon,

and W. F. Bleam. 2002. Zinc-sulfur and cadmium-sulfur association in

11 metalliferous peats: Evidence from spectroscopy, distribution coefficients, and

phytoavailability. Environ. Sci. Technol. 36:3683-3689.

8. Shen, Y. N., and R. Buick. 2004. The antiquity of microbial sulfate reduction.

Earth-Science Reviews 64:243-272.

9. Stackebrandt, E., D. A. Stahl, and R. Devereux. 1995. Taxonomic

relationships, p. 49-87. In L. L. Barton (ed.), Sulfate-reducing bacteria. Plenum

Press, New York.

10. Staker, E. V., and R. W. Cummings. 1941. The influence of zinc on the

productivity of certain New York peat soils. Soil Sci. Soc. Proc. 6:207-214.

11. Stevenson, F. J., and M. A. Cole. 1999. The sulfur cycle, p. 330-368, Cycles of

soil: Carbon, nitrogen, phosphorus, sulfur, micronutrients. Wiley, New York.

12. Tabatabai, M. A. 1994. Sulfur oxidation and reduction in soils, p. 1067-1078. In

P. J. Bottomley (ed.), Methods of soil analysis. Part 2, microbiological and

biochemical properties. Soil Science Society of America, Madison, Wis., USA.

Chapter 2

Sulfur-Reducing Prokaryotes in Surface Peats Subjected to Variable Redox Conditions

Carolina Yáñez, Soh-joung Yoon, Mary Ann Bruns, and Carmen Enid Martínez

Submitted for publication to Applied and Environmental Microbiology

ABSTRACT

Peats developed over the Lockport Dolomite formation of Western New York concentrate metals (such as Zn and Cd) by biogeochemical processes. The solubility and mobility of Cd and Zn can increase when the peatlands are drained, initiating oxidation of organic matter and sulfides and leading to phytotoxicity. Moreover, dissimilatory sulfur reduction reactions can control the mobilization and bioavailability of metals classified as chalcophiles (i.e., Ag, Cd, Hg, Zn) and can involve precipitation as metal sulfides and complexation with organic matter. Our objective was to study the diversity of microorganisms involved in sulfur reduction reactions in these peats. The diversity of prokaryotes involved in sulfur reduction was evaluated in oxic peat soils containing high natural levels of sulfur (5,800 to 11,000 mg kg-1), zinc (137.5 to 3,675 mg kg-1) and

cadmium (<1 to 4 mg kg-1). The peats were collected in July 2002 from an agricultural

field in the Manning peatland region of Western New York. Soils from this region

undergo seasonal redox fluctuations, being water-saturated during winter/early spring and

artificially drained in late spring prior to planting. We used most-probable-number

(MPN) enrichments, ribosomal intergenic spacer analysis (RISA), and PCR-based

detection of dsrAB genes to obtain qualitative information on dominant prokaryotic

populations and to estimate the proportion of sulfate-reducing prokaryotes (SRPs) in the

soils. Sulfur electronic oxidation states in these soils were assessed using X-ray

absorption near edge structure (S-XANES) spectroscopy, which showed that only a small

percentage of total soil S (≤10%) existed in the most oxidized form (i.e., sulfates). Much

14 higher percentages of soil S (50-63%) existed in the most reduced electronic oxidation states (i.e., sulfides, disulfides and thiols). Moreover, sulfonates (electronic oxidation state of +5) accounted for 12-32% of the total soil S. MPN enumerations indicated that

SRPs using acetate or lactate as electron donors represented only a small fraction (~103 g-

1 dry soil) of total cell densities as determined by direct microscopy (~109 g-1 dry soil).

Isolates obtained from MPN dilutions were phylogenetically affiliated with Clostridium

species and Desulfotomaculum guttoideum. Cloning and sequencing of RISA fingerprint

bands from MPN enrichments yielded sequences related to Clostridium sp.,

Desulfitobacterium sp., Caloramator sp., and Geosinus sp, all belonging to the

Firmicutes. No amplification was observed with archaeal 16S rRNA-based primers.

Bacterial RISA fingerprints of DNA extracts from all surface soils were highly similar

and DNA sequences revealed high taxonomic bacterial diversity. No dsrAB-PCR

products were obtained from any peat DNA extracts, suggesting that members of the five

currently recognized SRP taxa which possess these genes were absent or present in very

low numbers. With their high sulfonate content and variable redox conditions, we

propose that the Manning peat soils represent an environment with high discovery

potential for studying unknown pathways of sulfur cycling. Our microbiological and

spectroscopic data suggest that sulfur respiration in these soils may involve the use of

sulfonates as electron acceptors and may be driven primarily by members of the

Firmicutes.

INTRODUCTION

Geochemical transfers between the Lockport Dolomite Formation and overlying wetlands near the town of Manning in Western New York have resulted in peat soils that contain high levels of sulfur, zinc, and cadmium (7, 40). Soluble Zn is presumed to have entered these wetlands by means of drainage and groundwater emanating from the weathering of dolomitic rock and fossilized reef fauna containing unusually high concentrations of Zn (7). In this region (referred to as the Manning peatlands), phytotoxicity in vegetable crops was observed in the early 1940s (53), not long after the bogs were drained for agricultural purposes, and very high concentrations of Zn were measured in plant shoots (53). High soil concentrations of S, Zn and Cd and areas producing severe phytotoxicity have persisted for decades (40).

These artificially drained peat soils have undergone drastic seasonal changes in redox for the last 60 years. Although sphalerite nodules (ZnS) were initially identified in the bogs by Cannon (7), these have been weathered out from surface peat soils (39, 40).

Despite prolonged oxic conditions, especially in surface soils, 35-45% of the total sulfur was still present in the most reduced electronic oxidation states such as sulfides (R-S-R,

R-S-S-R, and metal sulfides) and thiols (R-S-H), as shown by sulfur X-ray absorption near edge structure (S-XANES) spectroscopy (40). Dissimilatory sulfur reduction during soil microbial activity may account for the persistence of reduced forms of sulfur in these surface peats. Moreover, dissimilatory sulfur reduction can exert control over the mobility and bioavailability of heavy metals (i.e., Zn and Cd) that tend to precipitate as

16 sulfides and/or form complexes with thiol functional groups of organic matter. These are compelling reasons to evaluate the diversity and abundance of prokaryotes involved in sulfur reduction in these oxic peat surface soils.

Diverse prokaryotes reduce sulfate as a terminal electron acceptor during anaerobic respiration. So far, five phylogenetic groups of sulfate-reducing prokaryotes

(SRP) have been identified based on rRNA sequence analysis (8, 52). These include the low G+C Gram positive spore forming SRB (Desulfotomaculum spp.); Gram-negative mesophilic members of δ-proteobacteria (Desulfovibrio spp.); thermophilic bacteria in the Thermodesulfobacterium division; thermophilic members of the Nitrospira division

(Thermodesulfovibrio spp.); and thermophilic archaeal (Archaeoglobus spp.). In addition to inorganic sulfate, soil bacteria have been shown to utilize organosulfur compounds such as sulfonates and sulfate esters as sulfur sources during assimilation (36-38).

Furthermore, Denger et al. (12) demonstrated that an isolate from untreated anoxic sludge was able to ferment sulfonates in laboratory incubations. Sulfur and other oxidized forms of sulfur can also be used anaerobically as terminal electron acceptors in respiration (14,

34, 46, 49). However, prokaryotes that reduce sulfur, sulfite or thiosulfate do not form phylogenetically coherent groups of bacteria or archaea (46).

Most studies of the biological sulfur cycle have been conducted in anoxic environments such as marine and freshwater sediments as well as in rice paddy soils where SRB play important roles in carbon and sulfur biogeochemical cycles (23, 27, 66).

The surface soils from the Manning peatland region are likely to represent a novel environment for investigating dissimilatory sulfur reduction since they have been subjected to more than 60 years of continuously variable oxic/anoxic conditions and

17 therefore to dynamic and multidirectional sulfur transformations. As a first approach to understanding the link between sulfur and metal (Zn and Cd) biogeochemical cycling in the surface soils from the Manning peatlands, we aimed to identify the dominant sulfate- reducing prokaryotic populations and estimate their abundance. Molecular (dsrAB amplification) and cultural approaches (enrichments and MPN enumeration) were used for the characterization of communities involved in the sulfur reduction process.

Microbiological data was coupled to the spectroscopic characterization (S-XANES) of the oxidation states of sulfur in the peats under investigation. Our objective was to elucidate the potential substrates involved in sulfur cycling in these sulfur-rich surface soils and the microorganisms driving the reactions.

MATERIALS AND METHODS

Soil samples and chemical analyses. Peat soils in the Manning, New York, region developed from coniferous (hemlock and pine) and hardwood (largely oak, but also maple, beech, elm and hickory) trees and are classified as typic haplosaprists of the

Carlisle Series (7, 24). Surface soil samples (0 - 5 cm) were collected by soil coring with an 8-cm-wide auger from a field planted with onion in the Manning peatland region of

Western New York (78º6.146'W, 43º9.470'N) in July 2002. The surface soils were collected along a gradient starting in an area where there was no plant growth (soil labeled “Peat 1”) and moving eastwards for about 8 meters to a zone of typical plant growth (soil labeled “Peat 8”). The soil samples were brought to the laboratory where they were kept at 4ºC for about 6 months. The gravimetric moisture content was

18 determined after drying the soils at 50°C for about 16 hours. Total elemental composition of bulk surface soils was determined by ICP analysis of HClO4/HNO3/H2SO4 digests

using 2 g of air-dried soil (22). Total sulfur in the soils was determined by the high-

temperature furnace combustion method (ASTM D4293-02) with infrared absorption and

soil organic matter content was determined by weight loss on ignition (43). Exchangeable

zinc was determined after adding 25 mL of a 0.01M Ca(NO3)2 solution to a centrifuge tube containing 2.5 g of soil (dry-weight basis). The soil-Ca(NO3)2 suspension was

shaken for 4 h in an end-over-end shaker, the suspension was then centrifuged for 10 min

at 15000 rpm (17542 g) and the supernatant filtered using a 0.22 µm membrane filter.

The concentration of Zn was measured by atomic absorption (Instrumentation

Laboratories Video 22 AA/AE spectrometer). The pH of the 0.01 M Ca(NO3)2 extract

was also measured. X-ray diffraction analyses have indicated that quartz and dolomite are

the only crystalline phases present in surface peats of this region (39, 40). Soil properties

are presented in Table 2-1.

Sulfur X-ray Absorption Near Edge Structure Spectroscopy (S-XANES). The

oxidation states in which sulfur (organic and inorganic forms) is present in these soil

samples was determined using S-XANES spectroscopy. The sulfur K-edge (2472 eV)

XANES spectra were collected at Beamline X-19A of the National Synchrotron Light

Source (NSLS), Brookhaven National Laboratory under standard operating conditions

(2.801 GeV and a current ranging from 250 to 100 mA). The monochrometer used in

these experiments consisted of two parallel Si {111} crystals with an entrance slit of 0.5

mm. Each soil sample was air-dried and ground before pressing it into a 0.5-mm thick

acrylic holder with a 2.5 µm thick Mylar film (Chemplex Industries, NY) window. The

19 spectra were recorded in fluorescence mode using a Stern-Heald ionization detector filled with He gas and positioned 90o to the incident beam. The monochrometer was detuned

70% at the S K-edge in order to reduce fluorescence induced by high-order harmonics.

The elemental S K-edge spectrum (assigned a value of 2472 eV) was used for energy

calibration. Scans ranged from 20 eV below to 50 eV above the S absorption edge with

0.2 eV step size.

The experimental S-XANES spectra were deconvoluted into pseudocomponents using the nonlinear least-square fitting routine SOLVER from MS-Excel. Sulfur-XANES data analyses for organic-rich surface soils are described by Martínez et al. (40) and additional details on the methodology are reported by Xia et al. (69). Our experimental

XANES spectra were fitted using a series of Gaussian peaks (labeled G1 to G6) that represent the s → p transitions (white line), and arctangent step functions that represent the transition of ejected photoelectrons to the continuum (step height or background). The energy positions (eV) of the Gaussian curves are used to identify the oxidation states of S present in the sample while the area of the Gaussian curves is used to calculate the percentage of the total S present at that particular oxidation state. The linear component of the spectral baseline was removed prior to fitting and the areas were corrected for the change in absorption cross-section with increasing oxidation state (69). We report electronic oxidation states, rather than formal oxidation states, that reflect the actual electronic density in the valence shell of sulfur. The electronic oxidation states of S in its most reduced forms (disulfides, sulfides, thiols) generally fall within the range of 0.1 to

0.8 depending on S bonding environment (S, H, C, metals). For S with valence ≥ 4 the

differences between the electronic and formal oxidation states are not significant because

20 of the electronegativity of the O atom. The accuracy limit of the estimates obtained by fittings of sulfur XANES spectra has been reported to be 5 to 10 % (20, 21, 57).

