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Photoferrotrophy and iron cycling in Woods Hole microbial mats

(or: My Summer Quest for Unusual and Possibly Imaginary Phototrophs)

Erin Banning

Microbial Diversity 2005

Abstract Photoferrotrophs, anoxygenic phototrophs that use ferrous iron as an electron

donor, may play an important role in the cycling of iron in aquatic and near-shore habitats around Woods Hole. This project has enriched for photoferrotrophs from a variety of sites around Woods Hole, which have displayed differences in the manifestation and detectability of pigments, the nature of oxidized iron precipitates produced, the cell morphologies that are detectable and optimum light treatments. In addition, it appears that iron-reducers may be closely associated with these iron-oxidizers, both in the natural environment and in enrichment cultures. The possibility is raised, based on culture-independent clone sequence analysis, that novel photoferrotrophs may have been recovered in the enrichment cultures. A completely unknown theoretical organism, an anoxygenic methanotrophic phototroph, or photomethanotroph, is also enriched for and may exist in the Woods Hole area. Introduction

Iron is the fourth most abundant element in the earth, and the second most abundant redox-active element in near-surface aquatic environments. Its cycling influences the flux and distribution of many other elements through a variety of processes, including adsorption onto ferric oxides (phosphates, many metals), precipitation as ferrous minerals (sulfides, carbonates, phosphates) and the fixation (during microbial oxidation) and remineralization (during microbial reduction) of carbon. In the presence of molecular oxygen, reduced ferrous iron rapidly oxidizes abiotically to its oxidized ferric state, which is highly insoluble. Many geochemical indicators suggest that free iron was much more common during earlier periods of the earth’s history when atmospheric oxygen was low or absent and may have played an important role in the development of early life. In addition, with only a few controversial exceptions, every contemporary known life form requires iron for its metabolism and biosynthesis at some level. For all these reasons, microbial communities involved in high-flux cycling of iron are of great interest.

During field excursions as part of the Microbial Diversity course in 2005 small patches were observed within the context of extensive sulfur-dominated phototrophic mats (at Lesser Sippewissett, Greater Sippewissett – see Fig. 1, and Trunk River – see Fig. 2) where iron oxides appeared to be particularly common. This raised the possibility that patches within an area generally dominated by sulfur cycling may be dominated by iron cycling microbial processes, such as heterotrophic iron reduction, neutrophilic iron oxidation and photoferrotrophy. Heterotrophic iron reduction results from microbial respiration using ferric iron (3+) as an electron acceptor, and has been found to be

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important in iron-rich anaerobic environments for organic matter degradation (Lovley 2000). Neutrophilic iron oxidation is carried out by autotrophic microbes at oxygen concentrations low enough that biological oxidation with oxygen as the electron acceptor can compete with abiotic oxidation (Emerson and Weiss 2004) – therefore it is usually found in steep oxygen gradients where soluble ferrous iron is mixing with oxidized fluids or gases. Photoferrotrophy is a microbial process by which ferrous iron is oxidized phototrophically without oxygen, and has been found to be carried out by a diverse group of anoxygenic phototrophs (Croal et al. 2004). Ferrous iron can also be oxidized using nitrate as an electron acceptor (Straub et al. 2004).

In light of this observation, a crude model of possible dynamics in an iron-dominated phototrophic mat was developed, shown in Figure 3. In this situation, either there is enough oxidized iron in the sediment column to deplete organic matter before sulfate reduction becomes significant or there is insufficient organic matter to deplete the available ferric oxides. Therefore, any sulfide diffusing upwards from deeper layers is precipitated in the form of iron sulfides before nearing the surface, and instead high levels of dissolved ferrous iron diffuse upward towards the oxic-anoxic transition. As ferrous iron diffuses upwards, it may be oxidized by anaerobic nitrate-reducing microbes. Once it enters the photic zone of the mat, it may be subject to photoferrotrophic oxidation, and at the oxic-anoxic transition neutrophilic iron oxidation can operate. Above that point, oxygen concentrations would be too high to allow biological iron oxidation to operate at circumneutral pH. The resulting iron oxide layer might be overlain by oxygenic photosynthesizers, such as cyanobacteria or eukaryotic algae. If this type of dynamic does exist in iron-dominated mats, it would be expected generate different gradients of key chemical parameters such as oxygen, sulfide and pH than those found in dominantly sulfur-cycling mats, and might be able to be identified using microelectrode profiles.

In addition, extremely stable and vigorous methane bubbling was observed in Trunk River, a site of intense phototrophic growth, as shown in Figure 4. There is no known evidence for the phototrophic oxidation of methane, termed photomethanotrophy, but if it were to exist an environment such as Trunk River’s would be extremely favorable. Methane has been theorized to have been common on the early Earth, and if photomethanotrophy exists it may have been an important process at that time.

