A BIODIVERSITY STUDY OF HIGH TEMPERATURE MUD POOL MICROBIAL COMMUNITIES: IMPLICATIONS OF REGIONAL/GEOGRAPHICAL ISOLATION AND ENDEMISM
by Benjamin R. Wheeler II
A thesis submitted to the Faculty of the University of Delaware in partial fulfillment of the requirements for the degree of Master of Science in Marine Studies
Winter 2006
Copyright 2006 Benjamin R. Wheeler II All Rights Reserved UMI Number: 1432290
Copyright 2006 by Wheeler, Benjamin R., II
All rights reserved.
UMI Microform 1432290 Copyright 2006 by ProQuest Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.
ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, MI 48106-1346 A BIODIVERSITY STUDY OF HIGH TEMPERATURE MUD POOL MICROBIAL COMMUNITIES: IMPLICATIONS OF REGIONAL/GEOGRAPHICAL ISOLATION AND ENDEMISM by Benjamin R. Wheeler II
Approved: ______S. Craig Cary, Ph.D. Professor in charge of thesis on behalf of the Advisory Committee
Approved: ______Nancy M. Targett, Ph.D. Dean of the Graduate College of Marine Studies
Approved: ______Conrado M. Gempesaw II, Ph.D. Vice Provost for Academic and International Programs
ii ACKNOWLEDGMENTS
The love and support of my family will always be my inspiration through everything that I do in life. Thank you Mom, Dad, Jim, and Mudder for always believing in me and for your overwhelming encouragement. A special thanks to Dr. Craig Cary for the opportunity to branch out into the realm of molecular biology and conduct my research in some of the most beautiful places on Earth. For those experiences, I will be forever grateful. Thank you to Dr. William Ullman and Dr. David Kirchman for their continued enthusiasm, support, guidance, and patience throughout my studies and work at CMS. For expert guidance and encouragement in learning the ins and outs of molecular biology, I would like to thank Dr. Kathy Coyne, Dr. Barbara Campbell, and Dr. Carol DiMeo. I would also like to thank the faculty, staff, and students of the Thermophile Research Unit, University of Waikato, New Zealand for their support and hospitality, especially Dr. Roy Daniel, Dr. Hugh Morgan, Tom Neiderberger, and Lee- Anne Rawlinson for their assistance in field sampling. Thank you to Karen Savidge for her time and patience in assisting me with nutrient analyses and Ken Voglesonger and Alf Harris for their assistance with ion, elemental, and SEM analysis. My years at CMS were blessed with the encouragement and support of a fantastic group of friends. My sincerest thanks go to Elizabeth and Sarah McCliment for their friendship and encouragement through both my graduate work and in life. Thanks to Justin Ossolinski, Tommy More, Ryan Dale, and Mia Steinberg for their friendship and countless hours of skimboarding. Thanks to Brandon Jones for his
iii friendship and comic relief throughout the trying moments of proposal writing. A special thanks goes to Steve and Becky Thur, Robert Trouwborst, Lindsay Kendall, and Mike League for being great friends and fantastic housemates. Thanks to Kevin Portune for countless hours of musical entertainment and to Karen Pelletreau for hours of conversation and encouragement through late nights of lab work. I am surrounded by a fantastic group of friends in Hawaii that have been a constant source of encouragement and make Hawaii truly an island paradise. Thanks to Paul Lethaby, Susan Curless, David Nichols, Tara Clemente, Rita Steyn, Christine Tallamy, Brian Glazer, Christine Pequignet, Michael Stat, and Jamie Becker. I would also like to thank Dr. Ruth Gates and the entire Gates Lab for their patience, encouragement, and support through my thesis writing and defense preparation. Thank you to Peggy Conlon, Debbie Booth, Connie Edwards, Julie Tigue, Doris Manship, Lisa Perelli, and the entire CMS support staff for always having the time to answer a thousand questions and for making sure I was always prepared for the next step. Throughout my life I have been blessed with the greatest friends anyone could ever wish for…thank you to Brantley Cauley, Jeff Waters, Saundra Butcher, Pedro de Cardenas, Matthew Bell, Cecile Deen, Mike Deen, Becky Holyoke, Amanda Whitmire, and Kitty Fielding for your unwavering love and friendship over the years. This work was supported by the National Geographic Society and the National Science Foundation. Student support provided by the Delaware Biotechnology Institute and the Graduate Research Trainee program.
iv TABLE OF CONTENTS LIST OF TABLES...... vii LIST OF FIGURES...... viii ABSTRACT...... x Chapter 1 - INTRODUCTION ...... 12 Background...... 12 Biodiversity Studies of High Temperature Ecosystems...... 13 Origins of Life...... 15 High Temperature Mud Pools...... 16 Geography and Hydrothermal Systems...... 17 Molecular Methodologies for Determination of Biodiversity ...... 19 Objectives...... 22 References...... 25 2 - ACCESSING THE BACTERIAL COMMUNITIES OF HIGH TEMPERATURE MUD POOLS: A COMPARATIVE STUDY OF REGIONAL AND GEOGRAPHICAL ISOLATION...... 30 Abstract ...... 30 Introduction...... 31 Materials and Methods...... 37 Sample collection, preparation and field measurements ...... 37 Nucleic acid extraction optimization and recovery of extracellular DNA...... 38 Geochemistry (New Zealand regional survey) and water content determination...... 40 Bacterial community fingerprinting analysis...... 40 Sequence analysis...... 42 Results...... 43 Field sampling and geochemistry ...... 43 Nucleic acid extraction optimization ...... 46 Bacterial community fingerprinting analysis...... 50 Discussion...... 59 Conclusion ...... 71 References...... 73 3 - PHYLOGENETIC ANALYSIS OF THE MICROBIAL COMMUNITY INHABITING A HIGH TEMPERATURE MUD POOL FROM THE TAUPO VOLCANIC ZONE, NEW ZEALAND...... 83 Abstract ...... 83 Introduction...... 84 Materials and Methods...... 87 Sample collection, preparation and field measurements ...... 87 Geochemistry ...... 88 Water content, elemental and particle size analyses...... 89
v Nucleic acid extraction...... 89 Community fingerprinting analyses...... 90 Clone library construction ...... 93 Results...... 95 Chemical, elemental, and particle size analysis...... 95 PCR-DGGE Bacterial and Archaeal community characterization...... 99 Bacterial and Archaeal community characterization of the Tokaanu mud pool ...... 102 Discussion...... 108 Conclusion ...... 120 References...... 122 4 - CONCLUSIONS AND FUTURE DIRECTIONS...... 130 Appendix...... 137
vi LIST OF TABLES Table 2.1 Collection sites and physical parameters of New Zealand high temperature mud pools sampled during this study...... 44 Table 2.2 Collection sites and physical parameters of Costa Rica, Lassen, and Yellowstone high temperature mud pools sampled during this study...... 45 Table 2.3 Chemical analyses of pore water from New Zealand mud pools...... 47 Table 2.4 Efficiency of nucleic acid extraction protocols on high temperature mud pool samples...... 49 Table 2.5 Sequence and BLAST results of excised bands from the DGGE analysis of New Zealand mud pool bacterial communities...... 55 Table 2.6 Sequence and BLAST analyses of excised bands from DGGE analysis of mud pool bacterial communities from New Zealand, Costa Rica, Yellowstone, and Lassen...... 57 Table 3.1 Oligonucleotides used for PCR...... 91 Table 3.2 Sample site overview and physical characterization of the Tokaanu Thermal Area (38° 58' S; 175° 45' E)...... 96 Table 3.3 Chemical analysis of pore water from the Tokaanu mud pool and its neighboring hot springs...... 97 Table 3.4 Ion chromatography of cations from the Tokaanu mud pool...... 98 Table 3.5 BLAST results of excised bands from the bacterial DGGE analysis of the Tokaanu mud pool and its neighboring hot springs...... 104 Table 3.6 Summary of bacterial 16S rRNA clone analysis from the Tokaanu mud pool...... 109 Table 3.7 Summary of archaeal 16S rRNA clone analyses from the Tokaanu mud pool...... 110 Table A.1 Commonly used abbreviations...... 138
vii LIST OF FIGURES Figure 2.1 A. DNA was exposed to low pH and high temperature (65º C) conditions for 1 hour, amplified using PCR, and run on a 1% agarose gel. 1: Control, 2: DNA, pH 3, 65º C, 3: DNA, 65º C, 4: DNA, pH3. B. DNA was exposed to neutral pH and high temperature (90º C) conditions for 10 minutes, amplified using PCR, and run on a 1% agarose gel. 1: T0, pH 8.0; 2: T0, pH 6.7; 3: pH 8.0, RT; 4: pH 6.7, RT; 5: pH 8.0, 90º C; 6: pH 6.7, 90º C...... 50 Figure 2.2 DGGE analysis of bacterial communities associated with 13 mud pools located in the Taupo Volcanic Zone, North Island, New Zealand. Numbers designate bands that were excised, re-amplified, and subjected to sequence analyses...... 52 Figure 2.3 Phoretix comparative analysis of New Zealand mud pool bacterial DGGE analyses. UPGMA dendogram displays similarities between the bacterial communities based on the presence and absence of bands in the profiles.54 Figure 2.4 DGGE analysis of bacterial communities from 16 mud pools from New Zealand, Costa Rica, Lassen, and Yellowstone. Numbers designate bands that were excised, re-amplified, and subjected to sequence analyses...... 56 Figure 2.5 Phoretix comparative analysis of New Zealand, Costa Rica, Lassen and Yellowstone mud pool bacterial DGGE analyses. UPGMA dendogram displays similarities between the bacterial communities based on the presence and absence of bands in the profiles...... 58 Figure 3.1 Elemental analysis of the solid phase component of the Tokaanu mud pool using KEVEX energy-dispersive elemental analysis...... 101 Figure 3.2 Particle size analysis of the Tokaanu mud pool solid phase using scanning electron microscopy...... 101 Figure 3.3 DGGE analysis of the bacterial communities associated with the Tokaanu mud pool and its neighboring hot springs. Numbers designate bands that were excised, reamplified, and subjected to sequence analyses...... 103 Figure 3.4 DGGE analysis of the archaeal communities associated with the Tokaanu mud pool and it neighboring hot springs...... 105 Figure 3.5 Phoretix comparative analysis of the bacterial DGGE analyses of the Tokaanu mud pool and its neighboring hot springs. UPGMA dendogram
viii displays similarities based on the presence and absence of bands in the profiles...... 106 Figure 3.6 Phoretix comparative analysis of archaeal DGGE analyses of the Tokaanu mud pool and its neighboring hot springs. UPGMA dendogram displays similarities based on the presence and absence of bands in the profiles..107 Figure 3.7 Bacterial phylogenetic tree of Tokaanu mud pool sequences. Phylogenetic tree as determined by neighbor joining analysis. Bootstrap values based on 100 replications...... 111 Figure 3.8 Archaeal phylogenetic tree of Tokaanu mud pool sequences. Phylogenetic tree as determined by neighbor joining analysis, Bootstrap values are based on 100 replications...... 112
ix ABSTRACT In assessments of global biodiversity, extreme environments like terrestrial and deep-sea hydrothermal venting systems have been the focus of much scientific research. Created by the interactions of crustal plates and “hot spot” eruptive events, these unique ecological niches have led to a wealth of information on the diversity and evolution of life on earth. The organisms that thrive in high temperature ecosystems have challenged our understanding of the physical and biochemical constraints on the upper temperature limits for life and stimulated new theories on the origins of life on earth and the possible existence of life on other planets. One terrestrial extreme environment that has remained relatively uncharacterized in terms of microbial biodiversity is the high temperature mud pool. While molecular and geochemical technologies, independent of traditional isolation and cultivation methodologies, have contributed immensely to the study of life in other high temperature systems like submarine hydrothermal vents and terrestrial hot springs, high temperature terrestrial mud pools have yet to be explored using these sophisticated tools. The aim of this research was to perform an intensive survey of the microbial diversity of high temperature mud pools on a regional and geographical scale using a combination of molecular and geochemical analytical techniques. Mud pool samples were collected from the thermal fields of the Taupo Volcanic Zone (TVZ), New Zealand (regional survey); Rincon de la Vieja National Park, Costa Rica; Lassen Volcanic Park, USA; and Yellowstone National Park, USA. Bacterial community analysis was performed using a combination of DNA fingerprinting and sequencing analyses.
x Community level analysis of high temperature mud pools showed the majority of the pools were dominated by members of the gamma Proteobacteria, the Cytophaga- Flavobacteria, and the alpha Proteobacteria. These bacterial signatures persisted across a broad range of physiochemically distinct mud pools. Although the dominant groups were consistent across the majority of the pools, the bacterial diversity observed in the DGGE fingerprinting analysis suggested physical isolation might dictate bacterial community diversity of individual pools over both a regional and geographical range. The second objective of this study was to determine if the mud pool communities support a unique endemic community when compared to hot spring systems. Samples were collected from a mud pool and several closely associated hot springs within the Tokaanu Thermal Area, New Zealand. DNA fingerprinting analysis of the bacterial and archaeal communities showed a unique microbial assemblage unique to the Tokaanu mud compared to neighboring hot water features. The mud pool was found to harbor a diversity of bacteria relating to the Firmicutes, epsilon Proteobacteria, gamma Proteobacteria, Thermus sp., and others from high temperature and deep subsurface environments. Signatures of the archaeal community associated with this pool revealed a closely related group of Archaea forming a unique clade of the Crenarchaeota. The strong similarities between these archaeal signatures and their relative distance from other characterized Crenarchaeotes suggested these microbes are endemic to the Tokaanu mud pool system.
xi Chapter 1 INTRODUCTION
Background Throughout geological history, hydrothermal systems have existed as surface indicators of the ever-changing and dynamic forces at work beneath the earth’s crust (Reysenbach and Cady, 2001). In localized regions, ground water or seawater is entrained through cracks in the Earth’s crust where it reacts with subsurface material, becoming super-heated, returning to the surface through venting systems. These hydrothermal vents occur both in marine and terrestrial environments. The hydrothermal fluid is generally rich in sulfur and metals at concentrations that greatly exceed those present in the original surface or sea waters (Jannasch, 1995). The origins of individual hydrothermal systems differ on a global scale due to their geographic and tectonic positioning. The terrestrial thermal features and deep-sea hydrothermal vents, created by the interactions of crustal plates and/or “hot spot” eruptive events, have been the focus of much scientific research in attempts to better understand the geological mechanisms driving these systems and the unique ecological niches created by subsurface forces. In 1897, the presence of life in the hot springs of Yellowstone was described to exist at temperatures in excess of 80ºC (Davis, 1897). The discovery of life at such high temperatures later initiated pioneering studies by Brock and colleagues into the identification and ecology of these thermophilic organisms (Brock, 1967; Bott and Brock, 1969; Brock and Darland, 1970). The discovery of deep-sea hydrothermal vents
12 in 1977 (Corliss et al., 1979) led to a rejuvenated interest and further attempts to gain an understanding of the physical and chemical interactions driving these high temperature systems and their associated biological communities. Studies of the organisms that thrive in high temperature ecosystems have led to new information on the diversity and evolution of life on earth, challenged our understanding of the physical and biochemical constraints on the upper temperature limits for life, and stimulated new theories on the origins of life on earth and the possible existence of life on other planets (e.g. Baross and Hoffmann, 1985; Pace, 1991).
Biodiversity Studies of High Temperature Ecosystems
In recent years, the assessment of global biodiversity has been the focus of much scientific research in an attempt to fully understand and preserve natural biological resources. These studies have shown that the dominant diversity of life resides in the microbial world, distributed through the three domains of life: Bacteria, Archaea, and Eucarya (Pace, 1997). Early studies of the microbial diversity of hydrothermal and other environments relied strongly on standard cultivation and isolation techniques. Even though the use of these methods proved successful at times, cultivation and isolation are estimated to exclude the majority of microorganisms present in natural environments (Amann et al., 1995). Developments in molecular genetic technologies have revolutionized our understanding of the diversity, distribution, and evolution of microbes in the environment and have lessened the need for cultivation in the study of microbial diversity (Pace et al., 1986; Ward et al., 1992). Molecular methodologies based on the small subunit of the rRNA (16S rRNA) gene are capable of revealing a wealth of previously unrecognized microbial diversity and novel phylogenetic lineages that
13 represent major components of global microbial assemblages (Barns et al., 1996; Hugenholtz et al., 1998; Takai and Sako, 1999). Phylogenetic analyses of complex environmental microbial communities indicate that the dominant proportion of resident Bacteria and Archaea are not represented in culture collections (Amann et al., 1995). These studies have provided insight into the widespread distribution of certain microbial lineages and more notably the evolutionary history of the prokaryotes (Woese, 1987). This same approach has been used in several high temperature systems with dramatic results that have implications pivotal to our understanding of the evolution of all the domains of life (Archaea, Bacteria, and Eukarya) and to the origins of life itself (e.g. Reysenbach et al., 1994; Reysenbach et al., 2000; Barns et al., 1994; Pace, 1991). Although novel thermophilic isolates from hydrothermal systems have increased exponentially in the past decade (Stetter, 1996a) and the upper temperature for life has been continually challenged (121°C, Kashefi and Lovley, 2003), the ecology and diversity of these high temperature microbial communities is relatively poorly studied. Analysis of rRNA sequences obtained from a hot spring in Yellowstone appear to branch below the bifurcation of the Crenarchaeota and Euryarchaeota kingdoms of the Archaea domain leading to the proposal of an entirely new kingdom, the Korarchaeota (Barns et al., 1996). Other potential members of the Korarchaeota have recently been reported from similar hot springs in Yellowstone (Reysenbach et al., 2000). More recently, identification of a new archaeal member, Nanoarchaeum equitans has led to the proposal of a new phylum within the Archaea, the Nanoarchaeota (Huber et al., 2002). The phylogenetic placement of thermophiles deeply within the tree of life has sparked theories into the role of thermophilic environments in the origins of life on earth and the possibility of life on other planetary bodies.
14 Origins of Life Two major theories on life’s origins have been proposed over the last two decades. The first theory is based on the hypothesis that life arose under “reduced, high temperature conditions” and the second is that life arose in a clay-rich environment. The “reduced, high temperature conditions” theory relies heavily on studies of deep-sea hydrothermal vents. Thermodynamic calculations have shown the abiotic synthesis of all 20 amino acids required for protein formation to be energetically favorable in “hot, moderately reduced, submarine hydrothermal fluids” compared to cooler, oxidized seawater (Amend and Shock, 1998). Early earth is believed to have been much warmer with an atmosphere probably rich in CO2 and H2 (Kasting, 1993). Given these conditions, life may have begun under conditions analogous to modern hydrothermal systems (Reysenbach and Cady, 2001). The deepest branching members of the Bacteria and Archaea based on the 16S rRNA tree of life are all thermophilic and many are chemolithoautotrophs (Stetter, 1996b; Woese et al., 1990). Although the role of chemoautotrophic thermophiles in the origins and evolution of life is still unclear (Doolittle, 1999; Galtier et al., 1999), there is strong evidence that suggests early life may have been chemosynthetic and may have required high temperatures for growth (Pace, 1997). The “clay hypothesis” theory focuses on the possible importance of clays in the abiotic synthesis and replication of simple organic molecules. In 1951, Bernal suggested clays as possible sites for the concentration and catalysis of organic molecules due to the enormous surface area and the commonality of replicating layers of clays (Bernal, 1951). Cairns-Smith in 1966 expanded on Bernal’s hypothesis by suggesting that the irregularities of clay crystal formation caused by ion substitutions may encode information that could replicate under the proper crystal growth conditions (Cairns-
15 Smith, 1966). This idea was based on observations that newly formed replicate layers could separate from the nucleating layer allowing the new surfaces to become adsorptive and catalytically functional (Cairns-Smith, 1966). The “clay hypothesis” does not rely heavily on temperature dependence, yet hydrothermal environments are important systems for the synthesis of clays from free ions due to the high mineral content of hydrothermal fluids and the presence of liquid water under conditions of high temperature at high pressure (Hartman, 1986). In light of these theories, the presence of the high temperature clay/fluid matrix in mud pools may present favorable environmental conditions for a more in-depth study of early life and evolution.
