MicroDiv2010 Author: Wang, Harris H.

A mini-project report for the 2010 Microbial Diversity Course at the Marine Biology Laboratory in Woods Hole, MA, USA

DNA subsistence, natural competency, and horizontal gene transfer in marine and soil organisms in the environment

Harris H. Wang, Ph.D.

Department of Genetics Harvard Medical School, Boston, MA, USA

Address: 77 Avenue Louis Pasteur, NRB Room 238, Boston, MA 02115, USA Email: [email protected] Phone: 617-432-6976

Course Instructors: Prof. Dan Buckley and Prof. Steve Zinder

Date: July 27, 2010

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MicroDiv2010 Author: Wang, Harris H.

ABSTRACT

DNA is the universal genetic material of cells, but can potentially serve also as a source of nutrients as it is rich in carbon, nitrogen, and phosphate. Utilization of DNA as nutrition requires importing the large DNA molecules into the cell through mechanisms akin to those of natural competency. This study aimed to enrich and isolate DNA-subsisting bacteria that are also naturally competent for their ability to uptake long pieces of DNA and utilize their genetic information through integration into the chromosome. DNA-subsisting bacteria were isolated both from soil and marine environments over the course of 3 weeks. Various physiological characteristics were measured including the potential for natural transformation. These enrichments and isolations are useful for developing new model organisms for which new genetic systems can be easily developed by reducing the barrier of introducing exogenous DNA into cells through natural mechanisms.

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MicroDiv2010 Author: Wang, Harris H.

INTRODUCTION

DNA is conventionally viewed as the essential information-carrying genetic material common to all organisms that exist in the intracellular space of cells. However, dissolved DNA is also prevalent in the environment, existing up to 0.2-50 micrograms per gram of soil and 0.2-44 micrograms per liter of marine water [1]. DNA is particularly rich in phosphate which is known to be a nutritional limitation in marine environments. It has been shown that up to 25% of all phosphates in some freshwater ecosystems are found in dissolved DNA [2]. Furthermore, DNA has a high turnover rate, which makes it an important factor in the carbon, nitrogen, and phosphate recycling system in nature [3, 4]. However, few evidences are found in the literature for the existence of oligotrophic organisms that can utilization DNA for nutrition in the environment. A survey of marine water revealed Vibrionales and Altermonodalecea that can subsist on DNA [5], but no such studies have been done for terrestrial bacteria. On the other hand, there is a wealth of literature describing the general phenomenon of natural competence in which extracellular DNA is taken up into the cell by membrane transporters [6]. These naturally competent cells are putatively thought to uptake extracellular DNA as a means for acquisition of novel trait that are encoded by the heterologous DNA. More likely than not, similar organisms tend to share more similarly useful genes, which is certainly the explanation for why some organisms such as H. influenza specifically uptake exogenous DNA containing sequence motifs similar to its own genome [7]. Genetic experiments have shown that competence genes are important in the utilization of DNA as nutrition in long-term stationary phase E. coli [8], providing further evidence that DNA utilization and natural competency may be linked in environmental organisms.

Understanding DNA recycling in the environment is an important step towards understanding the role of DNA not only as a source of genetic material but also as a source of nutrition and structural material as is sometimes found in biofilms. The ability to scavenge DNA for nutrition may also be associated with predation and cannibalism phenotypes of competing organisms under nutrient limiting environments. This study aims to investigate the prevalence of microbes that can subsist on DNA as it sole carbon source in both soil and marine environments by enrichment, isolation, molecular phylogenic analysis, and physiological measurements for DNA catabolism. Furthermore, we test the hypothesis that bacteria subsisting on extracellular DNA are more likely to be naturally competent and may be enriched for their ability to incorporate extracellular DNA into its genome. Since DNA transformation is one of the biggest barriers to developing genetic systems for new organisms, isolation techniques for naturally competent cells from the environment may be a useful strategy to accelerate development of recombinant technologies for these more tractable environmental organisms.

MATERIALS AND METHODS

Field sampling. Marine samples were obtained from the mouth of Trunk River which flows to the Atlantic Ocean near Falmouth, MA, USA. About 50ml water samples were taken at a depth of 0.5m. Soil samples were obtained from organic rich soils along a nearby hiking trail.

