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LSU Master's Theses Graduate School

2008 Characterization of the lucinid bivalve- symbiotic system: the significance of the geochemical on bacterial symbiont diversity and phylogeny Angela Marissa Green-Garcia Louisiana State University and Agricultural and Mechanical College, [email protected]

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Recommended Citation Green-Garcia, Angela Marissa, "Characterization of the lucinid bivalve-bacteria symbiotic system: the significance of the geochemical habitat on bacterial symbiont diversity and phylogeny" (2008). LSU Master's Theses. 1970. https://digitalcommons.lsu.edu/gradschool_theses/1970

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CHARACTERIZATION OF THE LUCINID BIVALVE-BACTERIA SYMBIOTIC SYSTEM: THE SIGNIFICANCE OF THE GEOCHEMICAL HABITAT ON BACTERIAL SYMBIONT DIVERSITY AND PHYLOGENY

A Thesis

Submitted to the Graduate Faculty of the Louisiana State University and Agricultural and Mechanical College In partial fulfillment of the Requirements for the degree of Master of Science

in

The Department of Geology and Geophysics

by Angela Marissa Green-Garcia B.S., University of Houston, 2003 May 2008

DEDICATION

This thesis is dedicated to my father, Rudi, who in and death, has been a driving force behind my pursuit of an education, and to my family, who has continuously supported me and encouraged me to achieve all of my goals. Thank you and I love you all.

ii ACKNOWLEDGEMENTS

I would like to thank Dr. Annette Engel for all her patience and help along this (what has

felt like) endless process of researching, writing, and presenting the thesis. She has served as a

constant source of support and guidance and her assistance has been invaluable. I could not have asked for a better mentor; she is someone I truly respect and admire. I would also like to thank

the other members of my committee, Dr. Laurie Anderson and Dr. Huiming Bao, for all of their

assistance. Also, thanks to Audrey Aronowsky for her help and direction, Maureen Thiessen for

her awesome dedication and work ethic, and to Wanda LeBlanc for her help with soil XRD analysis, her patience, and her smile.

In addition, I would like to thank my family, for all of the love and support that they have

given me throughout all these years working to get to this point. Jeannie, you are a fabulous mother and thanks for being my shoulder to lean on and a constant source of encouragement.

Angel, Tashi, and Adan thanks for making me strong. Ms. Lady G and Ms. Donna, thanks for all the encouraging kisses and all the snuggling. I love you all. Phroggie, thanks for the love and

always taking care of me.

For all of the amazing people I have met and shared this graduate experience with

…whew! I can’t believe it’s over. Issaku; it has been fun, you are like a brother. Thanks for all

the encouragement. , thanks for the pep; you are a great girl and have been a delightful

perky friend, as well as an amazing, positive voice through this experience. And, to TEAM

NORAD, thanks for all of the unforgettable experiences in the field, all the GP, and the endless

laughter. And, to anyone else I have not named, thank you.

Funding for the thesis was provided by the Geoscience Alliance to Enhance Minority

Participation (GAEMP) (NSF-OEDG #0303138) in the Department of Geology and Geophysics,

and Sigma Xi by a Grants-in-Aid Research award. Thank you.

iii TABLE OF CONTENTS

DEDICATION...... ii

ACKNOWLEDGMENTS ...... iii

LIST OF TABLES...... vi

LIST OF FIGURES ...... vii

ABSTRACT...... viii

INTRODUCTION ...... 1 Background...... 2 and Antiquity ...... 3 - Relationship ...... 4 Objectives and Hypothesis...... 6 Importance of Research ...... 8

MATERIALS AND METHODS...... 9 Sample Collection...... 9 Mineralogy by X-Ray Diffraction (XRD) ...... 10 Acquisition and DNA Extraction...... 12 Sediment and DNA Extraction ...... 13 Polymerase Chain Reaction (PCR) Amplification, Cloning, and Sequencing ...... 13 Sequence Analysis ...... 14

RESULTS ...... 16 Aqueous Geochemical Analysis ...... 16 Core Mineral Analysis ...... 16 Gill Endosymbiont Diversity ...... 18 Sediment Bacterial Diversity from Cedar Key, Florida...... 26 Microbial Diversity with Core Depth ...... 32

DISCUSSION...... 36 Habitat Geochemistry and Mineralogy...... 36 Intra- and Inter- Diversity...... 38 Free-living Bacterial Diversity ...... 39 Endosymbiont Sequence Diversity and Possible Metabolic Implications...... 41

CONCLUSIONS...... 43

REFERENCES ...... 45

APPENDIX A: CORE MINERAL ANALYSIS AND XRD...... 53

iv APPENDIX B: TAXANOMIC DISTRIBUTION OF BACTERIAL CLONES ...... 61

VITA...... 65

v LIST OF TABLES

1. Field geochemical data from Cedar Key, Florida...... 16

2. Cation concentrations and anion concentrations of ocean and pore water Cedar Key, Florida...... 17

3. Percentage abundances of minerals in core , Cedar Key, Florida ...... 17

4. Average genetic distance within host species form 16S rRNA gene sequences ...... 22

5. Average genetic distance between endosymbiont 16S rRNA gene sequences from different host taxa ...... 22

6. Distribution of 16S rRNA gene bacterial sequences from sediment cores at Cedar Key, Florida ...... 33

vi LIST OF FIGURES

1. Maps of sample collections sites...... 11

2. Rarefaction curves for endosymbiont 16S rRNA gene sequences ...... 20

3. Phylogenetic reconstruction based on Jukes-Cantor neighbor-joining analysis of 16S rRNA gene sequences of endosymbiont sequences retrieved from lucinid from Cedar Key and 16S rRNA sequences from sediments, representing the free-living habitat...... 21

4. Phylogenetic reconstruction based on Jukes-Cantor neighbor-joining analysis of gill 16S rRNA bacterial sequences belonging to previously identified g-proteobacterial groups. Sequences retrieved from hosts were collected from Cedar Key (CK), Lemon Bay (LB), and Jack Island (JI), Florida, and the Mouth of Pigeon Creek (MPC), The Bahamas...... 23

5. Phylogenetic relationship based on Jukes-Cantor neighbor-joining analysis of 16S rRNA gene sequences of sequences retrieved from lucinid gills from Jack Island, Florida, and the Mouth of Pigeon Creek, San Salvador, Bahamas...... 25

6. Rarefaction curves for the two core sequence sets from Cedar Key ...... 27

7. Bacterial diversity of each of the core sediment 16S rRNA gene sequence clone libraries, shown grouped with increasing depth (every 10 cm) ...... 35

vii ABSTRACT

Extensive characterization of a single lucinid bivalve habitat was conducted to

characterize the relationship between host bivalve and thiotrophic bacterial endosymbionts. For

lucinids, the ecological and evolutionary relationships between hosts and endosymbionts are

poorly understood. Reconstructing the evolutionary history of lucinid endosymbiosis, and the

geologic significance of the association, has been hampered by insufficient knowledge of

endosymbiont ecology and taxonomic diversity. Host ( nassula and

Phacoides pectinatus) were collected from Cedar Keys, Florida, within the top 15-20 cm of the

sediment in grass beds. PCR amplification and sequencing of bacterial 16S rRNA genes from

lucinid gills and sediment cores retrieved ~900 sequences. Based on comparative phylogenetic methods, gill endosymbiont sequences were most closely related to uncultured

Gammaproteobacteria associated with symbiosis, and specifically to lucinid endosymbionts (97-

99% sequence similarity) and not to free-living organisms. Not all gill sequences were

genetically identical, with intra- and inter-gill sequence diversity. Sediment diversity was high,

represented by 13 major taxonomic groups, including equally dominant , and Delta-

and Gammaproteobacteria. Other organisms included the , ,

Spirochetes, and . Rare (<0.3%) sequences from the sediment were related to lucinid gill endosymbionts. Results support the hypothesis that recruitment of free-living organisms is likely. Based on habitat geochemistry, however, the bacteria are constrained to reducing conditions, and this may be reflected in the habitat types colonized by the host. Habitat-host- symbiont diversity was evaluated from other locations from Florida and The Bahamas. 16S rRNA gene sequences retrieved from those hosts revealed that not all lucinid endosymbionts belong to the Gammaproteobacteria, because some sequences were most closely related to

viii Alphaproteobacteria. One sequence was most closely related to Methylobacterium spp., which may indicate that dual symbiosis (thiotrophy and methanotrophy) in lucinid bivalves may be possible. Together, these results are significant to paleoecological and evolutionary studies using lucinids in the record (e.g. isotope studies).

ix INTRODUCTION

Symbiosis is considered to be one of the most important driving factors for (Gil et al., 2004; Stewart et al., 2005). Symbiosis is a strictly inter-species association (Savazzi et al.,

2001), and symbiotic associations between bacteria and are wide-spread, from

mammalian or insect digestive systems to trophosome tissues in tubeworms (e.g., Fullarton et al.,

1995; Gil et al., 2004; Goffredi et al., 2004). The exact functional and evolutionary of many symbiotic relationships (e.g., species dependency, host specificity, toxicity effects within

the environment, etc.) remains unclear (Taylor and Glover, 1997; Distel, 1998). Symbiosis

among bacteria and eukaryotes may have allowed eukaryotes an evolutionary strategy to

overcome limited metabolic capabilities in extreme (Gil et al., 2004). Therefore, it is not

surprising that symbiotic bacteria- associations have been widely studied, especially

these associations relying on chemosymbiosis (e.g., Wiley and Felbeck, 1995; Gros et al., 1996;

Krueger et al., 1996; Distel, 1998; Savazzi, 2001; Gros et al., 2003a; 2003b; Imhoff et al., 2003;

Duperron et al., 2005; Gil et al., 2004; Stewart et al., 2005; Suzuki et al., 2005; Duperron et al.,

2006; Taylor and Glover, 2006; Caro et al., 2007; Duperron et al., 2007). In a chemosymbiotic

relationship, bacteria live inside of a host and metabolize reduced compounds, such as hydrogen

sulfide, and fix inorganic carbon to organic carbon that may or not be used by the host (Distel,

1998; Stewart et al., 2005). Symbiosis between and chemolithoautotrophic bacteria

was first discovered in the late 1970’s when productive were found at the deep-sea

hydrothermal vents, a place originally considered too extreme for life to thrive (Distel, 1998;

Arndt et al., 2001; Goffredi et al., 2004).

One symbiotic association that has received attention recently is between members of the

Lucinidae family of the and sulfur-oxidizing (thiotrophic) endosymbiotic bacteria

(Dando and Southward, 1986; CoBabe, 1991a; 1991b; Distel, 1998; Barnes and Hickman, 1999;

1 Clark and Moran, 1999). Chemosymbiosis in the Lucinidae Bivalvia was first reported in the

1980’s (Taylor and Glover, 2006). In this thesis, I continue to investigate this association by examining the genetic diversity and ecology of a variety of lucinid hosts and their bacterial endosymbionts from spatially separated locations, as well as the bacterial diversity of the host habitat (marine siliciclastic sediments) to understand symbiont ecology. The results from this study provide unique genetic evidence for bacterial endosymbiont diversity, as well as indicate that geography may play a role in the diversity of the bacteria-lucinid association.

Background

Chemosymbiosis is an ancient evolutionary mechanism found in some Bivalvia that has

been suggested as a pervasive evolutionary mechanism to allowing this group to thrive for >400

million years (Watson and Pollack, 1999, Fortey, 2000; Bailey et al., 2006). Chemosymbiosis

appears ubiquitous in the Lucinidae family. All lucinid species studied to date (>25) have

thiotrophic bacteria that symbiotically colonize the host gills (Taylor and Glover, 1997); no other

symbiotic association has been described for the lucinids (e.g., methanotrophy). Lucinid

bivalves live in a wide variety of habitats including fjords, shallow marine sea grass beds,

mangrove swamps, and deep-sea cold seeps and hydrothermal vent sites (e.g., Wiley and

Felbeck, 1995; Gros et al., 1996; Imhoff et al., 2003; Duperron et al., 2005), being situated at or

near the oxic-anoxic interface.