High Temperature Mud Pools One unique, globally-distributed, terrestrial geothermal feature that has remained relatively uncharacterized in terms of microbial biodiversity is the high temperature mud pool. Mud pools are hot, acidic pools of bubbling silts and clays formed by the dissolution of rock as steam rises from groundwater chambers deep below the Earth’s surface. The clay matrix is physically characterized by dramatic between pool variability in viscosity, temperature, and pH. These ecosystems represent a hot, reduced environment that when coupled with the microbial and chemical signatures may resemble features of primordial earth and could provide valuable insight into the origins and evolution of life on Earth. Although molecular and geochemical technologies, independent of traditional isolation and cultivation methodologies, have contributed immensely to the study of life in high temperature hydrothermal systems like submarine hydrothermal vents and terrestrial hot springs, high temperature terrestrial mud pools have yet to be explored using these sophisticated tools. The presence of a viscous, clay matrix coupled
16 with extreme physiochemical variability is unique to mud pool systems. Mud pool ecosystems may have been a prominent feature in the early evolution of the planet and may represent a primitive niche similar to that of early earth. As a possible analog to primordial earth environments, these dynamic systems may support a microbial community of deep evolutionary lineage and the utilization of highly specialized metabolic strategies. Previous studies have resulted in the isolation and cultivation of only a few extremophiles (e.g. Pyrococcus sp.) from mud pools (Stetter, 1986); cultivation independent molecular technologies have not been applied to mud pool environments. The lack of cultivation successes may be attributed to a poor understanding of the geochemical conditions under which these organisms thrive and the metabolic strategies utilized within these systems.
Geography and Hydrothermal Systems Some of the most significant regions of hydrothermal activity occur along zones of global plate subduction. Subduction zones are areas where two tectonic plates collide with one passing under the other. These zones are marked by high volcanic activity such as the Pacific “Ring of Fire.” The “Ring of Fire” is an arch of volcanic and seismic activity that stretches from New Zealand, along eastern Asia’s edge, across the Aleutian Islands of Alaska to the north, and south along the coasts of North and South America. Over 75% of the world’s active and dormant volcanoes lie within this zone. The thermal regions created by these plate margin events have been the focus of much research due to the abundance and accessibility of surface thermal features. The North Island of New Zealand, located directly above the southeastern rim of the “Ring of Fire” where the Pacific Plate slides beneath the Indo-Australian Plate, is a highly active site of volcanic and geothermal activity. Within the last million
17 years, most of New Zealand’s volcanism has occurred within the Taupo Volcanic Zone (TVZ), which includes three frequently active volcanoes (Ruapehu, Tongariro/Ngauruhoe, and White Island), and two of the world’s most productive caulderas (Okataina and Taupo). The TVZ is classified as being “extremely active” on the scale of global volcanism (Volcanoes in New Zealand: Institute of Geophysical and Nuclear Sciences Limited website, 2005). The TVZ consists of 17 geothermal fields containing hundreds of individual geothermal features. Hydrothermal activity in this region creates one of the highest concentrations of mud pools spread over a significant geographical area in the world. Other areas of significant volcanic and tectonic activity along the eastern edge of the “Ring of Fire” occur beneath Costa Rica and Lassen Volcanic National Park, California, USA. Beneath Costa Rica, the collision and subduction of the Cocos and Caribbean oceanic plates creates a highly-active system of volcanoes, ridges, and earthquake centers. One of the most active regions of terrestrial hydrothermal activity in Costa Rica is associated with the Rincon de la Vieja National Park in the northwestern region of the country. This isolated and highly active region contains several geothermal features including hot springs and several highly active mud pools. Lassen National Park, located in the western United States, is a compact area of volcanic and other thermal features. Situated on a vast lava plateau of isolated volcanic peaks, the Lassen peak is the southernmost volcano in the Cascade Range. The geothermal areas associated with Lassen peak are comprised of numerous bubbling mud pools, steaming fumaroles, and other boiling water features. One of the most interesting and unique areas of volcanic, tectonic, and hydrothermal activity in the world is Yellowstone National Park in the western United
18 States. One of only a few regions on the planet designated as a continental “hot spot,” magma lying merely a few thousand feet below the surface creates an abundance of geothermal activity and geological features. Yellowstone contains the world’s largest, most variable concentration of geothermal features including hot springs, geysers, steam vents, and mud pools. Yellowstone National Park is one of the most studied locations of geothermal activity in the world from both a geological and biological perspective.
Molecular Methodologies for Determination of Biodiversity A variety of molecular methodologies have been employed in recent years in studies of microbial biodiversity across a wide range of ecosystems, including high temperature environments. One of the significant advantages of these techniques lies in the ability to perform community level analyses of the microbial populations without the necessity of cultivation. Through the use of the polymerase chain reaction (PCR) (Saiki et al., 1988) and the subsequent cloning of particular genes of interest, comparisons can be made within and between communities (Pace et al., 1986). The phylogenetic characterization of microorganisms has been traditionally based on analysis of the 16S small subunit ribosomal RNA gene DNA sequence which contains a mosaic of highly conserved and variable regions (Woese, 1987). This method has become the most common technique for cataloging members of microbial communities (Fuhrman et al., 1994). It has been used successfully in the examination and analysis of prokaryotic communities of many different environments including marine environments (e.g. Giovannoni et al., 1990; Britschgi et al., 1991; DeLong, 1992), hot springs (e.g. Ferris et al., 1996; Ward et al., 1998), deep-sea hydrothermal vents (e.g. Campbell and Cary, 2001), deep-sea sediments (e.g. Vetriani et al., 1999), Antarctic coastal waters (e.g. Murray et al., 1998), and many others. Phylogenetic studies based on the 16S rRNA
19 gene can be performed at the species level or higher due to the gene’s structure of both variable and conserved regions (Woese, 1987). Typically, the construction of 16S rDNA clone libraries and screening with restriction fragment length polymorphism (RFLP) analysis can be utilized in estimating microbial community diversity and structure in environmental samples (Moyer et al., 1994); however, these methodologies can be expensive and tedious. Recently, a PCR-based comparative technique known as denaturing gradient gel electrophoresis (DGGE) has been employed in studies of microbial community structure in environmental samples (Myers and Maniatis, 1986; Muyzer et al., 1993). This DNA fingerprinting technique allows for the separation of PCR- generated DNA fragments (amplicons) from samples containing multiple templates. With DGGE, amplicons of the equal size containing a GC clamp are exposed to a gradient of increasing denaturant (40% formamide and 7M urea) and a constant high temperature (60°C). As the double-stranded amplicons move through the gradient of denaturant, portions of the double-stranded fragment begin to denature creating double- stranded and single-stranded regions within the same fragment. The conformational change in the amplicon’s structure slows the migration of the fragment through the gel. This separation is a factor of the melting properties of the individual nucleotide sequences of the amplicons (Fischer and Lerman, 1980; Myers et al., 1987) and the gradient of denaturant used in the gel. More traditional agarose or acrylamide gel electrophoresis separates amplicons based on fragment size. As the amplicon begins to partially melt, it will stall, resulting in a stable, unique, visible band within the gel profile. In studies of diverse environmental microbial communities using DGGE, a single gene from DNA extracted from the entire community is amplified using the PCR and
20 conserved primers. When subjected to DGGE, each unique amplicon will migrate independently of those from the total community. The migration and denaturation of these fragments create a multiple banding pattern or “fingerprint” of the total community of amplicons within the matrix of the gel. Amplification of the 16S rDNA gene using the PCR followed by DGGE (Muyzer and Ramsing, 1996) provides a “snapshot” fingerprint of the dominant members of microbial communities (Muyzer et al., 1993; Duineveld et al., 1998). This method of microbial diversity and community structure is very cost and labor effective compared to that involved in cloning and can provide resolution down to a single base pair variation (Fodde and Losekoot, 1994) between individual members of the community. The excision and sequencing of individual bands from DGGE gels can be performed for phylogenetic placement of the DNA. Cloning and DGGE are both powerful tools in assessing microbial diversity and community structure, yet, there are inherent biases associated with each method that include nucleic acid extraction efficiency, cloning efficiency, and the PCR amplification (Wintzingerode et al., 1997). Nucleic acid extraction methods differ in their ability to extract DNA and RNA from samples taken from specific environments and the cellular structure of organisms differs in their susceptibility to different extraction agents. In using methodologies based on the 16S rRNA gene such as DGGE, sequence heterogeneity within multiple copies of this gene within an individual microbe must be considered as this may result in inaccurate estimates of diversity and community structure in natural samples (Farrelly et al., 1995). Despite its effectiveness in the initial assessment of community structure and diversity, DGGE does exhibit some limitations. DGGE is limited in the ability to separate a large number of DNA fragments present in a
21 single sample and there exists the potential for comigration of different amplicons with closely related nucleotide sequences (Muyzer et al., 1993; Vallaeys et al., 1997). Other limitations associated with DGGE are the detection of heteroduplex molecules (base mismatches in double-stranded DNA) (Ferris and Ward, 1997) and sensitivity in the detection of minor community members (Muyzer, 1999). Although the PCR is widely used in molecular biology for the amplification and study of 16S rRNA genes from mixed template samples, variability in the efficiency of the PCR is often observed. The content and structure of the template used in the PCR reaction partially defines amplification efficiency. The use of templates that are G-C rich show higher binding efficiencies of the amplification primers, denaturation of template strand in the reaction tends to be more preferential to those containing low G-C content, the use of degenerate primers, and sequences adjacent to priming sites all effect the amplification reaction and other downstream processes (Suzuki and Giovannoni, 1996; Polz and Cavanaugh, 1998). There are also concerns that certain templates within a community sample may be more abundant or more easily amplified, thus dominating the reaction (Reysenbach et al., 1992; Suzuki and Giovannoni, 1996). Studies of PCR bias have defined some methods for minimizing these artifacts such as avoiding the use of degenerate primers, using high template concentration in reactions, combining replicate reactions before downstream use, and keeping the cycle number of the reaction low (Suzuki and Giovannoni, 1996; Polz and Cavanaugh, 1998).
Objectives The microbial diversity and community structure of high temperature mud pools has remained relatively uncharacterized prior to the present study. Previous work involved the use of traditional cultivation techniques and was met with very little success.
22 The research goal of this study was to perform an intensive survey of the microbial biodiversity of high temperature mud pools from four unique, geographically isolated hydrothermal systems using a combination of molecular and geochemical techniques. I hypothesize that high temperature terrestrial mud pools are possible analogs to the primordial earth environment and may harbor unique and deeply branching assemblages of Bacteria and the Archaea. The first phase of this research project was to perform a biogeographical assessment of the mud pool microbial communities from 4 field sites (Taupo Volcanic Zone (New Zealand), Rincon de la Vieja (Costa Rica), Yellowstone National Park (USA), and Lassen National Park (USA) using denaturing gradient gel electrophoresis (DGGE), geochemistry, temperature, and pH data. These 4 study sites are areas of significant hydrothermal activity and characterized by high concentrations of assessable mud pools. The goals of this research phase were:
1. To develop an effective protocol for the extraction of nucleic acids from high temperature mud pool samples and determine the longevity of exogenous inputs of DNA into these systems. 2. Use of DGGE to access the bacterial community structure and diversity of high temperature mud pools from 8 thermal regions within the Taupo Volcanic Zone (North Island, New Zealand) correlating the bacterial community structure of high temperature mud pools to the geochemistry and measurements of temperature and pH. 3. Compare the bacterial community structure and diversity of high temperature mud pools from 4 geographically isolated hydrothermal systems.
23 The second phase of this research project consisted of intensive sampling, molecular genetic, and geochemical characterization of a single mud pool and neighboring hot springs. The single pool displaying a unique Bacterial and Archaeal community (Tokaanu) was chosen for a more focused and in depth study. Molecular and geochemical comparative analyses were performed on this mud pool and neighboring hot springs to determine if the microbiology is endemic to the mud pool systems. The goals of this research phase were:
1. To characterize and compare the physiochemical environment of the Tokaanu mud pool and its neighboring hot springs. 2. Performing a direct comparison of the microbial communities (Bacteria and Archaea) of the Tokaanu mud pool and its neighboring hot springs. 3. Constructing bacterial and archaeal clone libraries of the 16S rDNA signatures from the Tokaanu mud pool for phylogenetic inference.
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26 Hugenholtz, P., C. Pitulle, K.L. Hershberger, and N.R. Pace. 1998. Novel Division Level Bacterial Diversity in a Yellowstone Hot Spring. Journal of Bacteriology. 180: 366-376. Jannasch, H.W. 1995. Microbial interactions with hydrothermal fluids, p. 273-296. In S.E. Humphris, R.A. Zierenberg, L.S. Mullineaux, and R.E. Thomson (ed.), In: Seafloor hydrothermal systems: Physical, chemical, biological, and geological interactions, vol. 91. American Geophysical Union, Washington, D.C. Kashefi, K. and D.R. Lovley. 2003. Extending the upper temperature limit for life. Science. 301: 934. Kasting, J.F. 1993. Earth’s early atmosphere. Science. 259: 920-926. Moyer, C.L., F.C. Dobbs, and D.M. Karl. 1994. Estimation of diversity and community structure through restriction fragment length polymorphism distribution analysis of bacterial 16S rRNA genes from a microbial mat at an active, hydrothermal vent system, Loihi Seamount, Hawaii. Applied and Environmental Microbiology. 60: 871-879. Murray, A.E., C.M. Preston, R. Massana, L.T. Taylor, A. Blakis, K. Wu, and E.F. DeLong. 1998. Seasonal and spatial variability of bacterial and Archaeal assemblages in the coastal waters near Anvers Island, Antarctica. Applied and Environmental Microbiology. 64: 2585-2595. Muyzer, G.E., C. de Waal, and A.G. Uitterlinden. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes encoding for 16S rRNA. Applied and Environmental Microbiology. 59: 695-700. Muyzer, G. and N.B. Ramsing. 1996. Molecular Methods to Study the Organization of Microbial Communities. Water Science and Technology. 32: 1-9. Muyzer, G. 1999. DGGE/TGGE a method for identifying genes from natural ecosystems. Current Opinion in Microbiology. 2: 317-322. Myers, R.M. and T. Maniatis. 1986. Recent advances in the development of methods for detecting single-base substitutions associated with human genetic diseases. Cold Spring Harbor Symposium for Quantitative Biology. 51: 275-284. Myers, R.M., T. Maniatis, and L.S. Lerman. 1987. Detection and localization of single base changes by denaturing gradient gel electrophoresis. Methods in Enzymology. 155: 501-527. Pace, N.R., D.A. Stahl, D.J. Lane, and G.J. Olsen. 1986. In: Marshall KC (ed) Current microbial ecology. Plenum Press, New York, pp. 1-55. Pace, N.R. 1991. Origin of life-Facing up to the physical setting. Cell. 65:531-533.
27 Pace, N.R. 1997. A Molecular View of Microbial Diversity and the Biosphere. Science. 276: 734-740. Polz, M.F. and C.M. Cavanaugh. 1998. Bias in template-to-product ratios in multitemplate PCR. Applied and Environmental Microbiology. 64: 3724-3730. Reysenbach, A.L., L.J. Giver, G.S. Wickham, and N.R. Pace. 1992. Differential amplification of rRNA genes by polymerase chain reaction. Applied and Environmental Microbiology. 58: 3417-3418. Reysenbach, A.L., G.C. Wickham, and N.R. Pace. 1994. Phylogenetic analysis of the hyperthermophilic pink filament community in Octopus Spring, Yellowstone National Park. Applied and Environmental Microbiology. 60: 2113-2119. Reysenbach, A.L., M Ehringer, and K. Hershberger. 2000. Microbial diversity at 83 ˚C in calcite springs, Yellowstone National Park: another environment where Aquificales and “Korarchaeota” coexist. Extremophiles. 4: 61-67. Reysenbach, A.L. and S.L. Cady. 2001. Microbiology of ancient and modern hydrothermal systems. TRENDS in Microbiology. 9: 79-86. Reysenbach, A.L. and E. Shock. 2002. Merging genomes with geochemistry at hydrothermal ecosystems. Science. 296:1077-1082. Saiki, R.K., D.H. Gelfand, S. Stoffel, S.J. Scharf, R. Higuchi, G.T. Horn, K.B. Mullis, and H.A. Erlich. 1988. Primer-directed enzymatic amplification of DNA with a thermostable DNA-polymerase. Science. 239: 487-491. Stetter, K.O. 1986. Diversity of extremely thermophilic archaebacteria. In Thermophiles: General, Molecular, and Applied Microbiology. Eds. T. D. Brock. John Wiley & Sons, New York, pp. 39-74. Stetter, K.O. 1996a. Hyperthermophilic prokaryotes. FEMS Microbiology Reviews. 18: 149-158. Stetter, K.O. 1996b. Hyperthermophiles in the history of life. Ciba Foundation. Symposium. 202: 1-10. Suzuki, M.T. and S.J. Giovannoni. 1996. Bias caused by template annealing in the amplification of mixtures of 16S rRNA genes by PCR. Applied and Environmental Microbiology. 62: 625-30. Takai, K. and K. Sako. 1999. A molecular view of archaeal diversity in marine and terrestrial hot water environments. FEMS Microbiology-Ecology. 28: 177. Vallaeys, T., E. Topp, G. Muyzer, V. Macheret, G. Laguerre, and G. Soulas. 1997. Evaluation of denaturing gradient gel electrophoresis in the detection of 16S rDNA sequence variation in rhizobia and methanotrophs. FEMS Microbiology- Ecology. 24: 279-285.
28 Vetriani, C., H.W. Jannasch, B.J. MacGregor, D.A. Stahl, and A.-L. Reysenbach. 1999. Population structure and phylogenetic characterization of marine benthic Archaea in deep-sea sediments. Applied and Environmental Microbiology. 65: 4375- 4384. Volcanoes in New Zealand. 2005. Institute of Geophysical and Nuclear Sciences Limited. 25 Nov 2005
29 Chapter 2 ACCESSING THE BACTERIAL COMMUNITIES OF HIGH TEMPERATURE MUD POOLS: A COMPARATIVE STUDY OF REGIONAL AND GEOGRAPHICAL ISOLATION
Abstract Studies of microbial biodiversity using molecular genetic techniques have revolutionized the field of microbial ecology and allowed for a more comprehensive understanding of microbial interactions with their local environment. The application of these sophisticated tools have allowed for the exploration and understanding of the evolution and origins of life on earth. Critical to these studies are the organisms and geochemical processes occurring in high temperature environments, like deep-sea hydrothermal vents and hot springs. High temperature mud pools present a unique, dynamic niche dominated by water-rock interactions, yet little is known about the microbial diversity of these unique ecosystems. The goal of this study was to explore the bacterial community structure of high temperature mud pools is relation to the local physiochemical environment from four geographically isolated hydrothermal systems. Mud pool microbial community structures were analyzed and compared using DGGE in correlation with characterization of the physical and chemical environment. Bacterial community analysis revealed unique bacterial communities dominated by thermophilic members of the gamma and alpha Proteobacteria across both regional and global ranges. These bacteria displayed high tolerance and adaptability to a range of physiochemical conditions. Domination of specific bacterial groups across a global
30 scale may have implications for an endemic mud pool bacterial assemblage. Future studies of the Archaea and direct comparisons between mud pools and other hot water features will assist in our understanding of these unique environments.