Media. For liquid media, NH4Cl (10 mM), Na2SO4 (1 mM), KH2PO4/K2HPO4 (1 mM), MOPS (10 mM), trace elements, and dissolved DNA were added to FW base (for cultivation of soil samples) or SW base (for cultivation of marine samples) to make FW broth (FWB) or SW broth (SWB), pH to 7.2 with NaOH, autoclaved for 15 minutes at 122 °C, and supplemented with pre- sterilized vitamin mix. The FW Base contains NaCl (17.1 mM), MgCl2 6H2O (1.97 mM), CaCl2 2H2O (0.15 mM), and KCl (6.71 mM) dissolved in MilliQ water. The SW base contains NaCl

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(342.2 mM), MgCl2 6H2O (14.8mM), CaCl2 2H2O (1.0 mM), and KCl (6.71 mM). Noble agar (15g/L) was added to the pre-autoclaved solution for making agar plates. Filter sterilized antibiotics were added as necessary: chloramphenicol (25 ug/ml), kanamycin (50 ug/ml), gentamycin (15 ug/ml), spectromycin (50 ug/ml), tetracycline (20 ug/ml), and ampicillin (100 ug/ml). Two forms of dissolved DNA were used: low-molecular weight (LMW) herring sperm DNA (Sigma D7290) and high molecular weight (HMW) double stranded DNA (salmon testes, sodium salt; Sigma D-1626) were used at final concentrations of 0.15 g/L to 1 g/L. The dissolved DNA is the sole carbon sources in the growth media. Salt Water Complete (SWC) rich medium was used for marine organism and nutrient Broth rich medium was used for soil organisms.

Enrichment. Liquid culture enrichment for bacteria that subsistent on DNA was performed using either FW media for soil sample or SWB media for marine sample supplemented HMW DNA. Simultaneously, control inoculums of FW or SWB media without dissolved DNA were also made. FW enrichment of soil sample was made by addition of 0.5g soil into FW media, vigorous vortexing for 5 minutes, settling for 1 hr, and inoculation of 100ul of the supernatant into 3mL of FW media with or without DNA. SW enrichment of marine sample was made by direct inoculation of 100ul of marine sample into SW media with or without DNA. Final DNA concentration in the enrichment was at 0.5-0.6 g/L. Samples were grown in the dark at 30 °C in a shaking incubator.

Isolation. Clones were isolated on FW or SW agar plates with and without dissolved DNA (0.15-0.5g/L). Isolates were obtained first on LMW DNA plates, but was subsequently switched to HMW DNA plates because subsistence on longer DNA fragments will increase the likelihood of enriching for naturally competent bacteria. Plates were incubated at 30 °C in the dark. Visible colonies generally formed in 3-5 days.

Growth curves, DNA quantification, microscopy. Growth curve experiments were conducted by measuring the optical density (OD 600nm) of enrichments at every ~12 hrs, blanked to a standard. Simultaneously, the extracellular DNA concentration in the growth media was determined on an UV-Vis Spectrophotometer (NanoDrop 2000c, Thermo Scientific, USA) by measuring the absorbance at 260 nm. Quality of the DNA reading was determined by the A260/280 ratio, which general varied between 1.8 and 2.0. Images of cell suspensions were taken on a Zeiss epifluorescent microscope (Axio) using the AxioCam MRc imaging system.

Clone library construction and phylogenetic identity of isolates. To determine the identity of individual organisms in the enrichment cultures, clone libraries of 16S rRNA genes were constructed from a genomic preparation (Ultraclean Soil DNA Isolation Kit, Mo-Bio Inc., USA) of the enrichment (1 ml sample), using bacterial PCR primers 8F and 1492R. The resulting PCR product was cloned into a vector for sequencing using the Invitrogen TOPO TA cloning Kit. A total of 96 clones were picked for each sample set. Four total sample sets were sent for Sanger DNA sequencing (Marine Biology Laboratory, Sequencing Facility): 1) enrichment on SWB media on HWM DNA, 2) enrichment on SWB media without DNA as control, 3) enrichment on FW media with HMW DNA and 4) enrichment on FW without DNA as control. Sequences were analyzed using the RDP web application (http://rdp.cme.msu.edu/). For identification of individual isolates (marine samples), individual colonies were picked into a 20ul lysis buffer 0.05% Triton X-100 and boiling at 105 °C for 5 minutes. One ul of the cell lysis product was used as template for PCR of the 16S rRNA gene using universal 8F and 1492R bacterial primers. The PCR consisted of initial denaturation at 95 °C for 5 min, 30 cycles of [46 °C annealing for 30 sec, 72 °C extension for 90 sec] and final extension at 72 °C for 5 min. The