In order to approach these ideas, the following research questions were developed:

• Are photoferrotrophs culturable from iron-rich areas around Woods Hole? • If they are, at what wavelengths do their pigments absorb the most at, as a proxy

for their ideal position in the light column? • Do they co-occur with iron-reducers? • Are there systematic chemical differences between sulfur-dominated and iron-

dominated microbial mats? Are photomethanotrophs culturable from Trunk River? Materials and Methods Enrichment culturing

A base seawater media stock of two liters was prepared anaerobically using a Widdell vessel, the base seawater recipe is in Table 1. It was designed to have no carbon

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source and only nutritional levels of sulfur (as both sulfate and thiosulfate, based on reports that some photoferrotrophs are unable to assimilate sulfate as a sulfur source (Straub et al. 1999)). The media was dispensed anaerobically under positive N2/CO2 pressure on the bench top into serum vials in 10 and 25 mL aliquots, depending on the size of the bottle, and the bottles stoppered and crimped. Excess media was stored in screwtop bottles for later use. For photoferrotroph media, bottles were injected with .1 mL of 200 mM Fe(II)/NTA (5.4 g FeCl2*6H2O and 4.0 g Nitrilotriacetic acid in 100 mL deionized H2O, mixed and filter-sterilized anaerobically in the glovebox) per mL of seawater media. For iron-reducers, seawater base media was dispensed on the benchtop into acid-washed Pfennig bottles (with about 5.5 mL of ~100 mM ferrihydrite solution – made by titrating a FeCl3 solution with 1 M NaOH to pH~7 while stirring and repeatedly washing by allowing ferrihydrite to settle and pouring off supernatant – and 2.75 mL 200 mM autoclaved sodium acetate already dispensed into each) until nearly full and capped with screw caps. Photomethanotroph media was prepared by dispensing base seawater media into large serum bottles (for larger headspace) under an N2/CO2 headspace and, after inoculation, injecting ~20 mL ultrapure methane with a gas-tight syringe.

Photoferrotroph bottles were inoculated in triplicate from water and sediment samples collected from: four sites at Trunk River (rusty pool 1, rusty pool 2, orange/pink/green veil and red-orange patch – shown in Fig.2), an iron oxide mat in the School Street parking lot (shown in Fig.5), the top of a Nalgene full of water taken from Trunk River and overgrown by purple and green sulfur bacteria, and three sites at the Parker River Wildlife Refuge (a dry iron mat, rusty mud and a shallow dusty pool in the parking lot). The three bottles from each inoculum were divided between three light regimes, incubated and shaken by hand at least once a day: incandescent (a 120V 40W soft white bulb), fluorescent (a Kitchen & bath FC8T9.KB 22W bulb) and infra-red (a 940 nm LEDTronics LED). As they developed significant amounts of iron oxides or color associated with pigments, milliliter aliquots were removed and transferred into fresh bottles – at which point wet mounts were made and examined under the microscope.

Iron reducer bottles were inoculated with three 9-mm sections of a 1-mL syringe core taken from an iron-oxide-rich mat at the Greater Sippewissett salt marsh, shown in Fig.1, one section per bottle, and incubated at room temperature in the dark. Syringe core was prepared by using scissors to cut off the tip of a sterile 1-mL syringe, inserting the syringe into sediment and drawing back on the plunger while twisting it in until full.

Photomethanotroph bottles were inoculated in duplicate from two bubbling regions at Trunk River (one of which is shown in Fig.4) in the same manner as the photoferrotroph bottles, except for the methane injection. The two bottles from each inoculum were divided between the fluorescent and infra-red light regimes, and an uninoculated methane-injected bottle was kept under the fluorescent light regime as a control. Upon the development of significant growth or color, one milliliter was transferred into a fresh bottle and a wet mount made and examined. The bottles were shaken at least daily. Anaerobic shake tubes

Anaerobic shake tubes were prepared to aid in isolation of photoferrotrophs from enrichments. A three-percent solution of SeaPlaque GTG agarose (low-melting) was made, washed once, autoclaved and kept fluid in a 70 degrees C water bath while being

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poured in 3 mL aliquots into acid-washed and autoclaved batch tubes. Base seawater media was prewarmed to about 45 degrees C in another water bath, and the agarose aliquots autoclaved. A heating block was filled with sand and preheated to 65 degrees C. When all ingredients were assembled, they were moved quickly into the glovebox under an N2/CO2/H2 atmosphere – seven agarose tubes and one batch of prewarmed seawater media at a time. Each tube had 6 mL of prewarmed media added and was quickly capped and returned to the sand bath. One milliliter of primary enrichment (Trunk River rusty pool infra-red and School Street lot fluorescent enrichments were each diluted to 10-7) was injected into one tube (the 10-1 tube), which was inverted gently to mix. In quick succession, one milliliter was removed from the 10-1 tube and added to the next, and the process repeated in series until the 10-7 tube was inoculated. The tubes were then removed from the glovebox, crimped and incubated under the appropriate light regime. The tubes were checked at least once every two days for colony development using the dissecting microscope. Upon colony development, colonies were picked under the dissecting microscope using a Pasteur pipet connected to a pipet tip (held in the mouth) by a flexible hose into 20 to 50 microliters of base seawater media or sterile phosphate buffer solution and resuspended for DNA extraction, wet mounts or inoculation into fresh media. DNA extraction and Clone Libraries

The MoBio Ultraclean DNA Kit for Soil was used in an attempt to extract amplifiable DNA from three- and six-mm sections of two 1-mL syringe cores taken in the Greater Sippewissett Salt Marsh. One core was taken within the iron-oxide-rich area, and the other about 10 cm away in an apparently more sulfur-dominated area. Both cores were kept on ice after collection and then frozen until extraction, at which point they were slowly extruded and each section knocked into a 1.5-mL Eppendorf tube in turn. The kit procedure was used without significant modification.