Introduction Hydrothermal venting systems have been indicative of the powerful and energetic forces occurring deep beneath the earth’s crust throughout geological history (Reysenbach and Cady, 2001). Important in the cycling of elements between the lithosphere, hydrosphere, and atmosphere during the early evolution of Earth (Des Marais, 1996), hydrothermal fluids flow from the surface environment and interact chemically with subsurface material taking on distinct geochemical and biological characteristics that result in large transfers of materials and energy (Farmer, 2000). As these superheated fluids return to the surface through terrestrial and deep-sea venting systems, they create some of the most thermally extreme habitable environments on the planet. High temperature venting environments present a great challenge to the organisms thriving in and around these ecosystems, leading to the evolution of unique mechanisms for survival under such harsh conditions (Childress, 1992). Implicated as potential modern analogs to the early earth environment (reduced, high temperature) and the phylogenetic placement of thermophiles deeply within the tree of life (Woese, 1987; Stetter, 1996), hydrothermal venting systems may be representative of the “ancestral niche” for the origin of life (Walter and Des Marais, 1993). The study of thermophilic environments have contributed to a wealth of new information into the diversity of bacteria capable of living at high temperatures (e.g. Barns et al., 1994; Reysenbach et al., 1994; Barns et al., 1996; Hershberger et al., 1996) while stimulating new theories of the origins of life on earth and the possible existence of
31 life on other planets (Baross and Hoffmann, 1985; Pace, 1991). Although the identification and isolation of novel thermophilic microorganisms from hydrothermal systems have increased exponentially in recent years (Stetter, 1996; Marteinsson et al., 2001; Vetriani et al., 2004; Voordeckers et al., 2005), the ecology and diversity of many high temperature environments and their resident microbial communities remain relatively poorly understood. Studying life in high temperature environments has been aided immensely by the development of modern molecular genetic-based technologies that allow microbial populations and their specific metabolic capabilities to be explored without the need for cultivation (e.g. Cary et al., 2004). Microbial biodiversity studies of high temperature environments have largely centered on terrestrial hot springs and deep-sea hydrothermal vents (e.g. Ward et al., 1998; Takai and Horikoshi, 1999; Reysenbach and Cady, 2001; Chapelle et al., 2002; Teske et al., 2002; Hoek et al., 2003, Keller and Zengler, 2004). One terrestrial hydrothermal ecosystem that has remained relatively uncharacterized in terms of microbial diversity is the high temperature mud pool (or mud pot). These pools of hot, homogeneously mixed, acidic, bubbling clays, silts, and volcanic ash are formed by the dissolution of rock by steam and hydrothermal fluids deep below the earth’s surface. Mud pools are found worldwide in areas of surface hydrothermal activity often exhibiting dramatic between pool variability in viscosity (via subsurface hydrothermal fluids and surface precipitation), temperature, and pH (Yellowstone website; personal observation). The chemically reduced, high temperature hydrothermal fluids and clay minerals comprising the mud pool ecosystem are key constituents in recent theories into the origins of life (Amend and Shock, 1998; Hartman, 1986). Thermodynamic calculations based on hydrothermal systems suggest that some complex organic
32 compounds are synthesized at high temperatures (Schulte and Shock, 1995). This was substantiated through experimental work with the synthesis of alcohols (Voglesonger, 1999) and the formation of oligopeptides (Imai et al., 1999) under simulated submarine venting system conditions. Integrations of the genetic evidence from all life on Earth suggest water-rock interactions in the presence of high temperatures were the earliest biological habitat (Shock, 1998). The volcanic clay-based matrix of mud pools may have implications on a theory that hypothesizes the potential importance of clays in the development of early biological systems. Proposed by Bernal in 1951 and expanded by Cairns-Smith in 1966, the “clay hypothesis” suggested clays as potential sites for the concentration, catalysis, and replication of organic molecules due to their enormous surface area in correlation with irregularities in clay crystal formation. Clay ion substitution during crystallization may encode information that could replicate under the proper crystal growth conditions (Cairns-Smith, 1966). There is also some evidence of the formation of clays from free ions due to the high mineral content of hydrothermal fluids and the presence of liquid water under conditions of high temperature at high pressure have also been documented (Hartman, 1986). Combining the presence of hydrothermal conditions and the structural framework of clays, high temperature mud pools may be representative of a unique niche harboring novel microorganisms of short, deeply branching phylogenetic lineages. Although a prominent surface feature in many areas of hydrothermal activity, little research has been conducted on characterizing the microbial communities of high temperature mud pool systems. Past investigations of the microbial communities of mud pools using traditional cultivation and isolation methodologies have resulted in the characterization of only a few microbes (e.g. Sulfolobus sp., Pyrococcus sp.) (Brock et
33 al., 1972; Stetter, 1986). Although effective at times, cultivation-based methodologies are unable to enrich and identify the majority of organisms present in the natural environment (Ward et al., 1990) and are limited by a dependence on the culture medium which may result in a biased, under representation of environmental microbial communities (Troussellier and Legendre, 1981). Developments in molecular genetic technologies have recently revolutionized our understanding of the diversity, distribution, and evolution of microorganisms in the environment and have lessened the need for cultivation (Pace et al., 1996; Ward et al., 1992). Molecular surveys based on the small subunit ribosomal RNA (16S rRNA) gene have revealed a wealth of previously unrecognized microbial diversity and new phylogenetic lineages (Amman et al., 1995; Ward et al., 1990; Giovannoni and Cary, 1993; Hugenholtz et al., 1998; Barns et al., 1996) and have been pivotal in understanding the evolutionary history of prokaryotes (Woese, 1987). Molecular-based technologies have been critical to studies of both terrestrial hot springs and deep-sea hydrothermal vents, where the use of cultivation and isolation has been problematic (e.g. Reysenbach et al., 1994; Reysenbach et al., 2000). Despite much success in other high temperature systems, molecular tools have not been applied to studying the microbial communities of high temperature mud pools. Molecular genetic fingerprinting techniques like denaturing gradient gel electrophoresis (DGGE) have proven successful when applied to community analyses of the natural environment (Fisher and Lerman, 1980; Myers and Maniatis, 1986; Muyzer et al., 1993), including both extremely high and low temperature environments (e.g. Ferris et al., 1996; Ward et al., 1998; Murray et al., 1998; Campbell and Cary, 2001). DNA fingerprinting techniques provide the ability to rapidly compare the dominant members of populations within an environmental microbial community. This method
34 allows for rough overall community level comparisons of a large number of samples with minimal DNA sequence retrieval independent of expensive cloning approaches. Now, these fingerprinting techniques are being utilized to study and compare microbial community composition and structure subjected to natural variations in the environment within and between habitats. Some intrinsic obstacles confront the application of molecular techniques to the study of high temperature mud pools. The solid matrixes of mud pools are rich in volcanic ashes, which have been reported to be high in silica (Baxter et al., 1999). These highly-charged silica particles have a high affinity for DNA (Vogelstein and Gillespie, 1979; Yamada et al., 1990; Zeillinger et al., 1993), making the recovery of nucleic acids problematic. Potentially low microbial biomass and the co-precipitation of inhibitors (e.g. humic acids, phenolic compounds, heavy metals, etc.) may disrupt downstream enzymatic processes (Tebbe and Vahjen, 1993; Wilson, 1997) necessitating the development of efficient nucleic acid extraction and recovery protocols specifically for mud pool samples. In order to effectively survey the microbial communities associated with high temperature mud pools using molecular technologies, methodologies must be developed and experimentally tested to ensure the most accurate, reliable, and consistent applications are employed in each individual step. In this study, we sought to characterize and correlate the physiochemical environment and bacterial community composition and structure of high temperature mud pools over both a regional (New Zealand) and geographical range of thermal systems. The Bacteria were the main focus of this initial comparative study due to the broad implementation of these methodologies in accessing microbial communities across many environments. Methods were developed for safe, aseptic collection of mud pool
35 samples, an efficient and reproducible method for the extraction of nucleic acids from these samples free of laboratory contamination, and determine the extent of exogenous inputs of DNA to the system. Using these methodologies, we performed community level comparisons of the bacterial communities of high temperature mud pools in correlation with the pool physiochemistry across a localized, relatively constrained zone of hydrothermal activity (Taupo Volcanic Zone (TVZ), New Zealand). Secondly, we performed bacterial community level analysis across 3 geographically isolated, distinct hydrothermal systems (Yellowstone National Park, USA; Lassen Volcanic National Park, USA; Rincon de la Vieja National Park, Costa Rica). Our results showed nucleic acids could be readily extracted from mud pool samples and the high temperature, acidic nature of most pools minimized signatures of exogenous, extracellular DNA within these systems. Analyses showed a broad range in the physiochemical nature of mud pools from the TVZ resulting in diverse bacterial communities. Although significant similarity existed between pools from different thermal areas both chemically and biologically, our analyses showed a high level of variability between individual thermal areas and within site heterogeneity of minor community members. A similar trend was observed for bacterial communities of mud pools from Yellowstone, Lassen, and the Rincon de la Vieja study sites. The dominance of 3 major bands across many profiles from different geographical sites was characterized as members of the alpha and gamma subdivisions of the Proteobacteria. These observations may be evident of endemic mud pool groups within the Proteobacteria. Although not examined here, the importance of the Archaea in high temperature systems is recognized and should be addressed in future studies of mud pool microbial communities.
36 Materials and Methods
Sample collection, preparation and field measurements Mud pool samples were collected from 24 pools from geothermal sites within the Taupo Volcanic Zone (13 pools) on the North Island of New Zealand in June of 2002, Rincon de la Vieja National Park (2 pools), Costa Rica in November of 2002, Lassen Volcanic National Park (2 pools), USA in July of 2003, and Yellowstone National Park (7 pools), USA in July of 2003. Samples were collected using a 5 meter retractable aluminum pool fitted with a stainless steel sampling arm configured to hold 4 sterile 50mL Falcon tubes and a HOBO H8 temperature data logger with an external probe (ONSET Scientific, Bourne, MA) for recording in-situ temperatures. Individual samples were taken and the temperature probe immersed for a minimum of one minute to allow the thermocouple to equilibrate and record multiple data points. Upon recovery, the pH of each sample was recorded with an Accumet portable AP5 pH meter fitted with an Accumet pH/Automatic Temperature Compensation probe (Fisher Scientific, Pittsburgh, PA). One sample tube from each pool (pH <6) was neutralized with 1 mL of a 10% (wt/v) sodium bicarbonate solution to minimize cell lysis during cooling (H. W. Morgan, personal communication). Pore water was collected on site by centrifugation (RCF = 1100 X g for 30 minutes) using a MobileSpin 12V field centrifuge (Vulcon Technologies, Grandview, MO). The pore water was filtered through a 0.2 µM Supor membrane Acrodisc syringe filter (Pall Corporation, Ann Arbor, MI), and placed into acid-washed polycarbonate vials. The resulting mud pool samples were stored at ambient temperature and pore water samples were frozen on dry ice for transport back to the laboratory.
37 Nucleic acid extraction optimization and recovery of extracellular DNA Due to the potential for co-precipitation of inhibitors and concern over the effects of high silica concentrations in the mud pool samples on nucleic acid extraction methodologies, several protocols were tested to ensure the most effective, consistent, contaminate-free method for the extraction of nucleic acids. The extraction protocols tested included the evaluation of three commercially available nucleic acid extraction kits and a modified cetyltrimethylammonium bromide-polyvinylpyrrolidone-β- mercaptoethanol (CTAB/PVP/β-ME) protocol (Coyne et al., 2001). The commercially available nucleic acid extraction kits evaluated in this study included: IsoQuick Nucleic Acid Extraction Kit (Orca Research, Inc.), QIAmp DNA Stool Mini Kit (Qiagen, Inc.), and the UltraClean Soil DNA kit (MoBio Laboratories, Inc.). All kit extractions were performed following the manufacturers’ protocols. Nucleic acid extractions were also performed on mud pool samples using a modified CTAB/PVP/β-ME extraction buffer and protocol (Coyne et al., 2001). Mud pool nucleic acid extracts were evaluated by quantification of nucleic acid yield via UV spectrophotometric scans, visualization of DNA on an agarose gel, success of PCR amplification of mud pool extract alongside a reagent blank, successful PCR amplification of small and large fragments of the 16S rRNA gene, and comparison of mud pool sample extract verse reagent blank via DGGE analysis. The CTAB/PVP/β-
ME extraction method proved to be the most efficient, clean, inhibitor-free protocol examined in this study and was thus used for all subsequent nucleic acid extractions of mud pool samples. Briefly, mud samples (4 g) were incubated with 10 mL CTAB buffer at 65ºC for 1 hour in a shaking water bath and then centrifuged for 1 min (2800 X g). The supernate was transferred to a new tube, an equal volume of chloroform: isoamyl alcohol (24:1) was added, and samples incubated at room temperature (RT) on an
38 inversion rocker for 20 minutes. Samples were centrifuged for 10 minutes (17000 X g) and the DNA in the aqueous phase was precipitated with an equal volume of isopropanol and 0.5 volumes of 5M NaCl at -80ºC for 2 hours. The precipitated DNA was centrifuged at 17000 X g for 20 minutes, the pellet washed with 70% ethanol, dried in a speed-vac at RT, and resuspended in 40 µL TE (10 mM Tris-Cl, pH 7.5; 1 mM EDTA).
The nucleic acids from each sample were quantified spectrophotometrically using a NanoDrop ND-1000 spectrophotometer at 260 nm (NanoDrop Technologies, Montchanin, DE). In order to address the potential for extracellular and exogenous DNA recovery from mud pool samples during the DNA extraction phase, experiments were designed to test the longevity of extracellular DNA under mud pool-like conditions. For these experiments, the wet weight of a volume of mud was determined followed by sterilization at 500°C overnight. The sterilized mud was reconstituted with sterile water, divided into three fractions of which one was titrated to pH 3.0, the second to pH 6.7, and the third to pH 8.0 for experimental use. For the initial experiment, 1g of mud was placed into 1.5 mL microfuge tubes (2 tubes of pH 3.0 mud and 2 tubes of pH 8.0 mud) and 500 ng of DNA was added to each tube of mud. Tubes were incubated under the following conditions for 1 hr: RT @ pH 3.0, RT @ pH 8.0, 65°C @ pH 3.0, and 65°C
@ pH 8.0). Samples were subsequently extracted using the CTAB extraction protocol described above and subjected to PCR amplification. Results of PCR were evaluated on a 1% agarose gel. A second experiment was performed to test the recovery of extracellular DNA was performed under higher temperature (90°C), near-neutral pH (pH
6.7) conditions for a maximum of 10 minutes. Setup was similar that described above and samples were incubated under the following conditions: RT @ pH 8.0 for 0 min, RT
39 @ pH 6.7 for 0 min, RT @ pH 8.0 for 10 min, RT @ pH 6.7 for 10 min, 90°C @ pH 8.0 for 10 min, 90°C @ pH 6.7 for 10 min. Samples were then extracted, amplified, and evaluated as described above.
Geochemistry (New Zealand regional survey) and water content determination Chemical analyses were performed on pore water samples for dissolved
+ - - 3- ammonium (NH4 ), nitrate + nitrite (NO3 + NO2 ), and orthophosphate (ΣPO4 ) using a Perstorp Analytical Flo-Through Analyzer (Perstorp Analytical, Wilsonville, OR).
+ Briefly, concentrations of NH4 were determined by the phenol hypochlorite method
- (Glibert and Loder, 1977; Grassfoff and Johansen, 1972). Concentrations of both NO3
- and NO2 were determined by the sulphanilamide/N(1-napthyl) ethylene diamine method
- - 3- following a cadmium reduction of NO3 to NO2 (Glibert and Loder, 1977). ΣPO4 concentrations were determined using the phospho-molybdenum blue method (Strickland and Parsons, 1972). Samples were run in duplicate, if possible, and dilutions were made to ensure measurement values were within the range of the standard curve for each analysis. The moisture content of the mud samples was determined using a standard thermogravimetric method (Wilson, 1971).
Bacterial community fingerprinting analysis The V3 region of the bacterial 16S rDNA genes of the mud pool microbial communities was amplified using the polymerase chain reaction (PCR) (Saiki et al., 1988) with primers 338F/GC clamp and 519RC as described by Muyzer et al. (1993). Reactions (50 µL) were performed using ~50 ng of template DNA and final concentrations of: 1X JumpStart PCR Buffer (Sigma, St. Louis, MO), 200 µM each deoxyribonucleoside triphosphate, 2mM MgCl2, 40 ng bovine serum albumin (BSA), 0.2
40 µM concentration of each primer, and 2 units of JumpStart Taq polymerase (Sigma).
The PCR reactions were performed on a PTC-200 Thermocycler (MJ Research, Inc., Waltham, MA) using the following hot start (D’Aquila et al., 1991), touchdown protocol (Don et al., 1991): 94ºC for 2 min, followed by 35 cycles of 94ºC for 1 min, 65-55ºC (- 0.5ºC per cycle) for 1 min, and 72ºC for 1min. The PCR products were stored at -20ºC until further analysis. Bacterial community composition was examined using denaturing gradient gel electrophoresis (DGGE) as described by Muyzer et al. (1993). Briefly, 20 µL of each PCR product (338FGC/519RC) was separated on an 8% acrylamide gel (37.5:1 ratio of arylamide-bisacrylamide) with a 25% to 65% gradient of denaturant (7M urea and 40% formamide) for 5 hours (130V) at 60ºC in 1X TAE buffer (40mM Tris base, 20 mM sodium acetate, 1 mM EDTA) using a DCode Universal Mutation Detection System (Bio-Rad, Hercules, CA). Gels were stained with ethidium bromide (500 ng/mL) for 30 minutes and destained in sterile water for 30 minutes. The gel was visualized using a UV transilluminator and digitally photographed using the AlphaImager 2000 imaging system (Alpha Innotech Corp., San Leandro, CA). Analyses of mud pool community samples run on DGGE gels was performed using the Phoretix 1D Version 5.10 software and the Phoretix 1D Version 1.13 Database (Nonlinear Dynamics Limited, UK). This software package was used for lane and band detection and lane relationship comparisons of amplicons for single and multiple gel comparisons. Individual bands, representing dominant amplicons within the gel profile common to multiple pools and those unique to individual pools, were excised and subject to sequencing and phylogenetic analyses.
41 Sequence analysis Individual DGGE bands were excised according to protocol reported by Individual DGGE bands were excised according to protocol reported by Campbell and Cary (2001) and subjected to reamplification (Muyzer et al., 1995; Murray et al., 1996) according to the PCR amplification conditions described above for 28 cycles. Reaction products were electrophoresed on a DGGE gel for confirmation of single bands. The PCR products of single bands were purified for sequencing using the GenElute PCR Clean-up Kit (Sigma-Aldrich Co., St. Louis, MO) according to the manufacturer’s instructions to remove excess primers and dNTPs and quantified by UV spectrophotometry. Purified products were subsequently sequenced with BigDye Terminator v1.1 Cycle Sequencing chemistry (Applied Biosystems Inc. (ABI, Inc.), Foster City, CA). Sequencing reactions (5 µL) were performed using 50 ng of purified template and 1.6 µmoles 519RC primer. Sequences were read using an ABI Prism 310
Genetic Analyzer automated sequencer (ABI, Inc.) Sequences were edited using Sequence Navigator and AutoAssembler software (ABI, Inc.). Sequence editing and alignments were performed using the SeqMan and MegAlign sequence manipulation programs (DNASTAR, Inc., Madison, WI). Sequences were subject to BLAST searches within the GenBank Database for comparison with submitted and characterized sequences. Final sequence alignments were created in the Genetic Data Environment (GDE version 3.2) (Smith et al., 1992) and phylogenetic relationships were determined to the subdivision level from the aligned sequences with Philip version 3.572 (Felsenstein, 1993). Neighbor-joining phylogenetic trees with 100 bootstrap replications were constructed alongside GenBank sequences for subdivision placement and relationship among known phylotypes.