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PCR products were screened by gel electrophoresis to confirm successful PCR amplification, treated by ExoSAP-IT kit, and sequenced by Sanger DNA sequencing.

DNA utilization plate assay. Utilization of DNA was determined on agar plates following a previous described method [5]. In brief, cells grown on plates with DNA substrates as the sole carbon source for 8 days prior was incubated in 600ng/ul of Ethidium Bromide in 1x Tris- acetate-EDTA (TAE) buffer for 15 minutes in the dark. The plates were then washed in fresh 1x TAE thoroughly prior to imaging on an UV-illuminated Gel Logic Imaging System. In general, a zone of clearance was observed for DNA-utilizing bacteria that secrete extracellular nucleases. As negative control, plates that do not contain dissolved DNA were used. As positive control, DNAaseI (20ul of 2500 units/ml) was added to plates that contained dissolved HMW DNA, incubated for 15 minutes, washed, EthBr stained, and assayed for DNA clearance.

Plasmid preparation. E. coli strains harboring plasmids (pRL27, pRL55, pRL59, pWM264, pDN18, pAH144, pCR4) containing antibiotic resistance cassettes (kanR, cmR, tetR, ampR, tetR, specR, ampR) were obtained from William Metcalf. The antibiotic cassettes on pRL55 and pRL59 are located on a transposon, which can randomly mobilize into the genome. Strains were grown up in the appropriate antibiotic and the plasmids were purified using a Qiagen Midi prep kit. The plasmids were resuspended in nuclease-free water and used for natural transformation assays.

Natural transformation assay. To determine the natural competence capacity of marine DNA- eating isolates, cells were first grown on SWC plates overnight. Using a multiplexed transformation method, clones were inoculated into a single tube containing SW broth (100ul). The tube was then split into two: HMW DNA + plasmids were added to one, and the other is left as a negative control. Equal amounts of plasmids were added (~1ug per plasmid) and supplemented with ~15ug of HMW DNA. Cells with or without HMW DNA+plasmids were incubated for 6 and 18 hours prior to plating 20 ul of the sample on to SWC plates that contained each of the antibiotics. From each of the antibiotic resistance plates, the number of resistance colonies was counted following incubation for 24-48 hours at 30 °C.

RESULTS

Isolation of DNA-subsisting bacteria from marine and soil environment

Individual bacterial isolates that subsist on DNA as its sole carbon source were successfully isolated from the marine environment. Plating on SWC agar produced ~130,000 viable colony forming units (CFUs) per ml of marine water. Plating on SWB+LMW DNA produced ~8,000 viable CFUs per ml. Plating on SWB+HMW DNA produced ~3,300 viable CFUs per ml. Plating on SWB (SW Broth, no carbon source) produced ~250 viable CFUs per ml suggesting that background growth was low. From this analysis, DNA-subsisting bacteria appear to exist in the marine environment at an abundance of 2.5-3.1% among culturable microbes with a slightly higher abundance of LMW DNA eaters (Figure 1a). Forty-eight colonies were isolated from the SWB+HMW DNA plate for further characterization including DNA utilization, antibiotic resistance, phylogenetic identification, natural competence assay.