A subset of the available DNA extracts from the cores was selected for PCR amplification: three sections comprising the topmost 9 mm of the iron-dominated core, one section taken from between 12 and 15 mm on the iron core and two sections from the sulfur-dominated core – from 6 to 9 mm and from 12 to 18 mm. Ten- and hundred-fold dilutions were used as template for a PCR reaction with the general bacterial primers 8F and 1492R with the following parameters: five minutes of denaturation at 95° C, 25 cycles of denaturing for 30 seconds at 95° C, annealing at 50° C for 30 seconds and extension for 1.5 minutes at 72° C followed by five minutes of final extension at 72° C. No PCR product was detected.

As an alternative DNA extraction method, the Epicentre soil DNA extraction kit was used to extract DNA from photoferrotroph enrichment cultures. Before extraction, three one-milliliter aliquots were taken from each of four enrichments (Trunk river rusty pool 1 IR, Trunk River rusty pool 1 IR first transfer, School Street lot FL, and Top of Trunk River Nalgene IR) and washed with sodium dithionate and ammonium oxalate to remove amorphous iron oxides. Fifty microliters of each solution (both ~100 mM) were added to each aliquot of culture and incubated for a few minutes at room temperature – some extracts showed marked clearing (in particular those that had the least amount of precipitate to begin with). The extracts were then centrifuged at maximum speed for 12 minutes, the supernatant discarded, and the pellets resuspended in phosphate buffer solution. The wash was repeated three times before moving the resulting cell suspension

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(resuspended in smaller volume) into the Epicentre soil kit with no further modifications to the procedure. After extraction, the extracts were directly used as templates of varying dilutions (2X, 1X, .5X) in a PCR reaction using the general bacterial primers 8F and 1492R under the same reaction conditions as above (except using 30 cycles). Four PCR products were obtained – the extract from Trunk River rusty pool 1 IR amplified at all dilutions and the School Street FL extract amplified at the 2X dilution. Immediately after PCR, aliquots of the Trunk River rusty pool 1 IR .5X dilution product and the School Street FL 2X product were cloned using the Topo TA kit. Forty-eight clones were picked for sequencing from each library, and the clone sequences aligned and analyzed using ARB. In addition, the 16S rRNA sequence of Thiodictyon strain Thd2 (GenBank accession number X78718) was downloaded from Genbank, added to the ARB database and aligned for comparison in phylogenetic trees.

In addition, three colonies were picked from the 10-7 dilution of each set of anaerobic agar shake tubes and resuspended in 20 µL of base seawater media. One microliter of P40 detergent solution was added to each aliquot and the aliquots incubated for several minutes in a 95° C water bath before being used as templates for direct PCR under the above conditions using 8F and 1492R primers. No PCR products were recovered. Pigment analyses

Three different procedures for pigment spectral analysis were attempted. First, one-milliliter aliquots of enrichment cultures were scanned in a spectrophotometer blanked with deionized water across the range from 400 to 1100 nm. Second, if no discernible spectrum was obtained, the aliquot was incubated with 125 µL each of 100 mM sodium dithionate and ammonium oxalate and then centrifuged to pellet the cells. The supernatant was removed and scanned in the spectrophotometer as above, while the remaining pellet was resuspended in phosphate buffer solution and again incubated with sodium dithionate and ammonium oxalate before a second centrifugation. The second supernatant and the resuspended pellet were scanned in the spectrophotometer.

Third, cultures that resisted pigment analysis via the above procedures were again aliquoted and the aliquots sonicated on ice three times for 30 seconds each at maximum output with 30 seconds of rest in between each sonication. The sonicated solution was directly read in the spectrophotometer, and then centrifuged for 10 minutes at 11000 rpm and the supernatant read in the spectrophotometer. Scanning Electron Microscopy

One-milliliter aliquots from both primary enrichment and first transfer of the Trunk River rusty pool 1 IR culture were gently filtered on Nucleopore membrane filters (pore diameter .2 µm) using a vacuum pump and then fixed for about 4 hours in a glutaraldehyde/formaldehyde fixative solution (see Table 2). The fixative was rinsed out with several changes of MilliQ water, and then dehydrated by replacing the last water rinse with acidulated 2,2-dimethoxypropane. The sample was agitated gently and allowed to stand for ten minutes before removal with a Pasteur pipet and replacement with absolute ethanol. The filters were then critical-point dried, mounted with double-sided tape on SEM stubs, sputter-coated and stored in a dessicator until examination using the Scanning Electron Microscope operated at 15 kV. DAPI staining and optical microscopy

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In an effort to determine microbes’ relationship with mineral precipitates in the photoferrotroph enrichment cultures, wet mount DAPI staining was used. A wet mount was made from a couple of drops of culture aliquot and one drop of 10 mg/mL DAPI solution added. The slides were incubated without a cover slip in the dark for three minutes before the addition of the cover slip and examination using epifluorescence microscopy. Microelectrodes