42 Results
Field sampling and geochemistry Samples were collected from 28 mud pools from four geographically separated terrestrial based hydrothermal systems. These included 13 pools from seven thermal regions within the Taupo Volcanic Zone (TVZ), New Zealand (Table 2.1), 2 pools from the Rincon de la Vieja National Park, Costa Rica, 3 pools from Lassen Volcanic Park, USA, and 10 pools from Yellowstone National Park, USA (Table 2.2). Samples were collected at points of active bubbling and gas emission for a minimum of one minute to obtain a stable temperature in-situ temperature reading. For the our regional survey of high temperature mud pools from the TVZ, in-situ temperature measurements were obtained during sample collection displaying a range from 72.7°C (WW Pool #2) to 97.7°C (KP Pool #2) (Table 2.1). pH samples were performed immediately following collection resulting in a broad range in pH from 2.6 (Mud Volcano) to 6.8 (Tokaanu Mud Pool) (Table 2.1). Moisture content of each sample showed a broad range from 55% (Devil’s Cauldron) to 84% (CM Red Pool) (Table 2.1). These measurements were also performed for the other sites in our geographical study (Table 2.2). In-situ temperature measurements and pH measurements for the 2 mud pool samples collected in Rincon de la Vieja displayed temperatures of 94.1°C and 97.7°C and pHs of 5.2 and 4.0 respectively (Table 2.2). The 3 pools in Lassen Volcanic Park displayed temperatures ranges of 85°C to 89.3°C and a pH range of 2.9 to 3.6 (Table 2.2). The 10 pools from Yellowstone National Park displayed a temperature range of 76.7°C to 108.7°C and a pH range of 2.1 to 4.9 (Table
2.2). Of particular note are the extremely high temperatures and low pHs of the Crater
43 Table 2.1 Collection sites and physical parameters of New Zealand high temperature mud pools sampled during this study.
Thermal Area Pool Abbreviation Lat; Long (°C) pH Content (%)
Hell's Gate Devil's Cauldron HG DC 38° 03' S; 176° 21' E 94.1 2.77 55.7 Hell’s Gate Mud Volcano HG MV 38° 03' S; 176° 21' E 95.9 2.60 65.2 Kuirau Park KP Pool #1 KP #1 38° 08' S; 176° 14' E 77.8 4.10 58.9 Kuirau Park KP Pool #2 KP #2 38° 08' S; 176° 14' E 97.7 5.60 71.3 Whakarewarewa WW Pool #1 WW #1 38° 09' S; 176° 15' E 83.8 2.90 70.5 Whakarewarewa WW Pool #2 WW #2 38° 09' S; 176° 15' E 72.7 2.90 57.4 Orakei Korako OK Pool #3 OK #3 38° 28' S; 176° 09' E 82.5 2.30 79.5 Orakei Korako OK Pool #4 OK #4 38° 28' S; 176° 09' E 92.4 2.30 76.4 Craters of the Moon CM Red Pool CM RP 38° 38' S; 176° 03' E 86.4 2.90 84.4 Craters of the Moon CM Grey Pool CM GP 38° 38' S; 176° 03' E 90.8 2.88 82.6 Waitaupo WTP Pool WTP 38° 20' S; 176° 22' E 89.3 2.85 68.6 Waitaupo WTP Mud Volcano WTP MV 38° 20' S; 176° 22' E 97.7 3.47 71.3 Tokaanu TK Pool TK 38° 58' S; 175° 45' E 90.8 6.77 83.8
44 Table 2.2 Collection sites and physical parameters of Costa Rica, Lassen, and Yellowstone high temperature mud pools sampled during this study.
Temperature Moisture Country/Site Thermal Area Pool Lat; Long (°C) pH Content (%)
Costa Rica Rincon de la Vieja RV #3 10° 02' N; 85° 05' W 94.1 5.15 67.4 Costa Rica Rincon de la Vieja RV #4 10° 02' N; 85° 05' W 97.7 4.00 76.4 Lassen, USA Sulfur Works SW 40° 29' N; 121° 30' W 85.1 2.97 66.8 Lassen, USA Boiling Springs BS #2 40° 29' N; 121° 30' W 87.8 3.62 92.3 Lassen, USA Devils Kitchen DK #1 40° 29' N; 121° 30' W 89.3 2.97 63.1 Yellowstone, USA Lower Geyser Basin LGB #1 44° 31' N; 110° 48' W 87.8 3.45 77.1 Yellowstone, USA Lower Geyser Basin LGB #2 44° 31' N; 110° 48' W 85.1 2.85 46.3 Yellowstone, USA Lower Geyser Basin LGB #3 44° 31' N; 110° 48' W 83.8 3.50 53.7 Yellowstone, USA Pots Hot Springs PHS #1 44° 26' N; 110° 34' W 76.8 4.86 68.4 Yellowstone, USA Mud Volcano MV #1 44° 37' N; 110° 26' W 77.8 3.76 79.3 Yellowstone, USA Crater Hills CH #1 44° 39' N; 110° 28' W 108 2.40 50.8 Yellowstone, USA Crater Hills CH #2 44° 39' N; 110° 28' W 104 2.07 30.2 Yellowstone, USA Artist’s Paint Pots APP #1 44° 41' N; 110° 44' W 80.1 3.26 63.9 Yellowstone, USA Fountain Flats FF #1 44° 33' N; 110° 53' W 85.1 3.06 47.6 Yellowstone, USA Fountain Flats FF #2 44° 33' N; 110° 49' W 83.8 3.15 48.6
45 Hills pools collected in Yellowstone National Park (Table 2.2). The moisture content of all the mud pool samples collected from these study sites was also determined for comparison. The Rincon pools showed moisture contents of 67 and 76%; the Lassen pools showed moisture contents of 63 to 93%; and the Yellowstone pools showed moisture contents of 30 to 79% (Table 2.2). For the New Zealand regional survey, a portion of the pore water from each mud pool sample was collected for chemical
+ analyses (Table 2.3). Ammonium (NH4 ) was high for all of the TVZ mud pools sampled with concentrations ranging from 34 µM to >1000 µM. Nitrate and nitrite
- - (NO3 + NO2 ) was extremely low in comparison for these pools ranging from 0.04 µM
3- to 1.53 µM. Phosphate (ΣPO4 ) concentrations ranged between 0.8 and 7.1 µM for the
3- TVZ mud pools with one exception. KP Pool #2 displayed a significantly higher ΣPO4 concentration (52.9 µM). Although variable between different thermal areas, we observed some degree of similarity between mud pools from the same thermal area.
Nucleic acid extraction optimization In order to determine the most effective, reproducible means of extracting high quality nucleic acids from mud pool samples, 4 extraction protocols were tested and evaluated (Table 2.4). The criteria used for determining nucleic acid efficiency and purity were the 260/280 ratio as examined by UV spectrophotometry, the positive amplification of a small DNA fragment (181 bp) using universal bacterial primers, the positive amplification of a large DNA fragment (1495 bp) using universal bacterial primers, and laboratory contamination via evaluation of a reagent blank extraction. Two of the commercially available kits, the QIAmp DNA Stool Mini kit and the UltraClean Soil DNA kit provided some degree of success in extracting DNA from mud pool
46 Table 2.3 Chemical analyses of pore water from New Zealand mud pools.
+ - - 3- Collection Site/ Temperature pH NH4 NO3 + NO2 PO4 Pool (°C) (µM) (µM) (µM) a
Hell’s Gate Devil’s Cauldron 94.1 2.77 >1000 0.51 3.01 Mud Volcano 95.9 2.60 >1000 0.79 2.92 Kuirau Park KP Pool #1 77.8 4.10 658 1.53 2.27 KP Pool #2 97.7 5.60 522 1.40 52.9 Whakarewarewa WW Pool #1 83.8 2.90 364 0.23 7.07 WW Pool #2 72.7 2.90 352 0.31 4.50 Orakei Korako OK Pool #3 82.5 2.30 60.5 0.56 1.95 OK Pool #4 92.4 2.30 34.5 0.35 1.29 Craters of the Moon CM Red Pool 86.4 2.90 698 0.93 1.30 CM Grey Pool 90.8 2.88 844 0.72 1.58 Waitaupo WTP Pool 89.3 2.85 416 0.85 0.89 WTP Mud Volcano 97.7 3.47 228 0.04 2.61 Tokaanu TK Pool 90.8 6.77 487 1.28 4.57
47 samples. Both of these methods produced amplifiable DNA in the < 200bp size range. However, the UltraClean Soil DNA kit yielded DNA of low purity based on 260/280 ratios and low molecular weight based on the inability to amplify larger fragments (> 1200bp). Reagent blanks from both these kits showed positive amplification using universal bacterial primers 338F GC/519RC, indicating the presence of bacterial contamination within the kit component reagents. The IsoQuick Nucleic Acid Extraction kit proved to be a poor extraction method for mud pool samples, yielding no quantifiable or amplifiable DNA. The modified CTAB/PVP/β-ME method proved to be the best extraction method utilized here by scoring favorably in all of the evaluation criteria. This method showed a high level of purity based on the 260/280 ratios, small and large gene fragments were easily amplifiable using universal bacterial primers, and the reagent blanks were found to be free of contamination. Replicate extractions using this protocol were amplified using bacterial DGGE primers and run on a DGGE gel to confirm the reproducibility of the DGGE banding profile. The CTAB/PVP/β-ME method was subsequently used for all mud pool nucleic acid extractions. Due to the potential inputs and subsequent recovery of extracellular and exogenous DNA not representative of viable, resident mud pool bacteria, we devised experiments to test the liability of exogenous, extracellular DNA under controlled, mud pool-like conditions (Figures 2.1A and 2.1B). The initial experiment showed that extracellular DNA could not be amplified using our standard protocol following incubation in a low pH (pH 3.0) buffer at a moderately high temperature (65°C) for a duration of 1hr (Figure 2.1A). When incubated in near neutral pH buffers (pH 6.7 and pH 8.0) at ambient and high temperatures (90°C) for a shorter duration (10min) (Figure
48 Table 2.4 Efficiency of nucleic acid extraction protocols on high temperature mud pool samples.
338F GC/519RC 27F/1522R 260/280 nma PCR Amplificationb PCR Amplificationc Contaminationd IsoQuick Nucleic Acid N/A _ _ N/A Extraction Kit (Orca Research, Inc.) QIAmp DNA Stool 1.6 + + + Mini Kit (Qiagen, Inc.) UltraClean Soil DNA kit 0.9 + _ + (MoBio Laboratories, Inc.)
CTAB/PVP/ -ME protocol 1.6 + + _ (Coyne et al., 2001) a 260/280 nm value based on quantification via UV spectrophotometry b PCR amplification of 181 bp region of the Bacterial 16S rRNA genes with primers 338FGC/519RC c PCR amplification of 1495 bp region of the Bacterial 16S rRNA genes with primers 27F/1522R d Presence of contamination based on comparison of 338FGC/519RC primed PCR product of extracted sample with a reagent blank via DGGE
49 MW 1 2 3 4 + - MW 1 2 3 4 5 6 + - A B
Figure 2.1 A. DNA was exposed to low pH and high temperature (65º C) conditions for 1 hour, amplified using PCR, and run on a 1% agarose gel. 1: Control, 2: DNA, pH 3, 65º C, 3: DNA, 65º C, 4: DNA, pH3. B. DNA was exposed to neutral pH and high temperature (90º C) conditions for 10 minutes, amplified using PCR, and run on a 1% agarose gel. 1: T0, pH 8.0; 2: T0, pH 6.7; 3: pH 8.0, RT; 4: pH 6.7, RT; 5: pH 8.0, 90º C; 6: pH 6.7, 90º C. 2.1B), no amplification was obtained regardless of pH. Any extracellular DNA appeared to be partially denatured and degraded under the high temperature, abrasive conditions.
Bacterial community fingerprinting analysis Bacterial community level analyses of mud pool samples were performed using DGGE. For the New Zealand regional survey of the TVZ, the bacterial communities of 13 mud pools were compared. The DGGE profiles of these 13 pools resulted in the detection of a total of 155 bands occupying 39 unique positions in the gel (Figure 2.2). Most notably was the presence of 3 dominant bands (designated 6, 7, 12; Figure 2.2) common across a majority of the pools with a higher degree of variation present in the minor amplicons. Band 6 was present in 6 of the 13 profiles, band 7 was present in 12 of the 13 profiles, and band 12 was present in 7 of the 13 profiles. The profiles of mud pools Whakarewarewa #1 and Waitaupo showed a more diverse banding
50 pattern over the other profiles. The Tokaanu mud pool profile displayed a unique banding profile with 4 dominating bands not observed in any of the other New Zealand mud pools examined. A UPGMA derived dendrogram of the DGGE banding profiles from these 13 mud pools was generated using similarity (presence or absence of band matches) across the mud pool profiles (Figure 2.3). Highest similarities were observed between pools from the same thermal area. Fourteen bands of representing dominant bands within the profiles were excised and sequenced (Table 2.5). Sequence analyses of the three dominant bands (designated 6, 7, 12; Figure 2.2, Table 2.5) were found to affiliate with uncultivated thermophilic and acidophilic Acinetobacters within the gamma Proteobacteria. Four bands (designated 1, 10, 13, 14; Figure 2.2, Table 2.5) were affiliated with the Cytophaga-Flavobacteria, 3 bands (designated 2, 4, 8; Figure 2.2, Table 2.5) were affiliated with Methylobacteria within the alpha Proteobacteria, two bands (designated 3, 11; Figure 2.2, Table 2.5) were representative of thermophilic members of the Firmicutes (Bacilli), and one band (designated 9; Figure 2.2, Table 2.5) was representative of thermophilic epsilon Proteobacteria. Overall, sequence analysis revealed a high representation of the Proteobacteria and Flavobacteria present in the New Zealand mud pool bacterial communities. Bacterial community level analysis using DGGE was also performed on mud pool samples from Rincon de la Vieja National Park, Costa Rica, Lassen Volcanic Park, USA, and Yellowstone National Park, USA. The samples run on this gel represented 2 mud pools from Rincon, 3 mud pools from Lassen, and 10 mud pools from Yellowstone. Due to the unique bacterial community associated with the Tokaanu
51 HG HG KP KP WW WW OK OK WTP CM CM TK DC MV #1 #2 #1 #2 #3 #4 WTP MV RP GP
11 9 7 5 12 1 6 13 10 2 8 14 3 4
Figure 2.2 DGGE analysis of bacterial communities associated with 13 mud pools located in the Taupo Volcanic Zone, North Island, New Zealand. Numbers designate bands that were excised, re- amplified, and subjected to sequence analyses.
52 mud pool sample, it was also run alongside these other pools for comparative purposes. The DGGE profiles from these 16 mud pools resulted in the detection of 125 bands occupying 19 unique positions (Figure 2.4). The banding profiles of the mud pools showed extreme heterogeneity between geographically isolated thermal sites with a higher degree of similarity between pools from the same thermal site. Three dominant bands (designated 4, 5, 8; Figure 2.4) were prevalent in the majority of mud pool samples from Yellowstone study sites. Bands 4 and 5 were present in 8 of the 10 Yellowstone profiles, while band 8 was seen in all 10 profiles from this thermal region. Band 8 was also observed in the profiles of the 3 mud pool samples collected from Lassen Volcanic Park. BLAST analysis of these dominant bands showed affiliation with thermophilic bacterium from sulfur-rich hot springs and hydrocarbon metabolizers within the gamma Proteobacteria (Table 2.6). Three other prominent bands (designated 3, 7, 11; Figure 2.4) were affiliated with uncultivated hot spring bacteria within the alpha Proteobacteria. Bands 10 and 12 were affiliated within the Cytophaga-Flavobacteria and band 6 was affiliated with hyperthermophilic members of the Thermotogales. The two mud pools from Rincon de la Vieja displayed almost identical DGGE profiles to one another yet unique in comparison to the other study sites. The profiles from the 3 Lassen pools displayed complex banding patterns with no particular bands dominating the profiles. Overall, the majority of variation between individual pools was most notable in the more minor bands. The banding profiles were used to create a UPGMA dendrogram (Figure 2.5). Similarity distance in the individual profiles showed the two mud pools from Rincon de la Vieja (RV #3 and #4) and 1 pool from Lassen (SW) to be quite unique in comparison to the other pools on the gel. This analysis also classifies the Rincon de la Vieja pools as having identical banding profiles.
53 CM GP CM RP WTP MV WTP KP #1 HG MV HG DC OK #4 OK #3 KP #2 WW #2 WW #1 TK
Figure 2.3 Phoretix comparative analysis of New Zealand mud pool bacterial DGGE analyses. UPGMA dendogram displays similarities between the bacterial communities based on the presence and absence of bands in the profiles.
54 Table 2.5 Sequence and BLAST results of excised bands from the DGGE analysis of New Zealand mud pool bacterial communities.
Length of Most closely related Band Sequence Accession no.a Phylum / Class Score Expected
1 120 AY711798 Cytophaga-Flavobacteria-Bacteroides 798 7e-58 2 117 AY453855 Proteobacteria / Alpha 220 5e-55 3 121 AJ229238 Firmicutes / Bacilli 248 3e-63 4 126 AY741717 Proteobacteria / Alpha 99 1e-18 5 122 AF500327 Proteobacteria / Gamma 230 7e-58 6 120 AJ633639 Proteobacteria / Gamma 214 3e-53 7 121 AF467302 Proteobacteria / Gamma 236 1e-59 8 117 AY453855 Proteobacteria / Alpha 220 5e-55 9 121 AY553059 Proteobacteria / Epsilon 220 5e-55 10 120 AY145539 Cytophaga-Flavobacteria-Bacteroides 287 3e-75 11 120 AY675242 Firmicutes / Bacilli 299 9e-79 12 119 Z93450 Proteobacteria / Gamma 276 1e-71 13 119 AY711798 Cytophaga-Flavobacteria-Bacteroides 272 2e-70 14 119 AY711798 Cytophaga-Flavobacteria-Bacteroides 272 2e-70 a Accession number of closest related sequence from the GenBank database.
55 RV RV DK BS LGB LGB LGB PHS MV CH CH APP FF FF #3 #4 SW #1 #2 #1 #2 #3 #1 #1 #1 #2 #1 #1 #2 TK
4
8 12
1 5 3 9 11
10 7 2 6
Figure 2.4 DGGE analysis of bacterial communities from 16 mud pools from New Zealand, Costa Rica, Lassen, and Yellowstone. Numbers designate bands that were excised, re-amplified, and subjected to sequence analyses.
56 Table 2.6 Sequence and BLAST analyses of excised bands from DGGE analysis of mud pool bacterial communities from New Zealand, Costa Rica, Yellowstone, and Lassen.
Length of Most closely related Band Sequence Accession no.a Division / Subdivision Score Expected
1 131 AY292867 Proteobacteria / Gamma 270 1e-69 2 139 AY463827 Proteobacteria / Gamma 287 4e-75 3 134 AY293404 Proteobacteria / Alpha 266 1e-68 4 128 AY298741 Proteobacteria / Gamma 347 5e-93 5 129 AY369139 Proteobacteria / Gamma 329 1e-87 6 142 U05660 Thermotogales 180 7e-43 7 134 AJ626893 Proteobacteria / Alpha 262 2e-67 8 130 AY327170 Proteobacteria / Gamma 309 1e-81 9 128 AY551938 Proteobacteria / Gamma 325 2e-86 10 140 AY712194 Cytophaga-Flavobacteria-Bacteroides 206 7e-51 11 135 AY293404 Proteobacteria / Alpha 262 2e-67 12 137 AY711798 Cytophaga-Flavobacteria-Bacteroides 285 2e-74 a Accession number of closest related sequence from the GenBank database.
57 TK PHS #1 BS #2 DK #1 FF #2 LGB #1 FF #1 MV #1 CH #2 CH #1 LGB #2 APP #1 LGB #3 SW RV #4 RV #3
Figure 2.5 Phoretix comparative analysis of New Zealand, Costa Rica, Lassen and Yellowstone mud pool bacterial DGGE analyses. UPGMA dendogram displays similarities between the bacterial communities based on the presence and absence of bands in the profiles.