Individual bacterial isolates that subsist on DNA as its sole carbon source were successfully isolated from the soil environment, although with more difficulty. Plating on NA agar produced ~88,000 viable CFUs per gram of soil. Plating on FWB+LMW DNA produced ~39,000 viable CFUs per gram of soil. Plating on FWB (FW Broth, no carbon source) produced ~42,000 viable

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CFUs per ml suggesting that background growth was rather high. It was difficult to determine the abundance of DNA-subsisting bacteria in the soil due to the high background level (Figure 1b). Nonetheless, isolates were successfully obtained from the FWB+LMW DNA plate because the colonies were much larger in size in comparison to background growth. The high background level is likely due to nutrient-rich soil micro-particles that are adsorbed to the cell, which supply nutrient for growth even under carbon limited conditions. Six colonies were isolated and re-streaked on FW+HMW DNA plates for DNA utilization assays and an unsuccessful attempt at phylogenetic identification by 16S rRNA profiling was performed.

Microscopic Visualization of Isolates and Dissolved DNA Degradation on Plates

Visualization of the isolates grown on FW+LMW DNA through the dissection microscope revealed mixed morphology, from round smooth colonies to fuzzy hyphal-structured colonies with dark pigmentation (Figure 2 a,b). Dark droplets were observed at the center of some colonies (Figure 2d) suggesting that these may be degradation droplets of solid agar, which the cells could be utilizing as a carbon source (these cells were subsequently shown to create DNA halos suggesting metabolism of dissolved DNA.

Plates that contained dissolved DNA could be quantitatively assayed for DNA concentration by EthBr staining. Incubation of post-8 day inoculated isolates on FW (Figure 3a) and SW (Figure 4) DNA containing plates showed distinct zones of clearance similar to a plaque assay. These results suggest that extracellular nucleases were excreted around the colonies for degradation of HMW DNA into LMW DNA for utilization. All 5 FW DNA-subsisting isolates showed zones of clearing although the morphology and growth rates among the isolates were drastically different. Control plates containing no DNA did not show zones of clearing with or without cells. Additionally incubation of FW+HMW DNA plates containing droplets of DNAaseI (Figure 3b) showed zones of clearing similar to the putative mode of DNA degradation by the cells. However on SW enriched marine isolate, some colonies did now show zones of clearing (Figure 4) suggesting that these were instead adopting an alternative strategy for DNA utilization, most likely from uptake of HMW DNA intact into the cell for further metabolism.

Quantification of DNA consumption in liquid enrichments

DNA consumption was quantified in liquid enrichments by using the NanoDrop and correlated to the optical density growth curves. The dissolved DNA concentration in liquid inversely tracked with the growth curve as expect, with decreasing dissolved DNA concentration with increased OD. Both the soil and marine enrichment produced turbid cultures after 24-48 hours (Figure 5). The optical density decreased over the 24-48 hours once the DNA concentration reached near zero levels. There was no background DNA degradation in the absence of addition of cells suggesting that the DNA is otherwise stable in solution over the growth period. The maximum density of the cells was higher in the SW enrichment compared to the FW enrichment (OD 0.2 vs 0.12).

Upon subsequent passaging, a more detailed growth curve and DNA consumption curve was generated (Figure 6) for which a doubling time of 7.0 hrs was determined for the FW sample and a doubling time of 6.3 hours was determined for the SW sample. Interestingly, the double time of DNA consumption for SW samples was 2.2 hrs versus 5.6 hrs for FW suggesting that the SW samples were eating the DNA at a faster rate while doubling at a moderately faster rate. A bulk estimation of the number of carbon molecules consumed per cell per doubling was 3.3 (±0.9) x 1010 carbon molecules for the FW enrichment and 1.5 x 1010 carbon molecules for the

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SW enrichment. In comparison, an E. coli cell is estimated to consume 7 x 1010 carbon molecules per doubling [9].

The liquid solution was run on an agarose gel to obtain a molecular weight size profile of the dissolved DNA. Dissolved DNA in FW broth in the absence of cells had a typical band size ~1- 5kb. The dissolved DNA in the FW soil enrichment showed DNA <1kb size after 60 hours of incubation, which further disappeared after 72 hours. The temporal change of DNA size is consistent with the observations from the DNA plate degradation assays in which DNAase is being secreted into the extracellular space to degrade HMW DNA into LMW DNA for consumption.

Microscopic imaging and community composition analysis of liquid enrichments.