The available Unisense oxygen/pH and Diamond General sulfide electrodes were used at the Greater Sippewissett Salt Marsh to obtain three profiles in and around an iron-oxide-rich location (shown in Fig.6) in an effort to compare iron-dominated to sulfur-dominated dynamics. Before going into the field, the oxygen/pH electrode was polarized in sparging water for more than two hours. The picoammeter was adjusted for each electrode to the appropriate polarity (+.08 V for the sulfide electrode, -.7 V for the oxygen/pH electrode) and the range adjusted. In addition, points on a standard curve for calibration of the sulfide electrode were prepared by anaerobically diluting a standard solution of NaS to 1, .5 and .25 mM. In the field, a garden cart was used to transport all the required equipment and serve as an operating platform in the marsh. Three profiles were obtained: the first, taken within the iron-rich area, penetrated to 9 mm taking five data points every 250 µm with a 15 second resting time after reaching each depth; the second, also in the iron-rich area, penetrated to nearly 15 mm taking three data points every 400 µm with the same resting time; and the third, taken outside of the iron-rich area about 10 cm away, penetrated to 15 mm and used the same parameters as the second profile. After returning from the field, additional calibrations of the data were performed using seawater boiled and cooled under N2/CO2 positive pressure, pH standard solutions and the sulfide standard curve. Methane analysis

In an effort to quantify any methane consumption in the photomethanotroph bottles, a Shimadzu GC-14A was used in FID mode to detect methane. A sterile syringe purged with N2/CO2 gas was used to withdraw about 400 µL of headspace gas from each photomethanotroph bottle and 100 µL of gas injected into the column. An uninoculated photomethanotroph bottle was also analyzed for methane concentration as a control. Results

Three main patterns emerged in the primary enrichments. First, several of the primary enrichments inoculated from Trunk River samples developed a strong green color under the incandescent light treatment and maintained a much lower level of precipitated material than the other bottles, as is shown in Figure 8A and 8C. A pigment spectra run on the primary enrichment (shown in Fig.7) revealed a strong peak at 758 nm in both of the Trunk River rusty pool primary enrichments incubated in front of an incandescent light bulb, along with a less distinct peak at 460 nm. These peaks indicate that the color is caused by green sulfur bacteria containing bacteriochlorophyll c. A wet mount was made and examined from the first transfer from the Trunk River rusty pool 1 incandescent enrichment showed a dominance of a mixed population of large (up to ~ 8 µm long, <1 µm in diameter) irregularly curved rods not seen in other enrichments, shown in Fig.9.

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Second, many of the enrichments under a variety of light treatments from a variety of inocula developed beige and deep orange colors and large amounts of oxide precipitates, as is visible in Figure 8A and 8B and tabulated in Table 3. Despite the efforts described above, no intelligible pigment spectra was able to be obtained from them – separating the cells from the iron oxides proved to be difficult. Figure 10 shows the various spectra obtained from different stages in the sonication and oxalate treatment procedures on the Trunk River rusty pool 1 infra-red primary enrichment. Two of these enrichments, as described above, had amplifiable DNA extracted from them and clone libraries were built for them. The DNA was extracted on July 26 from both enrichments, eight days after their inoculation. The School Street lot fluorescent library contained 34 clones that all group with high similarity to Shewanella putrefaciens (a heterotrophic iron-reducer), five clones that were highly similar to Aeromonas hydrophila (a common aquatic heterotroph that is often pathogenic), one clone that grouped closely with Rhodoferax ferrireducens (an iron-reducer) and Rhodoferax fermentans (a purple non-sulfur bacteria) and one clone that with Aquamonas fontana. The Trunk River rusty pool 1 infra-red library yielded 38 clones which for the most part group together in a new lineage within Thiomicrospira, a lineage of sulfur-oxidizers. In addition, it yielded one clone that also grouped within Thiomicrospira, most closely with T. crunogena, a nitrate-reducing hydrocarbon degrader, and one clone that grouped with a large number of uncultured Bacteroidetes. Figure 11 is a phylogenetic tree showing the relationships between the clone sequences from these two enrichments, selected clone sequences from a library constructed from the top of a Nalgene filled with Trunk River water and sediment (mentioned above) and known photoferrotrophs. Microscopically, the orange cultures appeared dominated by irregular aggregates of small cells, sometimes in close association with large masses of, presumably, iron oxides, as shown in Fig.12.

In addition, anaerobic shake tubes were inoculated in serial dilution from both the School Street FL and Trunk River rusty pool 1 IR primary enrichments, as described above. They yielded three different colony morphologies, shown in Fig.13, all greenish: large lumpy colonies (SchStFL), irregularly shaped colonies and small cigar-shaped colonies (TR rusty pool IR). Each colony morphology was resuspended and inoculated into fresh photoferrotroph bottles.

SEM imaging of filtered samples from both the Trunk River rusty pool 1 IR primary enrichment and its first transfer was also performed. A variety of cell morphologies and associations with oxide minerals was observed, shown in Fig.14. Only one definite cell was found from the transfer, also visible in Fig.14. The most common cell form was vibrioid, although straight rods of varying proportions were also observed at lower abundances and greater diversity.