58 Discussion High temperature mud pools are prevalent, common surface features of terrestrial hydrothermal activity, yet there is little known about the bacterial communities that reside in these dynamic, extreme ecosystems. The uses of isolation and cultivation- independent technologies to explore prokaryotic community structure within the mud pool system have been employed in past studies (e.g. Glamoclija et al., 2004). In the present study, we used more sensitive modern molecular techniques to obtain a more complete picture of the diversity of the bacterial communities of these high temperature mud pools over both a regional and geographically-isolated range of terrestrial hydrothermal systems. For the regional survey, characterization of the physiochemical environment was performed alongside bacterial community analyses in attempts to correlate the physical environment with structure of the resident bacterial community. In order to perform an effective survey of the bacterial communities of high temperature mud pools, an efficient nucleic acid extraction protocol had to be developed and tested on a range of mud pool samples. Because of the solid/fluid matrix and predominantly silica composition of the mud, methods for the extraction of DNA from soils were investigated for potential extraction methods. DNA extraction from samples containing soils and other particle-based medium have traditionally involved one of two methods, cell extraction or fractionation (separate cells from solid matrix prior to cell lysis) (Torsvik, 1980; Holben et al., 1988; Steffan and Atlas, 1988; Pillai et al., 1991; Jacobsen and Rasmussen, 1992) or direct lysis of cells within the solid matrix (Ogram et al., 1987; Steffan et al., 1988). Although both methods have been proven successful in the extraction of DNA from soil-based samples, direct lysis techniques have shown higher DNA yields and are considered to provide a less biased representation of
59 microbial community diversity (Holben et al., 1988; Leff et al., 1995; Steffan et al., 1988). Extraction efficiencies of direct lysis have been suggested to be > 60% of total bacterial DNA with recoveries of 86% to 100% having also been reported (More et al., 1994; Miller et al., 1999). This is compared to only about 25% to 50% of total bacterial community DNA for cell extraction/fractionation methods (Bakken and Lindahl, 1995). Based on previous studies, we decided to pursue direct lysis methods for the extraction of DNA from mud pool samples in order to minimize sample biases and to provide a more thorough evaluation of the indigenous bacterial communities. Four potential direct lysis extraction protocols were subsequently tested on mud pool samples collected in New Zealand including three chemical based protocols (IsoQuick Nucleic Acid Extraction Kit (Orca), QIAmp DNA Stool Mini Kit (Qiagen), and a CTAB/PVP/β-ME protocol) and 1 mechanical based protocol (UltraClean Soil
DNA Kit MoBio). DNA extraction from mud pool samples using the IsoQuick Nucleic Acid Extraction Kit yielded no quantifiable or amplifiable DNA. The failure of this method may have been attributed to the inability of the kit reagents to effectively lyse the microbial cells or more likely, the entire extracted DNA was lost by absorption to particles present in the sample. High losses of DNA during extraction have been reported for clay-containing soils due to strong binding of DNA with clays (Greaves and Wilson, 1969; Ogram et al., 1988; Frostegard et al., 1999). This binding has been observed to be significantly greater in soils with clays containing high allophane (a group of small clay minerals containing silica, alumina, and water (Parfitt 1990) content like many soils in New Zealand (Saggar et al., 1996; Lloyd-Jones and Hunter, 2001). The QIAmp DNA Stool Mini Kit proved to be an effective DNA extraction technique for mud pool samples in terms of relatively pure, amplifiable DNA. However, contamination
60 was discovered in reagent blanks of these kits upon amplification using universal bacterial primers. In this method, extracted DNA is purified and concentrated away from the extracted medium and reagents via a purification column. The observed contamination using this kit was consistent with a recent report noting the presence of bacterial DNA contaminates in the purification columns of Qiagen DNA extraction kits (van Der Zee et al., 2002). Another commercially available DNA extraction kit targeting soils via mechanical cell lysis (bead-beating), the UltraClean Soil DNA kit, was tested on mud pool samples. Test results showed recovery of DNA of low purity and low molecular weight as evidenced by the successful amplification of only small target fragments. Two of the major issues associated with bead-beating homogenization and extraction of DNA is the co-extraction of higher amounts of inhibitory humic acids (Leff et al., 1995; Ogram et al., 1987; Smalla et al., 1993) and the potential for sheering of the DNA (Leff et al., 1995). Reagent blanks of the UltraClean Soil kit’s components also revealed the presence of bacterial contamination. A fourth DNA extraction protocol based on a modified CTAB/PVP/β-ME protocol (Coyne et al., 2001) was tested on mud pool samples. DNA recovered using this method was of high purity, high molecular weight, and free of reagent bacterial contaminants. The use of CTAB in extraction buffers is highly beneficial due to the formation of insoluble complexes between CTAB, denatured proteins, polysaccharides, and other cellular debris (Saano et al., 1995). The addition of CTAB to the extraction buffer has also been reported to increase DNA recovery from soil samples and improve the purity of this DNA by binding inhibitory organic-based compounds (Knaebel and Crawford, 1995; Saano et al., 1995; Xia et al., 1995; Lee et al., 1996; Zhou et al., 1996; Chandler et al., 1997; Porteous et al., 1997). This CTAB protocol also contained an organic solvent extraction step using chloroform,
61 which has been reported to increase DNA yields over methods in which organic solvents are omitted (Miller et al., 1999). Our confidence in the effectiveness and performance based on our criteria of the CTAB/PVP/β-ME protocol in the extraction of DNA from mud pool samples led to its use in all subsequent extractions from mud pool samples. The co-extraction of unknown quantities of exogenous or extracellular DNA in conjunction with DNA from viable cells is common with environmental samples (Robe et al., 2003) and was an area of concern for the determination of true bacterial signatures from the mud pool samples. In order to address this issue, we experimentally tested the residence time of extracellular DNA under controlled, mud pool like conditions. The results of these experiments suggested that residence time for extracellular DNA under conditions of extreme temperature and/or pH is relatively short. The combination of moderately high temperatures (65°C) and low pH (pH 3.0) showed extracellular DNA degradation to occur in less than 1 hr as determined by amplifiability of the 16S rDNA. Our results also showed that extracellular DNA was degraded beyond 16S rDNA amplification in mud samples at extreme temperatures (90°C) independent of pH in <10 min. Although studies have shown that extracellular DNA in marine sediments can be up to 4.3 times higher than concentrations of DNA associated with total bacterial cells (Dell’Anno and Corinaldesi, 2004), other sediment studies have shown that recovered pools of extracellular DNA did not contain amplifiable 16S rDNAs (Corinaldesi, et al., 2005). The inability to amplify 16S rDNA signatures from our mud pool simulation experiments supported previous findings and instilled confidence in our obtaining 16S rDNA signatures from viable members of the mud pool bacterial community for our comparative and phylogenetic downstream analyses.
62 The temperature of 13 mud pools sampled from thermal fields in the TVZ, New Zealand, ranged from 77.8°C to 97.7°C and pH ranged from 2.3 to 6.77.
Measurements of pH for the individual mud pools showed similarity among the majority of mud pools collected from the same thermal field. However, this trend was not observed in measurements of pool temperature with the exception of the Hell’s Gate features. Various factors could potentially have had a large effect on these elements in any given thermal field including the source of hydrothermal fluids, the distance traveled form the source, localized mineralogy, and mixing with other groundwaters. The geothermal systems of the TVZ have been found to form two distinct groups, a high-gas,
CO2 rich system along the eastern rim and a low-gas system dominated by groundwaters to the west based on the chemical composition of the discharged fluids (Giggenbach, 1986; Giggenbach, 1992; Giggenbach, 1995). All of the thermal fields sampled for this study occupied intermediate locations within the TVZ, previously characterized as displaying low-gas, groundwater dominated fluids, with the exception of Waiotapu which has shown characteristics a high-gas, groundwater influenced system (Giggenbach, 1995). The acidic conditions of the majority of the mud pools from this survey are the result of CO2, H2S, and/or SO4 that rise from depth with steam and mix with shallower groundwaters (Henley and Ellis, 1983; Giggenbach, 1987). The Tokaanu mud pool from the Tokaanu Thermal area was the only near-neutral feature examined in our survey and was consistent with previous reports of an independent high temperature, near- neutral hydrothermal source discharging into this thermal field (Robinson and Sheppard, 1986).
+ Alongside measurements of temperature and pH, measurements of NH4 ,
- - 3- NO3 + NO2 , and ΣPO4 were performed for the 13 mud pool samples from the TVZ.
63 A high degree of variability was observed for these analyses both within site and across different thermal fields. However, individual pools within a given thermal field showed general trends in concentrations of these nutrients, mainly high micromolar
+ - - concentrations for NH4 and low micromolar concentrations of NO3 + NO2 across all
+ pools. High concentrations of NH4 concentrations have been observed for thermal fluids associated with hot springs from Yellowstone at micromolar to millimolar concentrations (Reysenbach and Shock, 2002). Analysis of the mud pool bacterial communities in correlation with these geochemical signatures showed similarity between
+ pools with NH4 concentrations > 600 µM Hell’s Gate Devil’s Cauldron and Mud Volcano, Kuirau Park pool #1, Craters of the Moon Red Pool and Grey Pool). The
+ variability in the temperature and pH of these high NH4 pools suggested the other
+ chemical elements might play a larger role in dictating community structure. Low NH4 concentrations were observed for mud pools from the Orakei Korako thermal field in relation to the other pools from the TVZ. These low concentrations are consistent with low concentrations observed by Rogers et al. (2000) for this thermal field. Similarities
+ were observed in the bacterial communities, NH4 concentrations (relative to other TVZ mud pools), and pH values for the Orakei Korako pools. Although the chemical signatures of the mud pools of the TVZ are highly variable, there was some evidence linking the bacterial community structure to the physiochemical environment. More in- depth analyses of other chemical species are needed for a better understanding of geochemical-bacterial interactions and metabolic potential in these systems. The assessment of the mud pool bacterial communities from these 13 TVZ mud pools was performed using DGGE. Profiles showed 3 dominating bacterial signature bands (designated 6, 7, 12; Figure 2.2) across a majority of the pools with
64 greater heterogeneity observed among minor community members. Taxonomically, the dominating bands were from three members of the gamma Proteobacteria. Members of the gamma Proteobacteria have been described in high temperature, low pH (pH 2.0) hot springs of Indonesia (Baker et al., 2001). The presence of these gamma Proteobacteria signatures across multiple pools suggests a tolerance across a range of temperature, pH, and chemical signatures. Studies of microbial communities from geothermal springs have shown tight coupling between the community structure and temperature and pH (Atkinson et al., 2000; Burton and Norris, 2000). While this may be true for the microbial community as a whole, our results suggested some bacteria or bacterial groups have a more loose association with the chemical environment. This observation became more prevalent as one group of gamma Proteobacteria, indicated “7” in the DGGE profile (Figure 2.2), was present in 11 of the 13 TVZ mud pool profiles. In fluctuating habitats, organisms that exhibit the ability to tolerate large variations in geochemistry and capable of utilizing a variety of electron donors and acceptors would have a competitive advantage (Reysenbach and Shock, 2002). Band 6, a dominant representative of the gamma Proteobacteria in our DGGE analysis, was present in 6 of the 13 pools from the TVZ. All 6 of these pools were collected from thermal areas (Hell’s Gate, Kuirau Park, and Waitapo) located in and around the extremely thermally active region of Rotorua.
+ These mud pools showed a 20°C temperature range, a pH range of 2.6 to 5.6, and NH4 concentrations of < 1000 µM to ~200 µM. The presence of this bacterial signature across all of these pools suggested a high tolerance for a range of environmental conditions and the proximately of this signature only in these pools may be an indication of a shared deep hydrothermal source undergoing physiochemical alteration as the fluids near the point of discharge. Additional analyses of the hydrothermal fluids beyond those
65 presented here are needed to fully explore the potential for the existence of shared hydrothermal source for the Rotorua region. Taxonomic analyses also revealed members of the Cytophaga- Flavobacteria present in pools of variable temperature and pH. Members of his group of bacteria (designated 1, 10, 13, 14; Figure 2.2) were seen in the high temperature, near neutral Tokaanu mud pool and the high temperature, low pH Craters of the Moon Grey Pool. Although three of four bands defined through BLAST were closest to the same database sequence, the migration of these bands on the DGGE gel suggested that these four bands represented 3 different members of the Cytophaga-Flavobacteria. Although members of the Flavobacteria have been described previously in a 90°C, pH 2 hot spring (Baker et al., 2001), our results indicated a tolerance for lower temperatures and near-neutral conditions for this subgroup. Members of the alpha Proteobacteria were also observed in the profiles of 4 of the TVZ mud pools. Indicated by bands 2, 4, and 8 (Figure 2.2), our analysis showed this group to be represented by two different members. The alpha Proteobacteria represented by band 4 was only observed in the high temperature, near-neutral TK Pool while those represented by bands 2 and 8 (likely the same band position for both) were observed in pools ranging from pH 2.9 to pH 6.8 and temperatures of 73°C to 98°C. The presence of members of this group across a range of temperature and pH regimes was consistent with other observations of the presence of the alpha Proteobacteria across a thermal gradient in hot springs (Hugenholtz et al., 1998) and gradients of temperature and pH in hot mud and Solfatara crusts (Johnson et al., 2003; Glamoclija et al., 2004). Bands 2 and 8 in the Whakarewarewa Pool #2 profile suggested the temperature of this pool (73°C) was not
66 as optimal as the higher temperatures of the other pools in which these bands were observed. Two bands (3 and 11; Figure 2.2) representing members of the Firmicutes were observed in our community profiles of pools from Tokaanu, Whakarewarewa (WW #1), Waitaupo (WTP), and Craters of the Moon (CM RP). The Firmicutes have been previously described in moderate and high temperature hot springs (e.g. Sharp et al., 1992; Baker et al., 2001), other moderate temperature, acidic geothermal features (Johnson et al., 2003), and subterranean hot springs (Baker et al, 2001). All of the mud
+ pools that had Firmicutes had NH4 concentrations above 350 µM. However, not all
+ pools with these NH4 concentrations and above showed the presence of this group. In the Whakarewarewa #2 mud pool, the most prominent band (band 9) in the community profile was an epsilon Proteobacterium. Members of the epsilon Proteobacteria have been previously described in other extreme environments like deep-sea hydrothermal vents (e.g. Haddad et al., 1995; Reysenbach et al., 2000) and hot springs (Huggenholtz et al., 1998) and may be involved in sulfate reduction (Phelps et al., 1998). Cladistic analysis of the TVZ mud pool profiles showed varying degrees of similarity both between and within a given thermal area (Figure 2.3). Nine of the 13 pools in this regional analysis grouped together to form a large cluster that was subsequently broken into three subclusters. One of the subclusters contained pools WTP and WTP MV, both located within the same thermal field in close proximately to one another. Although the physiochemical signatures measured for this study from these two pools were quite different, these pools probably shared the same source of hydrothermal fluid. The second subcluster contained pools HG DC, HG MV, CM RP,
+ and KP 1. These pools all showed NH4 concentrations > 650 µM. These were the
+ highest NH4 concentrations observed in the pools from this study. The third subcluster
67 of TVZ mud pools based on similarity of banding patterns contained pools KP 2, WW 2, OK 3, and OK 4. The OK 3 and 4 pools showed a higher degree of similarity to one another when compared to the other pools in this cluster. This was not surprising as
+ these pools are from the same thermal field and both exhibit very low NH4 concentrations in comparisons to the other TVZ pools sampled here. The loose similarity of the other two pools of varying physiochemical characteristics in this cluster suggested the presence of bacterial communities containing members with the ability to tolerate a broad range of temperatures, pH, and geochemical conditions. The three major outlying pools in this similarity index are the TK Pool and CM GP. The banding profiles of both of these pools showed unique structure compared to the other NZ pools. The distinct profile exhibited by the TK Pool was expected due to the near-neutral pH of this pool. Although, some similarities were seen across a majority of the mud pools from the TVZ, each individual pool displayed a distinct bacterial community different from the other pools. The presence of certain bacterial signatures across a broad range of mud pools displaying a broad range of physiochemical conditions was somewhat unexpected but may be contributed to the unique nature of the mud pool system. While the genetic diversity observed in hot springs correlates directly with the geochemical diversity of the sites (Reysenbach and Shock, 2002), the mud pools in our survey displayed a narrow range of dominant bacterial diversity over a diverse array of geochemical and physical parameters. Global survey: Along with the regional survey of the bacterial communities of high temperature mud pools from the TVZ, a geographical survey was performed over across three other continental hydrothermal systems. The purpose of this phase of our
68 study was for direct comparison of bacterial signatures and geochemistry of mud pools from isolated hydrothermal systems for predictions of physiochemical controls of community structure and endemism. The bacterial communities of mud pools from the thermal fields of Rincon de la Vieja, Lassen, and Yellowstone were compared on the basis of temperature, pH, and bacterial community structure via DGGE. Overall, the temperature and pH measurements for the mud pools collected from these sites showed less variability within a given thermal field than those observed for the pools in the TVZ. Samples have been collected for geochemical analyses from these sites and will supplement temperature and pH data for more in-depth characterization of the physiochemical environment. The DGGE bacterial community profiles of pools from different geographical sites, as expected, were not similar. However, sequence analysis of excised bands once again showed dominance by the same bacterial groups as seen in the TVZ. The gamma Proteobacteria were observed in all three geographical locations and dominated the profiles of mud pools from Rincon de la Vieja and Yellowstone The strong representation from this same group of bacteria was also seen in our regional study of mud pools from the TVZ as well as in previous studies of hot spring environments in Indonesia (Baker et al., 2001) and high temperature Solfatara crusts in Italy (Glamoclija et al., 2004). Signatures of the gamma Proteobacteria observed in the Lassen mud pool profiles were not as dominating as the same signatures in pools from the other two sites. The profiles from the Lassen pools showed high diversity among the bacterial community with no signatures dominating the community profile. Bands 3, 7, and 11 were noted in a majority of the mud pools from Yellowstone and the BS #2 pool from Lassen. These bands were representative of signatures from the alpha
69 Proteobacteria and were the second most dominant group observed in our community profile analysis. Strong signatures of members of the alpha Proteobacteria were also recognized in our TVZ survey, thermal mud from Solfatara, Italy (Glamoclija et al., 2004) and other studies of hot springs in Yellowstone (Hugenholtz et al., Johnson et al., 2003). The presence of a common bacterial signature in pools from both Lassen and Yellowstone might be indicative of a mud pool bacterium with a larger geographical range. Although unique, isolated hydrothermal sources fuel these two thermal regions, the close proximity of these two thermal areas on a geographical scale in comparison to the other sites may result in migration of species. Outside of this signature, the bacterial community profiles of the Lassen and Yellowstone systems are quite different. Cladistic analysis showed little similarity in community structure between these two systems. Faint signatures belonging to the Cytophaga-Flavobacteria were observed in several of the mud pool profiles from Yellowstone but do not appear in the profiles from Rincon or Lassen. Signatures from this group were quite prominent in our analysis of the mud pool communities in the TVZ and have also been reported in high, temperature low pH hot springs in Indonesia (Baker et al., 2001). Much like our observations for the New Zealand pools, the bacteria observed in these geographically isolated sites displayed a broad tolerance range in both temperature and pH. Although the populations of the individual sites and pools displayed a large degree of heterogeneity, especially among less dominant profile signatures, the same groups of dominant bacterial signatures were represented in all of the study sites. There appears to be a selective advantage for bacterial communities in mud pools that can tolerate drastic variations in the physiochemical environment. It is this adaptability to environmental change that has allowed these bacterial groups to colonize and flourish in this unique
70 high temperature niche. Cladistic similarity analysis of these 3 regions in conjunction with the TK Pool representative from the TVZ survey showed the pools to group into three clusters. The 2 pools from Rincon de la Vieja were shown to be identical for this similarity analysis of the bacterial community profile and clustered with the Lassen SW pool. The center cluster in Figure 2.5 only contained pools from Yellowstone. The third and final cluster contained the Lassen pools, DK 1 and BS 2, the TVZ TK pool, and the PHS 1 pool from Yellowstone. Observations of the community profiles from these geographically isolated sites, including the TK representative from the New Zealand study suggested the pools from a given geographical location were more related to one another than to pools of distant geographical range, despite the variation seen between the individual pools at the community level. Previous studies of hyperthermophilic archaea have shown there to be limited gene flow between microbial populations likely due to poor survivability limiting dispersal or competition from endemic species in a particular environment (Whitaker et al., 2003). Other evidence of geographic isolation of hot spring bacterial species has also been reported (Hudson et al., 1989; Saul et al., 1993; Williams et al., 1996; Peturdottir et al., 2000).