Bacterial enumerations and enrichments. Sulfate-reducing prokaryotes were estimated using the most probable number method (MPN). The basal medium

-1 composition was as follows (in g L ): MgSO4·7H2O (2.0 g); peptone (2.0 g); Na2SO4 (1.5 g); Beef extract (1.0 g); K2HPO4 (0.5 g); CaCl2·7H2O (0.13 g). Two media were

prepared, adding either sodium acetate (2.55 g L-1) or sodium lactate (3.5 g L-1) to the basal medium. The media were adjusted to pH 7.8 and distributed into 10-mL serum bottles. The bottles were closed with butyl stoppers (Bellco Biotechnologies, Vineland,

NJ), sealed with aluminum caps, made anoxic by N2 flushing, and autoclaved. A soil slurry was prepared (68) that served as inoculum for triplicate ten-fold MPN dilutions using samples peat 1, peat 2 and peat 8. Briefly, 2.5 g of peat and 0.5 g 1mm-acid-washed glass beads were mixed in 23.75 mL of a pH 7.2 phosphate buffer. Slurries were vortexed

at maximum speed for 5 minutes. Prior to inoculation, the following media constituents

were subsequently added from freshly prepared sterile solutions: Fe(NH4)2(SO4)2·6H2O

(3.92 g/100 mL) and sodium ascorbate solution (0.05 g/100 mL), 0.1 mL each.

Enrichments were incubated in the dark at 25°C for two months. Tubes with black precipitate due to sulfide production were considered positive for MPN calculations.

Calculation of MPN values and confidence intervals were performed using MPN tables

(68). A student t-test was used to determine whether the MPN values were significantly different (9). Isolates were obtained by spread plating and subsequently, streak purifying

MPN positive cultures using the same medium with 1.5% agar. Desulfovibrio vulgaris

21 ATCC 29579 was used as a positive control inoculum for demonstrating SRP growth in the acetate and lactate media.

Petri dishes were incubated under anaerobic conditions (glove box) for 1 month or until growth was observed. Total direct counts of bacteria in soil samples were obtained using epifluorescence microscopy (Olympus BX-60, 40X). Soil slurries were prepared as previously described in this section. A 1/100 dilution was prepared in the same buffer and

1 mL of soil suspension was stained with SYBR® green I (Cambrex; Rockland, ME) for

15 minutes. Counts were done with a hemocytometer in duplicate. Bacteria were counted

in 10 fields and the average per field (around 100 cells for each sample) was multiplied

by a conversion factor based on the hemocytometer.

DNA extraction. Microbial community DNA was extracted from ~0.5 g of peats

using the MoBio UltraClean™ Soil DNA Kit (Solano Beach, CA) according to the

manufacturer’s instructions. DNA concentration was estimated spectrophotometrically by

measuring the continuous absorbance from 230 to 300 nm (6). In addition, DNA from

positive MPN cultures was extracted using MoBio UltraClean™ Microbial DNA Kit

(Solano Beach, CA).

PCR amplification of dsrAB genes. PCR reactions were conducted using the set

of primers and PCR conditions described by Wagner et al. (64). This set consisted of

primers DSR1F (5’-ACSCACTGGAAGCACG-3’) and DSR4R (5’-

GTGTAGCAGTTACCGCA-3’) which amplifies a 1.9-kb fragment of the dsrAB gene.

The PCR reactions were prepared by mixing 15 pmol of each primer solution, 20 ng of

DNA, 2.5 µl of 10X PCR buffer, 1.25 µl of 10 mg/µl bovine serum albumine, 0.4 µl

dNTPs (25 mM each), and 1.5 U of Taq polymerase Eppendorf HotMaster in final

22 reaction volume of 25 µl. DNA from Desulfovibrio vulgaris ATCC 29579 was used as positive control.

RISA, cloning and sequencing. Soil microbial communities in peats 1 to 8 were studied by amplifying the spacer between the 16S and 23S ribosomal RNA genes. The bacterial primers used were 16S-1406f (5'-TGYACACACCGCCCGT-3') and 23S-115r

(5'-GGGTTBCCCCATTCGG-3') and have been previously described by Lane (33). The archaeal primers 16S-1214f (5'-GGTCAGYATGCCCCGAA-3') and 23S-46r (5'-

TCGGYGCCCGAGCCGAGCCATCC-3') were also tested (unpublished data).

Identification of isolates was assessed by amplification of nearly full-length 16S rRNA gene using primers 16S-27f (5'-AGAGTTTGATCMTGGCTCAG-3') and 16S-1492r (5'-

TACGGYTACCTTGTTACGACTT-3') (33). For PCR amplification, 15 pmol of each primer solution, 20 ng of DNA, 2.5 µl of 10X PCR buffer, 1.25 µl of 10 mg/µl bovine serum albumine, 0.4 µl dNTPs (25 mM each), and 1.5 U of Taq polymerase Eppendorf

HotMaster were combined in a final reaction volume of 25 µl. PCR reactions were setup in triplicate. After an initial denaturation step of 2 min at 94°C, amplification was carried out for 25 cycles, with each cycle consisting of 30 s at 94°C, 30 s at 55°C, and 1 min at

72°C. PCR products were visualized in a 1% agarose gel stained with ethidium bromide and triplicate reaction mixtures were pooled prior to electrophoresis. DNA fingerprints were obtained by separation of pooled amplicons using a 0.75 mm thick, 4% PAGE-TBE gel at 50 V for 15 hours and then stained with 0.01% SYBR-green I. Gels were photographed using an EpiChemi II digital camera and transilluminator (UVP Inc.,

Upland, CA). Selected bands were excised and DNA was extracted using the “crush and soak” method (48). The purified DNA was ligated into pCR®4-TOPO® and transformed

23 into ONE SHOT competent Escherichia coli cells following the manufacturer’s directions (TA Cloning System, Invitrogen). For isolates, PCR products from genomic

DNA were separated in a 1%-low-melting-point agarose gels and bands were excised for cloning and sequencing.

DNA sequences were obtained with an ABI Hitachi 3730XL DNA Analyzer at the Penn State’s Nucleic Acid Facility. The M13f and M13r primers were used to sequence both strands of the insert. Sequences from clones were edited to remove any remaining vector fragment and were assembled using SeqMan™ II software (DNASTAR

Inc., Madison, WI). Most similar 16S rRNA gene sequences in the GenBank database were retrieved using BLAST (2).

Phylogenetic trees were reconstructed by using Clustal X version 1.81 (60) and the neighbor-joining distance method with Jukes-Cantor correction (47). Maximum likelihood trees were obtained using the programs included in the PHYLIP package (18).

The bootstrap confidence levels were defined from 100 iterations of tree reconstruction

(17).

Nucleotide sequence accession numbers. Sequences were deposited in GenBank under accession numbers DQ491454 to DQ491482 for 16S rRNA-23S rRNA sequences and DQ479411 to DQ479420 for 16S rRNA sequences from isolates.

RESULTS

Chemical characteristics. All peats exhibited moderately acidic pH (4.9 to 5.9) and high (average 77%) soil organic matter content (Table 2-1). These peats possess

24 unusually high concentrations of sulfur, zinc, and cadmium compared to most soils

(Table 2-1). Zinc concentrations ranged between 137.5 and 3675 mg kg-1, in most cases exceeding the mean concentrations in soils (worldwide 10-300 mg kg-1; average 50 mg

kg-1) (1). Cadmium concentrations varied from below the limit of detection of the ICP (<

1 mg kg-1) to 4.0 mg kg-1. Sulfur concentration in surface peats ranged from 5,800 to

11,000 mg kg-1, greatly exceeding the typical range for soils (100 to 500 mg kg-1) (54).

The lowest, albeit elevated, sulfur concentrations were observed in Peat 1 and Peat 2, the soils closest to the dolomite outcrop and at the edge of the onion field (Table 2-1).

Whereas sulfur concentrations were relatively constant (~1%) in Peats 3 to 8, total soil Zn and Cd concentrations decreased from Peat 3 (no plant growth) to Peat 8 (typical plant growth). Decreased concentrations of total soil Zn and better plant growth were paralleled by decreases in the amount of salt-extractable Zn (exchangeable Zn) (Table 2-

1). Our chemical data (total soil Zn and salt-extractable Zn) did not explain why Peat 1 and Peat 2 supported no plant growth in the field. The lack of plant growth at this end of the sampling transect was perhaps the result of the soils being at the edge of the onion field, where they would have received more equipment traffic.

Table 2-1: Elemental concentrations (dry weight basis), soil organic matter (SOM), water content, exchangeable Zn, and pH of the peats (peats 1 to 8).

Soil Soil SOM Water S Zn Cd Fe Ca Exchangeable Zna pH (%) content (%) (mg kg-1) (mg kg-1) (mg kg-1) (mg kg-1) (mg kg-1) (%) Peat 1 5.89 79 52.6 0.58 137.5 1.25 11225 16925 0.20 (1.49) Peat 2 5.35 80 46.4 0.83 625 1.25 6600 23925 1.34 (2.14) Peat 3 5.36 78 77.0 1.04 3675 4.0 10800 18875 4.38 (1.19) Peat 4 5.21 77 77.3 1.10 3675 2.5 10250 16700 5.60 (1.52) Peat 5 4.98 52 73.5 1.06 2350 1.75 9100 21025 5.04 (2.14) Peat 6 5.02 84 69.7 1.07 1125 1.5 7900 16625 2.09 (1.86) Peat 7 5.02 82 51.3 1.10 350 < 1 4975 27500 0.89 (2.54) Peat 8 4.89 84 59.3 1.10 300 < 1 5225 17850 0.82 (2.75)

a Value in parenthesis represents the % of the total Zn in 0.01 M Ca(NO3) extracts.

Table 2-2: Percentage of the total soil sulfur at estimated electronic oxidation state for surface peats (peat 1 to 8).

Soil G1a G2 G3 G4 G5 G6 (0.09-0.1) (0.6-0.7) (1.4-1.5) (4) (5) (6)

Representative Structure

R-S-R R-S-R R-(SO)-R R-(SO2)-R R-SO3-H R-OSO3–H 2- 2- R-S-S-R R-S-S-R SO3 SO4 R-S-H R-S-H Peat 1 39.2 10.9 12.7 4.8 32.4 0.0 Peat 2 40.2 10.7 13.8 3.4 31.2 0.7 Peat 3 56.4 6.3 12.4 2.7 20.76 1.4 Peat 4 50.1 7.0 13.7 2.1 25.8 1.3 Peat 5 44.9 11.0 14.4 2.4 26.8 0.6 Peat 6 42.0 12.6 14.8 2.8 27.1 0.6 Peat 7 41.2 9.7 20.7 4.0 14.5 10.0 Peat 8 57.9 1.6 12.5 5.9 12.4 9.7

a Gaussian curve (G1, G2, etc.) and estimated electronic oxidation state (in parenthesis). The range of energy positions (eV) for each gaussian curve is as follows: G1 = 0.05-0.4; G2 = 1.2-1.4; G3 = 2.5-2.8; G4 = 6.1-7.7; G5 = 7.9-8.8; G6 = 9.5-10.0. 26

Spectroscopic (S-XANES) characterization of the oxidation states of sulfur. Spectral analyses indicate the presence of a range of S oxidation states in these soils (Fig. 2-1). Table 2-2 summarizes the results for all soils. Most of the sulfur (50-63%) was present in the most reduced electronic oxidation states (-0.09-0.7), which are represented by sulfide, disulfide and thiol chemical forms. Moreover, 12-32% of the total soil S had an electronic oxidation state of +5

(such as in sulfonate, R-S-O3-H), while ≤10% exists in the +6 electronic oxidation state (such as

2- in inorganic-sulfate (SO4 ) and/or organo-sulfate (R-O-S-O3-H)). These results agree with previously reported values for surface peats derived from the same parent material (40) and compare favorably to data for poorly drained soil, peat, wetlands and aquatic sediments (26, 42).

Enumeration of total bacteria and sulfate-reducers in soils. Total bacterial counts did not vary significantly (~109 cells/g dry soil) in Peat 1, Peat 2 and, Peat 8 (Table 2-3) and are

within the range described for organic soils (62). Growth of the indigenous SRP was observed up

to the 10-3 dilutions (but not in all triplicates) for both acetate and lactate enrichments in Peat 1,

Peat 2 and Peat 8. These numbers did not change after 4 and 6 months of incubation. The number

of culturable sulfate-reducing bacteria in these soils represented only a small percentage

(0.0001%) of the total microbial numbers observed by microscopy and its estimation did not

change with different electron donors (~103 cells/g dry soil).