Third, some of the Trunk River enrichments immediately precipitated black, presumably ferrous sulfide, particulates upon inoculation. This occurred in particular, as shown in Table 3 and Figure 8C, in the Trunk River rusty pool 2 and orange/pink/green veil enrichments. In the case of the Trunk River rusty pool 2 enrichment, the incandescent (and ultimately fluorescent) light treatments have resulted in green growth similar to that seen in several other Trunk River incandescent enrichments. However, the precipitation of sulfides undoubtedly represents a significant input of reduced sulfur into the media, which may be supporting phototrophic growth.

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In addition, it was noted that most of the cultures preferentially developed particulate masses on the sides of their bottles closer to the light. Every day the bottles were shaken to resuspend particulates and cells, and the next day particulates had almost always settled as close as possible to the light.

Several iron-reducer media bottles were inoculated, and after a week or so of incubation in the dark all inoculated bottles previously orange and rusty-colored precipitates became black and presumably reduced, as shown in Fig.15.

There was significant growth in the photomethanotroph bottles under fluorescent light which has transferred vigorously, as shown in Fig.16. A clean pigment spectrum was easily obtained without sonication from the Trunk River FL primary enrichment 2, shown in Fig. 17, with a major peak at 757 nm and another clear peak at 460 nm. This spectrum has an almost identical peak structure to the spectra produced by Trunk River incandescent photoferrotroph primary enrichments, also indicative of bacteriochlorophyll c in green sulfur bacteria. Upon microscopic examination, the first transfer from the same bottle showed that it is dominated by three morphological types of rods – a small class of about 2-3 µm long, a middle class about 4 µm long and slightly curved and a long class about 10 µm long, also slightly curved – all three are visible in Fig.18. The methane concentration does not appear to significantly vary across the time course of the measurements, as shown in Fig.19.

The profiles obtained from microelectrode profiling are shown in Fig.20. The pH does not appear to change significantly through any of the profiles, and pH measurements are incomplete due to a shortage of power inverter outlets at one point during the field measurements. Based on the best correction of the electrode measurements that could be executed, the profiles indicate free oxygen coexisting with free sulfide – a baffling and unlikely case. Discussion

Based on the diversity of culture types and clone sequences recovered from Trunk River and School Street iron mat enrichments, it seems likely that there is a high diversity of potentially novel photoferrotrophs in the Woods Hole area, with adaptations for different light niches.

Based on the high proportion of non-phototrophic clone sequences obtained from the photoferrotroph enrichments, it appears that the cultures had gotten overgrown and old by the time of DNA extraction. This is particularly evident for the School Street fluorescent enrichment, in which the clone library is overwhelmingly dominated by iron-reducers (Shewanella, etc.), which are probably using the biomass generated by iron-oxidizers to respire ferric iron. In fact, they may be the DAPI-stained cells evident closely associated with the surfaces of oxide aggregates, as in Fig.12, especially considering that many anoxygenic phototrophs have been found to stain poorly with DAPI. Had the time, extraction and cloning capabilities been available it would be interesting to test whether these secondary metabolizers are retained in subsequent transfers and isolates. The Rhodoferax clone sequence from this library is of great interest, since it is closely related to both an iron-reducer and anoxygenic phototrophs (Madigan et al. 2000; Jung et al. 2004). Two main hypotheses are evident: either the Rhodoferax is an iron-reducer and is secondarily respiring ferric oxides or it is a photoferrotroph still present in some numbers in the enrichment. If it does represent a novel photoferrotroph, it would be the first

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known amongst the β-Proteobacterial purple non-sulfur bacteria. Given that the sequence is closely related to a known iron-reducer (R. ferrireducens), it might seem that the strongest hypothesis is that it is an iron-reducer itself. However, it is nearly as closely related to non-iron-reducing phototrophs (which comprise the majority of the group) and some microbes have been shown to be capable of both autotrophic iron oxidation and mixotrophic iron reduction, Ferroglobus placidus for example, a hyperthermophilic archaea (Tor et al. 2001). This ability to switch metabolic lifestyles almost completely diametrically could be very advantageous for dealing with circumstances in which a critical resource becomes exhausted.

In the Trunk River rusty pool 1 infra-red enrichment, the community appears dominated by sulfur-oxidizers (Thiomicrospira). However, most of the clone sequences group somewhat separately from known Thiomicrospira species, and so may have a different physiology, especially considering that T. crunogena, which groups with the one clone sequence outside the new group, is described as a nitrate-reducing hydrocarbon-degrader. It may be that enough sulfide was introduced into the primary enrichment through its inoculum from sulfide-rich Trunk River to sustain a sulfide-oxidizing community for a substantial period of time, but the question of what is being used as an electron acceptor for sulfur oxidation must be faced – the cultures are maintained anaerobically and free nitrate is not an initial component of the media. Fixed nitrogen is added to the media in the form of ammonium chloride, which itself cannot be used as an electron acceptor. However, if the ammonia could be oxidized to nitrate using another electron acceptor, nitrate-reducing sulfur-oxidizers might be able to thrive for some time. In general, Trunk River does seem to be a promising site for photoferrotrophy based on the diversity of culture development and the clone sequences recovered from the general enrichment (top of Nalgene) that are related to varying degrees to known photoferrotrophs, one of which (Thiodictyon strain Thd2) was isolated from the Woods Hole area (Croal et al. 2004).

Of course, the interpretation of clone libraries built from old enrichments, especially primary enrichments, is problematic. It is likely that the enrichments have become overgrown with an undesired mixed population, or that the populations may have crashed as a result of exhausting a critical substrate in a batch culture situation such as this one.