The antiquity of lucinid symbioses, widespread geographic distribution of the lucinids,

habitat variety, and the fact that symbiosis has been found in all lucinid species make the

symbiotic relationship an intriguing system for study. As such, physiological, biochemical,

ecological, morphological, isotopic, enzymatic, microscopic, molecular and phylogenetic studies

have been conducted for more than two decades on these bivalves and their endosymbionts

(Wiley and Felbeck, 1994; Gros et al., 1996; Imhoff et al., 2003; Duperron et al., 2005; Stewart

2 et al., 2005; Suzuki et al., 2005; Duperron et al., 2006; Taylor and Glover, 2006; Caro et al.,

2007; Duperron et al., 2007).

Morphology and Antiquity

Lucinids are eulamellibranch bivalve mollusks, having a heterodont shell hinge, leaf-like gills, well-developed siphons, reduced guts, and large thick, fleshy, colored (yellow, black, grey) gills colonized by the endosymbionts (Allen, 1960; Distel and Felbeck, 1987; Wiley and Felbeck, 1995). The thiotrophic endosymbionts are housed in specialized cells called bacteriocytes (Duperron et al., 2005; Caro et al., 2007). Modern Lucinidae shells are aragonitic

(Taylor et al., 1969, Falini et al., 1996), having discoidal to subpherical shapes, and ranging in size from a few millimeters to >100 mm (Taylor and Layman, 1972; Taylor and Glover, 2007).

Lucinids exhibit cell types which are indicative of their symbiotic nature, including bacteriocytes, membrane whorls, but also intercalary cells (Distel, 1998; Savazzi, 2001). The shells contain lipids and stable isotope ratios that have been identified to indicate dietary preference, growth rate, sexual maturity, and ecological and environmental conditions (e.g., temperature and chemistry) (Jones et al., 1988; CoBabe and Pratt, 1995; Langlet et al.,

2002). In addition, internal features of their valves contain scars representing features linked to symbiosis (e.g., detached pallial vessel) (Taylor and Glover, 2006).

The earliest member of the Lucinidae, Iliona prisca, dates back to the (Taylor and Glover, 2006). Several other Silurian chemosymbiotic bivalve species (thyasirid, mytilid, and solemyid) are from habitats interpreted to be cold-seep sediments (Distel, 1998). Because all modern symbiont-bearing lucinids share similar morphological characteristics and living positions with I. prisca, this species is inferred to have also been a host for thiotrophic bacteria

(Taylor and Glover, 2006). Although only well-preserved shells, whose matrix was intact

3 (Distel, 1998), have been studied, δ13C isotope analyses of the matrix from the fossilized shells suggest chemosymbiosis did exist in ancient bivalves (CoBabe, 1991a; 1991b).

Host-Endosymbiont Relationship

Relationship to Environment

Host bivalves orient themselves in their habitat to ensure optimal substrate attainment for the endosymbionts, and to minimize the amount of sulfate lost to abiotic oxidation (Stewart et al., 2005). Consequently, lucinids predominantly occupy the oxic-anoxic interface allowing them to burrow into anoxic sediments so that the release of hydrogen sulfide from deeper sediment layers is enhanced (Felbeck et al., 1983; Savazzi, 2001). The hydrogen sulfide in the lucinid habitats is produced by free-living sulfate-reducing bacteria (Stewart et al., 2005).

Because lucinid endosymbionts are metabolically dependent on both sulfide content and dissolved concentrations, Arndt et al. (2001) show that an increase in the level of sulfide under periods of anoxia was due to a decrease in endosymbiotic metabolism. The movement and water pumping of the hosts within the habitat are more important to maintaining slightly oxygenated conditions so that the endosymbionts can be metabolically active. Heme are associated with the gill cytoplasm in symbiotic bivalves (Frenkiel et al., 1995; Kraus, 1995).

The purpose of these proteins is not fully understood, but they are inferred to bind oxygen to prevent spontaneous oxidation of sulfide (Frenkiel et al., 1995; Kraus, 1995). Sulfide oxidation by endosymbionts is vital to geochemical sulfur cycling within marine environments (Jorgenson and Bak, 1991; Craddock et al. 1995; Arndt et al., 2001). Thioautotrophic endosymbionts oxidize the sulfide for energy and fix CO2 into organic carbon compounds that can be used by the host (Krueger et al., 1996; Stewart et al., 2005). Organic nutrients are thought to be transmitted to the host via translocation, or by direct ingestion of the endosymbionts (Stewart et al., 2005); fatty acid profiles of the host that mimic those of the endosymbionts suggests that

4 latter (Pond, 1998; Suzuki et al., 2005), although previous carbon isotope studies could not conclusively show this (CoBabe, 1991a; 1995).

Endosymbiont Acquisition

The acquisition of the bacterial endosymbionts by the bivalve hosts has been intensely

studied (Gros et al., 1996; Distel, 1998; Peek et al., 1998; Fortey, 2000; Savazzi, 2001; Gros et

al., 2003a). Two mechanisms have been explained by Distel (1998) for the transmission of

bacteria to host. The first is reproductive, from parent to offspring during gametogenesis (Distel,

1998). The second mechanism is through environmental acquisition; this type of acquisition

indicates that the endosymbionts have the ability to live independent of their host (Distel, 1998).

For lucinids, previous research determined that the bacterial endosymbionts were free-living in

the host habitat (Gros et al., 1996; Peek et al., 1998; Fortey, 2000; Savazzi, 2001; Gros et al.,

2003a; Gros et al., 2003b; Caro et al., 2007), confounding the understanding of the degree to

which the host and endosymbionts depend on one another. Specifically, Gros et al. (1996) found

the environmental form of transmission for the host species, orbicularis, as the

symbionts were only found within the host after the larval growth stage. This starkly contrasts

what is known about other bacterial symbionts for other marine bivalves and with vertical

transmission where no free-living bacterial symbionts have been identified to date (Imhoff et al.,

2003).

Endosymbiont Identity

The thiotrophic endosymbionts of Lucinidae belong to a monophyletic group of

within the , the Gammaproteobacteria (Distel, 1998; Gros et al., Gros et al., 2003b). Distel et al. (1988) suggest that genetically identical thiotrophic species are found in multiple lucinid hosts, regardless of geographic location. However, these results are questionable because hosts and symbionts from the same locale were not examined (e.g., Gros et

5 al., 1996; Gros et al. 2003a; 2003b), and much of the species-level research for the bacteria had

been done with a limited number (8) of partial 16S rRNA gene sequences acquired from

different hosts (Distel et al., 1988; Distel et al, 1993; Durand et al., 1996; Distel et al, 1998).

Hence, endosymbiont genetic diversity has recently been questioned. Duperron et al. (2007)

identified multiple phylotypes of bacterial endosymbionts from the gills of the cold-seep ,

Lucinoma aff. kazani. The dominant 16S rRNA gene phylotypes belonged to the

Gammaproteobacteria, and were most closely related to an endosymbiont sequence from

‘Parvilucina’ costata, a shallow tropical lucinid. A second phylotype was most closely related to

Spirocheta coccoides, an autotrophic spirochete found in the hindgut of a termite. This work was the first to suggest that lucinid endosymbionts may be genetically diverse.

Moreover, endosymbiont diversity was studied from the nucleic acid content of cells separated in a flow cytometry study of (Caro et al., 2007). Caro et al. (2007) propose

that the host controls the entry and distribution of bacteria into and within the bacteriocytes.

They also determined that bacteria cell maturation, based on size and sulfur content, differed from the apical to basal tip of the cells within the bacteriocytes. This work suggests that up to seven genetically distinct subpopulations of varying size and intracellular nucleic acid content can exist within individual host specimens.

Objectives and Hypotheses

Symbiosis in the lucinid bivalves is one of the oldest identified symbiotic relationships

currently known. Unfortunately, given the recent research on endosymbiont diversity, the degree

of dependence between the endosymbiont and hosts is not well understood (Gros et al., 1996.

Gros et al.2003b; Duplessis et al., 2004). For example, characterizing the diversity and

distribution of free-living bacteria within a lucinid habitat will lead to a better understanding of

the degree of dependence between the host and bacterial species. Due to the high number of

6 lucinid species worldwide, and because at least some endosymbionts can exist outside of the host

(Gros et al., 1996; Gros et al., 2002a), it is possible that previous work overlooked host endosymbiont diversity and specificity of bacteria to specific hosts. Therefore, a detailed evaluation of the diversity and ecology of both endosymbionts and hosts, from one location, is needed before the evolutionary significance, and possible cophylogeny, of the thiotrophic symbionts and the bivalve species can be explored.

For my thesis, I had three main goals:

1) Define lucinid and bacteria habitat solid- and aqueous-phase geochemistry;

2) Characterize the bacterial diversity of the sediments where host organisms are found, as well

as from the gills of host taxa collected at one site;

3) Compare sediment and endosymbiont bacterial diversity to bacterial endosymbiont diversity

from different lucinid species collected from geographically separate locations.

For this work, I choose a siliciclastic sediment site off the coast of Florida, at Cedar Key, Gulf of

Mexico. Other comparative sites where lucinids were collected included Lemon Bay, Florida,

Jack Island, Florida, and San Salvador, The Bahamas.

The three hypotheses that I tested in the study were:

1) Hypothesis 1: Free-living symbiont diversity will be unique to specific sediment and water

geochemical conditions.

2) Hypothesis 2: Bacterial endosymbionts from lucinids at Cedar Key, Florida, will be closely

related to free-living bacteria from the same habitat.

3) Hypothesis 3: Bacterial endosymbionts from different species of lucinid hosts collected at

geographically separated locations will be genetically distinct from one another.

7 Importance of Research

Results from this research will lead to a more thorough understanding of the evolutionary and ecological relationship between lucinid bivalves and their thiotrophic symbionts. As this research is the first to characterize the bacterial diversity in the host habitat sediment, nearly exhaustive characterization of a single site will provide a much needed baseline for future research that should study different lucinid geochemical and geological habitats.

This research will also provide a background for modern day symbiotic relationships that can be applied to ancient symbiotic relationships.

8 MATERIALS AND METHODS

Sample Collection

Sediment, water, and lucinid samples were collected from Cedar Key, Florida (29º 7.457’

N; 803º 18.483’ W), located on the Gulf of Mexico side of the state (Figure 1). Some outer

islands and sand bars are just off of the coast at Cedar Key. This shallow water habitat supports

dense beds that are biologically active, especially with lucinid species (personal

communication, Harry G. Lee, Jacksonville Shell Club), and the targeted lucinid species were expected to be well represented within the first 10 cm of the seagrass beds. Species previously

retrieved from the area include Lucinisca nassula, pectinatus, and Stewartia floridana

(http://www.jaxshells.org/cedarkey.html).

Ten total individual lucinid specimens, including Lucinisca nassula and Phacoides pectinatus, were collected from Thalassia seagrass beds off of Atsena Otie Key near Cedar Key during a three day sampling period in June, 2006. Two sediment cores that were ~30 cm in

length and ~4 cm in diameter were collected in PVC tubes. The cores were collected from

within 1 m of the lucinid collection site, and were separated from each other by 1m. Cores were

stored at -20 oC until they were aseptically cut into 3 depth intervals each,~10 cm thick.

Filtered and raw pore fluids and ocean water were collected for geochemical analysis.

Pore fluids were sampled from the seagrass bed sediments at a depth of ~30 cm by low-flow

pumping through a mini-piezometer. Physiochemical parameters were determined on site

immediately using standard electrode methods (American Public Health Association [APHA],

1998), including temperature and pH on an Accumet AP62 meter with a double junction

electrode (Accumet, Fisher Scientific, USA), total dissolved solids (TDS) and temperature on a

YSI-85 meter (YSI, Inc., Yellow Springs, OH, USA), and other parameters (e.g., oxidation-

reduction potential on an Ultrameter).

9 Dissolved sulfide and trace-level dissolved oxygen (DO) was measured using the Methylene

Blue and Rhodazine D CHEMetrics (Calverton, VA) colorimetric methods, respectively, on a portable V-2000 multi-analyte photometer (APHA, 1998). Anions were measured by ion chromatography (IC) from 0.2 μm-filtered samples on an ICS-90 (Dionex, USA). Cations were measured from an acid-preserved 0.2µm-filtered sample by inductively coupled plasma mass spectrometry (ICP-MS) at the University of Texas at Austin (EPA method 9056; Manual SW-

846, Methods for Evaluating Solid Waste, Physical/Chemical Methods; http://www.epa.gov/epaoswer/hazwaste/test/main.html).