Table 2-3: Total cell number estimation and number of culturable sulfate-reducing bacteria in selected peats (peat 1, 2 and 8).

Total bacteria (cells g-1 dry soil) Culturable bacteria (MPN g-1 dry soil) Acetate Lactate Peat 1 1.50 x 109 1.48 x 103 2.94 x 103 (3.35 x 108)a (3.50 x 102 – 4.31 x 103)b (5.50 x 102 – 1.27 x 104) b Peat 2 1.01 x 109 3.73 x 103 1.65 x 103 (1.01 x 109) a (6.90 x 102 – 1.51 x 104) b (3.60 x 102 – 4.80 x 103) b Peat 8 1.35 x 109 5.60 x 103 8.94 x 103 (1.35 x 109) a (1.10 x 102 – 2.26 x 104) b (2.14 x 103 – 2.98 x 104) b

a Values in parentheses represent standard deviation. b Values in parentheses represent 95% confidence limits for MPN.

30 dsrAB gene detection and isolates. The occurrence of SRP in the surface peat soils was assessed by testing for the presence of dsrAB genes that encode dissimilatory sulfite reductases present in all known SRP (30, 64, 70). Reduction of sulfate was observed in MPN enrichments, indicating sulfate reduction activity. However, no PCR products were obtained from DNA extracted from unenriched peats, even after nested

PCR using previous PCR reactions as a template. These results suggest that SRP with homologous genes were absent or present in such low numbers that they could not be detected.

Colonies with different morphologies were selected to determine the identity of bacteria growing in SRP enrichments. Five bacterial strains were isolated from acetate enrichments and 5 strains from lactate enrichments (Table 2-4). Although culture media was confirmed to support the growth of Desulfovibrio vulgaris ATCC 29579, 16S rRNA sequencing of isolates’ genomic DNA indicated that they all belonged to the Firmicutes phylum (Fig. 2-2). Isolates with the same closest relatives were identified in both enrichment conditions and in different peat samples. Nearly full-length 16S rRNA gene sequences of CYP1 (Peat 1) and CYP9 (Peat 2), which were isolated in media containing acetate and lactate, respectively, were 99% similar to each other and most closely affiliated with Desulfotomaculum guttoideum (99% similarities). Pairwise 16S rRNA sequence similarities for CYP3 (Peat 2), CYP4 (Peat 8), CYP6 (Peat 1), and CYP11 (Peat

8) were all 99%, and these isolates were most closely related to Clostridium sp. ZIRB-1, an isolate from a metal-rich termite gut (63) (97-98% similarities). Overall, analysis of the phylogenetic relationships among isolates resulted in the identification of six ribotypes (subtype of a bacterial strain, based on the analysis of 16S rRNA genes) in the

31 peats (Fig. 2-2). Among these ribotypes, two were found in acetate (CYP2 and CYP5), two in lactate (CYP7 and CYP8), and two were common to both conditions. According to the phylogeny of the genus Clostridium (10), the ribotypes enriched within acetate were affiliated with Cluster XIVa and Cluster I, respectively. Those enriched with lactate were affiliated with Cluster XI and Cluster I, and the common ribotypes for acetate and lactate were affiliated with Cluster XI (CYP3, CYP4, CYP6, and CYP11) and Cluster I (CYP1 and CYP9). Maximum likelihood analysis generated a tree with identical topology and similar or higher bootstrap values obtained with neighbor-joining methods (data not shown).

The identification of Desulfotomaculum guttoideum as the close relative of isolates CYP1 and CYP9, when all other closest relatives were classified as Clostridium spp., appears to be consistent with its uncertain taxonomic assignment. Although it was originally classified within Cluster III of Desulfotomaculum spp., Stackebrandt et al. (51) subsequently pointed out its high relatedness to members of Clostridium cluster XIVa, including Clostridium sphenoides, Clostridium celerecrescens, Clostridium aerotolerans and Clostridium xylanolyticum (10). All these Clostridium spp., in addition to D. guttoideum, reduce sulfite and thiosulfate, but not sulfate. More recently, the ability of other Desulfotomaculum spp. to reduce sulfate has been called into question, with some members in Cluster I reported to have lost this ability in favor of syntrophy-based metabolism (25).

Table 2-4: 16S rRNA analysis of isolates obtained from enrichments with acetate or lactate in selected peats (peats 1, 2, and 8).

Most closely related Isolate Origin % Accession N° c bacterial sequences a similarity b Desulfotomaculum CYP1 Peat 1 SRB/acetate 99 DQ479411 guttoideum CYP9 Peat 2 SRB/lactate 99 DQ479419 (Y11568) Clostridium sp. CYP2 Peat 1 SRB/acetate 94 DQ479412 (AY221993) Clostridium sp. CYP3 Peat 2 SRB/acetate 98 DQ479413 (AY532163) CYP4 Peat 8 SRB/acetate 97 DQ479414 CYP6 Peat 1 SRB/lactate 98 DQ479416 CYP11 Peat 8 SRB/lactate 98 DQ479420 Clostridium tunisiense CYP5 Peat 8 SRB/acetate 99 DQ479415 (AY187622) Clostridium mangenotii CYP7 Peat 1 SRB/lactate 96 DQ479417 (M59098) Clostridium frigidicarnis CYP8 Peat 2 SRB/lactate 96 DQ479418 (AF069742)

a Accession number of closest relative is indicated in parenthesis. b Refers to percentage of similarity to related bacterial sequences. c Accession number refers to the unique identifier assigned in GenBank.

34 RISA. All eight surface peat soils produced similar RISA fingerprints, which suggested that dominant microbial populations in these surface peat soils were similar

(Fig. 2-3). The number of bands varied from 27 to 29, with 25 bands being common in all three fingerprints, and band sizes ranged from 400 bp bases to 1200 bp (Fig. 2-3). No amplification was observed when archaeal primers were used (data not shown).

BLAST analyses of 16S rRNA (~ 150 bp) bands (2-3 clones) common to Peat 1,

Peat 2 and Peat 8 fingerprints indicated that bacterial populations included representatives of Proteobacteria (Magnetospirillum sp.; Acetobacter sp.; Pseudomonas sp.; Candidatus Competibacter phosphatis; Nevskia sp.; Bradyrhizobium sp.; Frateuria sp.; Roseobacter sp.), Firmicutes (Bacillus sp.; Desulfitobacterium sp.), Acidobacteria

(Holophaga sp.) , and Verrucomicrobia (Opitutus sp.) (Table 2-5). Differences observed in cloned sequences from the same RISA band demonstrated that some of the bands (3, 4,

6) were heterogeneous (Table 2-5). Among the bands sequenced from DNA extracted directly from unenriched soils, only one showed similarity (91%) to a known SRB,

Desulfitobacterium sp. (Table 2-5). This band was obtained from the RISA fingerprint of peat 8. Sulfur-XANES analysis of Peat 8 indicated that ~10% of the total soil S exists as sulfates. These results are distinct from S-XANES analyses of Peat 1 and Peat 2 that showed <0.7% sulfate (Table 2-2). The availability of oxidized forms of sulfur (i.e., sulfate or sulfonate) appears to influence the bacterial populations involved in S cycling in these soils.

Enrichment cultures yielded RISA fingerprints that differ among peats 1, 2 and 8

(Fig. 2-4). As with sequences from isolates, most sequences belonged to Firmicutes which demonstrated that enrichment conditions were favorable for representatives of this

35 particular phylum (Table 2-6). In Peat 2 and Peat 8, however, enrichment with lactate selected for bacterial populations related to the δ-subgroup of Proteobacteria

(Desulfitobacterium hafniense), a bacterial division related to well-known SRBs.

Table 2-5: Sequence analysis of bands excised from RISA fingerprinting derived from bacterial 16S rRNA extracted from surface peat soils (peat 1, 2 and 8).

Band position Related bacterial sequences a % similarity b Accession N° c Soil band 1 Magnetospirillum magneticum 86 DQ491454 (AP007255) Soil band 2 Acetobacter oeni 86 DQ491455 (AY829472) Soil band 3 Bacillus sp. 88 DQ491456 (AY608947) Pseudomonas fluorescens 97 DQ491457 (DQ178235) Holophaga sp. 94 DQ491458 (AF385537) Opitutus sp. 98 DQ491459 (X99390) Candidatus Competibacter phosphatis 94 DQ491460 AY172151 Soil band 4 Nevskia ramosa 96 DQ491461 (AJ001343) Uncultured Acidobacteria bacterium 94 DQ491462 (AY697647) Bradyrhizobium japonicum 97 DQ491463 (X87272) Soil band 5 Frateuria aurantia 98 DQ491464 (AB091198) Soil band 6 Uncultured Roseobacter sp. 98 DQ491465 (AY627365) Magnetospirillum gryphiswaldense 87 DQ491466 (AM085146) Desulfitobacterium sp. 91 DQ491467 (AB194704)

Figure 2-4: RISA from SRB-specific enrichments. The intergenic spacer region was PCR amplified using bacterial primers and resolved on a non-denaturating 4% polyacrylamide gel. Three soils were studied: Peat 1 (lanes 1, 2, and 3), Peat 2 (lanes 4, 5, and 6) and Peat 8 (lanes 7 and 8). Acetate enrichments are shown in lanes 2, and 5; lactate enrichments are shown in lanes 3, 6, and 8. Unenriched soils are shown in lanes 1, 4, and 7. M, molecular weight marker. Bands selected for sequencing are marked with a number.

Table 2-6: Sequence analysis of bands excised from RISA fingerprinting derived from bacterial 16S rRNA extracted from enrichments.

Band position Related bacterial % similarity b Accession N° c sequences a Enrichment band 1 Clostridium tetani 97 DQ491468 (AE015945) DQ491469 Enrichment band 2 Desulfitobacterium 99 DQ491470 hafniense (AP008230) Enrichment band 3 Desulfosporosinus orientis 99 DQ491471 (Y11571) DQ491472 Enrichment band 4 Caloramator viterbiensis 95 DQ491473 (AF181848) Enrichment band 5 Desulfitobacterium 99 DQ491474 hafniense (AP008230) Enrichment band 6 Caloramator viterbiensis 95 DQ491475 (AF181848) Enrichment band 7 Desulfitobacterium 100 DQ491476 hafniense (AP008230) Enrichment band 8 Geosinus fermentans 100 DQ491477 (DQ145536) DQ491478 Enrichment band 9 Caloramator viterbiensis 96 DQ491479 (AF181848) Enrichment band 10 Geosinus fermentans 99 DQ491480 (DQ145536) DQ491481 Enrichment band 11 Clostridium tetani 98 DQ491482 (AE015927)

DISCUSSION

Draining of wetlands results in aerobic decomposition of organic matter, which is likely accompanied by the oxidation of sulfides to sulfates and pH reduction. However, after more than 60 years of exposure to variable and prolonged oxidation/reduction conditions, the surface peats from the Manning region of Western New York contain 50-

63% of the total sulfur in reduced forms. The presence, or persistence, of reduced forms of sulfur may be attributed to: (i) slow rates of oxidation resulting from restricted oxygen diffusion (during the dry season) into the interior of soil particles (50), (ii) blockage of thiol functional groups due to metal (Zn, Cd) bonding that retards or prevents its oxidation, (iii) microenvironments during the dry season and/or (iv) prolonged wet conditions prevailing in the field that can support dissimilatory sulfur reduction. We can infer, however, that some oxidation of reduced sulfur has occurred in the surface peats since a higher percentage (60-80%) of the total sulfur in deep soils is present in reduced forms and the pH of surface peats is lower than for deep peats (unpublished results).

Usually, the preferred source of sulfur for microorganisms is inorganic sulfur

(28). However, more than 90% of sulfur in soils is typically found in organic form, as sulfate-esters (oxidation state of +6) and carbon-bonded sulfur including sulfonates (58).