The development of green sulfur bacterial pigments in some of the photoferrotroph enrichments is suggestive that photoferrotrophs related to Chlorobium ferroxidans, the only known green sulfur photoferrotroph (Heising et al. 1999), may be growing in the bottles. Unfortunately time and resources did not allow quick response to the late development of green pigment in the Trunk River bottles in the form of DNA extraction and cloning. An alternative hypothesis that must be tested is that green sulfur bacteria are growing by oxidizing sulfide introduced into the media upon inoculation. However, the same kind of pigmentation might be expected to develop under the other light treatments as well if this were the case. The difference in visible particulate formation between the green and orange cultures also seems significant – raising several possibilities: that a green sulfur photoferrotroph is generating a different ferric iron product than in the other enrichments, which is more soluble; that it is complexing the ferric iron with some kind of siderophore; and that there is an iron reducer in close co-culture that is rapidly cycling the oxidized iron and preventing it from building up in

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culture. This last possibility is not outside the realm of possibility, since C. ferrooxidans has not been successfully isolated from a cocultured iron-reducer (Heising et al. 1999).

It is worth noting as well that at least circumstantially it appears that iron-reducers and photoferrotrophs may coexist in iron-rich mat systems, as iron-reducers were successfully enriched for from an iron-rich mat and so many iron-reducer clone sequences were recovered from the enrichment clone libraries.

In the future, FISH would be a useful tool for improving the ability to monitor the status of enrichments phylogenetically, at least by narrowing the component populations to general phylogenetic groups and overcoming the limitations of DAPI in staining phototrophic organisms with extensive intracellular membrane systems. In addition, moving to full pigment extractions using either acetone or methanol would be valuable – although the pigment is removed from its native cellular environment and loses any accessory absorptive adaptations it might have for a particular niche, at least phylogenetic identification on the basis of pigments would be facilitated and the pigments separated from iron oxides. Neither sonication and centrifugation or oxalate/dithionate washes appeared to be effective on their own in producing an interpretable pigment spectral signal.

Microelectrode profiling also proved to be inconclusive during this study. In hindsight, it appears that the oxygen electrode was not being properly calibrated. Other problems in the field that might have influenced the quality of the data include the dryness of the site at the time of sampling and the unknown magnitude of biological and chemical difference between the “iron-rich” and “sulfur-rich” portions of the site – a distinction made solely on the basis of surface accumulation of iron oxides and visible layering of sulfur phototroph pigment colors in different patches. An objective measure of iron concentrations across the site as well as a comprehensive culture-independent survey would have helped to assess whether this was in fact a workable site, or whether the gradient observed with the naked eye was superficial at best. In addition, the spread of measurements returned by the pH electrode essentially negated any ability to assess pH within the profiles.

Although there has been substantial growth in the photomethanotroph enrichments, it is at this point uncertain how that growth is being accomplished. It may be that enough sulfide was introduced with the inoculum to support sulfur phototrophic growth – although it would be expected to grow less well in transfers, not better as is being observed. Efforts to assess methanotrophy by monitoring methane consumption have been ineffective since the enrichment methodology was not designed with that in mind – at the high levels of methane introduced into the bottles’ headspace the procedural error of methane measurements by the available gas chromatograph is overwhelming – massive amounts of methane would have to be consumed extremely quickly to be unambiguously detectable. This issue can be addressed by setting up culture experiments across a gradient of methane concentrations, or by introducing less methane so that its consumption could be monitored. In addition, the use of 13C-labelled methane would permit stable-isotope probing to track what members of the enrichment community are consuming methane. Other components of the media, such as thiosulfate and sulfate, need to be controlled for as well and stoichiometric calculations performed to assess how much growth could be supported at their concentrations in the media.

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If, however, photomethanotrophs are in fact growing in the enrichment bottles the implications would be enormous. They might play an up until now completely unknown role in the global carbon cycle, and might have been extremely important during earlier periods of the earth’s history, when the atmosphere was more reducing than it is today. In addition, it would be extremely interesting to explore the question of how they are accomplishing the feat of activating and oxidizing methane in the absence of a strong oxidant such as oxygen.

Figures

Figure 1: The iron-oxide-rich patch on relatively high ground in the Greater Sippewissett Salt Marsh, with surface sand removed to show sulfide-rich sands and the thin accumulation of dense iron oxides at the surface. In contrast, the surrounding area was typified by green and purple sulfur bacterial layers. A metal spike was discovered buried underneath the patch, serving as a source of additional iron to the patch. Figure 2: Photos of sampling sites at Trunk River. A) Rusty pool 1. B) Rusty pool 2. C) Red-orange patch. D) Orange/pink/green veil.

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;

Figure 3: Cartoon showing crude model for the dynamics that might operate in an iron-dominated phototrophic mat. Oxygen is shown diffusing down from both the atmosphere and a layer of oxygenic photosynthesis on top of the mat. The oxygen is consumed via both abiotic and neutrophilic microbial iron oxidation in the underlying layers. Below the point at which all oxygen is absent but light is still penetrating at some wavelengths, photoferrotrophs are able to carry out iron oxidation. Below that, there might be anaerobic nitrate-reducing iron oxidation if nitrate levels are sufficiently high. The underlayers of the mat would be dominated by iron reducers growing heterotrophically and producing soluble ferrous iron which, if present in excess, would be expected to precipitate out all sulfide diffusing upwards from any zones of sulfate reduction below the mat.