Additional lucinid specimens, sampled by Dr. L. Anderson and Dr. A. Aronowsky for another project, were used as comparison species in this study (Figure 1). Three Lucinisca nassula host organisms were collected from Lemon Bay (26 º 55.952’ N; 82 º 21.254’ W), located on the Gulf of Mexico side of Florida, ~163 km from Cedar Key. One of these specimens was collected from nearshore sediments under Syringodium sea grass. The other two specimens were collected in an intertidal flat of Thalassia grass that was separated from the shore by a muddy channel.

Several Phacoides pectinatus host specimens were collected from the Atlantic side of Florida

(Figure 1), including one host from Jack Island (27 º 30.095’ N; 80 º 18.500’ W) in Ruppia maritima sea grass beds and macroalgae, and another specimen from Thalassia sea grass beds at the Mouth of Pigeon Creek, San Salvador, The Bahamas (23 º 57.810’ N 74 º 29.331’ W).

Taxonomic designations at the level in this thesis are after Taylor and Glover (2000).

Sediment Mineralogy by X-Ray Diffraction (XRD)

Two grams of sediment from each core interval were homogenized by mixing with a

Micronizing Mill (McCrone, USA) equipped with corundum micronizers. Six total sediment

10 samples were processed. Samples were homogenized for 3 min with 10 mL of 100% ethanol, and then centrifuged for 30 minutes to separate solid and suspended particle phases.

Jack Island Cedar Key (JI) (CK G1-9)

Lemon Bay (GA; BG)

San Salvador (MPC)

Figure 1: Maps of sample collections sites. Cedar Key and Lemon Bay, Florida, located on the Gulf of Mexico side, Jack Island on the Atlantic side of Florida, and San Salvador, located on the Atlantic side of The Bahamas. Maps were acquired and modified from www.awesomeflorida.com and www.fishforfun.com.

11 The supernatant was discarded and the remaining sediment slurries were dried overnight

at 60 ºC. Each dried sample was ground to a fine powder using a mortar and pestle. Samples

were analyzed by a BRÜKER Siemens D-5000 XRD using the operating setting of 40 kV and 30

milliamps. The XRD diffractograms of 27 of the most common sedimentary minerals were

compared for identification. XRD analysis were processed using JADE pattern processing software version 6.1 (MDI, USA). Minerals were identified based on diffraction peak

position; peak identification was checked by hand.

Gill Acquisition and DNA Extraction

Ten lucinid bivalve hosts were aseptically dissected to obtain their gills using a Heerburg

WILD M5A steereozoom microscope set at a magnification of 500x. One gill and one foot were

removed from each specimen. The second gill was frozen as reserve tissue. For each lucinid host

sample an entire gill, measuring between 5 and 10 mm in length (depending on individual) and

half of each foot sample were used.

DNA was extracted from each tissue sample using the Qiagen DNeasy extraction kit,

following manufacturer’s instructions. Briefly, each tissue sample was placed in 180 µL of

tissue lysis buffer and 20µL of concentrated proteinase K solution. The tissue was incubated at

55 oC overnight on a shaker table at 225 rpm. In order to completely digest the tissue, 200 µl of

AL buffer was added and samples were incubated at 70 oC in a water bath for 10 minutes. Two

hundred µl of 100% ethanol was added to each digest and the mixture was pipetted into the

DNeasy mini-spin column. DNA was eluted from the columns, following several steps of

centrifugation and the addition of DNeasy solutions. The quality and quantity of the DNA was

checked by running each extraction on a 1% TBE gel with ethidium bromide staining and using a

Nano Drop ND-1000 spectrophotometer. The amount of nucleic acids per sample ranged from

30 to 100 ng/µl. Resulting DNA was stored at -20ºC in TE.

12 Sediment DNA Extraction

Each of the sea grass sediment cores from Cedar Key were divided into three depth intervals (0-10 cm, 10-20 cm, and 20-30 cm). Approximately 3 g of sediment were aseptically removed from the center of the core for each sample. Two samples were used from each depth interval. The sediment was suspended into 5 ml sterile cell lysis buffer (10 mM Tris and 100 mM EDTA, pH 8.0) and shaken on a hand-shaker for 2 hr to disaggregate the material and wash cells from sediment surface. After shaking, 20µl of proteinase K were added to the sediments and the samples continued to shake at room temperature for 2 hr. Large particles in the suspensions were allowed to settle for 10 minutes, then ~3 mL of cloudy supernatant was taken and nucleic acids were extracted based on methods previously described by Engel et al. (2004)

and modified from the DNeasy extraction kit (Qiagen). Modifications included adding 5 mL of

fresh extraction buffer and 40 µl of additional proteinase K to the supernatant; samples were shaken at 225 rpm overnight at 55 oC; 750 µl of the sediment solutions were separated into

smaller tubes and 300µl of a protein precipitation solution (7.5 M NH4-acetate) was added; the

mixture was incubated on ice for 15 min and centrifuged; the supernatant was transferred to 95%

isopropanol and incubated on ice for 30 min to precipitate the nucleic acids. The pellet was

cleaned with 80% ethanol and eluted in 50µl of TE (10 mM Tris/ 0.5 mM EDTA) buffer, then

stored at -20ºC until use. The quality and quantity of the nucleic acids was checked by running

on a 1% TBE gel with ethidium bromide staining and using a NanoDrop ND-1000

spectrophotometer. The amount of nucleic acids per sample ranged from 100 to 1500 ng/µl.

Polymerase Chain Reaction (PCR) Amplification, Cloning, and Sequencing

Nearly full length 16S rRNA gene sequences from gills, core sediments, and foot tissues

were PCR amplified using the bacterial universal primers 8F (5’-AGAGTTTG ATCCTGGCTC-

AG-3’) and 1510R (5’-GGTTACCTTGTTACGACTT-3’) (Ausbel et al., 1990).

13 Amplification was performed with a MJ Research Dyad Disciple thermal cycler (Biorad, USA)

with 5 U/µl ABGene Taq DNA polymerase (ABGene, Thermo Fisher Scientific, USA), 10

mg/ml bovine serum albumin (BSA, Fisher Scientific), 10 mM dNTPs, and 10 mM MgCl2.

Optimal PCR conditions included an initial hot start at 94ºC for 4 min; denaturation at 94ºC for 1

min, primer annealing at 47ºC for 1 min, chain extension at 72 ºC for 3 min, this process was

repeated for 29 more cycles; and a final extension step of 20 min at 72 ºC. Amplified PCR

products of the correct size (~1500 bp) from each sample were purified by using a 0.7% TAE

low-melt agarose gel with a Wizard PCR prep DNA purification kit (Promega Corp., USA), following manufacturer’s recommendations. Concentrations of PCR products and purity were determined by spectrophotometry (NanoDrop).

Purified PCR products were cloned using TOPO Cloning Kit with the PCR2.1-TOPO® vector, according to manufacturer’s instructions (Invitrogen Corp., USA).

Optimal ligation reactions were achieved by overnight incubation at 14 ºC. Resulting clones were screened by PCR with the M13 forward (5’- GTAAAACGACGGCCAG-3’) and M13 reverse (5’- CAGGAAACAGCTATGAC -3’) vector primers (Invitrogen Corp., USA). Clones were checked for the correct size insert using 1% TBE gels with ethidium bromide staining.

M13 amplified clone sequences that were the correct length were diluted to between 80-

100 ng/µl and sequenced in both directions by capillary sequencing at the High-Throughput

Sequencing Solutions facility at the High-Throughput Genomics Unit (HTGU), Department of

Genome Science (http://www.htseq.org)., University of Washington, Seattle, Washington.

Sequence Analysis

Resulting sequences were assembled using Contig Express, a component of Vector NTI

Advance 10.3.0 (Invitrogen Corp., USA) (http://www.invitrogen.com/). Sequences were

14 subjected to BLAST Searches using GenBank (http://www.ncbi.nlm.nih.gov/) to determine

sequence similarities to cultured and not yet cultured organisms.

Nucleotide sequences were aligned using NAST (Nearest Alignment Space Termination)

in greengenes (DeSantis et al., 2006), and adjusted manually in BioEdit 7.0.4.1 (Hall, 1999). A

neighbor joining phylogeny with branch support from 500 bootstrap replicates was constructed

with the program MEGA (ver. 3.1) (Kumar et al., 2004) using the Jukes-Cantor model based on

concepts presented in Tamura et al. (2004). Distance matrix files were assembled from

nucleotide sequences in Phylip3.67 (Felsenstein, 2005) using the Jukes-Cantor model for

nucleotide substitution. Operational taxonomic units (OTUs) were defined and

comparisons were made using DOTUR (Distance Based Operational Taxonomic Units and

Richness Determination) (Schloss and Handelsman, 2005). OTUs (phylotypes) for the

sequences were determined based on nearest neighbor distance to 99% sequence identity. A

sequence identity of 99% was used to conservatively assign species level affinities.

To determine the coverage of the clone library, rarefaction curves were generated using

the approximation algorithm aRarefactWin (Analytic Rarefaction version 1.3, S. Holland;

http://www.uga.edu/strata/software/Software.html).

Accession numbers for 848 representative gene sequences (full and partial sequences) can be found in GenBank (http://www.ncbi.nlm.nih.gov/) from EU487786 to EU488633.

15 RESULTS

Aqueous Geochemical Analysis

Physicochemical conditions evaluated in the field for the ocean water and sediment pore fluids included temperature, pH, oxidation reduction potential (ORP), total dissolved solids

(TDS), dissolved oxygen concentration, and dissolved sulfide concentration (Table 1). The sediment pore fluids were more reducing, with lower DO and measurable sulfide. Cation and anion concentrations also were measured for the pore water and the ocean water (Table 2). All species were more concentrated in the pore water, except Na+ that was more concentrated in sea water. The pore fluids had higher sulfate concentrations than seawater.

Table 1: Field geochemical data from Cedar Key, Florida. Geochemical Parameters Ocean Water Core Fluids (surface) Temperature (°C) 29.6 30.1 pH 8.07 7.65 Oxidation Reduction Potential (mV) +67 -227 Total Dissolved Solids (ppt) 44.97 45.01 Dissolved Oxygen (mg/L) 7.60 2.03 Dissolved Sulfide (mg/L) 0 1.57

Core Mineral Analysis

Cores were collected from the lucinid site, and were separated from each other by 1m.

The core sediments consisted predominantly of sand-sized grains. Bulk XRD analysis of the sediments from both cores indicated the dominance of quartz, with rarer undifferentiated clays, pyrite, , and silicate minerals (Table 3). The two cores had slight mineralogical variations with depth. For example, core 2 had more pyrite at the 0-10 cm interval compared to Core 1, which had more calcite.

16

Table 2: Cation concentrations (ppm) and anion concentrations (ppm) of ocean and pore water at Cedar Key, Florida. Chloride concentrations were within seawater concentrations, but were removed from the waters prior to anion analysis. + + 2+ 2+ 2+ 2- - Sample Description Na K Ca Mg Sr Fe Mn Al Si SO4 NO3 CDK-06-001 Ocean water 9086.9 303.7 279.3 669.3 4.2 0.005 0.006 0.005 0.6 3493.4 11.0 CDK-06-002 Pore Fluids 8763.7 336.4 544.6 1215.2 9.4 0.5 0.14 0.2 6.1 3635.2 9.2

Table 3: Percentage abundances of minerals in core sediments, Cedar Key, Florida Site Depth (cm) Quartz Clay Calcite Pyrite Plagioclase Dolomite K-feldspar Anhydrite Core #1 0-10.5 95.4 2.33 0.61 0.55 0.18 0.6 0.6 0.11 10.5-21 96 1.84 0.71 0.62 0.13 0.086 0.56 0.08 21-31.5 95.4 1.2 1.03 0.91 0.12 0.52 0.64 0.12

Core #2 0-10.5 95.5 1.44 0.67 0.8 0.12 0.98 0.4 0.07

10.5-21 94.5 2.27 0.09 1.6 0.19 0.13 0.97 0.19

21-31.5 95.1 2.34 0.09 1.44 0.24 0.08 0.59 0.1

17 Gill Endosymbiont Diversity

The 16S rRNA gene sequences of bacterial populations from lucinid gills were evaluated at three sampling levels to test my research hypotheses. First, a single individual, as well as multiple lucinid individuals of the same species, were evaluated from one site (Cedar Key) to assess intra- species diversity. Second, lucinids from different species at the same location were evaluated to assess inter-species diversity. Third, various lucinid taxa from multiple geographic areas were compared to assess if endosymbiont bacterial species varied by geography (Cedar Key, Lemon Bay, and Jack Island, Florida, and San Salvador, The Bahamas) (Figure 1). To investigate if the bacterial endosymbionts were also free-living (e.g., Gros et al., 1996; Distel, 1998), the 16S rRNA gene sequence diversity of sediments from the lucinid habitat were examined from Cedar Key, Florida.