Conclusion The data presented here shows that amplifiable DNA can be directly extracted from high temperature mud pool samples. Due to the low temperature, acidic nature of the pools sampled in this study, the contribution of exogenous, extracellular DNA to the extracted pool is minimal. Our analyses of the mud pools from the TVZ, New Zealand showed a range of chemical and bacterial signatures across the different pools and thermal fields. Although differences were observed at the community level, thermophilic members of the gamma Proteobacteria, the Cytophaga-Flavobacteria, and
71 the alpha Proteobacteria dominated the majority of the pool bacterial profiles and displayed a tolerance of fluctuations in the physiochemical environment. Analysis of the bacterial community of high temperature mud pools from other globally distributed hydrothermal systems showed similar results to our regional New Zealand survey with members of the gamma Proteobacteria and alpha Proteobacteria dominating these systems over a broad range of physiochemical environments. The diversity of the profiles of the individual pools over both a regional and geographical range suggested physical isolation plays a role in bacterial community diversity. Future mud pool microbial community studies should focus on signatures from the Archaea in order to gain a complete picture of the microbial community. Also, direct comparisons between mud pools and neighboring thermal features may lead to further implications for endemism of microbial species to the mud pool environment.
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82 Chapter 3 PHYLOGENETIC ANALYSIS OF THE MICROBIAL COMMUNITY INHABITING A HIGH TEMPERATURE MUD POOL FROM THE TAUPO VOLCANIC ZONE, NEW ZEALAND
Abstract The microbial communities (Bacteria and Archaea) and physiochemistry of the high temperature, near-neutral Tokaanu mud pool and its neighboring hot springs were examined using a combination of molecular and chemical analyses. Community level analyses were performed using DGGE fingerprinting methods and 16S rDNA sequence analysis. Measurements of the physiochemical environment of the Tokaanu pools showed reduced, near-neutral conditions across the majority of thermal features. Bacterial community analysis of these pools showed a unique bacterial assemblage within the Tokaanu mud pool compared to nearby hot springs. These unique signatures were affiliated with uncultivated members of the Flavobacteria and other bacteria associated with hot springs, hydrocarbon seeps, and subsurface gold mine environments. Archaeal community analysis showed a tight group of seemingly closely related archaea unique to the Tokaanu mud pool profile compared to the other Tokaanu features. Phylogenetic analyses of the bacterial and archaeal 16S rDNAs yielded 12 unique bacteria and 13 unique archaea from the Tokaanu mud pool. The bacteria were affiliated most closely with characterized thermophilic members of the Firmicutes, epsilon Proteobacteria, and Thermus sp. Analysis of the archaea revealed a unique clade with the Crenarchaeota. The tight grouping of this clade between characterized Pyrobaculum
83 sp. and Ignicoccus sp. within the Crenarchaeota and the high degree of within pool similarity suggest a new, unique group of Crenarchaeotes endemic to the Tokaanu mud pool system.
Introduction Recent developments in molecular phylogenetic technologies have revolutionized our understanding of the diversity, distribution, and evolution of microbes in the environment and have lessened the need for cultivation in the study of microbial diversity (Pace et al., 1986; Ward et al., 1992). These molecular-based studies have dramatically contributed to our knowledge of the widespread distribution of specific microbial lineages and provided insight into the evolutionary history of all prokaryotes (Woese, 1987). The use of molecular genetic tools has proven particularly successful in studies of microbial life in extreme environments, specifically high temperature environments like terrestrial hot springs and deep-sea hydrothermal vents (Ward et al., 1998; Takai and Horikoshi, 1999; Campbell and Cary, 2001; Reysenbach and Cady, 2001; Chapelle et al., 2002; Teske et al., 2002; Hoek et al., 2003, Keller and Zengler, 2004). The molecular studies of extreme high temperature ecosystems have led to the proposal of a new kingdom, the Korarchaeota, within the Archaeal domain (Barns et al., 1996) and have continued to provide insight into the upper temperature limit for life (Kashefi and Lovley, 2003). One globally distributed, terrestrial extreme ecosystem that has remained poorly characterized in terms of microbial diversity using molecular genetic technologies is the high temperature mud pool. Formed by the dissolution of rock as steam and superheated fluids rise from deep below the Earth’s surface, these hot, often acidic pools of bubbling silts, clays, and volcanic ash are physically characterized by dramatic
84 between pool variability in viscosity, temperature, and pH. A common feature worldwide in areas of surface hydrothermal activity, mud pool ecosystems represent an environment reminisant of the hot, moderately reduced, hydrothermal conditions that have been proposed as favorable to the abiotic formation of some amino acids (Amend and Shock, 1998). Clays have been proposed as an important component in the abiotic synthesis and replication of simple organic molecules suggesting that ‘life arose from clay’ (Hartman, 1986). Considering these environmental conditions and evidence that the deepest branching members of the Bacteria and Archaea (based on the 16S rRNA tree of life) are all thermophilic and many are chemolithoautotrophic (Stetter, 1996b; Woese et al., 1990), we predict that mud pools may harbor previously undescribed microbes of deeply branching phylogenetic lineages and may provide new insight into the origins and evolution of life through unique metabolic and geochemical interactions. Although the role of chemoautotrophic thermophiles in the origins and evolution of life is still unclear (Doolittle, 1999; Galtier et al., 1999), there is strong evidence that suggests early life may have been chemosynthetic and may have required high temperatures for growth (Pace, 1997). One globally significant site of volcanic and surface hydrothermal activity, located directly above the southeastern rim of the “Ring of Fire” where the Pacific Plate slides beneath the Indo-Australian Plate, is New Zealand’s North Island. The North Island’s most active zone of surface activity is marked by the Taupo Volcanic Zone (TVZ), which extends between Mount Ruapehu in the southwest of the Tongariro Volcanic Center into the Bay of Plenty and White Island to the northeast (Gamble et al., 1993). As the youngest and most active site of volcanism in New Zealand (Wilson et al., 1995), this zone consists of approximately 20 thermal fields containing hundreds of hot
85 water surface features including hot springs, geysers, mud pools and sinter deposits (Risk et al., 2002). In a previous study, we utilized a multifaceted molecular and geochemical approach to compare the Bacterial communities of mud pools from 7 thermal fields within the TVZ. Of the13 mud pools sampled from the TVZ, our initial Bacterial community analyses revealed a single mud pool in the Tokaanu Thermal Area that exhibited a unique physiochemical environment supporting a very distinct Bacterial community in comparison to other pools sampled within the TVZ (See Chapter 2). Located near the southern rim of the TVZ along the southern edge of Lake Taupo, the Tokaanu-Waihi thermal field is comprised of three main zones of activity: Tokaanu, Waihi and Hipaua and remains poorly explored compared to other thermal fields within the TVZ (Risk et al., 2002). The Tokaanu Thermal Area contains hot springs, thermal ground, a geyser, and a small mud pool occurring at the base of the Tihia volcano (Risk et al., 2002; personal observation). As described in previous study (Chapter 2), the Tokaanu mud pool in the Tokaanu Thermal Area has high temperature (90°C), near-neutral pH (pH 6.8) conditions resulting in a unique bacterial community assemblage compared to other mud pools from the TVZ. Due to these observations, the Tokaanu mud pool was chosen for a direct comparison of the geochemistry and microbial communities of a high temperature mud pool and its neighboring hot springs. For this study, we collected mud pool and hot spring samples from the Tokaanu Thermal Area. We utilized a suite of molecular, geochemical, and field analyses to describe the unique microbial assemblages and geochemical environment associated with the Tokaanu pools. Microbial community analyses of the Tokaanu mud pool and the 4 neighboring hot springs demonstrated each to contain unique Bacterial and Archaeal communities. Bacteria associated with the
86 Tokaanu mud pool were closely related to known sequences from the Clostridium- Bacillus subphylum of gram-positive bacteria and anaerobic thermophiles. The archaeal community of the Tokaanu mud pool resulted in the identification of a unique clade from the Crenarchaeota.
Materials and Methods
Sample collection, preparation and field measurements Environmental samples were collected from 1 mud pool and 4 neighboring clear hot spring pools (38° 58' S; 175° 45' E) located in the Tokaanu Thermal Area on the North Island of New Zealand in August of 2003. Mud pool samples were collected using a 5-meter retractable aluminum pool fitted with a stainless steel sampling arm configured to hold 4 sterile 50mL Falcon tubes and a HOBO H8 temperature data logger with external probe (ONSET Scientific, Bourne, MA) for in-situ recording of temperatures. Each pool was sampled for one minute to allow the thermocouple to equilibrate and record multiple data points. Upon recovery, the pH of each sample was recorded with an Accumet portable AP5 pH meter fitted with an Accumet pH/Automatic Temperature Compensation probe (Fisher Scientific, Pittsburgh, PA). Pore water was collected on site by centrifugation (RCF = 1100 X g for 30 minutes) using a MobileSpin 12V field centrifuge (Vulcon Technologies, Grandview, MO). The pore water was filtered through a 0.2 µM Supor membrane Acrodisc syringe filter (Pall Corporation,
Ann Arbor, MI), placed into acid-washed polycarbonate vials, and frozen on dry ice. Mud pool samples were stored at ambient temperature for transport back to the laboratory.
87 Hot spring pool samples were collected from pools in the Tokaanu Thermal Area that were located in close proximity to the Tokaanu mud pool. Sterile 60 cc syringes (Becton Dickinson and Company, Franklin Lakes, NJ) were used to filter approximately 120 mL of each sample through a 0.2 µM Supor membrane Acrodisc syringe filter (Pall Corporation). Filters were sealed and frozen in dry ice for transport back to the laboratory. The filtrate was collected and frozen for nutrient and chemical analysis as described previously. Measurements of temperature and pH were performed at the point of collection for each hot spring sample. All samples (mud pool, hot spring pool, and pore water) were stored at -80º C in the laboratory for further analysis.
Geochemistry Chemical analyses were performed on pore water and hot spring pool filtrate
+ - - samples for dissolved ammonium (NH4 ), nitrate + nitrite (NO3 + NO2 ), and
3- orthophosphate (ΣPO4 ) using a Perstorp Analytical Flo-Through Analyzer (Perstorp
+ Analytical, Inc., Silver Spring, MD). Briefly, concentrations of NH4 were determined by the phenol hypochlorite method (Glibert and Loder, 1977; Grassfoff and Johansen,
- - 1972). Concentrations of both NO3 and NO2 were determined by the sulphanilamide/N(1-napthyl) ethylene diamine method following a cadmium reduction of
- - 3- NO3 to NO2 (Glibert and Loder, 1977), and ΣPO4 concentrations were determined using the phosphor-molybdenum blue method (Strickland and Parsons, 1972). Samples were run in duplicate, if possible, and dilutions were made to ensure measurement values were within the range of the standard curve for each analysis.
88 Water content, elemental and particle size analyses The moisture content of the mud samples was determined using a standard thermogravimetric method (Wilson, 1971). Analysis of some of the major cations
+ + + + 2+ (lithium (Li ), sodium (Na ), ammonium (NH4 ), potassium (K ), magnesium (Mg ), and calcium (Ca2+) in the pore water from the Tokaanu mud pool was performed using a Dionex ICS-2000 Ion Chromatography system (Dionex Corporation, Sunnyvale, CA) equipped with an IonPac CG16 guard column and a IonPac CS16 cation-exchange column (Dionex Corporation). The eluent used was methanesulfonic acid and calibration was done using external standards. For particle size determination and elemental analyses, a thin layer of the mud sample was spread onto an EM stub and allowed to air dry. The stub was subsequently coated with platinum and imaged using a Hitachi S4100 field emission scanning electron microscope (FESEM) (Hitachi High Technologies America, Inc., Pleasanton, CA). Elemental composition of the mud sample was performed with a KEVEX energy-dispersive elemental analyzer (Thermo Electron Corporation, San Jose, CA) and IXRF software (IXRF SYSTEMS, Inc., Houston, TX) according to manufacturer’s instructions.
Nucleic acid extraction Nucleic acids were extracted using a modified CTAB (cetyltrimethylammonium bromide-polyvinylpyrrolidone- -mercaptoethanol) extraction protocol (Murray and Thompson, 1980; Coyne et al., 2001). Briefly, mud samples (4 g) were extracted with 10 mL CTAB extraction buffer (100mM Tris-HCl [pH 8.0], 20 mM EDTA [pH 8.0], 1.4 M NaCl, 2% CTAB, 2% PVP (polyvinyl pyrrolidone), 0.2% - mercaptoethanol) at 65ºC for 1 hr in a shaking water bath. The extraction was then
89 centrifuged for 1 min (2790 X g). The supernate was transferred to a new tube, an equal volume of chloroform: isoamyl alcohol (24:1) added, and samples incubated at room temperature (RT) for 20 min on an inversion rocker. Following a 10 min centrifugation (17000 X g), the aqueous phase was transferred to a new tube and DNA precipitated with an equal volume of isopropanol and 0.5 volumes of 5M NaCl at -80ºC for at least 1 hr. The precipitated DNA was pelleted by centrifugation (17000 X g) for 20 min, washed with 70% ethanol, dried in a speed-vac at RT, and resuspended in 40 µL TE (10mM
Tris-HCl, 1mM EDTA, pH 8.0). For the extraction of hot spring filtered water samples, 3 mL of CTAB buffer was loaded into a 3cc syringe and injected into the filter assembly until full. The filter was sealed, placed into a plastic bag with the syringe of buffer attached, and submerged in a 65ºC water bath for 1 hr. Following the incubation, the remaining extraction buffer was pushed through the filter into microfuge tubes (0.6 mL per tube). The remainder of the extraction protocol was performed as described above. Nucleic acids from each sample were quantified spectrophotometrically using a NanoDrop ND- 1000 spectrophotometer at 260 nm (NanoDrop Technologies, Montchanin, DE).
Community fingerprinting analyses The bacterial and archaeal communities were initially examined using denaturing gradient gel electrophoresis (DGGE) (Muyzer et al., 1993) in order to obtain a snapshot fingerprint of the microbial populations present within the individual sample DNA pools. For bacterial DGGE analysis, the V3 region of the bacterial 16S rDNA genes present in the mud pool and hot spring samples was amplified using the polymerase chain reaction (PCR) (Saiki et al., 1988) with primers 338F/GC clamp and 519RC as described by Muyzer et al. (1993) (Table 1). For
90 Table 3.1 Oligonucleotides used for PCR.
Primer Sequence (5’ → 3’) Domain Reference
338FGC CGCCCGCCGCGCGCGGC Bacteria Muyzer et al., 1993 GGGCGGGGCGGGGGCAC GGGGGGCCTACGGGAGG CAGCAG 519 RC ATTACCGCGGCTGCTGG Bacteria Muyzer et al., 1993 27F AGAGTTTGATCMTGGCTCAG Bacteria Lane, 1991 907 R CCGTCAATTCMTTTRAGTTT Bacteria Lane et al., 1985 1522R AAGGAGGTGATCCANCCRCA Bacteria Lane, 1991 347FGC CGCCCGCCGCGCCCCGCG Archaea Raskin et al., 1994 CCCGTCCCGCCCCCGCCA CGGGGCGCAGCAGGCGCG 519RA GGTATTACCGCGGCGGCTG Archaea Muyzer et al., 1993
Arch21F TTCCGGTTGATCCYGCCGGA Archaea Delong, 1992 Arch958R YCCGGCGTTGAMTCCAATT Archaea Delong, 1992
DGGE analysis of the archaeal community, mud pool and hot spring samples were amplified using a nested PCR approach with16S rDNA specific archaeal primers 21F and 958R (Delong, 1992), then archaeal DGGE primer 347F/GC clamp (Raskin et al., 1994) and 519RA (Muyzer et al., 1993) (Table 3.1). Reactions (50 µL) were performed using ~50 ng of template DNA and final concentrations of: 1X JumpStart PCR Buffer
(Sigma, St. Louis, MO), 200 µM each deoxyribonucleotide triphosphate, 2mM MgCl2, 40 ng bovine serum albumin, 0.2 µM concentration of each primer, and 2 units of
JumpStart Taq polymerase (Sigma). The PCR reactions were run on a PTC-200 Thermocycler (MJ Research, Inc., Waltham, MA) using the following hot start (D’Aquila et al., 1991), touchdown protocol (Don et al., 1991) for DGGE primer sets:
91 94ºC for 2 min, followed by 35 cycles of 94ºC for 1 min, 65-55ºC (-0.5ºC per cycle) for 1 min, and 72ºC for 1min. Initial archaeal PCR reaction: 94ºC for 2 min, followed by 35 cycles of 94ºC for 1 min, 55ºC for 1 min, and 72ºC for 1 min, and 72ºC extension for 5 min. This reaction was then precipitated with a half volume of 30% polyethylene glycol (PEG)/1.5M NaCl to remove excess primers prior to DGGE PCR amplification. For microbial community DGGE analysis, PCR products were separated on an 8% acrylamide gel (37.5:1 ratio of arylamide-bisacrylamide) with a 25% to 65% gradient (for Bacteria) or 40% to 80% gradient (for Archaea) of denaturant (7M urea and 40% formamide). All gels were run for 5 hr (130V) at 60ºC in 1X TAE buffer (40mM Tris base, 20 mM sodium acetate, 1 mM EDTA) on a DCode Universal Mutation Detection System (Bio-Rad, Hercules, CA). Gels were stained with ethidium bromide (500ng/mL) for 30 minutes and destained in sterile water for 30 minutes. Gels were visualized using a UV transilluminator and digitally photographed using the AlphaImager imaging system (Alpha Innotech Corp., San Leandro, CA). Comparisons of mud pool and hot spring community samples run on DGGE gels were performed using the Phoretix 1D Version 5.10 software and the Phoretix 1D Version 1.13 Database (Nonlinear Dynamics Limited, UK). This software package allowed for lane and band detection and lane relationship comparisons of amplicons for single and multiple gel comparisons. These comparisons were used to create cladistic maps of relationships between the individual pools. Individual bands, representing dominant amplicons within the gel profile common to multiple pools and those unique to individual pools, were excised and subject to further sequence analyses. Select bacterial DGGE bands were excised and reamplified according to a protocol described by Campbell and Cary (2001) and subjected to reamplification
92 according to the PCR amplification conditions described above. Reaction products were electrophoresed on a DGGE gel for confirmation of single bands. The PCR band products were purified for sequencing using the GenElute PCR Clean-up Kit (Sigma- Aldrich Co., St. Louis, MO) according to the manufacturer’s instructions and quantified by spectrophotometry. Purified products were subsequently sequenced with BigDye Terminator v1.1 Cycle Sequencing chemistry (Applied Biosystems Inc. (ABI), Foster City, CA). Sequencing reactions (5 µL) were performed using 50 ng of purified template and 1.6 µmoles 519RC primer according to the manufacturer’s instructions.
Clone library construction 16S rRNA genes of the Tokaanu mud pool microbial community were amplified using primers 27F and 1522R (Lane, 1991) for Bacteria and primers 21F and 958R for Archaea (Table 3.1) under the conditions described above using the following amplification protocol: 94ºC for 2 min, followed by 35 cycles of 94ºC for 1 min, 55ºC for 1.5 min, and 72ºC for 3 min, and 72ºC extension for 5 min. All PCR products were evaluated by electrophoresis on agarose gels following amplification and stored at -20°C until further analysis. The PCR products of the 937 bp region of the Archaeal 16S rRNA (primers 21F and 958R) and the 1495 bp region of the Bacterial 16S rRNA (primers 27F and 1522R) from the Tokaanu mud pool were ligated into a TOPO TA pCR 2.1 cloning vector (Invitrogen Life Technologies, Carlsbad, CA), transformed into TOP 10 chemically competent E. coli cells (Invitrogen Life Technologies), and grown according to the manufacturer’s instructions. Ninety-six clones from each were subjected to PCR amplification using primers M13 Forward/M13 Reverse and compared using restriction fragment length polymorphisms (RFLP) analysis with site-specific restriction enzymes (Hae III and Rsa I for Archaea and Rsa I and HinP I for Bacteria).