Fe(II)

Light

O2

S2-

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Figure 4: Photograph of vigorously bubbling area at Trunk River, actively emitting methane gas without requiring any perturbation.

Figure 5: Photograph of area of School Street iron mat on the edge of the School Street parking lot sampled for enrichments.

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Figure 6: Photograph of electrode profiling sites in the Greater Sippewissett Salt Marsh – each profile is marked with the appropriate number, with 1 and 2 within the iron patch and 3 outside of it.

Pigment spectra for Incandescent Trunk River cultures

0

0.5

1

1.5

2

2.5

3

400 500 600 700 800 900 1000 1100

wavelength (nm)

abso

rban

ce

Trunk River rusty pool 1 Incandescent

Trunk River rusty pool 2 Incandescent

Figure 7: Graph showing the pigment spectra collected from the two Trunk River rusty pool primary enrichments incubated in front of an incandescent bulb. The strong peak at 758 nm and the less distinct one at 460 nm are diagnostic of bacteriochlorophyll c, common in green sulfur bacteria.

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Figure 8: Photographs of photoferrotroph bottles from three inocula with light treatments set next to each other. A) Trunk River rusty pool 1. B) School Street lot iron mat. C) Trunk River rusty pool 2.

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Figure 9: Photomicrograph at 1000X magnification of a mixed population of variably curved rods from the first transfer from the Trunk River rusty pool 1 incandescent primary enrichment. Scale bar is 10 µm.

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Figure 10: Pigment spectra of the Trunk River rusty pool 1 infra-red primary enrichment after a variety of procedures intended to improve spectral recovery. The top dark blue line (TRrd1 IR after sonication) was collected after sonication of a culture aliquot, and the magenta line (TRrd1 IR after sonication and centrifugation) was obtained after centrifugation of the sonicated aliquot and scanning of the supernatant. The yellow line (TRrd1 IR initial) was obtained by scanning the culture aliquot without modification, and the light blue line (TRrd1 IR pellet after oxalate) was obtained from the resuspended pellet after two oxalate treatments. The purple line (TRrd1 IR first supernatant) was obtained from the first supernatant from the oxalate wash.

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Figure 11: Maximum likelihood tree using a filter constructed using all full sequences on the tree except for the cyanobacterial outgroup. Each group’s phylogenetic affiliation is shown, known photoferrotrophs are designated in bold and known iron-reducers are designated by italics. For the groups that have a large number of clone sequences grouping closely together, the number of clones is noted to the right of a representative sequences or three (in the case of Shewanella). The School Street FL clones are highlighted in green, the Trunk River rusty pool 1 IR clones are highlighted in orange and the selected clones from the top of the Trunk River Nalgene are highlighted in purple.

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Figure 12: Phase contrast and epifluorescence photomicrographs of wet mounts from the Trunk River rusty pool 1 infra-red primary enrichment (top) and first transfer (bottom). Scale bar on top is 20 µm, scale bar on bottom is 10 µm.

Figure 13: Photomicrographs of colonies in anaerobic shake tubes (top) and of resuspended cells from each colony morphology (bottom). A) Typical colony morphology in School Street FL 10-6 tubes at 5X, about 400 µm in diameter. B) Cell aggregate from colony in A. C) Both small (designated by black arrows) and large colony morphologies in Trunk River rusty pool 1 IR 10-6 tubes at 5X, the large colony morphology is about 100 µm wide. D) Cell aggregates typical of the small colonies from C. E) Cell aggregates found in larger colony from C. All scale bars are 5 µm.

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Figure 14: SEM images taken of filtered cells and particulates from Trunk River rusty pool 1 IR primary enrichment (upper left and right, lower right) and first transfer (lower left).

Figure 15: Photo of inoculated iron-reducer culture on the left, with clearly dark precipitates, and an uninoculated control bottle on the right, with orange colored iron oxides. The culture was inoculated from a section of a 1-mL syringe core taken from a high ground, iron-rich area at the Greater Sippewissett Salt Marsh.

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Figure 16: From left to right: Trunk River FL primary enrichment 1; Trunk River FL primary enrichment 2; Trunk River FL first transfer 2; Trunk River IR primary enrichment 2; Trunk River IR primary enrichment 1.

Pigment spectrum of primary enrichment from Trunk River

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Figure 17: Pigment spectrum obtained from Trunk River FL primary photomethanotroph enrichment 2. The spectrum shows a major peak at 757 nm and a clear secondary peak at 460 nm, indicative of bacteriochlorophyll c, characteristic of green sulfur bacteria.

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Figure 18: Two different phase contrast photomicrographs of the same microscopic field under slightly different focus. On the left the two smaller classes of rods are visible (~2-3 µm and 4 µm) and on the right a smaller number of the large class of rods is visible (10 µm long). Scale bars are 10 µm.

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Figure 19: Graph of three time points of methane concentration from the Trunk River fluorescent enrichments and the uninoculated control bottle. Error bars are one standard deviation, the 11- and 25-hour time-points are the average of triplicate measurements.