Sediment mineralogy and fluid geochemistry were also compared from each sediment sample to understand possible ecological relationships to bacterial community structure and metabolic activities. No amplified bacterial 16S rRNA gene sequences were retrieved from lucinid foot samples.

DOTUR analysis of all full-length 16S rRNA endosymbiont sequences from all hosts (214 total ) grouped in 169 phylotypes at 99% relatedness. At 97% relatedness, all sequences grouped into 12 OTUs. According to Stackebrandt and Goebel (1994), 95% sequence identity could roughly correspond to genus-level relationships; 97% sequence identity could correspond to species-level relationships. Assigning OTUs based on 99% sequence identity was done to be conservative of species-level relationships. The DOTUR results imply that there is greater diversity among the lucinid endosymbionts than previously known (Distel et al., 1998), and that there are possibly more than one species of bacteria capable of being a lucinid endosymbiont.

18 Rarefaction curves estimated if the bacterial diversity retrieved in the clone libraries was representative of the full diversity for the samples. Results from all of the gill sequences show that the coverage of the clone libraries was nearing saturation with respect to the number of phylotypes defined at 99% sequence identity after screening >200 clones from 9 individuals (Figure 2, blue curve). If sequence diversity was low, then a curve would be a nearly horizontal line, as additional clones would represent previously sampled phylotypes. Alternatively, the somewhat steep curves for the separate Cedar Key libraries (Figure 2, pink curve) and the gill clone libraries from the other sites from all sequences (including non-gammaproteoabcterial sequences; Figure 2, orange curve) indicate that phylotype diversity for the gill endosymbionts is much higher than previously considered. These curves indicate that diversity was not sampled completely. More clones may need to be screened in order to achieve full coverage of phylotype diversity.

A total of 65 gill endosymbiont 16S rRNA gene sequences, retrieved from three lucinid

L.nassula host individuals collected at Cedar Key. All of the bacteria belonged to the

Gammaproteobacteria (γ-Proteobacteria) (Figure 3). Symbiont sequences were distantly related to previously published symbiont sequences from other lucinid hosts (Distel et al., 1988; 1993, Durand and Gros, 1996; Arkawa et al., 2006), including Codakia orbicularis, L. nassula, Stewartia floridana, and ‘Parvilucina’ costata (95-99% sequence identity). DOTUR results indicated that all

65 endosymbiont sequences grouped into one phylotype at 95% similarity; at 99%, there were 19 phylotypes.

All of the endosymbiont sequences from Lemon Bay (LB) were closely related to endosymbiont sequences previously retrieved from C. orbicularis at 99% sequence similarity (93% sequence coverage) (Distel et al., 1988) (Figure 4). Mean sequence distances calculated for the LB sequences using MEGA showed that endosymbiont sequences were genetically identical (Table 4).

19 40

35

30

25

20

15

10 All gill endosymbiont sequences

(99%) phylotypes of Number 5 Cedar key gill endosymbiont sequences Other sites gill endosymbiont sequences 0 0 50 100 150 200 250

Number of clones

Figure 2: Rarefaction curves for endosymbiont 16S rRNA gene sequences based on OTU designation at 99%. The pink curve is for γ-proteobacterial sequences from Cedar Key. The other sites includes non-γ-proteobacterial sequences (orange curve), but the combined dataset (blue curve) is only for endosymbionts belonging to γ-Proteobacteria from all hosts.

Forty-one full length 16S rRNA sequences were retrieved from the host P. pectinatus from the Mouth of Pigeon Creek (MPC). Of the sequences, 39 grouped into a clade closely related (99% sequence identity, 93% coverage) to gammaproteobacterial endosymbiont sequences previously retrieved from P. pectinatus [AJ581863] (Williams et al., 2004) (Figure 4). One sequence (MPC2-

46) was closely related to the α-Proteobacteria, Methylobacterium sp. [AB220088] (99% sequence identity, 93% coverage) (Figure 5). The remaining sequence (MPC2-26) was closely related to the

Bacteroidetes, Muricea elongota [DQ917900] (Figure 5) and (84%). Mean distances were calculated for all of the γ−Proteobacteria sequences using MEGA the MPC endosymbionts were

98% related (Table 4).

Fourteen 16S rRNA gene endosymbiont sequences retrieved from the host bivalve species

P. pectinatus collected at Jack Island (JI), had the most distance of the lucinid individuals sampled.

BLAST indicated that half of the bacterial sequences belonged to the α-Proteobacteria class

20 57 CK_G9 12 [EU487832] CK_G9 15 [EU487835] CK_G9 8 [EU487829] CK_G9 1 [EU487822] CK_G1 1 [EU487786] CK_G5 1 [EU487797] CK_G9 3 [EU487824] CK_G9 4 [EU487825] CK_G1 6 [EU487790] CK_G9 16 [EU487836] CK_G9 5 [EU487826] Figure 3: Phylogenetic relationship based on CK_G9 21 [EU487840] 63 CK_G9 2 [EU487823] Jukes-Cantor neighbor-joining analysis of 16S CK_G9 24 [EU487843] CK_G9 7 [EU487828] CK_G9 31 [EU487849] rRNA gene sequences of endosymbionts sequences 80 CK_G9 19 [EU487838] CK_G9 28 [EU487847] retrieved from lucinid gills from Cedar Key and 96 CK_G5 5 [EU487801] CK_G5 8 [EU487804] 16S rRNA sequences from sediments, representing CK_G5 9 [EU487805] 64 CK_G9 6 [EU487827] the free-living habitat. Beggiatoa alba was used as CK_G1 10 [EU487794] CK_G5 6 [EU487802] an outgroup. Bootstrap percentages were obtained CK_G9 17 [EU487837] CK_G9 14 [EU487834] using 500 replicates and values >50% are CK_G1 7 [EU487791] 53 CK_G1 11 [EU487795] indicated. Sequences labeled “G” are gill CK_G9 23 [EU487842] CK_G9 29 [EU487848] 64 CK_G9 22 [EU487841] sequences and “C” are sequences retrieved from CK_G9 27 [EU487846] CK_G9 20 [EU487839] the cores. Modified published lucinid genera CK_G5 10 [EU487806] 86 CK_G5 11 [EU487807] names to follow Taylor and Glover (2000). CK_G1 4 [EU487788] CK_G9 25 [EU487844] CK_G5 3 [EU487799] CK_G5 12 [EU487808] CK_G7 1 [EU487809] CK_G9 9 [EU487830] CK_G1 9 [EU487793] CK_G5 7 [EU487803] CK_1C3_51 [EU488029] CK_G9 13 [EU487833] CK_G9 10 [EU487831] CK_1C1 40 [EU487897] CK_G9 26 [EU487845] CK_G8 8 [EU487812] CK_G1 8 [EU487792] CK_G8 18 [EU487821] CK_G9 32 [EU487850] CK_G8 13 [EU487817] CK_G8 1 [EU487810] CK_G8 10 [EU487814] CK_G8 15 [EU487819] CK_G8 9 [EU487813] 93 CK_G1 5 [EU 487789] CK_G8 17 [EU487820] 99 CK_G8 14 [EU487818] CK_G5 4 [EU487800] CK_G8 11 [EU487815] CK_G8 5 [EU487811] CK_G8 12 [EU487816] 81 CK_G1 3 [EU487787] CK_G5 2 [EU487798] CK_G1 12 [EU487796] 63 Stewartia floridana gill symbiont [AB238979] ‘Parvilucina’ costata gill symbiont [L25712] 82 aequizonata gill symbiont [AY536232] Codakia orbicularis gill symbiont [M99447] 58 82 Lucinisca nassula gill symbiont [X95229] Escarpia spicata endosymbiont [AF165909] 99 CK_1C4 7 [EU488099] CK_1C4 27 [EU488053] 93 Uncultured gamma proteobacterium from Antarctica Sva0864 [AJ240988] 99 Uncultured gamma proteobacterium from Antarctica Sva0857 [AJ240969] 60 99 Uncultured gamma proteobacterium from Antarctica Sva1046 [AJ240991] CK_1C2 67 [EU487971] CK_1C4 49 [EU488077] Lucinisca pectinata symbiont [X84980] 57 clone [AM086643] 65 CK_1C4 36 [EU488063] 99 North Sea Sediments Gammaproteobacteria clone [DQ351741] Uncultured gamma proteobacterium from Antarctica Sva0115 [AJ240974] Olavius algarvensis sulfate-reducing endosymbiont [AF328857] 97 99 Marine sediment clone 4aFS [AM039662] CK_1C4 12 [EU488037] 99 CK_2C6 17 [EU488492] 99 clone IE094 [AY605145] 99 Spirochaeta bajacaliforniensis clone DSM16054 [AF328857] 0.02 estimated distance Seafloor methane hydrate clone [AB177212] 73 97 Chloroflexi clone LAS-B16N [AF513960] 99 CK_2C3 41 [EU488179] mangrove soil clone [DQ811879] 99 Dehalococcoides clone BHI80 [AJ431247] 99 Chloroethene-contaminated sludge pond clone [AF388531] 99 CK_1C2 23 [EU487925] 99 Ridge flank crustal fluids clone [AY869674] Beggiatoa alba [L40994] 21

(Figure5). Five of the remaining sequences were identified as γ- Proteobacteria, and two sequences were affiliated with Spirochetes. Distances calculated for only the γ-Proteobacteria sequences using MEGA however, showed that the JI endosymbionts had 99%similarity within group (Table

4).

Endosymbiont 16S rRNA sequences from host bivalves collected on the Gulf of Mexico side of Florida (LB and CK) formed a separate clade from the endosymbiont sequences collected on the

Atlantic side of Florida (JI and MPC), with strong bootstrap support (Figure 4). Mean distances calculated between the sequence groups using MEGA indicated that LB endosymbiont sequences shared 97% sequence similarity with sequences from CK and only 94% sequence similarity with endosymbiont sequences from the MPC and JI (Table 5). Endosymbiont sequences from MPC were 98% similar to JI sequences and only 93% similar to CK sequences (Table 5).

Table 4: Average genetic distance within host species for γ-proteobacterial endosymbiont16S rRNA gene sequences. Host Specimen Collection Site Host Species Avg. distance within sequence group Lemon Bay, Gulf of Mexico Coast, Florida Lucinisca nassula 0

Mouth of Pigeon Creek, The Bahamas Phacoides pectinatus 0.02

Jack Island, Atlantic, Florida Phacoides pectinatus 0.01

Cedar Key, Gulf of Mexico coast, Florida Lucinisca nassula 0.01

Table 5: Average genetic distance between γ-proteobacterial endosymbiont 16S rRNA gene sequences from different host taxa Site Lemon Bay Pigeon Creek Jack Island Cedar Key Lemon Bay, Florida -- 94% 94% 97% Mouth of Pigeon Creek, Bahamas 0.06 -- 98% 93%

Jack Island, Florida 0.06 0.02 -- 94%

Cedar Key, Florida 0.03 0.07 0.06 --

22

Figure 4: Phylogenetic reconstruction based on Jukes-Cantor neighbor-joining analysis of gill 16S rRNA bacterial sequences belonging to previously identified γ-proteobacterial groups. Sequences retrieved from hosts were collected for Cedar Key (CK), Lemon Bay (LB), and Jack Island (JI), Florida, and the Mouth of Pigeon Creek (MPC), The Bahamas. Beggiatoa alba was used as an outgroup. Bootstrap percentages obtained using 500 replicates and values >50% are indicated. LB sequences not shown in the tree are represented by accession numbers EU488499-EU488592. Modified published lucinid genera names to follow Taylor and Glover (2000).