Soil bacteria are able to utilize organosulfur compounds including sulfonates and sulfate esters as sulfur sources for assimilation (28) or for bacterial survival in sulfur-deficient soils (41). Experiments using pure SRB strains in laboratory incubations have also demonstrated that sulfonates (isethionate, cysteate, and taurine) can act as terminal

41 electron acceptors for growth by sulfate- and sulfite-reducing bacteria with the resultant production of hydrogen sulfide (36-38). In our work, S-XANES indicated that a large fraction of the total soil sulfur is found in the +5 oxidation state, suggesting that sulfonates can participate in and support sulfur cycling in these peat soils. Sulfate- reducers have been enumerated using MPN in such diverse environments as sea sediments, aquifers, mine tailings, and ricefields where the conditions are suitable for their development, with populations ranging from 102 to 107 cells/g dry weight soil or

sediment (3, 5, 19, 65, 67). In the surface soils from the Manning peatlands, the number

of SRP was in the order of 103 cells/g dry soil. SRP counts in the same order of

magnitude were reported by Escoffier et al. (16) using ricefield soils (102 to 104 cells/g dry soil). Low counts of SRP in surface soils could be explained by the presence of oxygen. Peters at al. (44) demonstrated that strictly anaerobic bacteria such as methanogenic, sulfate-reducing, and homoacetogenic bacteria are present in oxic soils and can be cultured. After enrichment, these soils presented lower anaerobic cell counts than those from typical anoxic habitats; for example, the number of sulfate- reducing bacteria varied from 4 to 96,000 cells g-1 dry soil. Sulfate-reducing bacteria have oxygen-

protecting enzymes such as superoxide dismutase and catalase that permit their survival

under oxic conditions (11). Anaerobic microniches within soil particles might explain their presence in otherwise apparent oxic environments. Moreover, SRB have developed strategies for protection against oxygen, forming aggregates upon exposure (31).

Specific environmental conditions can contribute to the selection of distinct

groups of sulfate-reducing bacteria (13, 35, 45). Leloup et al. (35), for example, reported

that in freshwater sediments most sequences were affiliated with Desulfotomaculum

42 species (Gram positive) while in a mixing zone between marine and freshwaters sequences belonged to the Desulfobacterales (Gram negative) order. Furthermore, low

G+C Gram positive bacteria were important populations in rice paddies, possibly due to their ability to form spores which could render them resistant to redox changes resulting from seasonal flooding (55, 56, 67). Surface peat soils in the present study harbored principally members related to the phylum Firmicutes, specifically Clostridium spp.

These soils underwent repeated wet/dry cycles and contained unusually high concentrations of sulfur. In contrast to the “classical” SRP that have been isolated from sediments with sustained low redox and saturation, the populations in these surface peats represent SRP that are selected as a result of fluctuating redox conditions.

Clostridium spp. are not considered “classical” sulfate-reducing organisms.

However, clostridia are known to be ubiquitous in soils. Escoffier et al. (15) reported

Clostridium strains capable of reducing thiosulfate but not sulfate, suggesting a significant role of this genus in S cycling in ricefield or wetland soils. Some of the strains isolated from our surface peats share this ability (data not shown), confirming the potentially important role of Firmicutes in S cycling in soils subjected to variable redox conditions. Denger et al. (12), on the other hand, demonstrated that fermentation of taurine is catalyzed by a strictly anaerobic, Gram-positive, spore-forming strain

GKNTAU. Besides the fermentation of sulfonates, the authors also reported thiosulfate as a novel fermentation product. The production of thiosulfate coupled to its use as an electron acceptor by some bacterial strains suggest a possible pathway for autochthonous soil bacteria to dissimilate oxidized forms of sulfur other than sulfate.

43 One genetic characteristic of Clostridium spp. that may contribute to their abundance and survival in oxic samples is high copy number of rRNA operons. The copy number of rRNA operons per bacterial genome varies from 1 to as many as 15 and the greatest numbers of rRNA operons per genome are found in spore-forming bacteria from soils. Klappenbach et al. (29) have suggested that this feature reflects ecological strategies of Clostridium sp. to adapt to stressful conditions, such as high metal concentrations. This genetic characteristic might explain the prevalence in our isolates of representatives of the Firmicutes.

In unperturbed organic soils, the community genome size may be equivalent to

6000–10000 Escherichia coli genomes, while the corresponding diversity in agriculturally disturbed or heavy metal-polluted soils may only be equivalent to 350–

1500 genomes, as shown by DNA re-association kinetic experiments (61, 62). Qualitative comparisons of bacterial communities using RISA indicated a high degree of diversity in all surface peats studied. It is worth mentioning that soil bacterial populations that represent less than 1% of the total bacterial community will not give rise to DNA bands in RISA fingerprints (4). Since these soils contained 1 x 109 cells/g, populations of SRB

would need to be at densities of 1 x 108 cells/g or greater to be detectable in RISA.

One likely explanation for the recovery of different types of sequences from soils

and enrichments was the neutral pH of the enrichment media, which contrasted with the

moderately acidic pH of the soils, and thus resulted in low outgrowth of resident SRP

acclimated to lower pH. The pH contrast and long incubation of the enrichments were

likely explanations for the lack of overlap between sequences from direct-amplified DNA

and the enrichments. For example, an isolate from the Peat 8 enrichment had an rRNA

44 sequence with 99% similarity to that of Clostridium tunisiense, which grows between pH

5.5 - 8.7 with an optimum of 7.8 (59). Such neutrophilic populations are not likely to be present in high numbers in the peats of this study. Another explanation for low SRP estimates was inappropriate electron donors. Although lactate can be used as an electron donor for sulfate reduction by most species (Desulfobacter and some Desulfobacterium species are exceptions) and acetate is used by many sulfate reducers capable of complete oxidation of the electron donor (including strains of the genus Desulfobacter and some

Desulfotomaculum strains) (8), these substrates may not have been optimal for supporting the outgrowth of SRP in the surface peats of the Manning region. In their study of SRP in subsurface mine biofilms, for example, Labrenz and Banfield (32) used enrichment media containing pyruvate, which was not used in our study.

CONCLUSIONS

This study reports the bacterial diversity and sulfate-reducing populations in surface peat soils developed over the Lockport Dolomite formation of Western New

York. Our results provide several lines of evidence to suggest that the sulfur cycle in these soils is driven by Gram-positive bacteria related to Clostridium spp. Sulfur-XANES spectroscopy indicate that sulfur species with an estimated electronic oxidation state of

+5 (i.e. sulfonates, R-SO3-H) comprise a substantial fraction of the total sulfur in these peats and might provide important substrates for dissimilatory metabolism. However, the

use of sulfonates for dissimilatory respiration needs to be confirmed by testing isolates

obtained from these peats. Soils from the Manning peatland region appear to represent a

45 novel habitat for sulfur reduction and populations: substrates, pathways and products may differ from well-studied anoxic environments. Further studies are necessary to enhance our understanding of terrestrial sulfur cycling in the soils from the Manning peatland region and its relation to metal mobility and bioavailability.

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Chapter 3

Novel Functional Genes and Enzymatic Pathways Involved in Dissimilatory Reduction of Sulfur Compounds in Agriculturally Drained Peats

Carolina Yáñez, Mary Ann Bruns, and Carmen Enid Martínez

To be submitted for publication to FEMS Microbiology Ecology 58

ABSTRACT

Agriculturally drained peatlands near the town of Manning in Western New York present new opportunities for studying dissimilation of sulfur compounds. These peatlands, which developed over Silurian Age Lockport Dolomite, are rich in sulfur and metals. For more than 60 years, they have been subjected to seasonal changes in redox conditions, being water saturated in winter and tile-drained in late spring. This study was based on our previous findings that although surface peat soils exhibited sulfate reduction, microbial community DNA extracted from soils or enrichment cultures yielded no PCR amplification products for dissimilatory (bi)sulfite reductase (dsrAB) genes. Our failure to detect these genes, possessed by all recognized groups of dissimilatory sulfate- reducing prokaryotes, was consistent with recovery solely of members of the Firmicutes phylum from enrichment cultures. To extend our understanding of sulfur cycling in these surface peat soils, we investigated the presence of other genes coding for enzymes known to be involved in dissimilatory and assimilatory sulfate reduction. We used PCR amplification to detect the presence of genes encoding adenosine-5’-phosphosulfate reductase (apsA) in community DNA extracted from peat soils. Additional primers were developed for genes encoding asrA, the anaerobic sulfite reductase subunit A genes which mediate dissimilatory sulfur reduction in Salmonella typhimurium, Clostridium spp., Thermoplasma acidophilus, and Thermococcus litoralis. Soil enrichments were prepared with media targeting diverse groups of sulfate reducers and Clostridium spp., and DNA was extracted from positive enrichment cultures. PCR amplification of apsA

59 indicated presence of these genes in all soils tested. Phylogenetic associations of cloned sequences showed that apsA sequences had diverged from those of known, cultured

SRPs. Although asrA genes were not detected in direct soil DNA extracts, they were detected in enrichments from all soils tested and in genomic DNA of one Clostridium sp. isolated from one of the soils. Phylogenetic analysis of asrA sequences revealed an extensive distribution and diversity of this gene. Our results suggest that indigenous populations in these organic soils have functional genes for sulfate reduction that differ from those of well-known SRPs or they might use alternative pathways for sulfur dissimilation.

INTRODUCTION

Peat soils developed over the metal-rich Lockport Dolomite formation of Western

New York contain high levels of sulfur, zinc and cadmium (7, 28, 29, 44). In the 1940’s, not long after these peatlands were drained for agricultural purposes, peat soils sampled near the town of Manning, NY, were found to be phytotoxic to crops (41). At that time, high concentrations of Zn (exceeding 500 mg kg-1, dry weight basis) were measured in

plant shoots. Recent studies have demonstrated the persistence of severe phytotoxicity

and high concentrations of S, Zn, and Cd in the peats (29, 44). Martínez et al. (29)

demonstrated that 35-45% of the total sulfur in these soils was present in the most

reduced electronic oxidation states such as sulfides (R-S-R and metal sulfides) and thiols

(R-S-H). Although Yáñez et al. (44) demonstrated sulfate-reducing activity in oxic soils

sampled from the surface of an agricultural field, zinc sulfides (ZnS) appeared to have

been weathered out from these soils during the 60 years following drainage (28, 29).

Despite sulfate-reducing activity observed in enrichment cultures from these soils,

microbial community DNA extracted from soils or enrichments yielded no PCR

amplification products for dissimilatory (bi)sulfite reductase (dsrAB) genes, although these genes are found in all recognized groups of sulfate-reducing prokaryotes. Using molecular and cultural approaches, the authors provided evidence that sulfur cycling in these metalliferous soils was mainly driven by members of the Firmicutes phylum. The authors proposed that these oxic soils represent a novel environment for investigating

61 microbially mediated reduction of sulfur that may involve oxidized forms of S such as sulfonates.

Sulfate respiration represents one of the most ancient metabolic pathways. Isotope fractionation studies indicate that sulfate-reducing activity dates back through the

Proterozoic and into the Archaean (6). Dissimilatory sulfate reduction is carried out by phylogenetically diverse prokaryotes (1, 33, 35), many of which are deeply rooted in the universal tree of life (40, 43). Sulfate reduction is recognized as occurring intracellularly, requiring that sulfate be transported into the cell by an energy-requiring but reversible process (9). In the cytoplasm, sulfate is converted to adenosine-phosphosulfate (APS).

Dissimilatory sulfate reduction involves two enzymes present in all known sulfate- and sulfite-reducing prokaryotes (SRPs) to date (17, 24, 43). Adenosine-5’-phosphosulfate reductase (EC 1.8.99.2) catalyzes the reduction of APS to sulfite. Dissimilatory sulfite reductase (EC 1.8.99.3) catalyzes the six-electron reduction of sulfite to sulfide, which is the central energy-conserving step in both sulfite- and sulfate-reducing prokaryotes (31,

35). The genes coding for adenosine-5’-phosphosulfate (apsA) and dissimilatory sulfite reductase (dsrAB) have been used to infer evolutionary history of anaerobic sulfate

(sulfite) respiration (21, 25, 26, 43) and diversity of uncultured SRP populations (3, 8, 11,

13, 27, 30, 32, 34, 39). It has been demonstrated that these genes are subjected to lateral gene transfer (17, 25, 46). Comparisons of phylogenetic trees for DsrAB and ApsA have suggested the occurrence of frequent lateral gene transfers between Gram-positive and thermophilic SRPs (17, 25, 46).

Dissimilation of sulfur compounds for energy has also been described in non- sulfate reducers (4). Anaerobic sulfite reductases with dissimilatory activities have been

62 found in Salmonella typhimurium and some Clostridium species (18, 19, 23). These enzymes, together with the dissimilatory and assimilatory sulfate reductases, have a common ancestor and their functional diversification is proposed to have preceded the divergence of Bacteria and Archaea (12). These enzymes are trimers formed by a ferredoxin protein (asrA gene), a flavoprotein (asrB gene) and a siroheme-binding protein (asrC gene) (23). In the genomes of Thermoplasma acidophilum and

Thermococcus litoralis, both sulfur-reducing thermophilic archaea, dsrAB genes have not been found (36, 37). Instead, genes encoding for proteins similar to AsrA and AsrB of

Salmonella spp. are present (36, 37). These findings suggest that other prokaryotes may have this pathway for sulfur dissimilation and that their roles in sulfur cycling may not be fully appreciated.