93 Libraries of unique ribotypes were constructed for both the bacterial and archaeal 16S rRNA genes from the Tokaanu mud pool based on the RFLP analyses. Unique clones were sequenced in the forward and reverse directions using BigDye Terminator v1.1 Cycle Sequencing chemistry (ABI, Inc.). Sequencing reactions (10 µL) were performed using 50 ng of purified template and 3 µmoles of the respective primer (27F/907R for Bacteria; 21F/958R for Archaea). All sequencing was performed using the BigDye Terminator Cycle Sequencing Ready Reaction chemistry (Applied Biosystems, Inc. (ABI), Foster City, CA) according to the manufacturer’s instructions on an ABI Prism 310 Genetic Analyzer DNA sequencer (ABI, Inc.). Sequences were edited using Sequence Navigator and AutoAssembler software (ABI, Inc., Foster City, CA). Initial sequence alignments were performed using Clustal W analysis (Thompson et al., 1994) in DNASTAR’s MegAlign sequence manipulation program (Lasergene, Inc., Madison, WI). Sequences were subject to BLAST searches within the GenBank Database for comparison with submitted and characterized sequences. Final sequence alignments were created in the Genetic Data Environment (GDE version 3.2) (Smith et al., 1992) and phylogenetic distance relationships were inferred (only to subdivision for DGGE stabs) from alignments created alongside characterized sequences from the GenBank Database using the Olson method (Olsen, et al., 1988). Neighbor-joining distance trees with 100 bootstrap replications were constructed alongside GenBank sequences for subdivision placement and relationship among known phylotypes using Philip version 3.572 (Felsenstein, 1993).
94 Results
Chemical, elemental, and particle size analysis The mud pool showed a significantly higher temperature (90.8°C) than neighboring hot spring pools (60.2°C – 85.0°C) (Table 3.2). The near neutral Tokaanu mud pool and hot spring pools TOK 2, TOK 6, and TOK 10A ranged between 6.5 and 6.8. Hot spring pool TOK 7 was the exception with a more acidic pH of 5.7.
+ Concentrations of NH4 were significantly higher in the mud pool (486 µM) when compared to the hot spring pools (54-119 µM) (Table 3.3). Additionally, concentrations
- - 3- of NO3 + NO2 and PO4 were low in the mud pool at 1.3 µM and 4.6 µM respectively, in comparison to the surrounding hot spring pool samples which ranged from 2.2 –
- - 3- 119.0 µM for NO3 + NO2 and 8.1 – 36.1 µM for PO4 . Considering these data, the physiochemical environment of the Tokaanu mud pool showed the highest degree of similarity to the TOK 10A hot spring. This hot spring was also its closest neighboring thermal feature. In addition, cation analyses of pore water (Table 3.4) from the Tokaanu mud pool displayed concentrations of sodium (Na+), potassium (K+), and calcium (Ca2+). Elemental composition of the solid fraction (Figure 3.1) of the Tokaanu mud pool using energy-dispersion showed large aluminum (Al) and silica (Si) peaks along with smaller peaks for iron (Fe), magnesium (Mg), and oxygen (O). Scanning electron microscopy analysis (Figure 3.2) of the particulate fraction of the mud pool matrix revealed a large particle size range (< 2 µM to > 12 µM) with varying morphology from small spheres to larger fibrous, tube-like particles (Figure 3.2).
95 Table 3.2 Sample site overview and physical characterization of the Tokaanu Thermal Area (38° 58' S; 175° 45' E).
Temperature Moisture Sample Name Thermal Feature (°C) pH Content (%)
TOK 2 Hot spring 85.1 6.75 100 TOK 6 Hot spring 67.1 6.57 100 TOK 7 Hot spring 60.7 5.72 100 TOK 10A Hot spring 60.2 6.74 100 TOK MP Mud pool 90.8 6.77 83.8
96 Table 3.3 Chemical analysis of pore water from the Tokaanu mud pool and its neighboring hot springs.
+ - - 3- Temperature NH4 NO3 + NO2 PO4 Sample Name (°C) pH (µM) (µM) (µM)
TK Mud Pool 90.8 6.77 487 1.28 4.57 TOK 2 Hot Spring 85.1 6.75 119 19.0 28.9 TOK 6 Hot Spring 67.1 6.57 54.0 119 36.1 TOK 7 Hot Spring 60.7 5.72 ------TOK 10A Hot Spring 60.2 6.74 211 2.17 8.09
97 Table 3.4 Ion chromatography of cations from the Tokaanu mud pool.
Concentration in Tokaanu Mud Pool Cation (parts per million [ppm])
Li+ 14.68 Na+ 1034 + NH4 10.71 K+ 77.56 Mg2+ 7.538 Ca2+ 71.30
98 PCR-DGGE Bacterial and Archaeal community characterization Analysis of the Tokaanu bacterial DGGE gel resulted in detection of 64 bands at 31 different positions (Figure 3.3). The bacterial amplicons from the Tokaanu samples revealed significant variation between the banding patterns of the mud pool in comparison to the hot spring samples. Of the 13 bands present in the Tokaanu mud sample, the four most prominent bands (Figure 3.3) were observed that were not present or present as minor bands in the hot spring pool profiles. Similarly, the four most prominent bands (Figure 3.3) in the hot spring pool samples profiles were not observed in the Tokaanu mud pool profile. The TOK 2 hot spring DGGE profile displayed 4 unique bands not observed in the profiles of any of the other Tokaanu pools sampled in this study. Twelve bands were excised and successfully sequenced from the Tokaanu bacterial DGGE profiles (see number and position in Figure 3.3; Table 3.5). Bands 5 and 6 were similar (93% and 97% respectively) to Methylobacterium sp. and bands 7 and 8 were similar (96% and 94% respectively) to Thermus sp. In addition, bands 2 and 3 were highly similar (98% and 97% respectively) to uncultivated eubacteria from hydrocarbon seep sediments and deep subsurface gold mine environments. Other sequenced bands from the Tokaanu bacterial DGGE profile corresponded loosely to characterized sequences from saline and highly radioactive extreme environments. Archaeal DGGE community analysis of the Tokaanu pools resulted in the detection of 40 bands in 26 positions and revealed a similar comparative trend to that observed for the bacterial DGGE analysis. Banding patterns observed for the Tokaanu mud pool showed 4 unique, dominate bands (Figure 3.4) not observed in the hot spring profiles. TOK 2 and TOK 6 hot spring profiles were also unique in comparison to the
99 Figure 3.1 Elemental analysis of the solid phase component of the Tokaanu mud pool using KEVEX energy-dispersive elemental analysis.
100 Figure 3.2 Particle size analysis of the Tokaanu mud pool solid phase using scanning electron microscopy.
101 other hot spring profiles and to one another. The most dominant band in the mud pool DGGE profile is not present in any of the hot spring profiles while there was a high degree of profile similarity between the hot springs. Attempts to excise and reamplify the bands from the Tokaanu archaeal DGGE gel were unsuccessful and so necessitated construction and sequencing of a full 16S rRNA gene library. Bacterial and archaeal DGGE community profiles of the Tokaanu pools were subjected to Phoretix analysis for profile comparison and UPGMA derived dendrograms were generated using similarity (presence or absence of band matches) between the pools (Figures 3.5 and 3.6 respectively). Cladistic analysis of the bacterial profiles grouped the Tokaanu mud pool loosely with hot springs TOK 6 and 7 while hot springs TOK 10A and TOK 2 were the most distant (Figure 3.5). Analysis of the archaeal profile revealed strong similarities between the hot springs with the Tokaanu mud pool being the most distant of all the pools examined here (Figure 3.6).
Bacterial and Archaeal community characterization of the Tokaanu mud pool Following initial microbial community structure analysis using DGGE of the Tokaanu pools the archaeal and bacterial communities of the Tokaanu mud pool were examined in finer phylogenetic detail. Bacterial and archaeal rDNA were successfully amplified, cloned, and libraries screened for unique operational taxonomic units (OTUs) using RFLP analysis. Ninety-six clones were screened for both the Bacteria and Archaea resulting in 12 bacterial OTUs and 13 archaeal OTUs. A representative of each RFLP-defined OTU was sequenced from both the 3′ and 5′ ends. The bacterial OTUs were in three known classes of the Eubacteria, including the Firmicutes (7 OTUs), the Deinococci (2 OTUs), and the gamma Proteobacteria (1 OTU). The remaining 2 OTUs
102 TOK MP TOK 2 TOK 6 TOK 7 TOK 10A
1 10 11 12 2
3 4 5 6
7 9 8
Figure 3.3 DGGE analysis of the bacterial communities associated with the Tokaanu mud pool and its neighboring hot springs. Numbers designate bands that were excised, reamplified, and subjected to sequence analyses.
103 Table 3.5 BLAST results of excised bands from the bacterial DGGE analysis of the Tokaanu mud pool and its neighboring hot springs.
Most closely related sequence match Percent Sequence Band (accession number) Score Similarity Length Origin
Band 1 uncultured Flavobacteria 230 96 148 Salt marsh (AY711798) Band 2 uncultured bacterium D48 226 98 130 Gold mine (AF337870) Band 3 uncultured bacterium BPC060 272 97 164 Hydrocarbon (AF154081) seep sediment Band 4 uncultured bacterium MTU-25 54 90 75 Thermophilic (AY099249) bioleaching Band 5 uncultured Methylobacterium 99.6 93 98 Hot spring (AY569289) (pink mat) Band 6 Methylobacterium sp. 4BR9 248 97 140 Radioactive (AY792012) water Band 7 Thermus W28 A.1 214 96 139 Hot spring (L10068) Band 8 Thermus W28 A.1 202 94 158 Hot spring (L10068) Band 9 uncultured bacterium ALPHA8 155 91 147 Salmonid gill (AY494638) Band 10 uncultured bacterium TTMF57 194 92 157 Gold mine (AY741701) borehole water Band 11 uncultured eubacterium env.OPS 17 176 94 117 Thermal stream (AF018199) Band 12 uncultured bacterium B02-T2-2 170 91 153 UMTRA site (AY995088)
104 TOK MP TOK 2 TOK 6 TOK 7 TOK 10A
Figure 3.4 DGGE analysis of the archaeal communities associated with the Tokaanu mud pool and it neighboring hot springs.
105 TOK 10A
TOK 7
TOK 6
TOK MP
TOK 2
Figure 3.5 Phoretix comparative analysis of the bacterial DGGE analyses of the Tokaanu mud pool and its neighboring hot springs. UPGMA dendogram displays similarities based on the presence and absence of bands in the profiles.
106 TOK 10A
TOK 7
TOK 6
TOK 2
TOK MP
Figure 3.6 Phoretix comparative analysis of archaeal DGGE analyses of the Tokaanu mud pool and its neighboring hot springs. UPGMA dendogram displays similarities based on the presence and absence of bands in the profiles.
107 were epsilon Proteobacteria (Table 3.6). Initial BLAST analysis of the 13 archaeal OTUs resulted in similarities most closely related to Pyrobaculum calidifontis and other uncultivated Crenarchaeotes from other high temperature hydrothermal environments (Table 3.7). Individual neighbor-joining distance trees were constructed comprised of bacterial and archaeal 16S rRNA gene sequences isolated from the Tokaanu mud pool and a subset of related sequences obtained from the GenBank database (Figures 3.7 and 3.8 respectively). This analysis of the bacterial community from the Tokaanu mud pool placed sequences into 3 distinct groups (Figure 3.7). Group I was most closely related (74-98%) to previously described thermophilic gram-positive bacteria (i.e. Bacillus sp.) and ammonium-dependent gram-positive bacteria (i.e. Ammoniphilus sp.). Group II grouped loosely (75-96%) with characterized thermophilic, anaerobic, hydrogen oxidizers and Group III displayed some relation (75-99%) to known Thermus species (Figure 3.7). Phylogenetic analyses of archaeal sequences (Figure 3.8) obtained from the Tokaanu mud pool grouped 11 of the 13 sequences into a unique clade of organisms within the Crenarchaeota with no known affiliate (Figure 3.8). This unique clade showed an 88-99% within clade identity and was flanked within the archaeal tree by published sequences of Ignicoccus sp. and Pyrobaculum sp. Clone TMP A1 displayed moderate degree of similarity to Ignicoccus islandicus (92%) and clone TMP A12 showed some affiliation to Pyrobaculum calidifontis (96%).
Discussion Although a globally distributed, prevalent surface feature of terrestrial hydrothermal activity, little is known about the microbial communities inhabiting high temperature mud pools. Previous studies of high temperature mud pools based on
108 Table 3.6 Summary of bacterial 16S rRNA clone analysis from the Tokaanu mud pool.
Clone Closest GenBank Percent Accession Type Database match Score identity number Class
TMP B11 Thermus sp. 1239 98 L09671 Deinococci T351 TMP B10 Thermus 1263 99 L09667 Deinococci filiformis TMP B4 eubacterium 791 93 AF050591 Firmicutes WCHB1-84 TMP B5 eubacterium 799 94 AF050591 Firmicutes WCHB1-84 TMP B3 Clone 013B04 720 89 CR933129 Firmicutes B SD P15 TMP B1 S17sBac21 1015 95 AF299126 Epsilon Proteobacteria TMP B12 S17sBac21 982 95 AF299126 Epsilon Proteobacteria TMP B6 Bacterium 1316 99 AJ295684 Firmicutes IrT-RS2 TMP B7 Ammoniphilus 1176 97 Y14579 Firmicutes oxalaticus TMP B2 Desulfitobacterium 1017 94 U68528 Firmicutes chlororespirans TMP B8 Desulfitobacterium 1063 94 AF357919 Firmicutes sp. Viet-1 TMP B9 Klebsiella oxytoca 1289 99 AJ871860 Gamma Proteobacteria
109 Table 3.7 Summary of archaeal 16S rRNA clone analyses from the Tokaanu mud pool.
Clone Closest GenBank Percent Accession Type Database match Score identity number
TMP A9 clone CH8 3a ARC 1274 94 AY672492 16SrRNA 9N EPR TMP A13 clone CH8 3a ARC 854 96 AY672492 16SrRNA 9N EPR TMP A4 clone CH8 3a ARC 1291 95 AY672492 16SrRNA 9N EPR TMP A11 clone CH8 3a ARC 1285 94 AY672492 16SrRNA 9N EPR TMP A10 clone CH8 3a ARC 801 94 AY672492 16SrRNA 9N EPR TMP A5 clone CH8 3a ARC 1247 94 AY672492 16SrRNA 9N EPR TMP A12 Pyrobaculum 1498 88 AB078332 calidifontis TMP A6 clone CH8 3a ARC 1165 93 AY672492 16SrRNA 9N EPR TMP A3 clone CH8 3a ARC 1250 94 AY672492 16SrRNA 9N EPR TMP A8 clone CH8 3a ARC 1346 94 AY672492 16SrRNA 9N EPR TMP A1 clone pCIRA-S 1273 94 AB095128 16SrRNA TMP A7 clone CH8 3a ARC 1251 86 AY672492 16SrRNA 9N EPR TMP A2 clone CH8 3a ARC 1322 94 AY672492 16SrRNA 9N EPR
110 Sulfolobus solfataricus (X03235)
TMP B3
99 TMP B4
TMP B5
TMP B7 81 Ammoniphilus oxalaticus (Y14579) Group I TMP B6
60 Bacillus thermoamylovorans (AJ586361)
91 TMP B2
TMP B8
Desulfitobacterium frappieri (U40078)
Rhodovulum sulfidophilum (D16422)
95 Nitrosomonas nitrosa (AJ298740)
TMP B9
Escherichia coli (AJ605115)
Caminibacter hydrogeniphilus (AJ309655) 54 100 TMP B1 Group II
TMP B12
TMP B10
100 TMP B11 Group III
Thermus filiformis (L09667) 0.1
Figure 3.7 Bacterial phylogenetic tree of Tokaanu mud pool sequences. Phylogenetic tree as determined by neighbor joining analysis. Bootstrap values based on 100 replications.
111 Korarchaeota SRI-306 (AF255604)
Thermodiscus maritimus (X99554)
Hyperthermus butylicus (X99553) Pyrolobus fumarii (X99555)
66 Staphylothermus hellenicus (AJ012645) Desulfurococcus saccharovorans (X99558) Ignicoccus pacificus (AJ271794) 50 Ignicoccus islandicus (X99562) TMP A1 TMP A7 TMP A4 85 TMP A3
TMP A8 Crenarchaeotal TMP A6 clade from the TMP A5 75 Tokaanu mud pool TMP A2 54 TMP A9 TMP A11 TMP A13 72 TMP A10 TMP A12 100 Pyrobaculum calidifontis (AB078332) Pyrobaculum islandicum (L07511) 0.1
Figure 3.8 Archaeal phylogenetic tree of Tokaanu mud pool sequences. Phylogenetic tree as determined by neighbor joining analysis, Bootstrap values are based on 100 replications.
112 isolation and cultivation technologies have resulted in the characterization of only a few extremophiles (e.g. Pyrococcus sp. and Desulfotomaculum sp.) from these systems (DeRosa et al., 1975; Zillig et al., 1980; Stetter, 1986; Huber et al., 2000a; Huber et al., 2000b; Goorissen et al., 2003). While the use of these traditional methods has resulted in an exponential increase in thermophile isolates in recent years (Stetter, 1996), they still exclude a large portion of the resident bacterial and archaeal assemblages within a given environment (Amann et al., 1995; Ward, 1998; Vetriani et al., 1998). Molecular genetic technologies provide a more comprehensive alternative to isolation/cultivation based research and have been used with much success in many high temperature environments (e.g. Pace, 1991; Barns et al., 1994; Reysenbach et al., 1994; Reysenbach et al., 2000). Despite the success of the use of these methodologies, their use in accessing the microbial diversity has eluded high temperature mud pools. This study used molecular genetic methodologies to compare the microbial community structure of a high temperature mud pool with nearby hot water features. Our goals were three-fold: to directly compare the Tokaanu mud pool with a similar hot spring based on physiochemical characteristics; to assess the microbial community structure of the Tokaanu mud pool and nearby hot springs using DNA fingerprinting; and to perform a more in-depth characterization of the bacterial and archaeal assemblages of Tokaanu mud pool. Our data confirms the presence of a unique and dominant microbial assemblage associated with the Tokaanu mud pool, which was absent from the neighboring hot springs. Analyses of the physiochemical characteristics of the Tokaanu mud and hot springs were critical to understanding how the local environment of the individual features might be structuring the microbial assemblages of the mud pool and hot spring
113 systems. The near-neutral pH, high temperature waters we observed for the Tokaanu pools are consistent with previous observations of the Tokaanu thermal system (Robinson and Sheppard, 1986). In a previous study of 13 high temperature mud pools from the TVZ, the Tokaanu mud pool was characterized by close to mean range
+ - - 3- measurements of temperature and concentrations of NH4 , NO3 + NO2 , and ΣPO4 , yet it was the only near-neutral mud pool observed during this study (Chapter 2). In the
+ present study, the Tokaanu mud pool showed elevated temperature and NH4
- - concentrations and deplete NO3 + NO2 concentrations in comparison to its neighboring hot springs (Table 3.3). Hot spring TOK 2 was most similar to the mud pool in terms of temperature (85°C) while TOK 10A hot spring was most similar based on the chemical signatures. Considering these observations and the close proximity of TOK 10A hot spring to the mud pool, it is likely that the same hydrothermal source fluids feed these
+ two pools. The elevated NH4 concentrations and elevated temperature of the mud pool compared to the TOK 10A hot spring were probably aided by the high-suspended particle loading of the mud pool. The solid phase fraction of the Tokaanu mud pool was examined in more detail in attempts to understand the biological and geochemical significance of the clay matrix. The solid fraction ranged in size from >12 µm to <2 µm in size (Figure 3.2).