23 LB_GB72-36[EU488558], LB_GA82-18 [EU488516], LB_GA8-21 [EU488527] Representative gill sequences from 3 host bivalves, nassula. Lemon Bay, Gulf of Mexico coast of Florida 94 clone sequences

CK G1 10 [EU487794] Cedar Key, Florida; L. nassula Stewartia floridiana gill symbiont [AB238979] ‘Paryilucina’ costata gill symbiont [L25712] Lucinisca nassula gill symbiont [X95229] Lucinoma aequizonata gill symbiont [AY536232] Codakia orbicularis gill symbiont [M99447] CK G8 10 [EU487814] CK_G8_14 [EU487818] CK G8 17 [EU487820] CK G8 9 [EU487813] CK_G5_4 [EU487800] CK_G8 15 [EU487819] CK_G8 13 [EU487817] CK_G8_11 [EU487815] CK G8 5 [EU487811] CK_G8_12 [EU487816] CK_G8_ 17 [EU487820] CK_G1_4 [EU487788], CK_G9_12 [EU487832], CK_G5_1 [EU487797]

14 clone Representative sequences from 3 hosts; L. nassula CK G1 5 [EU487789] sequences CK_G8_8 [EU487812], CK_G9_16 [EU487836], CK_G5_5 [EU487801] Representative gill sequences from 3 hosts; L. nassula 38 clone sequences MPC2 41 [EU488600] Mouth of Pigeon Creek, Bahamas; P. pectinatus Phacoides pectinatus symbiont [X84980] MPC2 48 [EU488604] MPC2 22 [EU488615] MPC2 39 [EU488630] MPC2 14 [EU488608] MPC2 10 [EU488605] MPC2 7 [488597] MPC2 44 [EU488602] JI4 19 [EU487860] Jack Island, Florida; P. pectinatus MPC2 17 [EU488611] MPC2 34 [EU488625] MPC2-24 [EU488616], JI4-17 [EU487858] Representative gill sequences from 2 hosts; P. pectinatus 25 clone sequences Olavius algarvensis sulfate-reducing endosymbiont [AF328857] MPC2 32 [EU488623] The Bahamas; P. pectinatus Beggiatoa alba [L40994]

0.02 Estimated

24 Spongiobacter nickelotolerans [AB205011] Endozoicimonas elysicola (sea ) clone [MKT110] 100 Muricea elongata ongiobacter clone ME19 [DQ917863] 94 Australia Great Barrier clone [AY700607] 7 Cystodytes dellechiajei (sea squirt) clone [DQ860081] JI4 16 [EU487857] 73 62 100 JI4 12 [EU487853] 74 JI4 15 [EU487856] JI4 17 [EU487858] 50 100 Microbulbifer JAMB A94 [AB160954] Pelagiobacter variabilis Ni2088 [AB167354] 59 99 Pseudomonas DW101 [AJ534849] 66 Pseudomonas Z49zhy [AM411057] Balneatrix alpica [Y17112] γ-Proteobacteria 99 Polychlorinated dioxin dechlorinating microbe [AB187503] Heavy metal polluted marine sediment clone [AY665404] 100 68 CK_G5_ 6 [EU487802] CK_G9_4 [EU487825] 98 γ−Proteobacteria 100 CK_G1_3 [EU487787] CK_GA8_2_7 [EU488505] gill endosymbionts 100 CK_GB7_2_13 [EU488572] 94 MPC 2 33 [EU488624] Phacoides pectinatus gill symbiont [X84980] 75 terraeregina gill symbiont [U62131] phillipiana gill symbiont[L25711] 69 71 Endosymbiont Ridgeia piscesae[L25711] 100 Endosymbiont of [AY129116] Caulobacter MCS6 [M83811] 98 56 100 Whalefall clone [131718] Antarctic lake water clone ELB16 005 [DQ015795] Citromicrobium [JL1041] 65 JI 4 3 [EU487864] JI4 14 [EU487855] 73 82 JI4 13 [EU487854] 78 JI4 4 [EU487851] JI4_1 [EU487863] 100 JI4_2 [EU487862] Travertine hot spring clone SM1D09[AF445675] 98 Caedibacter caryophilus [AY753195] α-Proteobacteria 100 Caedibacter caryophilus [AJ238683 ] 67 Candidatus Odyssella thessalonicensis [AF069496] Devosia clone DS8 [EF584042] Rhodopseudomonas palustris BisB18[CP000301] 100 Rhodopseudomonas palustris BisB18 [CP000301] Rhodopseudomonas palustris BisB18 [NC_007925] 90 Methylobacterium variabile MPC2_46 [EU488632] 100 67 Methylobacterium PB142[AB220088] Methylobacterium PB140 [AB220086] 100 Methylobacterium PB145 [AB220090] 84 MPC2_26 [EU488633] 100 Muricea elongata Flavobacterium clone BM [DQ917900] Antarctic lake ice cover clone ANTLV2 F0 [DQ521522] Bacteroidetes 56 Gyrosigma fasciola [AF514847] Haslea salstonica [AF514854] 73 Haslea wawrikae [AF514855] Haslea nipkowii [AF514850] Spirochaeta africana [AF165909] 100 JI4 18 [EU487859] Spirochetes 100 JI4 20 [EU487861] 99 Olavius loisae endosymbiont [|AF104472] Cytophaga BHI80 86 [AJ431239] 0.02 86 Cytophaga BHI80 3 [AJ431238] 87 Marine hydrocarbon seep sediment clone [DQ004680] Thermus aquaticus [X58340]

Figure 5: Phylogenetic relationship based on Jukes-Cantor neighbor-joining analysis of 16S rRNA gene sequences of endosymbionts sequences retrieved from lucinid gills from Jack Island, Florida, and the Mouth of Pigeon Creek, San Salvador, Bahamas. Beggiatoa alba was used as an outgroup. Bootstrap percentages were obtained using 500 replicates and values >50% are indicated. Modified published lucinid genera names to follow Taylor and Glover (2000).

25 Sediment Bacterial Diversity from Cedar Key, Florida

A total of 634 clones, comprising 13 major taxonomic groups, were retrieved from all of the core sediment samples at Cedar Key (Table 6). Sixty-one percent of the sequences were affiliated with the Proteobacteria division (Table 6). α−Proteobacteria were the dominant class (43%) followed by δ−Proteobacteria (11%), γ−Proteobacteria (3%), β−Proteobacteria (2%),

ε−Proteobacteria (2%). Other major phyla represented were the Chloroflexi (17%),

Acidobacterium (4.2%) Firmicutes (3.7%), (3.7%), Candidate divisions (2.6% ),

Actinobacteria (2.3%), Bacteroidetes (2.0%), Spirochete ( 1%), Deferribacteris (1.1%),

Nitrospirae (0.31% ), and Verromicrobium and Thermotogales (0.15%). Figure 6 shows the rarefaction curves for both of the core sediment sequence datasets; Core 1 had four clone libraries and Core 2 had five. The combined datasets represent 648 full length 16S rRNA gene sequences.

Phylotypes (OTUs) were defined at 99% sequence identity. The steep curves for Core 1 show that the diversity of bacteria is higher than retrieved in Core 2 (having a less steep curve). The curves also show that diversity was not fully sampled, and future work should increase the number of sequences per sample in order to reach full coverage.

Taxonomic Descriptions

Alphaproteobacteria

Most of the 16S rRNA from the sediment sequences were affiliated with the α-

Proteobacteria (Table 6). The largest clade consisted of 209 sequences and was most closely related to Rhizobium spp. [DQ873663], free-living gram-negative soil bacteria that are usually associated with infection of legume roots and nitrogen fixation (Long, 1996). Free-living rhizobia subsist on dead organic matter and do not fix nitrogen (Burdass, 2002). Other α-proteobacterial

26 250 Core 1 95% sequence similarity Core 1 97% sequence similarity Core 2 95% sequence similarity 200 Core 2 97% sequence similarity

150

100 OTUs observed

50

0 0 50 100 150 200 250 300 350 400 Clones sequences

Figure 6: Rarefaction curves for the two core sequence sets from Cedar Key. Core saturations are shown at 97% (dark pink, dark green) as well as saturation at 95% (light pink and light green). sequences were closely related to sequences represented by the families Hyphomonadaceae and

Rhodobacteraceae. These groups are typically found in marine environments and exhibit a wide variety of morphological, physiological, and biological features (Lee et al., 2005). Three sequences were related to Hyphomonas polymorpha [DSM2665]. Cultured hyphomicrobia are facultative methylotrophs.

Eight sequences formed a clade with Rhodopseudomonas rhenobacensis [AB087719], a nitrate-reducing, non-sulfur bacterium isolated from a eutrophic pond in Germany (Hougardy et al.,

2000). These bacteria are rod-shaped cells that are capable of growth both photosynthetically under anaerobic conditions and non-photosynthetically under aerobic conditions using a variety of organic compounds as metabolic substrates (e.g., formate, acetate, pyruvate) (Hougardy et al., 2000).

Deltaproteobacteria

Eleven percent of the sediment sequences were affiliated with the δ−Proteobacteria (Table

6). All of these sequences clustered most closely with anaerobic sulfate-reducing, iron-reducing, or

27 syntrophic bacterial species. Eighteen sequences were related to Syntrophus aciditrophicus

[NC_00759], a rod-shaped, gram-negative bacterium that anaerobically recycles organic substrates to carbon and methane (McInerney et al., 2007). Thirteen sequences were most closely related to an uncultured δ−Proteobacteria isolated from an intertidal mud flat in the Wadden Sea, Germany

[AY771946] (Mussman et al., 2005). Five sequences formed a clade with an uncultured

δ−Proteobacteria from Gulf of Mexico seafloor sediments from a hydrate system in the northern

Gulf of Mexico continental slope [AY542227] (Mills et al., 2005).

Gammaproteobacteria

Approximately 3% of the sequences were affiliated with the γ−Proteobacteria. Sequences grouped into two distinct clades related to Thialkalivibrio thiocyanodenitrificans [AY360060], an obligate chemolithoautotrophic sulfur-oxidizing bacteria (Sorokin et al., 2004), and to Thiothrix spp. [EF641919], a filamentous gram-negative bacterium that oxidizes reduced sulfur compounds and is commonly found in sulfide-containing natural waters (Meyer et al., 2007). Nine of the sequences were closely related to lucinid or other metazoan symbionts within the γ-Proteobacteria.

Only two of the sequences were identical to the CK 16S rRNA gene sequences retrieved in this study (Figure 3), representing 0.3% of the sediment sequences. The other sequences were related to previously retrieved sequences from Arctic sediments (Ravenschlag et al., 2000).

Epsilonproteobacteria

Two percent of the sequences were affiliated with the ε−Protebacteria. Two distinct clades were represented. Four of the sequences grouped with Sulfurimonas denitrificans [NC_007575], a sulfur-oxidizing chemolithoautotrophic bacterium that oxidizes sulfide and thiosulfate and commonly associated with hydrothermal vent communities (Madigan et al., 2002). The remaining two sequences grouped with Sulfurovum lithotrophicum [AB091292], an obligately aerobic, sulfur-

28 and thiosulfate- oxidizing bacterium commonly associated with deep sea cold seeps and hydrothermal vents (Inagakai et al., 2004; Suzuki et al, 2005).

Betaproteobacteria One clone sequence was most closely related to Leptothrix sp. str S11 [DQ241397], a filamentous bacterium typically found in iron-rich waters (Madigan et al., 2005).

Acidobacteria

The Acidobacteria comprise a poorly understood division of bacteria (Sabree et al., 2006).

Approximately 4.2% of the core sediment sequences were related to the Acidobacteria.

Acidobacteria are abundant in soil and alkaline conditions, and some are thought to be photosynthetic in the presence of oxygen (Sabree et al., 2006).

Actinobacteria

About 2.3% of the sequences were related to Actinobacteria. These bacteria are common in soil, and are almost all anaerobic (Ventura et al., 2007). Actinobacteria are gram-positive and are morphologically, physiologically, and metabolically diverse (Ventura et al., 2007). They are common decomposing bacteria that play an integral part in the carbon cycle (Conn, 2005). These bacteria produce many different enzymes and secondary metabolites, such as antibiotics, which are used for drugs (Conn, 2005).