In this study, we investigated the diversity of genes encoding enzymes known to be involved in dissimilatory sulfate and sulfite reduction in surface soils from the

Manning peatlands. Our approach was based on detection by PCR amplification of dsrAB, apsA, and asrA genes in unenriched and enriched peat soils. We designed oligonucleotide primers for the detection of asrA genes and determined the phylogenetic associations of selected clones for each gene tested. The data contribute to a better understanding of biological sulfur cycling in these sulfur- and metal-rich surface peat soils.

63 MATERIALS AND METHODS

Peat samples. Peat soils in the Manning, New York, region are classified as typic haplosaprists of the Carlisle Series (7, 20). Surface soil samples (0 - 5 cm) were collected by soil coring with an 8-cm-wide auger from a field planted with onion (78º6.146'W,

43º9.470'N) in July 2002. The surface soils were collected along a gradient starting in an area where there was no plant growth (soil labeled “Peat 1”) and moving eastwards for about 8 meters to a zone of typical plant growth (soil labeled “Peat 8”). The soil samples were kept at 4ºC prior to analysis. Elemental composition, organic matter content, and other chemical properties of these soils are reported in Chapter 2 (Table 2-1).

Enrichment cultures. To assess the presence of classical SRPs in these peats samples, enrichments targeting different groups of sulfate-reducing bacteria were prepared from Peats 1, 2, and 8. Four different culture media were used: Medium 63 containing lactate for Desulfovibrio spp., Medium 193 containing acetate for

Desulfobacter postgatei, Medium 194 containing propionate for Desulfobulbus spp., and

Medium 63a containing pyruvate and acetate for Desulfotomaculum spp. (Deutsche

Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany, http://www.dsmz.de). Slurries were prepared by resuspending 10 g of soil in 95 mL 0.1% sodium pyrophosphate with 1 g of sterile, 1-mm, acid-washed glass beads. The slurries were vigorously mixed by hand and then placed on a shaker at 200 rpm for 20 minutes

(45). Five mL of the slurry were used as inoculum for each medium. Media without soil inocula and with heat-treated soil (30 min, 120°C) were used as controls. Enrichments were incubated in the dark at 25ºC for two months or until growth was observed. An

64 additional medium (Reinforced Clostridial Medium, BD Diagnostic Systems) supporting growth of Clostridium spp. was also used.

DNA extraction. Microbial community DNA was extracted from soils and positive enrichment cultures using the MoBio UltraClean™ Soil DNA kit and MoBio

UltraClean™ Microbial DNA Kit (Solano Beach, CA), respectively. DNA was visualized by electrophoresis in ethidium-bromide-stained 1% agarose gels in Tris-acetate-EDTA buffer. DNA purity and concentrations were estimated by spectrophotometry measuring the continuous absorbance from 230 to 300 nm (5).

PCR amplification of dsrAB and apsA. Amplification of dsrAB genes was performed using the primers (DSR1F and DSR4R) and conditions described previously

(43, 44). An ~390-bp fragment of apsA was amplified by PCR using primers described by Friedrich (17). Forward (APS-FW 5’-TGGCAGATMATGATYMACGG-3’) and reverse (APS-RV 5’- GGGCCGTAACCGTCCTTGAA -3’) primer solutions at 15 pmol each were mix with 20 ng of DNA, 2.5 µl of 10X PCR buffer, 1.25 µl of 10 mg/µl bovine serum albumin, 0.4 µl dNTPs (25 mM each), and 1.5 U of Taq polymerase Eppendorf

HotMaster in final reaction volume of 25 µl. PCR reactions were performed as follows: an initial denaturation step (2 min, 94°C) was followed by 30 cycles of denaturation (30s,

94°C), annealing (45 s, 60°C), and extension (60 s, 72°C) and a final extension step (7 min, 72°C). Genomic DNA from Desulfovibrio vulgaris ATCC 29579 was used as positive control.

Anaerobic sulfite reductase subunit A genes (asrA). Available AsrA protein sequences were retrieved from the PEDANT database (http://pedant.gsf.de) and from the

National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov). These

65 included protein sequences from Clostridium perfringens (NC_003366), Clostridium acetobutylicum (AE001437), Clostridium tetani (AE015927), Clostridium botulinum

ATCC3502, Clostridium difficile (630), Salmonella typhimurium (NC_003197), and

Thermoplasma acidophilum (AL445063). Sequences were aligned using ClustalX (42) and alignment was visually inspected for conserved regions. Areas of ambiguous alignment were not included in the analyses. Protein sequences selected for primer design were manually converted to nucleotide sequences using the degenerate code for amino- acids. The forward primer, asrAF (5’-ATGGGDACNAATMAARCWGA-3’) and the reverse primer asrAR (5’-CCRCANCCNACRCACATATG-3’), hybridize at positions

440 to 459 and 932 to 953 of Salmonella typhi (Fig. 3-3). Heated cell suspensions from pure cultures CYP1-CYP12, which were obtained from peat enrichments in a previous study (43), were used as templates to test the primers. PCR master mix was prepared using 15 pmol of each primer, 20 ng of DNA, 2.5 µl of 10X PCR buffer, 1.25 µl of 10 mg/µl bovine serum albumin, 0.4 µl dNTPs (25 mM each), and 1.5 U of Taq polymerase

Eppendorf HotMaster, combined in a final reaction volume of 25 µl. Amplifications were carried out using an initial denaturation step of 2 min at 94°C followed by 30 cycles of 30 s at 94°C, 30 s annealing at 51 °C and 1 min extension at 72°C.

Cloning and sequencing. PCR products were loaded into low-melting point agarose gels, and DNA bands were excised for purification, ligation into pCR®4-

TOPO®, and transformation into ONE SHOT competent Escherichia coli cells (TA

Cloning System, Invitrogen). For each soil sample exhibiting positive PCR amplification, three clones were randomly selected for sequencing of their inserts with an ABI Hitachi

3730XL DNA Analyzer. The M13f and M13r primers were used to sequence both strands

66 of the inserts. Sequences from clones were edited and assembled using SeqMan™ II software (DNASTAR Inc., Madison, WI).

Phylogenetic analysis. Nucleotide sequences from clones and their closest relatives determined by BLAST (2) were aligned using the program ClustalX (42). The phylogenetic trees were constructed using the neighbor-joining distance method with

Jukes-Cantor correction (38). In order to determine confidence values for individual branches of final tree, bootstrap analysis (100 replications) was applied (15). Maximum likelihood analyses were also done with the dnaml program in PHYLIP (16), which generated trees with comparable topology and bootstrap values.

Nucleotide sequence accession numbers. The sequences obtained in this study are available from the GenBank nucleotide sequence database under accession numbers

DQ995757 to DQ995779 for asrA and DQ995780 to DQ995800 for apsA.

RESULTS AND DISCUSSION

Enrichments. In our previous work sulfate-reducing activity was observed in peats 1, 2 and 8, but 16S rRNA sequences from soil enrichments and cultured isolates were unrelated to classical sulfate-reducing prokaryotes (44). Nevertheless, our data showed that members of the Firmicutes were involved in sulfur transformations in these surface soils. After an incubation of 2 months at 25°C in the dark, only some enrichments yielded growth: Peat 1 (Medium 63 for Desulfovibrio spp.); Peats 2 and 8 (Medium 63 for Desulfovibrio spp and Medium 63a for Desulfotomaculum spp.). No growth was

67 observed in enrichment media 193 (for Desulfobacter spp.) or 194 (for Desulfobulbus spp.) in any of the soils tested, even after 4 and 6 months of incubation.

dsrAB genes and apsA diversity in surface peats. For all eight samples tested, template DNAs extracted directly from soils and used in PCR with apsA primers yielded amplicons of the expected size (~390 bp) (Fig. 3-1). The phylogenetic associations of apsA clones are shown in Figure 3-2. All cloned sequences but one formed a single group separated from other apsA sequences of historically recognized SRP strains. Cloned sequences from a single peat sample did not group together. Overall, these sequences varied sufficiently to generate three clear clusters. Previous results obtained in our laboratory (44) showed that despite the observance of sulfate-reducing activity, a key enzyme in sulfate respiration (43), dsrAB, was not detected by PCR in any of the soils tested. Using DNA extracted from positive SRP enrichments and pure cultures, we failed to amplify dsrAB genes (data not shown). These observations suggest that SRPs present in these soils harbored functional genes distinct from those of classic SRP groups.

Figure 3-2: Neighbor-joining phylogenetic tree showing the associations for apsA genes. Cloned sequences were denoted indicating soil origin (S, 1 to 8) and clone analyzed (CL). Tree was rooted with Archeoglobus fulgidus. The numbers on the branches are the results of 100 bootstrap replicates and indicate the frequency with which the sequences grouped together in the way shown.

70 The use of apsA amplicons as markers for sulfate-reduction has been reported in other studies on detection of uncultured sulfate reducers (11, 39). It has been assumed that this enzyme can perform the reverse reaction in some sulfur oxidizers. But these assumptions have been challenged by the results of Dahl et al. (10), where the apsA- mutant Allochromatium vinosum (formely Chromatium vinosum) showed no significant effect on its ability to oxidize sulfite. Hipp et al. demonstrated that the genes for adenosine-5’-phosphosulfate (apsA) and sirohaem sulfite reductase (dsrAB) from the sulfur-oxidizing phototrophic bacterium A. vinosum are true homologues of their counterparts in SRPs (21). These observations provide evidence for a protogenotic event in the divergence between the oxidative and the reductive modes of dissimilatory sulfur metabolism (21).

asrA gene. An alignment of seven sequences for asrA genes was used to develop

PCR primers (Fig. 3-3). Genomic DNA from Clostridium subterminale ATCC 25774 was used as a positive control for asrA gene detection (14) , and it generated PCR products of expected length (~500 bp) (Fig. 3-4). Amplification products were not obtained from the genomic DNAs of Escherichia coli or Desulfovibrio vulgaris.

Table 3-1: List of bacterial strains used in this study.

Strain Related bacterial sequences a % similarity b Accession N° c CYP1 Desulfotomaculum guttoideum 99 DQ479411 (Y11568) CYP2 Clostridium sp. 98 DQ479412 (AY221993) CYP3 Clostridium sp. 98 DQ479413 (AY532163) CYP4 Clostridium sp. 97 DQ479414 (AY532163) CYP5 Clostridium tunisiense 99 DQ479415 (AY187622) CYP6 Clostridium sp. 98 DQ479416 (AY532163) CYP7 Clostridium mangenotii 96 DQ479417 (M59098) CYP8 Clostridium frigidicarnis 96 DQ479418 (AF069742) CYP9 Desulfotomaculum guttoideum 99 DQ479419 (Y11568) CYP11 Clostridium sp. 98 DQ479420 (AY532163)

72 In testing primers for asrA, no amplification products were obtained with DNA extracted directly from Peats 1, 2, or 8 (data not shown). However, PCR products of appropriate lengths were obtained with DNA from enrichments from all three soils and which were positive for sulfate reduction (Fig. 3-4). Isolates (Table 3-1) previously obtained from these surface soils (44) were analyzed for asrA, dsrAB and reduction of sulfate. After PCR, only one isolate, denoted CYP5 with 16S rRNA sequence 99% similar to that of Clostridium tunisiense, showed an amplicon for asrA (Fig. 3-4). None of the isolates reduced sulfate or gave positive results for dsrAB amplification.

Interestingly, CYP5, but none of the other isolates, also possessed the ability to reduce thiosulfate. These results suggest that the asrA reduction pathway is not distributed among all clostridia and may be restricted to specialized Clostridium strains capable of reducing thiosulfate.

Phylogenetic associations of asrA genes are presented in Figure 3-5. Cloned sequences did not group together based on source soils. However, cloned sequences from enrichments grown in Reinforced Clostridium Medium did group together, indicating that this medium selected for closely related organisms carrying asrA genes. Cloned sequences obtained from the enrichment media selective for Desulfovibrio spp. and for

Desulfotomaculum spp., on the other hand, appeared in different groups within the tree.

The lack of availability of additional asrA sequences in databases prevented the establishment of a clear phylogenetic association for each cloned sequence.

The pathway of the anaerobic sulfite reductase has not been elucidated but it has been demonstrated that it is used for dissimilation in Salmonella typhimurium (22, 23).