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Figure 20: Microelectrode profiles of points from Fig.6. Error bars are one standard deviation around the mean, based on either five measurements per point (Profile 1) or three measurements per point (Profiles 2 and 3).

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Table 1: Base seawater media recipe (1 L) Combine 1 L 1X SW base with 5 mL 1 M NH4Cl, 10 mL 150 mM K-phosphate, 2.5 mL 1 M MOPS (6.8), 1 mL HCl-chelated trace elements, .25 mL 1 M NaSO4 in a Widdell vessel. Autoclave and cool under N2/CO2. Add .25 mL of 12-vitamin solution, .25 mL of Vitamin B-12 solution, 7.5 mL of 1 M NaHCO3, 12.5 mg 3-(3,4-dicyclorophenyl)-1,1-dimethylurea and 50 mL 4 mM Na2S2O3. Dispense media anaerobically into serum bottles, stopper and crimp.

Table 2: Glutaraldehyde/formaldehyde fixative solution, pH 7.0 Buffer, .2 M: 4.28 g Na-cacodylate 50 mL MilliQ water ca. 8 mL HCl, .2 M add Qwater to 100 mL and adjust to desired pH with the HCl Fixative: 50 mL buffer, .2 M 8 mL glutaraldehyde in water (final conc. will be 2%) 4.5 mL formaldehyde, 37% If necessary, adjust osmolarity with NaCl Add Qwater to 100 mL

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Results of photoferrotroph enrichments as of 7/27

Table 3: Report of culture color upon shaking and color of particulates, if any, within the bottle prior to shaking, of all primary enrichments. Date of inoculation is noted with each inoculum. References Croal, L., C. Johnson, et al. (2004). "Iron isotope fractionation by Fe(II)-oxidizing

photoautotrophic bacteria." Geochimica et Cosmochimica Acta 68(6): 1227-1242. Emerson, D. and J. Weiss (2004). "Bacterial iron oxidation in circumneutral freshwater

habitats: Findings from the field and the laboratory." Geomicrobiol J 21(6): 405-414.

Heising, S., L. Richter, et al. (1999). "Chlorobium ferrooxidans sp nov., a phototrophic green sulfur bacterium that oxidizes ferrous iron in coculture with a "Geospirillum" sp strain." ARCHIVES OF MICROBIOLOGY 172(2): 116-124.

Jung, D., L. Achenbach, et al. (2004). "A gas vesiculate planktonic strain of the purple non-sulfur bacterium Rhodoferax antarcticus isolated from Lake Fryxell, Dry Valleys, Antarctica." ARCHIVES OF MICROBIOLOGY 182(2-3): 236-243.

Lovley, D. R. (2000). Fe(III) and Mn(IV) Reduction. Environmental Microbe-Metal Interactions. D. R. Lovley. Washington, DC, ASM Press: 3-30.

Inoculum incandescent fluorescent Infra-red (940 nm) Trunk River rusty pool 1 inoculated 7/18

Green with small amount orange-brown particulates

Orange with orange and green particulates

Deep orange with gray-green and orange particulates

Trunk River rusty pool 2 inoculated 7/18

Green with some black particulates

Black with black particulates

Black with black particulates (no visible growth)

School Street lot iron mat inoculated 7/18

Orange-gray with green-gray and orange particulates

Orange with orange and pale green particulates

Yellow-beige with gray-green and orange particulates

Trunk River orange/pink/green veil inoculated 7/18

Black with black floating particulates

Black with black particulates

Black with black particulates

Trunk River red-orange patch inoculated 7/18

Gray and orange with orange-tinged gray-green particulates

Beige-orange with gray-green and orange particulates

Beige-orange with floating beige particulates

Top of Nalgene from Trunk River inoculated 7/18

Green with some filaments and orange-brown particulates

Beige-orange with orange and gray-green particulates

Creamy orange with pale particulates

Parker River Fe mat inoculated 7/18

Orange-beige with orange and beige particulates

Beige with orange and beige particulates

Light beige with green and gray floating particulates

Parker River rusty mud inoculated 7/23

Some dark particulates

No visible change after inoculation

No visible change after inoculation

Parker River lot pool inoculated 7/23

Ring of dark orange against bottom, tan

No visible change after inoculation

No visible change after inoculation

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Madigan, M., D. Jung, et al. (2000). "Rhodoferax antarcticus sp nov., a moderately psychrophilic purple nonsulfur bacterium isolated from an Antarctic microbial mat." ARCHIVES OF MICROBIOLOGY 173(4): 269-277.

Straub, K., F. Rainey, et al. (1999). "Rhodovulum iodosum sp. nov, and Rhodovulum robiginosum sp. nov., two new marine phototrophic ferrous-iron-oxidizing purple bacteria." Intl J Syst Bacteriol 49: 729-735.

Straub, K., W. Schonhuber, et al. (2004). "Diversity of ferrous iron-oxidizing, nitrate-reducing bacteria and their involvement in oxygen-independent iron cycling." Geomicrobiol J 21(6): 371-378.

Tor, J., K. Kashefi, et al. (2001). "Acetate oxidation coupled to Fe(III) reduction in hyperthermophilic microorganisms." Appl Environ Microbiol 67(3): 1363-1365.