Bacteroidetes

Bacteroidetes are rod-shaped, gram-negative bacteria found in a variety of habitats, including marine sediment and sea water. These bacteria are predominantly anaerobes (Madigan and Martinko, 2005). Bacteroidetes comprise 2.0% of the core sediment sequences. About half of the sequences grouped with bacterial sequences belonging to Cytophaga spp. [AB015532,

AJ431254], deep sea sediment clones. Cytophaga are gliding bacterial group found in soils rich in

29 organic matter at neutral pH, such as aerobic bottom sediments and algal mats. The remaining sequences grouped with environmental bacterial sequences associated with mangrove soils collected from the Natural Nature Reserve, Hainan Island, China [DQ811911, DQ811905] (Yan et al., 2006).

Chloroflexi

Seventeen percent of the sequences retrieved from the core sediments belonged to the

Chloroflexi phylum (Table 5). Chloroflexi, or green non-sulfur bacteria, are filamentous prokaryotes typically found in marine microbial mats (Madigan et al., 2005). They are anoxygenic phototrophs capable of photoheterotrophy and chemoorganotrophy (Madigan et al., 2002). Most sequences were closely related to environmental sediment clones, including the mangrove soil clone

MSB-5G10, isolated from mangrove soil in China [DQ811879].

Planctomycetes

Sequences represented by the Planctomycetes accounted for3.7% of the bacterial sequences retrieved from the core sediments. Planctomycetes are budding bacteria with a protein stalk lacking peptidoglycan (Fuerst, 2001). These chemoorganotrophic bacteria are found in diverse environments, including marine and soil habitats (Fuerst, 2001).

Firmicutes

Firmicutes are also gram-positive bacteria that have rod- or cocci-shaped cells. Nearly 4% of the sediment sequences were related to the Firmicutes. Sequences grouped into three clades, represented by two groups of Firmicutes, the clostridia and the bacilli. Clostridium litorale DSM

5388 [X77845] is a strictly anaerobic fermentative bacteria that forms endospores. It is commonly associated with extreme conditions, such as the human gut or sulfate-reducing marine

(Zhang and Fang, 2001; Ley et al., 2006). The second clade was represented by the Ruminococcus

30 sp. str. 14531 [AJ315979], a bacterium within the bacilli. Bacilli are obligate or facultative aerobes commonly found free-living in soil, as well as natural flora in the human gut (Ley et al., 2006). The third clade was associated with Alkakiphilus crotonatoxidans [AF467248], a strictly anaerobic, chemoorganotroph isolated from a methanogenic environment (Coa et al., 2003).

Spirochetes

Spirochetes are slender, mobile, flexible and coiled, gram-negative bacteria found in aquatic environments (Madigan et al., 2005). Less than 2% of the clone sequences from the sediment were associated with the spirochetes. These bacteria have been previously found in marine, hydrogen sulfide-containing anaerobic habitats (Madigan and Martinko, 2005). They can also be found free living as anaerobic or facultative aerobic bacteria, and pathogens. Most of the sequences grouped in a clade represented by the environmental spirochete [AY605139] retrieved from a microbial mat in the Ebro and Camargue delta, Spain (Gurrero et al., unpublished). The remaining sequences grouped in a clade with Spirochaeta halophila [M88722], a facultative anaerobe found in Hunter

Hot Spring, Oregon (Paster et al., 1991).

Nitrospirae, Defferribacteres, , and Thermotogales

Both Nitrospirae and Deferribacteres are newly identified phyla within the bacterial that little is known about these groups (Madigan et al., 2005). Nitrospirae and Defferibacteres each accounted for <1% of the clones. Nitrospirae are nitrogen-oxidizing bacteria capable of chemolithoautotrophy or chemoorganotrophy. These bacteria are commonly found in a variety of habitats, (marine and fresh water, deltas, and aquariums) and temperatures (Daims et al., 2001;

Madigan and Martinko, 2005). Deferribacteres are obligate anaerobes that utilize various electron acceptors, such as sulfur, nitrate, and fumorate (Janssen, 2002). Rare clones from the sediment were affiliated with the Verrucomicrobia (0.15%) and Thermotogales (0.15%). Verrucomicrobia

31 are warty-shaped bacteria that are aerobic to facultative aerobic, and are found in marine environments (Madigan et al., 2005). Thermotogales are strict anaerobes, commonly characterized as sulfate-reducing bacteria (Madigan et al., 2005).

Candidate Divisions

Seventeen core sediment sequences were identified as belonging to the proposed bacterial lineages within the Candidate Divisions OP8, OP11, and OD1. The sequences grouped with sequences from the Ocean Drilling Program (ODP), collected from methane hydrate bearing sub seafloor sediments from Peru and the Cascadia Margins (Inagaki et al., 2006).

Microbial Diversity with Core Depth

Core sediment samples were compared at ~10.5 cm intervals to determine changes in diversity with depth (Figure 7). Core 1 was dominated by δ-Proteobacteria and Chloroflexi in the first ~30 cm. Between 10-20 cm, ε-Proteobacteria and Planctomycetes were more prevalent.

Between 30-40 cm, Actinobacteria and Planctomycetes were dominant, while the relative abundance of Chloroflexi decreased. At 40-50 cm depth, the relative abundance of δ-

Proteobacteria decreased with an increase in Chloroflexi, γ-Proteobacteria, and α-Proteobacteria.

α−Proteobacteria were the dominant bacteria at all depths from Core 2 (Figure 7), especially at ~40 cm where this group accounted for ~75% of total diversity. Chloroflexi and δ-Proteobacteria were abundant in the top ~20 cm. At 20-40 cm, the abundance of δ-Proteobacteria decreased but at 40-

50 cm it increased again. ε-Proteobacteria were not present at any depth in Core 2. Firmicutes were not retrieved at the shallow depth in Core 2, but increased in abundance between 30-40 cm.

32 Table 6: Distribution of 16S rRNA gene bacterial sequences from sediment cores at Cedar Key, Florida Total # PHYLUM CK 1C-1 CK 1C-2 CK 1C-3 CK 1C-4 CK 1C-5 CK 2C-6 CK 2C-5 CK 2C-4 CK 2C-3 CK 2C-2 Clones Core Depth (cm) 0-10.5 0-10.5 10.5-21 10.5-21 21-31.5 0-10.5 0-10.5 10.5-21 10.5-21 21-31.5

Proteobacteria 17 34 23 31 4 15 30 105 122 21 402

Acidobacteria 4 2 3 2 0 0 2 7 2 5 27

Actinobacteria 1 2 2 4 1 0 1 1 2 1 15

Bacteroidetes 1 0 3 5 0 0 1 1 1 1 13

Chloroflexi 19 26 10 14 2 5 8 11 10 3 108

Planctomycetes 1 6 2 4 3 0 1 4 2 1 24

Firmicutes 4 2 1 4 0 1 1 8 3 0 24

Spirochaetes 0 2 0 2 0 1 0 1 0 1 7

Deferribacteres 2 1 1 1 0 0 0 2 0 0 7

Nitrospirae 0 0 1 0 0 0 0 0 1 0 2

Verrucomicrobia 0 0 1 0 0 0 0 0 0 0 1

Thermotogae 0 0 1 0 0 0 0 0 0 0 1

Candidate Divisions 2 3 3 2 1 1 1 2 2 0 17

Total # sequences 51 78 51 69 11 23 45 142 145 33 648

Total # full length sequences 51 78 51 69 11 23 45 142 145 33

33 The microbial communities between the two cores had a 0.93 Sorenson Index of Similarity

(calculated by EstimateS), meaning that all of the major taxa were represented in both corees, even though the relative abundances of each major taxonomic group were different. Visual inspection of core distribution versus chemistry and mineralogy of the cores shows no statistical difference.

Further statistical testing is needed to determine any differences.

34 CK 1C-1 CK 2C-6 th p α-Proteobacteria β-Proteobacteria γ-Proteobacteria

0-10 cm de δ-Proteobacteria CK 1C-2 CK 2C-5 ε-Proteobacteria Acidobacteria Actinobacteria Bacteroidetes Chloroflexi Planctomycetes Firmicutes Deferribacteres CK 1C-3 CK 2C-4 Nitrospirae Verrucomicrobia Candidate Division th p

CK 1C-4 CK 2C-3 10-20 cm de th p CK 2C-2 Figure 7: Bacterial CK 1C-5 diversity of each of the core sediment 16S rRNA gene sequence

20-30 cm de clone libraries, shown grouped with increasing depth (every 10 cm).

35 DISCUSSION

Despite the numerous investigations describing the symbiotic association between lucinid hosts and their thiotrophic bacterial endosymbionts (e.g., Wiley and Felbeck, 1994; Gros et al.,

1996; Krueger et al., 1996; Distel, 1998; Imhoff et al., 2003; Duperron et al., 2005; Stewart et al.,

2005; Suzuki et al., 2005; Duperron et al., 2006; Taylor and Glover, 2006; Caro et al., 2007;

Duperron et al., 2007), many questions still remain regarding bacterial endosymbiont phylogenetic diversity, host-bacterial species specificity, and the diversity and availability of endosymbionts in host habitats. This study aimed at addressing these issues by examining the diversity among bacterial endosymbionts within hosts of the same species from the same habitat, as well as among different host species from the same habitat, and from different geographically separated host species. To my knowledge, this is the first study to examine the bacterial diversity of a lucinid habitat, which includes the identification of free-living bacteria belonging to the

Gammaproteobacteria and possibly related to lucinid endosymbiont sequences. The main study site was the sea grass and silicate sediments habitat at Cedar Key, Florida. For Cedar Key, habitat parameters such as geochemistry and mineralogy were assessed to evaluate possible relationships among the Cedar Key habitat, host taxa, and their endosymbiont species. Results indicate that the bacterial endosymbionts from lucinid bivalves are not all genetically identical across different host species in the same or different habitats, within the same host species from the same or different habitats, or even within an individual host .

Habitat Geochemistry and Mineralogy

Shallow marine lucinids require a unique geochemical habitat, not unlike living at deep-sea vent systems. Previous work from shallow marine environments did not assess the environmental habitat parameters for lucinids and their symbionts, and determine if there was a

36 connection between the environmental conditions and the host species or endosymbiont species.

Lucinid hosts must position themselves in the sediment to balance dissolved oxygen and dissolved sulfide concentrations (Felbeck et al., 1983; Savazzi, 2001). Because sulfide oxidation can occur spontaneously in the presence of high concentrations of dissolved oxygen, host bivalves live at or near the oxic-anoxic interface (Distel, 1998; Stewart et al., 2005). The dissolved oxygen concentration within the Cedar Key sediment pore waters was 2.03 mg/L (Table 1), which is high enough that some dissolved sulfide could spontaneously oxidize (Frenkiel et al., 1995; Kraus,

1995). However, because the sediment had high concentrations of dissolved sulfide, conditions within the sediment were still conducive for thiotrophy. Slight variations in pyrite abundance with depth (Table 3), but with slightly more pyrite from the deeper sediment, may indicate changing reducing conditions with depth, but may also correlate to the zones where the flux of dissolved sulfide through the sediments is greatest. With little to no iron in the sediment, formation of pyrite would decrease and would increase the amount of dissolved sulfide. Based on the depths that lucinid hosts were collected (within the first 10 cm of the sediment surface), the hosts may position themselves above the depths where the potential for pyrite mineralization is greatest.

The type of bacteria present in any environment, and available for acquisition for symbiosis, is likely controlled by the distinct geochemistry and mineralogy of the sediment habitat. The mineralogy of the Cedar Key sediments showed that the dominant mineral was quartz, as would be expected in a siliciclastic environment (Table 3). Lemon Bay is also a siliciclastic setting, and this could be a reason why the sequence similarities between the CK and LB endosymbiont sequences was higher than those from JI and MPC, whose habitats are dominated by (Table 5). A habitat study in a dominated environment would be necessary to determine whether the free-living symbionts grouped with the endosymbionts from host bivalves found there. Until that

37 work is done, this study of the microbial diversity from a lucinid sea grass siliciclastic sediment habitat (Cedar Key) is unique A comparative habitat study would also be important to test whether or not the acquisition of endosymbionts depends on the availability of bacteria within the habitat, or if acquisition is related to specificity for the host species. If the free-living symbionts do not exist in carbonate sediments, then this could indicate that the endosymbionts must be acquired from another source, such as free-living cells from seawater.