Microscopy of the liquid enrichments of both the soil and marine samples revealed a diverse set of microorganisms. Dominant representative morphologies are shown in Figures 8 and 9. Some large cell consortium or clumps were observed for both enrichments, while other free living individual cells were observed. The rosebud-like cell morphologies appear to be those of Roseobacter (Figure 9) which was confirmed by the 16S profiling of 8 of 10 clones of the SW+DNA isolates classifying in the Rhodobacteraceae group (Figure 10).

Community composition analysis by 16S profiling of the clone libraries of SW-DNA and SW+DNA enrichments revealed very different organisms, with one dominated by Vibrionales and the other by Methylophaga. Due to technical error in labeling the sequencing plate, it was uncertain which clone library corresponded to the SW-DNA enrichment and which corresponded to the SW+DNA enrichment. It was likely that the Methylophaga group corresponded to the DNA subsisting enrichment (SW+DNA samples), but definitive claims require further verification by re-sequencing. However, the SW+DNA isolate 16S sequences did not overlap with either of the SW enrichment clone libraries, generating only α-proteobacteria not seen in the enrichment libraries. For the FW clone libraries, FW+DNA enrichment was dominated by Pseudomonadales, which was not seen in the FW-DNA enrichment. Sequences with the highest blast hits belonged to Acinetobacter calcoaceticus and Pseudomonas stutzeri, which are both known naturally competent bacteria.

Antibiotic Resistance Profile and Natural Transformation

Antibiotic sensitivity profile was determined for 96 direct isolates from the soil and marine samples for kanamycin, gentamycin, spectromycin, chloramphenicol, and tetracycline. In general, many isolates (>30%) were resistant to kanamycin, which may be due to the antibiotic concentration used. Most strains were sensitive to the five antibiotic tested, with a few having two resistances (Figure 12 a, b). The resistance profile of the soil and marine samples did not differ (Figure 12 d). The resistance profile of the 48 marine DNA-subsisting isolates from SW isolates did not differ from those of the 96 direct SW isolates (Figure 12 c, e). Isolates were most sensitive to tetracycline and chloramphenicol.

Twenty-four DNA-subsisting SW isolates were used to test for their ability to naturally transform DNA. A multiplex reaction was conducted in which the isolates were picked from colonies into a pooled tube containing SW broth with and without HMW DNA+plasmids. The cells were incubated for 6 and 18 hours at 30 °C prior to plating on various antibiotic plates. Plating of the pooled cell population on antibiotic resistance plates revealed a handful of antibiotic resistance cells (Table 1). After 6 hours of incubation, overnight cmR antibiotic plates showed 2 positive colonies for cells+DNA and 3 colonies for cells-DNA. On the other hand, 1 and 3 colonies were

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MicroDiv2010 Author: Wang, Harris H. observed on specR and gentR plates respectively for the cells+DNA incubation, while no colonies were observed for the cells-DNA incubation. However, lawns of colonies were found on the tetR and ampR plates for both cells-DNA and cells+DNA. After 18 hours of incubation, 161 cmR and 26 specR cells were found on the cells+DNA incubation and none on the cells-DNA incubation. Repetition of the experiment with a short 2 hr incubation produced 1 colony on each of the cells+DNA and cells+DNA incubations.

CONCLUSION AND DISCUSSION

In this study, I successfully enriched and isolated bacteria that subsist on DNA as their sole carbon source. Microscopy and phylogenetic analysis of the SW isolates revealed that the cells were Roseobacter. Community analysis of the enrichment cultures revealed different populations between the with- and without-DNA samples. For the FW soil enrichment, there was an enriched population of Pseudomonadales. Prior literature has shown that these organisms tend to be naturally competent. For the SW marine enrichment, there was a mix-up of the samples during sequencing that precluded definitive identification of whether the enriched Vibrionales or the Methylophaga belonged to the SW+DNA sample (since the individual SW isolates revealed α-proteobacteria Roseobacter, instead of the γ-proteobacteria Vibrio or Methylophaga enrichments). The physiological growth characteristics as determined by OD measurements and DNA consumption assays confirmed that these cells are indeed subsisting on DNA, with further proof from the zones of clearance on the DNA plate assays. Transformation of the cells with broad spectrum plasmids showed preliminary evidence that some of the strains are naturally competent for uptake of DNA, although these individuals were not isolated due to the time constraint of this project.