Elemental analysis of these particles showed the presence of high amounts of silica (Si) and aluminum (Al) (Figure 3.1). Also observed was the presence of Na and to a lesser extent Ca (Table 3.4) in conjunction with the Si and Al. This is consistent with other descriptions of the elemental composition of soils rich in volcanic ash (e.g. Nanzyo, 1999). These analyses confirm our predictions of high silica concentrations for the Tokaanu mud pool based on previous reports of silica-rich soils in New Zealand (Saggar
114 et al., 1996; Lloyd-Jones and Hunter, 2001). The potential for high silica content and its affinity for the binding of DNA in mud pool samples was addressed in a previous study for optimization of DNA extraction methods (See Chapter 2). The charged surfaces of these clays could be of benefit to the resident microbes through particle attachment associations and/or metabolic catalysis. These charged surfaces might also compete with resident microbes through the scavenging of nutrients and e-donors rendering them biologically unavailable. The intense sheer forces associated with the mixing of the semi-fluid clay matrix might be a major selective factor in the microbial diversity of the system. Microbe/particle interactions within mud pool environments are area certainly worthy of future research focuses. For our assessment of the microbial communities of the Tokaanu pools, we utilized the DNA “fingerprinting” technique, DGGE. This method provides a “snapshot” fingerprint of the dominant members of microbial communities (Muyzer et al., 1993; Duineveld et al., 1998) and is very cost and labor effective compared to other methods. Its sensitivity can provide resolution down to a single base pair variation (Fodde and Losekoot, 1994) between individual members of the community and excised band sequencing can provide valuable information to infer phylogenetic placement. DGGE is somewhat limited in the ability to separate a large number of DNA fragments present in a single sample with the potential for comigration of closely related nucleotide sequences (Muyzer et al., 1993; Vallaeys et al., 1997) and lacks some sensitivity in the detection of minor community members (Muyzer, 1999). This method was well suited for the present study as these high temperature systems generally support low diversity communities and it allows for direct microbial community structure intercomparison of the different Tokaanu thermal features.
115 Initial DGGE assessment of the Tokaanu pools bacterial assemblages showed the mud pool to contain a unique community (Figure 3.3). These observations were somewhat expected due to the presence of a semi-solid medium coupled with the higher temperatures, reduced chemical environment observed for the Tokaanu thermal system. The bacterial profiles of hot springs TOK 10A, TOK 7, and TOK 6 shared a high degree of similarity in the observed dominant banding patterns. Although individually distinct from a chemistry standpoint, the similarities in temperature and pH of these pools may be the controlling factor. The TOK 2 hot spring, however, showed a very distinct bacterial community profile. This may be the result of this pool having a significantly higher temperature and lower pH when compared to the other hot springs in this study. These observations are consistent with studies of the microbial communities from other geothermal springs (Atkinson et al., 2000; Burton and Norris, 2000). Sequence analysis of DGGE bands 1-5 isolated from the bacterial profile of the Tokaanu mud pool revealed sequences related to uncultivated Flavobacteria and uncultivated, unclassified members of the bacteria from hot springs, hydrocarbon seep sediments, and gold mine bore hole waters (Table 3.5). Interestingly, band 10 isolated from the bacterial profile of hot spring TOK 10A (also present in TOK 7 profile) was also related to an uncultivated eubacterium from gold mine bore hole waters. The presence of bacterial signatures relating to those from deep subsurface environments suggested these pools are fed from the same hydrothermal fluid source and there may have been transport of bacteria into these pools by means of the hydrothermal fluids, at least on a local scale. Subsurface hydrothermal fluids are an ecological niche allowing for the potential distribution of thermophilic microbes through subsurface transport (Marteinsson et al., 2001). Sequence analysis of bands 6, 7, and 8 were representative of Methylobacterium
116 sp. and Thermus sp. and were observed as dominant bands in the profiles of hot springs TOK 6, TOK 7, and TOK 10A (Table 3.5). The lack of these strong signatures in the Tokaanu mud pool and TOK 2 hot spring suggests these pools may exceed the upper temperature limit for these bacterial species. The archaeal community of the Tokaanu mud pool was different from that of the hot spring communities (Figure 3.4). Much like the trends observed in the bacterial DGGE analyses, most the bands detected in the mud pool and TOK 2 hot spring profiles are absent from the profiles of the other hot springs. As previously observed, the shifts in these profiles coincided with significantly higher temperature for these two features. The similarities of the archaeal profiles for TOK 7 and TOK 10A appeared to support the prediction that these pools shared a common hydrothermal source fluid with potential for microbe transport through the subsurface. Comparative analysis of all the archaeal communities observed via DGGE showed the Tokaanu mud pool at the greatest distance in terms of the banding pattern similarity (Figure 3.6). The tight archaeal DGGE banding pattern and distance from other Tokaanu features inferred by the community level analysis suggested the mud pool archaeal community was composed of unique, highly related group of archaea. Several attempts to excise and amplify bands from the archaeal analysis failed. Bacterial and archaeal DGGE community analyses of the Tokaanu mud pool showed unique microbial signatures in comparison to neighboring hot springs. Although there were notable variations in temperature, pH, and chemistry between the sampled features, the delineating parameter was the solid/liquid mixed matrix of the Tokaanu mud pool. These clay particles can affect metabolic activity of microbes due their affinity for the absorption of microbes and their nutrient substrates, changing the
117 kinetics of metabolic reactions (Stotzky and Rem, 1966; Filip, 1973, Kunc and Stotzky, 1974; Kunc and Stotzky, 1977; Kunc and Stotzky, 1980; Smith et al., 1992; Bengtsson et al., 1993). The seemingly harsh presence of solid particles in suspension may provide several advantages for resident mud pool microbes. The susceptibility of microbes to sheering forces could provide a means of selectivity eliminating more fragile microorganisms (Bulut, et al., 1999). It has also been suggested that microbial adhesion to particles may help reduce predation, increase resistance to toxins, and may even provide a carbon source (Bright and Fletcher, 1983; Diab and Shilio, 1988; Bar-or, 1990; Bossier and Verstraete, 1996). Other studies have shown the presence of clay minerals to positively affect the capability of microbial cells to metabolize certain compounds (e.g. Hwang and Tate, 1997). Taking all of these possibilities under consideration, it is possible that the microbes inhabiting the Tokaanu mud pool have found a preferential niche over the seemingly less strenuous environment of a hot spring pool. In attempts to better understand and assess the microbial community structure of the Tokaanu mud pool, a more extensive sequence analysis was performed. The phylogenetically distinct OTUs of the bacterial clone library constructed from the Tokaanu mud pool were affiliated with Thermus sp., members of the Firmicutes, and γ
Proteobacteria. This is consistent with one study where a member of the Firmicutes, Desulfotomaculum solfataricum, was observed and isolated from a solfataric mud pool in Iceland (Goorissen et al., 2003). Other members of the γ Proteobacteria have been detected in fumeroles near mud pools, in Solfatara Crater, Italy (Glamocija et al., 2004) and in bacterial community analyses of Indonesian hot springs (Baker et al., 2001). The Tokaanu mud pool Bacteria formed three distinct groups among characterized Firmicutes, ε Proteobacteria, and Thermus sp. Group I bacterial OTUs from the
118 Tokaanu mud pool grouped most closely with known anaerobic, gram-positive eubacterial from the Clostridium-Bacillus subphylum. Baker et al. (2001) also reported similar extremely thermophilic Bacilli sequences from a high temperature acidic and near-neutral Indonesian hot springs to affiliate with the extreme thermophiles, Bacillus
+ caldovelox and Bacillus thermoleovorans. Considering the high concentrations of NH4 observed for the Tokaanu mud pool and the grouping of these phylotypes among known
+ NH4 dependent anaerobes (Ammoniphilus oxalaticus), a large percentage of the resident
+ bacterial community of the Tokaanu mud pool may require the high NH4 concentrations observed for their metabolism. Although sulfur species were not measured, representatives grouping among known sulfate-reducers suggested there is some sulfate- reduction based metabolism present among the Tokaanu mud pool bacterial community. A small number of bacterial clones grouped within known members of the thermophilic, anaerobic, hydrogen-oxidizing, ε-Proteobacteria (Caminibacter hydrogeniphilus). This group of ε-Proteobacteria uses elemental sulfur and nitrate as electron acceptors, producing hydrogen sulfide and ammonia (Alain et al., 2002). The presence of this
+ metabolic group may contribute to the high concentrations of NH4 and the deplete
- concentrations of NO3 observed in the Tokaanu mud pool chemical signatures. A third grouping of bacterial clones obtained from the Tokaanu mud pool fell closely to an aerobic bacterium, Thermus filiformis. The clones sequenced from archaeal library contained sequences closely related to previously characterized members within the Crenarchaeota. The Crenarchaeota is a small kingdom within the domain of the Archaea, containing representatives with the highest temperatures for growth (Stetter, 1996). Metabolically, members of this kingdom utilize aerobic and anaerobic metabolism that includes the
119 reduction of sulfur or nitrate and the oxidation of sulfur and hydrogen. Members of the Crenarchaeota have been found to be a common constituent of archaeal communities in many extreme high temperature environments including subterranean hot springs in Iceland (Marteinsson et al., 2001), high temperature subsurface fissure waters from gold mines (Takai et al., 2001), deep sea hydrothermal vents (e.g. Takai et al., 2000), mid- ocean ridge subseafloor habitats (Huber et al., 2002), and cold seeps (Knittel et al., 2005). The Tokaanu mud pool archaeal phylotypes grouped tightly together into a single unique clade distantly outgrouped by characterized members of the Crenarchaeota, Pyrobaculum sp. and Ignicoccus sp. This observation was quite significant as this clade falls directly between characterized Crenarchaeotes from submarine hydrothermal venting systems (Ignicoccus) and continental hot springs and solfataras (Pyrobaculum). Although mud pools are linked to continental hydrothermal activity, the mineral-rich clay matrix may have some similarities to the mineral-rich sulfide-based precipitates that form deep-sea vent chimneys. The tight grouping of the 13 Tokaanu mud pool archaeal phylotypes and their relative distance from other Crenarchaeotes suggested these mud pool archaea are a novel clade of Crenarchaeotes bridging the phylogenetic gap between deep sea and terrestrial species. This novel clade of archaeal sequences is not represented in the neighboring hot pools which suggests that these may represent organisms not only be unique to Tokaanu but to the other pools of New Zealand. The existence of this unique archaeal clade provides strong evidence for the existence of an endemic assemblage of Crenarchaeota within the Tokaanu mud pool environment.
Conclusion The data presented here suggests that the Tokaanu mud pool is a very complex environment supporting a unique microbial assemblage distinctly different from
120 other neighboring hot water features. This resident microbial community residing in its high temperature, near-neutral clay/fluid based matrix uses the dynamic characteristics of this system to obtain a metabolic and survival advantage over other hot spring systems microorganisms. The Tokaanu mud pool was found to harbor a diverse array of bacteria relating to the Firmicutes, epsilon Proteobacteria, gamma Proteobacteria, Thermus sp., and others from high temperature and deep subsurface environments. The archaeal community associated with the Tokaanu mud pool belonged to a unique group of microbes tightly affiliated within the Crenarchaeota. The strong relationship between the mud pool archaeal signatures and their relative distance from other characterized Crenarchaeotes may be representative of a community of microbes within the Archaeal domain that are unique to the Tokaanu mud pool system. Future research of the microbial assemblages of other mud pools and nearby hot springs may provide more evidence for microbial endemism in the mud pool environment.
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129 Chapter 4 CONCLUSIONS AND FUTURE DIRECTIONS
The use of molecular genetic technologies has been critical to the understanding of global biodiversity and the evolution of all domains of life. The value of these techniques has been increasingly evident in studies of extreme ecosystems like high temperature environments. These molecular studies of high temperature systems have largely focused on deep-sea hydrothermal vents and terrestrial hot spring environments. Although a common surface feature of continental hydrothermal systems, the high temperature mud pool has remained relatively uncharacterized using molecular- based technologies. These systems present a unique ecosystem of dynamic water-rock interactions coupled with a highly variable physiochemical environment that may present a modern analog to the early earth conditions. The aim of this study was an assessment of the microbial diversity of high temperature mud pools from 4 distinct, geographically isolated hydrothermal systems through the development of effective sampling and DNA extraction techniques and advanced molecular and geochemical technologies. We hypothesize that high temperature terrestrial mud pools are possible analogs to the primordial earth environment and may harbor unique and deeply branching assemblages of Bacteria and the Archaea. The initial phase of this study sought to address the following questions:
• Can extraction nucleic acids be effectively extracted from high temperature mud
130 pool samples?
• What is the longevity of exogenous, extracellular inputs of DNA into the mud pool system?
• How do the bacterial community structure and diversity of high temperature mud pools compare across 8 thermal regions within the Taupo Volcanic Zone, New Zealand?
• What are the similarities/differences of bacterial community structure and diversity of high temperature mud pools from 4 geographically isolated hydrothermal systems?
• How does bacterial community structure of high temperature mud pools correlate with geochemistry and measurements of temperature and pH?
The second phase of this research focused on a direct comparison of the microbial communities and geochemistry of the unique Tokaanu mud pool and nearby hot springs by addressing the following research questions:
• Are there correlations between the physiochemical environment of the Tokaanu mud pool and its neighboring hot springs?
• How do the microbial communities (Bacteria and Archaea) of the Tokaanu mud pool compare with communities from nearby hot springs?
• Based on 16S rDNA signatures from the Tokaanu mud pool, how does phylogenetic inference of mud pool signatures affiliate with other known microbial lineages?
In attempts to gain a more thorough understanding of high temperature mud pool biodiversity, I set out to characterize the bacterial communities of mud pools from
131 the TVZ, New Zealand; Rincon de la Vieja National Park, Costa Rica; Lassen Volcanic Park, USA; and Yellowstone National Park, USA. Attempts were also made to correlate these bacterial community structure data with the physiochemical environments of these unique systems. In order to assure implementation of the most effective tools for this study, methods were developed and tested for the effective extraction of DNA directly from mud pool samples representative of the entire community free of laboratory and environmental contamination (exogenous and extracellular DNA signatures). The results confirmed that that high quality, relatively pure, amplifiable DNA could readily be extracted directly from mud pool samples. Through rigorous laboratory testing and comparative analyses we were able to move forward with confidence that the microbial signatures observed in our community level analyses were indeed representative of resident, viable microbial cells. Based on our observations of the longevity and amplifiability of exogenous, extracellular DNA subjected to mud pool-like conditions, it was determined that residence time for these DNAs was extremely short due to the combined degradive forces of high temperature, acidic conditions, and the intense abrasive sheer forces present in mud pools. Analyses of 13 mud pools from the TVZ showed a broad range of physical, chemical conditions and associated bacterial assemblages across the individual pools. This variability was observed among pools from the same thermal region as well as across the entire range of thermal fields comprising the TVZ hydrothermal complex. Although quite different on a finer scale, general trends in the chemical signatures for
+ - - NH4 and NO3 + NO2 were observed for pools within a given thermal area suggesting chemical alteration of a shared hydrothermal fluid source might be altered by the localized effects of mineralogy and mixing with meteoric ground waters as it nears the
132 point of discharge. The dominant bacterial community members of the TVZ mud pools were affiliated with thermophilic and acidophilic Acinetobacters (gamma Proteobacteria). Other dominant signatures from these pools were affiliated with the Cytophaga-Flavobacteria and Methylobacterial members of the alpha Proteobacteria. A high degree of heterogeneity was observed among the minor members of the mud pool bacterial communities that included affiliations with thermophilic Firmicutes (Bacilli) sp. and members of the epsilon Proteobacteria. The physiochemistry and bacterial community profile of the Tokaanu mud pool showed very unique signatures in comparison to the other TVZ fields. The high temperature, near-neutral pH conditions of this pool likely selects for a unique microbial assemblage and was noted as an area of interest for future research. Bacterial community level analysis was also performed on mud pools from other geographically isolated areas of continental hydrothermal activity: Rincon de la Vieja National Park, Costa Rica; Lassen Volcanic Park, USA; and Yellowstone National Park, USA. Analysis of these communities showed a high degree of heterogeneity in the mud pool community profiles across the geographical boundaries. Some similarity was observed between pools from within the same thermal region, however, the structure of the individual pools bacterial communities were quite unique. Affiliations for the dominant community members from these mud pool systems were strong among sulfur and hydrocarbon metabolizing members of the γ Proteobacteria and members of the α Proteobacteria associated with hot spring environments. Other signatures from these pools included affiliations with thermophilic Flavobacteria and hyperthermophilic members of the Thermotogales. The dominance of these groups was consistent with findings from dominant bacterial signatures from TVZ mud pools. This
133 dominance may be attributed to the adaptability of these groups of bacteria to thrive in the extreme fluctuations in the physiochemistry of these mud pool environments. Although there is a small degree of similarity between pools from the same thermal field, the variation seen across the individual pools on both a localized and large geographical scale suggested the subsurface conduit network and localized physical strata near the point of discharge created unique mud pool environments even among pools in close proximately to one another and sharing a single deep hydrothermal source. The variability of the physiochemical environment of the individual pools may lead to unique, localized microbial assemblages. More thorough studies of the microbial assemblages, including the Archaea, and more extensive geochemical characterization are needed to fully understand these communities and the forces driving their community structure. Based on these initial surveys, research efforts were then focused on an intercomparison of the microbial community structure and physiochemical environment of the high temperature, near-neutral Tokaanu mud pool in comparison to nearby hot spring features. Analyses of the physiochemical parameters revealed reducing, near- neutral conditions across all of the Tokaanu pools. The higher temperature of the Tokaanu mud pool might be the result of the insulating properties of the clay/fluid matrix. Similarities in community structure were observed between the Tokaanu mud pool and the TOK 10A hot spring. These similarities may be the result of a shared source of hydrothermal fluids. Fingerprinting analyses of the microbial community structures of the Tokaanu features showed the mud pool to be quite unique in comparison to the hot spring pools. Analysis of dominant bacterial bands from these profiles showed strong affiliation with the Methylobacterium (mud pool) and Thermus sp. (hot springs). In contrast, the archaeal community of the mud pool was a less
134 diverse, unique community in comparison to the other pools. Finer scale phylogenetic analyses were performed for the mud pool bacterial and archaeal communities for better resolution of these unique communities. These analyses revealed 12 unique bacterial OTUs affiliating with the Firmicutes, Deinococci, gamma Proteobacteria, and sequences from an uncultivated, unclassified group within the eubacteria. Phylogenetic analyses of these sequences showed 3 distinct groupings of the Tokaanu mud pool bacterial OTUs among thermophilic Bacilli (Group 1), epsilon Proteobacteria and thermophilic H2 oxidizers (Group 2), and Thermus sp. (Group 3). Phylogenetic analysis of the Tokaanu mud pool archaeal community revealed 13 unique OTUs all grouping tightly within the Crenarchaeota. These archaeal OTUs formed a unique clade loosely affiliating with Pyrobaculum sp. (hot springs and solfataras) and Ignicoccus sp. (deep-sea hydrothermal vents) of the Crenarchaeota. The high degree of within clade similarity of the Tokaanu mud pool archaeal signatures and the absence of these signatures in the other Tokaanu features suggested this archaeal clade may be endemic to the Tokaanu mud pool system. The recognition of a potentially endemic clade of microbes associated with the Tokaanu mud pool is suggestive of a wealth of previous unrecognized microbial diversity in the mud pool systems. To fully understand and grasp the extent of mud pool microbial diversity and endemism, future research efforts need to focus on intense geochemical and mineralogical characterization to better understand the cycling of nutrients, elements, and metabolites through the system. Also, much focus should be applied to the characterization of other archaeal assemblages across continental and global boundaries. Along with the archaea, future studies should explore the potential presence of members of the Nanoarchaea within these unique systems. Another area of
135 future interest would be the molecular exploration of microbial metabolism within the mud pool environment. Coupled with intense geochemical analyses, metabolic characterization may reveal unique metabolic pathways and signatures coupled with the presence of a high temperature clay matrix. Lastly, attempts should be made to isolate and culture organisms from mud pool environments. Recently, a thermophilic archaean grouping within the crenarchaeotal Tokaanu mud pool clade has been isolated and grown in culture by members of the Thermophile Research Unit, University of Waikato, New Zealand. Full characterization of this novel isolate will lead to a better understanding of the biological and metabolic potential of the Tokaanu mud pool.
136 Appendix A ABBREVIATIONS
137 Table A.1 Commonly used abbreviations.
Abbreviation Phrase
CTAB/PVP/β-ME cetyltrimethylammonium bromide- polyvinylpyrrolidone-β-mercaptoethanol
DGGE denaturing gradient gel electrophoresis OTU operational taxonomic unit PCR polymerase chain reaction RFLP restriction fragment length polymorphism rRNA ribosomal RNA TVZ Taupo Volcanic Zone
138