Intra- and Inter-Species Endosymbiont Diversity

Previous research suggested that all lucinid bivalve taxa contained the same species of bacterial endosymbionts (e.g., Gros et al., 2003). Distel et al. (1988; 1994) and Gros et al. (2003) suggest that endosymbionts are species-specific to their hosts. I hypothesized that bacterial endosymbionts from hosts collected at geographically separated locations would be genetically distinct. One way I tested this hypothesis was by evaluating the bacterial diversity of different host taxa from the same habitat. However, because DNA quality and quantity was low from hosts other than L. nassula, I was unable to determine whether the endosymbionts found in multiple species from the same habitat were the same or different. With the 16S rRNA gene sequences I retrieved from Cedar Key from three L. nassula hosts, DOTUR analysis demonstrated that there was only one phylotype at 95% sequence identity. This indicates that the L. nassula hosts from Cedar Key contain endosymbionts from the same genus. Comparing all of the gammaproteobacterial sequences at 99% sequence identity, however, revealed that 19 phylotypes were identified, suggesting that endosymbiont identity varies among host organisms, even within the same habitat (Figure 6).

Similarly, the endosymbionts do not always belong to the same bacterial phylotype within the same host organism (e.g., clones beginning with the label “CK_G8”) (Figure 3) or within multiple hosts of the same species (e.g., clones beginning with the label “CK_G8” versus

38 “CK_G5”) (Figure 3, Table 4). These results are consistent with the recent study of Caro et al.

(2007) that found that up to seven genetically distinct subpopulations of endosymbionts, of varying size and intracellular nucleic acid content, were present within individual host specimens.

When geography was examined as an influencing factor that may control bacterial endosymbiont diversity, sequences from host bivalves taken at geographically similar locations grouped more closely than those sequences from bivalves collected at distant geographic locales

(Table 5). Endosymbiont sequences from host bivalves collected on the Gulf of Mexico side of

Florida (CK and LB) grouped into a separate clade compared to the endosymbiont sequences collected on the Atlantic side of Florida (JI and MPC), with high bootstrap support (Figure 5).

Coincidentally, these clades also correlated to the same host taxa (e.g., L. nassula from CK and LB and P. pectinatus for MPC and JI). Therefore, it is difficult to determine whether endosymbiont phylotype diversity is controlled more by geography, habitat type (e.g., siliciclastic versus carbonate), or if diversity is associated with host taxa specificity. Further research should be done to determine whether or not it is host species or geography that might have a greater influence on endosymbiont species diversity. It is possible that various metabolic requirements for both the endosymbiont and host, and the variety of habitats that these organisms live in, have caused genetic variations and adaptations among endosymbionts (Caro et al., 2007). Regardless of these limitations, it is clear that bacterial endosymbionts associated with lucinids are not all the same species within a single host, across hosts of the same species, or across different species of hosts.

Free-living Bacterial Diversity

One of my research goals was to characterize the bacterial diversity within the lucinid free- living (sediment) habitat. I hypothesized that the free-living bacterial diversity would reveal sequences related to the endosymbionts. Thirteen major taxonomic phyla were retrieved from the

39 lucinid sea grass bed sediments at Cedar Key. Although a thorough study of the bacterial diversity from seagrass sediments has been done, the overall sequence diversity among bacteria was comparable to the types of microbial groups previously found in marine sediments (e.g.,

Ravenschlag et al., 2001). Core 1 and 2 were separated by 1m, and the communities in the two cores were very similar (0.93 for the Sorenson Index of Similarity), but the distributions of the group varied. Core 1 was dominated nearly equally by δ-Proteobacteria and Chloroflexi while

Core 2 was dominated by α-Proteobacteria (Figure 7).

The most significant result from studying the sediment was that 0.3% of the sequences retrieved from all 856 clone sediment sequences were related to sequences from the Cedar Key gill gammproteobacterial endosymbionts (Figure 3). One sequence [CK_1C1_40] was retrieved from

0-10.5 sediment depth, while the other [CK_1C3_51] was retrieved from 10.5-21 cm depth. Lucinid hosts were collected from within the top 10 cm of the sediment surface. Gros et al. (2003a) suggest that this group is free-living in sediments hosting lucinids. Others have suggested that there are free- living bacteria are environmentally acquired (Gros et al., 1996; Peek et al., 1998; Fortey, 2000;

Savazzi, 2001; Gros et al., 2003b; Caro et al., 2007). Therefore, the low representation of these bacteria in sediment may be due to a high recruitment rate by host bivalves, which would keep the overall abundance of free-living cells low in the lucinid habitat. But, finding sequences closely related to the gammaproteobacterial endosymbionts in the sediments does not necessarily mean that the endosymbionts are metabolically active while free-living, as their distribution in marine sediments and their metabolic role in sediment have not been established.

The similarity between some free-living symbiont sequences to endosymbiont sequences at

CK, as well as the fact that endosymbiont sequences were found most closely related to sequences from hosts collected at the same locale, could suggest that endosymbiont diversity is not regulated

40 by host specificity, but rather by environmental availability. This could also be supported by the presence of significantly different endosymboints (e.g., α-Proteobacteria) in host species found in distinctly different habitats (e.g., carbonate or silisiclastic).

Future research could include the characterization of another habitat similar to Cedar Key, but also a distinctly different habitat (e.g., carbonate) to determine the presence of free-living endosymbionts. Future research should also include an investigation of the changes in free-living bacterial diversity through time, as well as bivalve abundance and determine the changes in bacterial diversity with changes in bivalve presence, or even seasonal changes. Disturbance, such as hurricanes, can also redistribute bacteria within sediment, as well as lucinids, and may impact diversity through time.

Endosymbiont Sequence Diversity and Possible Metabolic Implications

Retrieving clone sequences from lucinid gills that were not related to Gammaproteobacteria was an unexpected result. Although recent research by Caro et al. (2007) suggest that individual lucinid gills can contain up to seven different bacterial subpopulations (from C. orbicularis), and

Duperron et al. (2007) found non-gammaproteobacterial sequences in the gill of a deep sea clam, no studies have retrieved 16S rRNA gene sequences from shallow marine lucinids (i.e. not deep sea) that were not Gammaproteobacteria. In addition to gammaproteobacterial sequences, a bacterial sequence was retrieved from a P. pectinatus host collected from the Mouth of Pigeon Creek, The

Bahamas [MPC-2_46] that grouped most closely with an Alphaproteobacteria, a Methylobacterium spp. clone (Figure 5). Also from P. pectinatus from Jack Island, FL, 16S rRNA sequences were related to both γ- and α-Proteobacteria (Figure 5). These results are unique to this thesis, as no study has uncovered non-gammaproteobacterial diversity for the shallow marine lucinids. Hosting both γ- and α-Proteobacteria could be due to the environment in which the host lived, or could be a

41 trait specific to P. pectinatus. These results may indicate dual symbiosis (thiotrophy and methanotrophy) in some lucinid bivalves. Dual symbiosis has previously been found in the

Mytilidae from deep sea vent systems (Distel et al., 1995). Further work is underway to address possible dual symbiosis for the MPC and JI individuals, as well as to sequences more clones from lucinids hosts to achieve more coverage in clone diversity. The presence of dual symbiosis may suggest either an evolutionary shift toward utilization of multiple metabolic pathways, or possibly an ancient metabolic pathway that is being abandoned in other lucinids through genetic alteration in order to have sulfide oxidation be the dominate endosymbiotic pathway.

42 CONCLUSIONS

Characterization of a single lucinid habitat, including aqueous and sediment geochemistry, as well as the gill endosymbiont and free-living bacterial diversity, has allowed us to better understand the functional and evolutionary relationship between bacterial endosymbiont and their hosts. Previous research indicated that all endosymbiont sequences belonged to a single phylogenetic clade within the γ−Proteobacteria (Distel, 1998; Gros et al., 2003). With recent evidence suggesting that diversity may be greater than expected (Caro et al., 2007; Duperron et al.,

2007), I determined from 16S rRNA gene surveys that lucinid bivalve hosts do not always contain the same species of bacterial endosymbiont, and that endosymbiont sequences are not identical within the same species, across multiple hosts within the same environment, across host species within the same geographic locale, or across geographic locales.

The results from this study indicate that endosymbionts in lucinid bivalves form several distinct phylogenetic clades (Figure 4 and 5). This bacterial diversity may indicate that endosymbionts are not host specific. The presence or absence of free-living symbionts in a lucinid habitat, and especially those symbionts that could potentially become thiotrophic gill endosymbionts, may play an important role for host specificity for symbiont acquisition. However, based on the geochemical variation of the habitat, metabolic requirements (i.e., habitat conditions) of the free-living symbionts will also influence potential symbiont diversity. It is also possible that environmental stresses to the host lucinids could provoke the utilization of multiple metabolic bacterial types within the gills. Therefore, the effects of habitat geochemistry could explain the possible existence of dual symbiosis in one lucinid host, but more research needs to be done to determine conclusively if dual symbiosis in some species of lucinids is actively occurring.

43 It has been suggested that isotope studies of bivalve shells may be related to changes in environmental parameters, dietary preference, growth rate, and sexual maturity (Jones et al., 1988;

CoBabe and Pratt, 1995; Langlet et al., 2002; Leitard and Pierre, 2006). Bacterial metabolism may also be recorded in the stable isotope composition of hosts. However, given that there is significant bacterial endosymbiont diversity, isotopic ratios obtained in ancient bivalve shells may not reflect the true nature of a lucinid’s metabolic history (e.g., Peng et al., 2007).

Lastly, results from this study will lead to a more thorough understanding of the evolutionary and ecological relationships between lucinid bivalves and their thiotrophic endosymbionts. This research is the first to characterize the bacterial diversity of a lucinid host habitat, and specifically a nearly exhaustive study of a single site. My research will also provide a background for modern day symbiotic relationships that can be applied to ancient symbiotic relationships.

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49 Ley, R.E., Turnbaugh, P.J., Klein, S., Gordon, J.I., 2006, Microbial ecology: Human gut microbes associated with obesity: Nature, v. 444, p. 1022-1023.

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52 APPENDIX A CORE MINERAL ANALYSIS AND XRD

1. XRD Mineral Abundance Reports for All Core Samples

2. Core 1 Mineral Peak Comparison

3. All Core Mineral Peak Comparison

53

LSU Geology & Geophysics Dept. 9/24/2007 - 10:23:27.99 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-1b.raw

QUARTZ 96.4972 CALCITE 0.6103 DOLOMITE 0.1466 ANHYDRITE 0.0901 K-FELDSPAR 0.4084 PLAGIOCLASE 0.0983 PYRITE 0.2405 CLAY 1.9087

LSU Geology & Geophysics Dept. 12/04/2007 - 10:25:28.50 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-1b.raw

QUARTZ 94.4842 CALCITE 0.5975 DOLOMITE 0.1436 ANHYDRITE 0.0882 PYRITE 0.2355 CLAY 1.8688 ARAGONITE 2.5821

LSU Geology & Geophysics Dept. 9/24/2007 - 10:24:36.81 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-2b.raw

QUARTZ 94.1685 CALCITE 0.0776 DOLOMITE 1.0481 ANHYDRITE 0.1233 K-FELDSPAR 0.7594 PLAGIOCLASE 0.2448 PYRITE 0.8345 CLAY 2.7438

54

LSU Geology & Geophysics Dept. 12/04/2007 - 10:26:39.46 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-2b.raw

QUARTZ 93.6360 CALCITE 0.0771 DOLOMITE 1.0422 ANHYDRITE 0.1226 PYRITE 0.8298 CLAY 2.7283 ARAGONITE 1.5641

LSU Geology & Geophysics Dept. 9/24/2007 - 10:25:19.05 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-3b.raw

QUARTZ 95.3693 CALCITE 1.3409 DOLOMITE 0.0757 ANHYDRITE 0.0534 K-FELDSPAR 0.7651 PLAGIOCLASE 0.1161 PYRITE 0.8096 CLAY 1.4699

LSU Geology & Geophysics Dept. 12/04/2007 - 10:27:15.55 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-3b.raw

QUARTZ 94.8244 CALCITE 1.3333 DOLOMITE 0.0752 ANHYDRITE 0.0531 PYRITE 0.8050 CLAY 1.4615 ARAGONITE 1.4475

55

LSU Geology & Geophysics Dept. 9/24/2007 - 10:25:56.95 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-4b.raw