Further work will be needed to confirm the natural transformation phenotype. Ideally, the community analysis should be done on the enrichments after further passaging as it would reduce the background DNA levels of dead cells from the initial inoculum. My 16S clone libraries were generated on passage 1 enrichments due to time constraints of this project. Further plate- based isolations of individuals are needed from the liquid enrichments.

In general, this project demonstrates the feasibility of isolation and enrichment of soil and marine DNA-eating bacteria with the potential to be naturally competent. These organisms may serve as new model organisms and workhorses for recombinant technology as DNA transformation by natural systems is an important barrier for developing whole suites of downstream genetic tools that may be valuable for future geneticists, molecular biologists, and synthetic biologists.

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Figure 1. (a) Plate counts of environmental samples grown on SWC plates, SW+LMW DNA plates, SW+HMW DNA plates, and SW only plates. (b) Plate counts of environmental samples grown on Nutrient Agar plates, FW+LMW DNA plates, and FW only plates.

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a b

10x 80x c d

10x 80x e f

10x 25x

Figure 2. Images of three DNA-subsisting isolates from soil in low (a, c, e) and high (b, d, f) magnification.

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Figure 3. DNA utilization plate assay by EthBr staining. (a) FWB+HMW DNA plates incubated with soil isolates for 8 days, stained by EthBr and visualized on UV imager. (b) Control plates, (top) FWB only plate, (middle) FWB+HMW DNA plate incubated with two drops of DNAaseI, (bottom) FWB only plate with cells.

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Figure 4. DNA utilization plate assay showing isolates that grow and produce clearing (left) and isolates that grow but does not produce DNA clearing suggesting uptake of intact DNA.

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a. soil

b. marine

Figure 5. DNA utilization and growth curves of liquid enrichments for soil (FW) and marine (SW) samples. Light blue solid line shows DNA concentration in FW/SW without DNA and without cells; dark blue solid line shows DNA concentration in FW/SW with DNA and without cells. Light orange solid line shows DNA concentration in FW/SW without DNA and with cells; dark orange solid line shows DNA concentration in FW/SW with DNA and with cells. Light blue dotted line shows OD600 measurement in FW/SW without DNA and without cells; dark blue dotted line shows OD600 measurement in FW/SW with DNA and without cells. Light orange dotted line shows OD 600 measurements in FW/SW without DNA and with cells; dark orange dotted line shows OD 600 measurements in FW/SW with DNA and with cells. (a) Time course of DNA utilization (left axis, solid lines) of soil sample as determined by UV absorbance at 260nm of liquid medium. Growth curve (right axis, dotted lines) of soil sample was determined by OD 600 in liquid medium. (b) Marine samples with time course of DNA utilization (left axis, solid line) and growth curves (right axis, dotted line).

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a. b.

Figure 6. DNA consumption (solid line) and OD measurement (dotted line) of (a) soil and (b) marine samples after second serial liquid enrichment passage.

Figure 7. DNA size distribution in liquid media of FW enrichment. FW+D-C denotes FW with DNA without cells, 60 hrs post-inoculation; FW+D+C denotes FW with DNA with cells after 60 hrs and 72 hrs. High molecular weight double stranded DNA appears to be degraded to lower molecular weight DNA after 60 hrs post-inoculation, suggesting extracellular DNAase activity.

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Figure 8. Representative epifluorescent microscopic images of FW soil enrichments.

Figure 9. Representative epifluorescent microscopic images of SW marine enrichments.

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SW-DNA enrichment (80 clones) SW+DNA enrichment (85 clones) Proteobacteria Proteobacteria Gammaproteobacteria Gammaproteobacteria Oceanospirillales Alteromonadales Oceanospirillaceae Ferrimonadaceae Marinomonas 1 Ferrimonas 1 Unclassified Oceanospirillaceae 1 Alteromonadaceae Vibronales Glaciecola 2 Vibrionaceae Marinobacter 1 Vibrio 15 Unclassified Alteromadaceae 1 Thiotrichales Listonella 8 Piscirickettsiaceae Unclassified Vibrionaceae 54 Methylophaga 55 Unclassified Gammaproteobacteria 1 Unclassified Gammaproteobacteria 25

SW+DNA isolates (10 clones) Proteobacteria Alphaproteobacteria Rhodobacterales Rodobacteraceae Ruegeria 8 Unclassified Rhodobacteraceae 2

Figure 10. Classification of 16S rRNA clone library of SW without DNA and SW with DNA enrichments and DNA-subsisting isolates from SW with DNA plates.