Harrison et. al. (19) purified and characterized an inducible sulfite reductase from

73 Clostridium pasteurianum, showing that the sulfite reduction for energy-yielding processes is not restricted to Salmonella spp. Our results indicate the presence of the asrA pathway in additional bacterial groups (Fig. 3-5). The wide distribution of asrA sequences suggests that this enzyme plays an important functional role. The activity of asrA may be an important contributor to terrestrial sulfur dissimilation processes, and its role in the sulfur cycle needs to be clarified.

74 Cont.

Figure 3-3: DNA sequence alignment used for the design of primers for asrA genes. Sequences were aligned using Clustal X and 75 screened by visual inspection. Conserved region (black boxes) served as template for design of degenerated primers.

78 CONCLUSIONS

Surface soils from the Manning peatlands represent a novel environment for studying microbial reduction of sulfur compounds. These peats are subjected to drastic changes in redox conditions and differ from permanently anoxic environments recognized as SRP habitats. Prokaryotes in these soils possess functional genes that differ from those found in historically recognized SRPs from marine sediments, rice paddies, and high-temperature environments. New sequences for apsA genes were detected in these peats. Moreover, alternative pathways for reduction of sulfite involving asrA genes appear to be important in these surface peat soils populated by Firmicutes members. Our results suggest that indigenous populations in these peat soils might use alternative pathways for sulfur dissimilation. Additional efforts should be made to demonstrate the prevalence and importance of asrA genes in the terrestrial sulfur cycle.

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Chapter 4

Sulfate-Reducing Bacteria Populations in Sulfur-Rich Peat Soils as a Function of Soil Depth and Moisture Content

ABSTRACT

Peat deposits that overlie the mineral bed of Lockport Dolomite in Western New

York contain elevated concentrations of some metals (Zn and Cd) and sulfur. In this work, we describe the composition of bacterial communities, enzymes involved in sulfate respiration, and sulfur redox species as a function of soil depth and moisture content of drained agricultural peats. Three intact peat cores were collected during dry

(July 2002) and wet (March 2004) seasons along a Zn phytotoxicity gradient, starting in an area where there was no plant growth and moving eastwards to an area of typical growth. The undisturbed soil columns (6 total) were kept at 4ºC until analyses when they were cut open and sectioned into 5 cm intervals. DNA was extracted from selected sub- samples (surface, deep and bottom soils) and ribosomal intergenic spacer analysis (RISA) was used to study bacterial communities. The dissimilatory sulfite reductase gene

(dsrAB), a key enzyme in sulfate respiration, was amplified by PCR to assess the presence of sulfate-reducing prokaryotes (SRPs). Bacterial communities differed among the soil cores, between seasons, and with depth. The dsrAB genes were detected only in sub-samples deeper than 45 cm in both dry and wet seasons. Furthermore, S-XANES analyses show that 40-80% of the total sulfur in these peats exists in reduced forms (e.g., sulfides and/or thiol groups) while 3-20% exists in oxidized forms (e.g., sulfate). Clone libraries demonstrated that novel sequences of dsrAB genes are present in soil samples with highly elevated levels of S and Zn. Our results suggest the presence of unrecognized

SRPs that may be specific to these metalliferous soil environments. The distribution and

89 activity patterns of SRPs may be relevant to understanding biogeochemical processes occurring in these metal- and sulfur- rich peat soils.

INTRODUCTION

Peat soils developed over the metal-rich Lockport Dolomite formation of Western

New York State contain high levels of S, Zn and Cd (4, 22). In the peatlands of this region, referred to as the Manning peatlands, phytotoxicity in vegetable crops was observed in the early 1940s (33), not long after the bogs were drained for agricultural purposes, and very high concentrations of Zn (exceeding 500 mg/kg, dry weight basis) were measured in plant shoots (33). More recently, Martínez et al. (22) demonstrated that very high total soil concentrations of Zn, Cd and S and severe phytotoxicity have persisted for decades since the initial studies (4, 33). More than 60 years of artificial

(seasonal) drainage have subjected these soils to drastic redox fluctuations. In spite of periods of prolonged oxic conditions, spectroscopic analysis using S-XANES (sulfur-X- ray absorption near edge structure) spectroscopy showed that 35-45% of the total sulfur was still present in the most reduced electronic oxidation states such as sulfides (R-S-R and metal sulfides) and thiols (R-S-H). Analysis of sulfate-reducing prokaryotes (SRPs) in surface soils from this region revealed that members of the phylum Firmicutes rather than “classical” SRPs might be responsible for driving biologically-mediated reactions in the sulfur cycle (42). This evidence suggests that peat soils from the Manning region represent a novel environment for SRPs. Further analyses of spatial and seasonal variations in sulfate reduction in these sulfur-rich soils are necessary to enhance our understanding of terrestrial sulfur cycling in these soils and its relation to metal mobility and bioavailability.

91 Sulfur-XANES has been used to identify and quantify sulfur species in bulk soils

(12, 25) and in humic substances extracted from marine sediments (39) and soils (24, 32,

41). These studies have demonstrated the predominance of reduced (thiol/sulfide) and oxidized forms of sulfur. However, oxidized forms of sulfur with an electronic oxidation state of +5 (sulfonates) are generally higher than sulfur species with an electronic oxidation state of +6 (sulfates). While marine sediments are characterized by high concentrations of iron, peat soils generally contain much lower Fe concentrations, ranging from 0.05 to 0.99 % (8, 29, 34). These chemical characteristics make pyrite the most common authigenic sulfide mineral in marine and lacustrine sediments but this would not be the case in peat soils.

Dissimilatory microbial sulfate reduction is an important metabolic activity in many reduced environments and it is an essential step in the sulfur cycle (9). This reaction is exclusively mediated by a group of diverse microorganisms, known as sulfate reducing prokaryotes (SRPs). SRPs are found in five bacterial and two archaeal phyla (5,

26). Sulfate reduction by SRPs occur in reducing environments where redox potential is below about -100 mV and sulfate is available (35). The end product of dissimilatory sulfate reduction is hydrogen sulfide (H2S). In the presence of sufficiently high

concentrations of some metals (chalcophiles such as Ag, Cd, Hg, Zn) the precipitation of

metal sulfides is predicted to occur. It is for this reason that SRPs may have an important

role in the coupled biogeochemical cycling of metals and sulfur. For example, sphalerite

(ZnS) formation at low-temperature has been reported to occur in natural biofilms by

aerotolerant SRB (15) thus demonstrating that microorganisms control metal

concentrations in groundwater. Although Zn and Cd are not redox sensitive elements, the

92 solid phases that can potentially retain Zn and Cd in the soils under investigation (e.g., Zn and Cd sulfides and Zn and Cd complexed by organic matter) respond to changes in redox status and microbial activity, which can influence their speciation and solubility.

Soil cores collected from the Manning peatland region of Western New York were used in this study. The aim of our research was to determine the influence of water content (redox status) in microbial populations and functionality and hence the bioavailability and mobility of Zn and Cd in these soils. We combined several approaches, including ribosomal intergenic spacer analysis (RISA), dsrAB detection (key enzyme in sulfate respiration), and the determination of S oxidation states to evaluate microbial community composition and sulfate reduction. The results contribute to increased understanding of sulfur cycling in these metalliferous sulfur-rich environments and to the identification of the key players mediating sulfate–reduction reactions in soils.

MATERIALS AND METHODS

Sampling of soil cores and chemical analyses. Intact soil cores were colleted in

Western New York in July 2002 (dry season) and March 2004 (wet season) from an agricultural field planted with onion in the Manning peatland region. The soil cores were collected along a gradient (visual inspection) starting in an area (approximately

78º6.146'W, 43º9.470'N) where there was no plant growth (first core, labeled “High Zn”) and moving eastwards for about 10 meters (second core, labeled “Medium Zn”), and for about 15 additional meters (third core, labeled “Low Zn” ). The soil cores labeled “High

Zn”, “Medium Zn” and “Low Zn” reflect the relative concentration of Zn in surface

93 peats. Three soil columns were pulled from the soil each season, for a total of six soil columns. The undisturbed soil cores were brought to the laboratory where they were kept at 4ºC until they were cut open and sectioned (5 cm intervals) for microbiological, chemical and spectroscopic analyses. The gravimetric moisture content was determined after drying the soils at 50°C for about 16 hours. Total elemental composition of bulk soils (sub-samples from 5-cm sections) was determined by ICP-AES analysis of

HClO4/HNO3/H2SO4 digests (acid ashing method) using 2 g of air-dried soil. Total sulfur

in the bulk soils was determined by the high-temperature furnace combustion method with infrared absorption and soil organic matter content was determined by weight loss on ignition. Soil pH was measured in 2:1 solution (10 mL 0.01M KNO3) to soil (5 g)

suspensions using a pH electrode. Soil properties are presented in Figure 4-1 and

Tables 4-1 and 4-2.

While chemical analyses were performed in all 5-cm sections from soil cores,

spectroscopic and microbiological analyses were performed at three different depths. The

5-cm sections that underwent spectroscopic and microbiological analyses are labeled

“surface”, “deep”, and “bottom” soils. The surface, deep, and bottom soils are indicated

with an open circle in Figure 4-1 and the soil depth and sample ID are detailed in

Tables 4-1 and 4-2. Deep soils occurred at intermediate depths of intact soil cores and have the highest concentration of Zn. Bottom soils represent the deepest peat sample before reaching the underling marl, and surface soils are the 0-5cm section of the soil core.

Sulfur X-ray Absorption Near Edge Structure Spectroscopy (S-XANES). The oxidation states in which sulfur (organic and inorganic forms) is present in our soil

94 samples was determined using X-ray absorption near edge structure (XANES) spectroscopy. The sulfur K-edge (2472 eV) XANES spectra were collected at Beamline

X-19A of the National Synchrotron Light Source (NSLS), Brookhaven National

Laboratory under standard operating conditions. Sample preparation and analysis have been described elsewhere (22, 42).

DNA extraction. DNA was extracted from peat soils (~0.5 g) with MoBio

PowerSoil™ DNA Isolation Kit (Solano Beach, California) according to the manufacturer’s instructions. DNA was visualized by electrophoresis in a 1% agarose gel in Tris-acetate-EDTA buffer. DNA concentration was estimated spectrophotometrically by measuring the continuous absorbance from 230 to 300 nm (3).

RISA. Soil microbial communities were studied by amplifying the intergenic spacer region between the 16S and 23S ribosomal RNA genes. The bacterial primers used were 16S-1406f (5'-TGYACACACCGCCCGT-3') and 23S-115r (5'-

GGGTTBCCCCATTCGG-3') and have been previously described (16). For PCR amplification, 15 pmol of each primer solution, 20 ng of DNA, 2.5 µl of 10X PCR buffer,

1.25 µl of 10 mg/µl bovine serum albumine, 0.4 µl dNTPs (25 mM each), and 1.5 U of

Taq polymerase Eppendorf HotMaster, were combined in a final reaction volume of 25

µl. PCR reactions were setup in triplicates. After an initial denaturation step of 2 min at

94°C, amplification was carried out for 25 cycles, with each cycle consisting of 30 s at

94°C, 30 s at 55°C, and 1 min at 72°C. PCR products were visualized in a 1% agarose gel stained with ethidium bromide and triplicate reaction mixtures were pooled prior to fingerprint analysis. RISA fingerprints were visualized by separation of the PCR-

95 amplified DNA fragments using 2.5% NuSieve GTG agarose and then stained with ethidium bromide.

PCR amplification of dsrAB genes. Primers used for the amplification of dsrAB genes were described by Wagner et al. (40). An approximately 1.9-kb dsrAB fragment was amplified from peat DNA using the degenerate primers DSR1Fmix (equimolar mixture of DSR1F, DSR1Fa, DSR1Fb, DSR1Fc, and DSR1Fd 10 pmol/µL each) and

DSR4Rmix (equimolar mixture of DSR4R, DSR4Ra, DSR4Rb, DSR4Rc, DSR4Rd and

DSR4Re 10 pmol/µL each). Amplification of genes was performed with 20 ng of DNA,

15 pmol of each primer mix solution, 2.5 µL of 10X PCR buffer, 1.25 µL of 10 mg/µL bovine serum albumine, 1 µL dNTPS (10 mM each), and 1.5 U of Taq (Eppendorf Hot

Master) combined in a final volume of 25 µL. Thermal cycling was carried out by using an initial denaturation step at 94°C for 2 min, followed by 30 cycles of denaturation at

94°C for 40 s, annealing at 54°C for 40 s, and elongation at 72°C for 90 s. The cycling was completed by a final elongation step at 72°C for 10 min. Positive (Desulfovibrio vulgaris ATCC 29579) controls were included in all PCR amplifications. PCR products were visualized by agarose (1%) gel electrophoresis. PCR were setup in duplicates and positive reactions were pooled for further analyses.