QUARTZ 96.6339 CALCITE 0.0799 DOLOMITE 0.0941 ANHYDRITE 0.1080 K-FELDSPAR 0.3463 PLAGIOCLASE 0.1279 PYRITE 0.4024 CLAY 2.2076

LSU Geology & Geophysics Dept. 12/04/2007 - 10:27:53.45 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-4b.raw

QUARTZ 89.5173 CALCITE 0.0740 DOLOMITE 0.0871 ANHYDRITE 0.1000 PYRITE 0.3728 CLAY 2.0450 ARAGONITE 7.8037

LSU Geology & Geophysics Dept. 12/04/2007 - 10:28:41.12 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-5&6b.raw

QUARTZ 94.6428 CALCITE 1.0301 DOLOMITE 0.5203 ANHYDRITE 0.1237 PYRITE 0.9071 CLAY 1.1878 ARAGONITE 1.5883

56

LSU Geology & Geophysics Dept. 9/24/2007 - 10:26:35.18

------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\1c-5&6b.raw

QUARTZ 95.4378 CALCITE 1.0388 DOLOMITE 0.5247 ANHYDRITE 0.1247 K-FELDSPAR 0.6378 PLAGIOCLASE 0.1239 PYRITE 0.9147 CLAY 1.1978

LSU Geology & Geophysics Dept. 12/04/2007 - 10:29:27.26 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\2c-1&2b.raw

QUARTZ 90.7266 CALCITE 0.6359 DOLOMITE 0.9324 ANHYDRITE 0.0687 PYRITE 0.7645 CLAY 1.3677 ARAGONITE 5.5042

LSU Geology & Geophysics Dept. 9/24/2007 - 10:27:11.15 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\2c-1&2b.raw

QUARTZ 95.5077 CALCITE 0.6694 DOLOMITE 0.9815 ANHYDRITE 0.0723 K-FELDSPAR 0.4037 PLAGIOCLASE 0.1209 PYRITE 0.8048 CLAY 1.4398

57 LSU Geology & Geophysics Dept. 9/24/2007 - 10:27:44.05 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\2c-3&4b.raw

QUARTZ 95.1197 CALCITE 0.0873 DOLOMITE 0.0800 ANHYDRITE 0.0991 K-FELDSPAR 0.5884 PLAGIOCLASE 0.2387 PYRITE 1.4436 CLAY 2.3432

LSU Geology & Geophysics Dept. 12/04/2007 - 10:30:07.52 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\2c-5&6b.raw

QUARTZ 95.6710 CALCITE 0.0945 DOLOMITE 0.1351 ANHYDRITE 0.1912 PYRITE 1.6136 CLAY 2.2946 ARAGONITE 0.0000

LSU Geology & Geophysics Dept. 9/24/2007 - 10:28:18.38 ------X-RAY DIFFRACTION : MINERAL CONCENTRATION REPORT

Library File: mineral.lib XRD Data File: c:\progra~1\xrdfil\2c-5&6b.raw

QUARTZ 94.5640 CALCITE 0.0934 DOLOMITE 0.1335 ANHYDRITE 0.1890 K-FELDSPAR 0.9691 PLAGIOCLASE 0.1879 PYRITE 1.5949 CLAY 2.2681

58

59

60 APPENDIX B TAXONOMIC DISTRIBUTION OF BACTERIAL CLONES

1. Taxonomic Distribution of Bacterial Clones

61

Phylogenetic Representative Sequence Closest Relative % No. Affiliation [Accession Numbers] [Accession Numbers] Similarity Clones α-Proteobacteria CK_2C3_13 [EU488151] Agrobacterium sp. PB [AF482682] 99 247 Uncultured alpha proteobacterium; trichloroethene-contaminated site CK_2C2_6 [EU488118] [AF529121] 99 1 Hyphomicrobium sulfonivorans CK_1C3_23 [EU488001] strain CT [AY468372] 91 2 Rhodobacter sphaeroides CK_2C3_84 [EU488222] [EU410423] 98 4 Sinorhizobium sp. R-25067 CK_2C3_105 [EU488243] [AM084031] 99 1 Erythrobacter piscidermidis; CK_2C3_103 [EU488241] [ EU127294] 99 3 Uncultured alpha proteobacterium [DQ811854]; [AM084883]; CK_1C4_4 [EU488067] [DQ985050 ] 97 3 Alpha proteobacterium ML42 CK_1C4_45 [EU488073] [AJ315683] 99 3 Sphingomonas elodea (Pseudomonas CK_2C3_61 [EU488199] sp. ATCC 31461) [AF503278] 99 1 Alpha proteobacterium EU26 CK_2C4_100 [EU488389] [AY428761] 97 3 CK_1C4_20 [EU488046] Methylocystis sp. 5FB1 [AJ868421] 98 3 Sphingomonas sp. JSS-26 CK_2C3_57 [EU488195] [AF131296] 98 1 Sphingomonas sp. AC83 CK_2C5_13 [EU488444] [AJ717392] 93 7 β-Proteobacteria CK_1C3_45 [EU488023] Leptothrix sp. S1.1 [DQ241397] 99 1 γ-Proteobacteria CK_1C1_26 [EU487882] Methylocaldum sp. E10a [AJ868426] 87 7 CK_1C1_40 [EU487897] Methylobacter sp. 5FB [AJ868427] 95 1 CK_1C4_10 [EU488035] Pseudomonas sp. 7040 [AM111036] 99 4 CK_1C2_54 [EU487957] Pseudomonas jessenii [AF068259] 98 2 Uncultured clone Belgica2005/10- CK_1C3_46 [EU488024] 140-11[DQ351790] 99 1 L..pectinata symbiont 16S rRNA CK_1C3_6 [EU488031] gene [X84980] 99 1 Lucina nassula gill symbiont CK_1C3_51 [EU488029] [X95229] 97 1 Delta proteobacterium S2550 δ-Proteobacteria CK_1C1_7 [EU487909] [AF148141] 98 43 Uncultured delta proteobacterium CK_1C1_25 [EU487881] clone MSB-5D12 [DQ811831] 99 21 Uncultured delta proteobacterium CK_1C5_3 [EU488106] clone SK11 [AY771946] 99 3 Desulfobulbus mediterraneus CK_1C4_69 [EU488098] [AF354663] 95 1 Olavius algarvensis sulfate-reducing CK_2C2_2 [EU488114] endosymbiont [AF328857] 99 14

62 Desulfoarculus baarsii DSM 2075 CK_2C4_12 [EU488301] [AF418174] 100 6 Shigella flexneri 2a str. 2457T, CK_2C3_7 [EU488145] complete genome [AE014073] 100 2 Epsilon proteobacterium E9S37-1 ε-Proteobacteria CK_1C4_9 [EU488101] gene [AB175511] 92 11

Acidobacteriaceae bacterium CH1 Acidobacteria CK_1C1_6 [EU487908] 1 [DQ355184] 91 9 Uncultured Acidobacteria bacterium CK_1C3_12 [EU487990] clone VHS-B4-69 [DQ395006] 98 5 Uncultured Acidobacteria/Holophaga group bacterium clone Cart-N4 CK_2C3_5 [EU488143] [AY118153] 95 5 Uncultured Acidobacteria bacterium CK_2C4_9 [EU488298] isolate 5g10 [AY177760] 91 7 Arthrobacter monumenti Actinobacteria CK_1C1_1 EU487865 [AJ315070] 83 1 CK_1C4_16 [EU488041] Arthrobacter pigmenti [AJ639827] 84 7 Uncultured actinobacterium CK_1C2_75 [EU487980] [AM259897] 93 7 Flavobacterium sp. Sun4 Bacteroidetes CK_1C1_39 [EU487896] [AF349726] 99 6 Salinibacter ruber strain M8 CK_1C4_42 [EU48070] [AF323501] 98 1 Bacteroidetes bacterium clone CK_1C3_19 [EU487997] Belgica2005/10-130-16 [DQ351761] 99 3 Bacteroidetes bacterium SC-1 clone CK_2C4_39 [EU488328] 89.1[DQ357758] 97 2 Dehalococcoides sp. FL2 Chloroflexi CK_1C1_11 [EU487867] [AF357918] 84 94 Uncultured Chloroflexi bacterium CK_1C3_47 [EU488025] clone GoM HDB-03 [AY542195] 90 17 Uncultured Chloroflexi bacterium CK_1C3_4 [EU488018] clone VHS-B3-80 [DQ394964] 98 1 Planctomycetes CK_1C1_31 [EU487888] Planctomycete A-2 [AM056027] 95 13 Uncultured planctomycete clone CK_1C4_32 [EU488059] SC1-36 [DQ289921] 92 5 CK_1C4_18 [EU488043] P,.maris 16S rDNA [X62910] 75 3 Uncultured planctomycete clone CK_2C4_117 [EU488406] MSB-5C12 [DQ811900] 100 3 Firmicutes CK_1C1_9 [EU487911] Bacillus sp. BR [AM050346] 98 4 Tepidimicrobium ferriphilum strain CK_1C4_66 [EU488095] SB91 [AY656718] 98 4 Abiotrophia defectiva strain CK_1C4_19 [EU488044] [AY879308] 98 1 Clostridium lactatifermentans CK_1C2_24 [EU487926] [AY033434] 84 2 CK_1C2_60 [EU487964] Bacillus sp. TX3 [AB043863] 100 1 Uncultured Clostridia bacterium CK_2C3_67 [EU488205] [AY370633] 97 1

63 Clostridium sp. MDA2315 CK_2C4_14 [EU488303] [AY238334] 92 10 Spirocheaetes CK_1C4_13 [EU488038] Spirochaeta sp. Buddy [AF357916] 99 5 Uncultured spirochete clone MS12- CK_1C2_70 [EU487975] 6-B11[AM712343] 99 2 Uncultured Deferribacteres clone Deferribacteres CK_1C1_18 [EU487873] MSB-4E9 [DQ811935] 100 7 Uncultured Nitrospirae bacterium Nitrospirae CK_1C3_34 [EU488012] gene [AB176701] 93 1 CK_1C1_12 [EU487868] Nitrospina sp. 3005 [AM110965] 99 2 Uncultured Nitrospirae bacterium CK_2C3_25 [EU488163] clone MSB-4D12 [DQ811893] 85 1 Uncultured Verrucomicrobia clone Verrucomicrobia CK_1C3_50 [EU488028] LD1-PA26 [AY114317] 91 1 Uncultured Thermotogales bacterium Thermotogae CK_1C3_33 [EU488011] clone B17_otu14 [DQ097679] 96 1 Candidate Candidate division OP8 bacterium Divisions CK_1C4_64 [EU488093] clone VHS-B3-2 [DQ394925] 99 14 Uncultured candidate division OD1 CK_2C4_133 [EU488422] bacterium clone [AY193175] 89 2 Candidate OP11 MSB-3A10 CK_2C3_119 [EU488257] [DQ811940] 100 1

64 VITA

Angela Marissa Garcia was born in September, 1978, to Jeannie Green and Rudolph (Rudi) Garcia in Idabel, Oklahoma. She is the only girl among four children. She has one older brother, Angel

Garcia, and two younger brothers, Anastacio Garcia and Adan Garcia. Angela was raised in

Houston, Texas, where she graduated from Bellaire High School. She worked for NASA as a coordinator and director of an Urban Outreach Program in Earth and Space Science while pursuing her bachelor’s in biology at the University of Houston-Downtown. She was a member of the

Scholar’s Academy and a recipient of the Office of Naval Research, USFDA, and Shell Mentor scholarships. Angela began her pursuit of a master’s degree at Louisiana State University (LSU) in the Fall of 2005, where she was a member of the Geoscience Alliance to Enhance Minority

Participation (GAEMP) program in the Department of Geology and Geophysics. Angela attended her first Geological Society of America conference in October, 2005, where she presented a poster about the GAEMP program titled: “Evolution and Geomicrobiology: An Essential Teaching Tool for GAEMP”. In the summer of 2006, Angela was a field camp instructor at the LSU geology field camp in Colorado Springs, Colorado. She has presented posters about her thesis research at the

Geological Society of America’s annual meeting in Denver, Colorado, in October, 2007, as well as the American Geophysical Union’s annual meeting in San Francisco, California, in December,

2007.

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