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Figure 11. Classification of 16S rRNA clone library of FW without DNA and FW with DNA enrichments.

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a. Ab resistance (soil)

d. direct isolation

b. Ab resistance (marine)

e. DNA eating isolation

c. Ab resistance (marine)

DNA eaters

Figure 12. Comparison of antibiotic resistance profile of direct soil and marine isolates with isolates that can subsist on DNA. (a) Resistance counts among 5 tested antibiotics from soil. (b) Resistance counts from marine environment. (c) Resistance counts of DNA-subsisters from marine environment. (d) Distribution of antibiotic resistance in soil (orange bar) and marine (blue bar) from direct isolation among 96 clones each. (e) Distribution of antibiotic resistance of DNA- subsisters from marine environment among 48 clones.

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6hr +DNA -DNA cmR 2 3 specR 1 0 Trial #1 gentR 3 0 ampR + + tetR + + 18hr +DNA -DNA cmR 161 0 Trial #1 specR 26 3 gentR 0 0 2hr +DNA -DNA Trial #2 cmR 1 1 specR 1 1

Table 1. Summary of cell counts on antibiotic plates for cells that have putatively uptaken exogenous plasmids and conferred resistance to antibiotic. Around 400 million cells were spread per plate.

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ACKNOWLEDGEMENTS

I would like to first thank the course directors Profs. Dan Buckley and Steve Zinder for organizing this amazing microbial diversity course. I would like to thank all the TAs who have provided assistance throughout this project, with special thanks to Chuck Pepe-Raney for assistance during clone library construction, sequencing, and community analysis and course coordinator Rebekah Ward for acquiring all the necessary reagents used in this project. I would also like to thank Bill Metcalf for his many great lectures and sharing the plasmids which were used here. I also want to give thanks to all my classmates for making this course a wonderful, educational, and fun experience. I want to thank James Hendriksen especially for helping me acquire the inoculation samples, making the antibiotic plates, performing the direct isolation of bacteria to the antibiotic plates, as well as making many valuable suggestions throughout this project. I would also like to thank my advisor Prof. George Church at Harvard Medical School for allowing me to attend this course as well as financial support from the MBL for making this experience possible. To these wonderful and generous people and organizations, I am eternally grateful.

REFERENCES

1. Dell’Anno A. and Danovaro R. (2002) Quantification, base composition, and fate of extracellular DNA in marine sediments, Limnol Oceanogr 47(3): 899-905. 2. Siuda W, Güde H. (1996) Determination of dissolved deoxyribonucleic acid concentration in lake water. Aquat Microb Ecol 11:193-202. 3. Dell’Anno A and Danovaro R (2005) Extracellular DNA Plays a Key Role in Deep-Sea Ecosystem Functioning. Science, 309:2179. 4. Paul JH, Jeffrey WH, DeFlaun MF. (1987), Dynamics of extracellular DNA in the marine environment. Appl. Environ. Microbiol., 53(1):170-179. 5. Lennon J. (2007) Diversity and Metabolism of Marine Bacteria Cultivated on Dissolved DNA. Appl Environ Microbiol. 73(9): 2799-2805. 6. Lorenz M G, Wackernagel W. (1994) Bacterial gene transfer by natural genetic transformation in the environment. Microbiol Mol Biol Rev. 58(3): 563-602. 7. Sisco KL, and Smith HO. 1979. Sequence-specific DNA uptake in Haemophilus transformation. Proc. Natl. Acad. Sci. USA, 76: 972-976 8. Finkel SE, Kolter R. (2001). DNA as a nutrient: A novel role for bacterial competence gene homologs. J. Bacteriol. 183: 6288-6293. 9. http://bionumbers.hms.harvard.edu/bionumber.aspx?s=y&id=103010&ver=4

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