Cloning of dsrAB genes and RFLP analysis. Prior to cloning, the PCR amplification products were purified by low-melting-point agarose (1.5%) gel electrophoresis and the bands of the expected size (~ 1.9 kb) were excised from the gel.

The purified DNA was ligated into pCR®4-TOPO® and transformed into ONE SHOT competent Escherichia coli cells as recommended by the manufacturer (TA Cloning

System, Invitrogen). Clones were screened for the correct insert by PCR reactions using

96 M13F (5′-GTAAAACGACGGCCAG) and M13R (5′-CAGGAAACAGCTATGAC) primers. Amplified clones with a 1.9-Kb insert were digested using the restriction endonuclease HhaI (5 ′- G CG▼C) (5U). Digestions were performed in 2 µl 10× RE buffer C, 0.2 µl acetylated BSA (10 µg/µl), 5 µl DNA, and sterile deionized water for a final volume of 20 µl. After incubation for 1.5 h at 37°C, the digested DNA fragments were separated by 2.5% agarose gel electrophoresis.

Sequencing and phylogenetic analysis. Cloned DNA with unique digestion patterns were selected for sequencing. The T3 and T7 primers were used to sequence both strands of the insert. Sequences were edited to remove any remaining vector fragment and were assembled using SeqMan™ II software (DNASTAR Inc., Madison,

WI). To construct phylogenetic trees based on the nucleotide alignments, sequences were aligned using Clustal X version 1.81 (36) and the neighbor-joining distance method with

Jukes-Cantor correction (28). The bootstrap confidence levels were defined from 100 iterations of tree reconstruction (10). Maximum likelihood analyses were also done using the programs included in the PHYLIP package (11). The topology and bootstrap values were similar to those obtained with neighbor-joining analysis.

Nucleotide sequence accession numbers. The sequences determined in this study have been deposited in the GenBank database under accession numbers DQ855242 to DQ855261.

Confocal Scanning Laser Microscopy (CSLM). Undisturbed peat particles were placed on Lab-Tek™ chambered coverglass (Nalge Nunc International, Rochester, NY) and embedded in 1% agarose. The embedded peat particles were visualized using a blue laser (488 nm) to detect autofluorescence produced by mineral particles (ZnS) within the

97 peat. Peat particles were subsequently stained using LIVE/DEAD® BacLight™

(Invitrogen Corporation, Eugene, OR), and after reaction for 15 minutes the particles were examined directly with an Olympus FluoView 300 confocal scanning laser microscope. Samples were examined using three excitation laser lines: red (633 nm), blue

(488 nm), and green (543 nm). Images were viewed with an UplanFL 40X objective

(numerical aperture 0.75). The z direction images (representing particle thickness) were acquired at 1 µm intervals within the peat particles. Images (stained and not stained) were processed and analyzed using the FluoView software.

RESULTS

Chemical characteristics and sulfur speciation in soil cores. Unusually high concentrations of sulfur (4,200 to 40,500 mg kg-1), zinc (564 to 71,175 mg kg-1) and cadmium (<1 to 310.7 mg kg-1) were present in the soil cores (Table 4-1, Table 4-2,

Fig. 4-1). Zinc and sulfur concentrations and soil pH, however, varied with depth in soil cores collected in both dry and wet seasons (Fig. 4-1). Total concentrations of Zn and S generally increase with depth in the soil profiles until the marl layer is reached, where Zn and S concentrations decrease in this mineral layer underlying the peat (Fig. 4-1). While soil Zn concentrations at 0 to ~30 cm depth decreased from the “High Zn” (center of the highly phytotoxic area) to the “Low Zn” (typical growth area) sampling sites, soil S concentrations remained relatively unchanged (Fig. 4-1). This results in much lower Zn/S ratios in the “Low Zn” area, which is a potentially important factor limiting Zn toxicity

(most of the Zn present could be bonded to sulfide). Soil pH typically increased with

98 depth and ranged from 4.8 to 5.5 in surface (0-30 cm) soils and from 5.5 to 6.9 in subsoils. Although most soils were very high in organic matter content (average 71%), several soils (particularly the bottom soils) exhibited lower values, likely a result of physical mixing of the peat with the underlying marl (Table 4-1 and Table 4-2). In general, the results presented in Figure 4-1 suggest that after more than 60 years of exposure to variable and prolonged oxidation/reduction conditions, aerobic decomposition of organic matter, which is likely accompanied by the oxidation of sulfides to sulfates, metal (Zn) release and pH reduction has occurred in the surface soils of the Manning peatland region of Western New York. It is worth mentioning that the highest Zn concentrations are accompanied by the highest S concentrations (samples labeled W8 and D14 in Table 4-1 and Table 4-2). Furthermore, ZnS had been identified in these soils (unpublished data) which suggest the re-immobilization of Zn in deep soils with high water content, presumably by microbial activity.

0 10 High Zn Dry Season 20 Medium Zn Low Zn 30 40 50 60 Depth (cm) Depth 70 80 90 0 0 0 0 0 00 00 00 0 0 0 0 10000 20000 30000 40000 5.0 5.5 6.0 6.5 7.0 0 0 0 0 0 0 0 3 6 9 0 12000 20 40 60 8 Total Zn (mg kg-1) Total S (mg kg-1) Soil pH

0 10 Wet Season 20 30 40 50 60 Depth (cm) 70 80 90 0 0 0 0 10000 20000 30000 40000 5.0 5.5 6.0 6.5 7.0 00 0 3000 6 9000 2000 5 0000 0000 0000 1 1 2 4 6 80000 Total Zn (mg kg-1) Total S (mg kg-1) Soil pH

Figure 4-1: Variations in Zn and Cd concentrations and pH values in soil cores (high, medium and low zinc) collected during dry (top) and wet (bottom) seasons. 99

Table 4-1: Elemental concentrations (dry weight basis), soil organic matter (SOM), water content, and pH of peat samples collected during dry season (July 2002).

Columns Depth and Sample pH SOM Water Total S Total Fe Total Zn Total Cd ID (%) content (mg kg-1) (mg kg-1) (mg kg-1) (mg kg-1) (%) Surface (0-5 cm) 5.3 81.6 59.1 11100 6850 9150 21.25 D30 High Zn Deep (40-45 cm) 5.4 81.7 183.4 22400 6400 32450 70 D38 Bottom (65-70 cm) 6.4 67.2 261.5 11800 13500 12900 42.5 D43 Surface (0-5 cm) 5.3 82.6 55.4 11300 9025 2650 4.25 D18 Medium Zn Deep (40-45 cm) 5.5 82.1 319.1 11000 5200 37900 90 D26 Bottom (50-57 cm) 5.6 33.9 111.4 8300 14725 9800 24.75 D28 Surface (0–5 cm) 4.8 83.2 54.9 11900 8275 725 1.75 D1 Low Zn Deep (65-70 cm) 6.0 76.0 300.7 39800 11800 71175 152.5 D14 Bottom (75-80 cm) 5.2 85.2 55.2 10200 18000 11025 62.5 D16

100

Table 4-2: Elemental concentrations (dry weight basis), soil organic matter (SOM), water content, and pH of peat samples collected during wet season (March 2004).

Columns Depth and Sample pH SOM Water Total S Total Fe Total Zn Total Cd ID (%) Content (mg kg-1) (mg kg-1) (mg kg-1) (mg kg-1) (%) Surface (0–5 cm) 5.3 81.8 39.0 9700 10092 6312.3 20.2 W1 High Zn Deep (35-40 cm) 5.8 75.3 160.2 40500 4455 41196.9 310.7 W8 Bottom (55-60 cm) 6.9 79.3 186.5 13600 9318 4765.5 48.3 W12 Surface (0-5 cm) 5.1 50.1 38.9 10000 19682 1657.2 3.7 W13 Medium Zn Deep (40-45 cm) 5.8 73.9 134.6 23300 15721 8974.5 85.6 W21 Bottom (50-55 cm) 5.8 50.3 168.8 18600 7314 8669.8 217.1 W23 Surface (0-5 cm) 5.2 82.7 32.2 8500 9236 564.0 bdl W24 Low Zn Deep (40-45 cm) 5.4 80.2 260.6 26600 6789 6203.4 6.9 W32 Bottom (65-70 cm) 5.7 31.1 43.5 4200 9162 2536.6 16.2 W37

bdl= below detection limit

101

102 Sulfur-XANES results indicated that sulfur exists in a wide range of oxidation states in these soils. Figure 4-2 presents the S-XANES results for surface, deep, and bottom soils from soil cores collected during dry and wet seasons. The most striking result was that the highest percentage (60 to 86%) of sulfur was in reduced species (such as sulfides and thiols) in deep soils where the water content was generally very high

(Figure 4-2). Deep soils also contained the lowest percentages of oxidized sulfur species

(3 to 10% sulfonates and 3 to 7% sulfates). Although containing lower water content than all deep soils and most bottom soils (Table 4-1 and Table 4-2), sulfur in surface soils included 50 to 57 % reduced sulfur species (Figure 4-2). The percent S in the +5 oxidation state (as in sulfonates) in surface peats ranged from 8 to 13% while 13 to 16% occurs in the most oxidized, +6, oxidation state (as in sulfates). Again, samples D-14 and

W-8 showed the most extreme results with more than ¾ of the total sulfur present in the most reduced chemical forms.

Dry Season

High Zn Medium Zn Low Zn sulfide/thiol sulfonate sulfate Surface

Deep

Bottom

0 204060800 204060800 20406080 % of S in various oxidation states

Wet Season

High Zn Medium Zn Low Zn

Surface

Deep

Bottom

0 204060800 204060800 20406080 103 % of S in various oxidation states

Figure 4-2: Percentage of total sulfur at estimated electronic oxidation state. Most reduced (sulfides and thiols), +5 and +6 oxidation states are represented. 104 RISA and dsrAB gene detection. The RISA fingerprints obtained using bacterial rDNA primers suggested high microbial diversity in the peats (Fig. 4-3). Many bands were observed in the size range 500 to 1300 bp. Some of the bands were present at all depths (surface, deep, and bottom soils) in a single core, however, those bands were not detected in all cores. For example, the 2 bands forming a “600-650 bp doublet” were characteristic of the Medium Zn core at all depths and seasons while the 900-950-bp bands appeared only in the Low Zn core during wet season. Variations in the composition of the bacterial community seemed to be affected by the moisture content (different fingerprint patterns in dry season compare to wet season; band pattern variations with depth) and by the total concentration in metals (differences in fingerprints between the three cores in each season). Other environmental factors, such as pH and temperature, may also influence the prevalence and activity of some bacterial populations. Overall, the results suggest that microbial composition of these soils is very dynamic.

Sulfate-reducing prokaryotes constitute a heterogeneous group, including members of several phyla and domains, preventing the use of the 16S rRNA gene as a molecular marker (5, 26). However, the use of dsrAB genes encoding the dissimilatory sulfite reductase, a key enzyme in dissimilatory sulfate reduction, has shown to be an appropriate marker (13, 14, 40). The detection of dsrAB was in close relation to the depth of the sample analyzed. An expected c.a. 1.9 kb amplicon was observed in deep and bottom soils (samples deeper than 45 cm) (Fig. 4-3). No amplification of dsrAB was obtained when DNA from surface soils was used.

105 Figure 4-3: Polymerase chain reaction analysis in soil DNA extracted from soil cores from dry and wet seasons. Samples are separated per season and per soil cores (high, medium and low zinc). Soil section analyzed are indicated as surface (S), deep (D) and bottom (B). A) RISA from peat soils obtained with primers specific for Bacteria. B) Detection of dsrAB genes. Expected size for PCR product is indicated. Products for dsrAB genes selected for clone libraries are marked (*). 106 dsrAB diversity in deep peat soils. Two deep peat samples, D-14 and W-8, were selected for the construction of clone libraries for dsrAB genes. The chemical data for samples D-14 and W-8 are presented in Table 4-1 and Table 4-2, respectively. Clone libraries were designated D and W for dry (D-14) and wet (W-8) season samples, respectively. Ninety-six clones were randomly collected for each library. Among these, only 84 for D-14 and 75 for W-8 had an insert of the expected size (1.9 Kb). After restriction fragment length polymorphism (RFLP) analysis, clones were grouped in 7

RFLP patterns for D-14 and 13 for W-8. A total of 27 clones represented by the 20 RFLP patterns were sequenced. After BLAST analysis (1), all clones from dry season and 58 from wet season contained dsrAB sequences.

The phylogenetic affiliations of dsrAB nucleotide sequences are shown in

Figure 4-4. Sequences grouped in two clusters, one of which consisted exclusively of sequences from wet season samples. None of the clusters is closely related to any cultured sulfate-reducing prokaryote lineage, forming deeply branching evolutionary lines of descent. All dsrAB nucleotides sequences with ≥97% similarity were grouped into an operational taxonomic unit (OTU). A total of 10 OTUs were produced for both libraries. One OTU (OTU 10) was present in both peat samples. Eight OTUs contained sequences from the W-8 sample and only one OTU (OTU 8) contained sequences from the dry-season (D-14) sample. Clones in OTUs 1, 2, 3, 4, 5, and 10 were related to clones obtained from acidic fens (21, 30). The remaining OTUs related to clones retrieved from acidic fens (21) as well as to clones derived from a uranium mill tailing site (6).

107

Figure 4-4: Phylogenetic tree showing the affiliation of dsrAB clones from peat samples W-8 and D-14. Tree was created after comparisons of dsrAB nucleotide sequences (sequences longer than 1700 bp with exception of AY015594 and AY015596; accession numbers in parenthesis) using neighbor-joining analysis. The dsrAB tree was rooted with Thermodesulfovibrio islandicus and bootstrap values were calculated after 100 resamplings. Bar corresponds to 10 nucleotide substitutions per 100 sequences positions.

108 CSLM. Confocal scanning laser microscopy showed that bacteria are present throughout peat soil particles (Fig. 4-5). However, bacteria are concentrated in some areas rather than being homogenously distributed within soil particles. ZnS autofluorescence and bacteria are co-localized in some regions (Fig. 4-5).

109 DISCUSSION

Peatlands from the Manning region represent a novel environment for the study of sulfur transformations. These soils have been subjected to drastic changes in redox conditions since they were reclaimed for agricultural purposes. Seasonal drainage likely results in organic matter decomposition, oxidation of sulfide to sulfate and pH reduction.

Sulfur speciation data showed that a large amount of reduced sulfur is still present in soil profiles in both seasons. The relative proportion of total S that is reduced increased with depth, with higher concentrations in samples from the deep sections (up to 80%). These results differ from previously published data (22, 42), where up to 60% of the total sulfur in surface peat soils was present in reduced forms. The difference in reduced sulfur content might be explained by oxidation of sulfur in surface peats. Variations in soil pH indicated that acidification has occurred in the surface layer of the profiles, confirming the oxidation of organic matter.

Prevalence of low redox and saturation conditions contribute to the selection of microorganism capable of carrying out anaerobic respiration. Results for detection of dsrAB genes showed that SRP populations are selective for locations where anoxic conditions prevail. Our observations compare favorably to previously reported results where no dsrAB genes were amplified in either unenriched (32) or enriched (31) surface soils collected during dry season from a different location within the Manning peatland region. Higher concentrations in reduced forms of sulfur were accompanied with high concentrations in Zn, Cd and S. The concentrations in the surface layer (down to 30 cm) in every column were constant for every parameter measured. We assumed that these

110 results reflect mixture of soils produced by plowing. Heavy metals seemed to be concentrated in deep samples presumably by downward movement due to oxidation of organic matter and metal sulfide forms initially present in surface soils. The solubility of metals may be controlled by sulfide under reducing conditions. Depending on the redox potential of the soil, Zn, Cd and Fe sulfides can be formed. However, based on the solubility products for CdS (-log KSO 36.1), ZnS (-log KSO 27.0) and FeS (-log KSO 18.1), and on the fact that these peat soils contain low concentrations of Fe (Table 4-1 and

Table 4-2), FeS is not likely to be present in these soils (23). Moreover, SEM and spectroscopic (EXAFS) analyses of deep peats (D-14 and W-8) have shown that ZnS and

CdS, and not FeS, are present in these peat soils (unpublished data).

Yáñez et al. (42) reported that sulfonates comprised an important fraction of the total sulfur in surface peat soils of this region. The authors suggested that these sulfur forms could be important substrates in biological sulfur cycling. In deep soils, relative concentrations of sulfonates and sulfates are lower than those found in the surface. These results indicate that microbiological populations inhabiting these deep peats might be responsible of the decrease in concentration of these oxidized sulfur forms. The ability of certain sulfonates to serve as terminal electron acceptors for anaerobic respiratory growth of various classic sulfate-reducing bacteria had been reported (18-20). In laboratory incubations, cells adapted to grow on sulfonate continued to utilize sulfonate when equimolar amounts of sulfate were introduced (20). In contrast, cells grown on sulfate did not use sulfonates and continued to consume sulfate (20). These observations suggest that sulfur reduction reactions in these metalliferous soils might be conducted by specialized groups of microorganisms dissimilating sulfonates and/or sulfates.

111 RISA has been successfully used to compare microbial diversity in soil (2, 27), overcoming the problems of bias which occur with microbial culturability from natural environments. Soils are complex environments, containing more than 109 bacterial cells and possibly thousands of different species in one gram (37, 38). Previous results obtained in our group have demonstrated that dominant microbial populations in surface peat soils were similar (42). In contrast, bacterial communities in deep peat soils seemed very different. RISA fingerprints showed the bacterial populations inhabiting these peats are very dynamic. Changes in RISA profiles were observed along soil depth and between soil cores thus suggesting water and metal (Zn and/or Cd) content may influence the bacterial populations present in these soils. Further studies are necessary to elucidate the phylogenetic affiliation of dominant and stable microorganisms in soil cores.

Clone libraries showed that sulfate-reducing prokaryotes in deep peats comprised distinct populations in wet and dry seasons. Genetic diversity of cloned dsrAB sequences was higher in wet-season than in dry-season samples, which differed only in water content. The proportions of reduced (sulfides, thiols) and oxidized (sulfate) forms of sulfur were very similar between W-8 and D-14. The differences in redox conditions might influence the selection of specialized bacterial populations and therefore, the selection of different SRP. Seasonal variations in SRP populations have been reported in deep sediments of a freshwater lake (17). Using oligonucleotide probes complementary for 16S ribosomal RNA, the authors demonstrated that Desulfobulbus spp. showed relatively high abundance during almost all of the study periods, while Desulfobacterium spp. and Desulfovibrio spp. exhibited low relative abundance.

112 We can suggest two situations to explain the higher diversity in wet soils. First,

SRB populations present in wet season decline to non-detectable levels during the dry season. These populations might re-inoculate the soil during wet season by suspended cells present in the entering water. Another possibility is to suggest that OTU8 is a permanent population present at similar levels in wet and dry seasons while other populations increase and become dominant during wet season.

Wet season clones formed 9 OTUs, contrasting to only two OTUs found in dry season sample. After calculation of homologous coverage for clone libraries, we established that 97% and 84% of diversity was covered for D-14 and W-8, respectively

(31). These results indicate sulfate-reducing bacteria inhabiting these peat soils belong to new phylogenetic groups or phyla and they are specific to these metalliferous peat soils.

Some of the cloned sequences were related to other sequences retrieved from acidic fens

(approximately pH 4) characterized by low concentrations of sulfate (AY167468,

AY167479, AY167472, AM179478, AM179501) (21, 30). These observations suggest that these organic-rich environments might represent habitats for little-studied sulfate– reducing prokaryotes. These sequences are related to OTU5 and OTU10.

Loy et al. (21) demonstrated that their clones contained new functional DsrAB because they contained the Cys motif Cys-X5-Cys and Cys-X3-Cys, characteristic of sulfite reductases (7). The Cys motif is present in the deduced amino acid sequences from the newly described dsrAB genes, suggesting the presence of new organisms capable of dissimilating sulfate or sulfite in these metalliferous organic-rich soils.

113 CONCLUSIONS

This study shows the influence of temporal (wet and dry seasons) and spatial (soil cores) variability in sulfate reduction in soils from the Manning peatland region of

Western New York. In these peats, most (over 50%) of the total sulfur exits in the most reduced oxidation state (sulfide), principally in deeper soils. High concentrations of sulfur are accompanied with high concentrations of metals, suggesting an association between sulfur, cadmium, and zinc. RISA analyses demonstrated that the diversity of bacterial populations was influenced by seasonal changes (water content) and transect location (metal content). Clone libraries for dsrAB genes showed that SRP populations differ between seasons. New sequences of dsrAB genes were found in samples of dry and wet seasons and they were not affiliated to any sequence of cultured SRP. These results suggest that not yet described sulfate reducing prokaryotes are present in these peat soils and they could be specialized colonists of these metalliferous environments. Overall, our results suggest that the distribution and activity patterns of novel populations of SRPs in these metalliferous soils may be relevant to understanding biogeochemical processes occurring in these metal- and sulfur- rich peat soils.

114

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Chapter 5

General Conclusions

122 Sulfur exists in a number of oxidation states, from -2 to +6. Sulfur species undergo diverse transformations, many of them driven by microbial activity and thus becoming key reactions in sulfur cycling. The understanding of key processes in the sulfur cycle and how they respond to environmental factors is important for predicting the impact of changes induced naturally or anthropogenically in the global sulfur cycle.

We hypothesized that bacterial populations involved in S cycling were present and active in naturally metalliferous soils of the Manning peatland region and that their distribution, abundance and diversity vary with soil depth and seasonal changes. Soils from the Manning peatland region are naturally rich in S, Zn and Cd, and they are subjected to drastic changes in redox conditions. Our findings indicated sulfate-reducing activity in surface and deep soils from the Manning peatlands. The populations involved in the dissimilation of sulfur compounds were not related to well known sulfate-reducing prokaryotes. Members of the phylum Firmicutes are suspected to drive dissimilatory reactions in surface soils. In deep soils, however, analyses of dsrAB genes revealed that new prokaryotes respiring sulfate are present in soil profiles and their diversity is influenced by seasonal changes such as fluctuations in water table.

Enzymes involved in dissimilatory sulfate reduction were found in surface and deep peats (Fig. 5-1). New sequences for apsA and asrA genes were obtained in surface peat, suggesting the presence of new functional genes and/or pathways in these metalliferous peats.

123

Figure 5-1: Schematic representation of the dissimilatory sulfate reduction and the enzymatic pathways found in soils from the Manning peatland region of Western New York.

124 A great percentage of the total sulfur was found in the most reduced forms

(sulfides and thiols) and it seemed associated with higher concentration in Zn and Cd.

These observations suggest a linkage between microbial activity and the precipitation of metal sulfides in deep soils with high water content and very high concentrations of Zn and S. Co-localization of ZnS and bacteria was observed using confocal laser scanning microscopy. Sulfonates were an important fraction of the total sulfur, particularly in surface soils. Pathways involving dissimilation of sulfonates might be important reactions supporting sulfur cycling in these organic- and metal-rich surface soils.

The results obtained from this research suggest that peat soils from the Manning region represent a novel environment for the study of biological reduction of sulfur compounds. Future research in this area should address the isolation and identification of novel species of SRPs and the evaluation of their capacity to reduce oxidized forms of sulfur in addition to sulfate.

VITA

Carolina Elvira María Yáñez Prieto

Carolina Yáñez Prieto was born in Concepción, Chile in July 14, 1969. She got her B.A. degree in Biochemistry from the University of Concepción (Chile) in 1996 with a thesis titled

“Study of cadmium resistance in gram-negative bacilli of environmental origin”. After graduation, she spent a few years working as a microbiologist in clinical laboratories before deciding to go back to school. Carolina obtained a scholarship from the French Government and left Chile in 2000. She earned a Masters degree in Chemistry and Microbiology of Water from the Université Henry Poincaré in Nancy (France) in 2001 with her thesis “The effect of iron oxides on bacterial growth in drinking water distribution systems”. While living in France, she worked for the Commissariat à l’Energie Atomique (French Atomic Energy Commission) in a project titled “Study of bio-physico-chemical mechanisms of transport of As-Fe complexes in As- polluted underground waters in Bangladesh”. In January 2003, Carolina and her Russian husband moved to State College. Not long after her arrival, she started to work as a research assistant in the project “Microbial processes and populations as related to zinc, cadmium, and sulfur speciation in natural metalliferous soil environments” under the supervision of Dr. Carmen Enid

Martínez in the Department of Crop and Soil Sciences at The Pennsylvania State University. In

September 2003, encouraged by Dr. Martínez, she enrolled in the Ph.D. program in Soil Science.

During her graduate career, she published journal articles and presented her research at national and international professional meetings. After 6 years of intensive and enriched experiences,

Carolina is returning to Chile where she plans to start her career as an environmental microbiologist.