Clonothrix fusca Roze 1896, a Filamentous, Sheathed, Methanotrophic γ-Proteobacterium

Sampling and growth media.Samples were taken monthly from the filters of an artesian well water pumping system in a farm in Salento (Apulia, Italy) over a period of 1 year (November 2005 to October 2006). The filters were found to be obstructed by a thick population of filamentous, sheathed bacteria with whitish-beige to yellowish-brown color.

Flavobacterium sp. strains AWR-3 and AWR-4 were isolated on ATCC media 1103 and 1503 (ATCC, Manassas, VA) after incubation at 28°C for 72 h. Bacillus sp. strain AWR-5 was isolated on ATCC medium 3 after incubation at 28°C for 24 h. The enrichment media for C. fusca were formulated using DSMZ medium 569 (http://www.dsmz.de/) that was diluted 1:5 (569-0.2X) as a base. The composition of 569-0.2X is 0.1 g liter⁻¹ KNO₃, 0.02 g liter⁻¹ MgSO₄·7H₂O, 0.002 g liter⁻¹ CaCl₂·2H₂O, 0.023 g liter⁻¹ Na₂HPO₄, 0.007 g liter⁻¹ NaH₂PO₄, 0.1 mg liter⁻¹ FeSO₄·7H₂O, 0.5 μg liter⁻¹ CuSO₄·5H₂O, 1.0 μg liter⁻¹ H₃BO₃, 1.0 μg liter⁻¹ MnSO₄·5H₂O, 7 μg liter⁻¹ ZnSO₄·7H₂O, 1 μg liter⁻¹ MoO₃. The medium was adjusted to pH 5.0. Samples were incubated under a methane (0.1 to 10%)/air atmosphere at 10°C in serum bottles with aluminum-paper covers placed in closed glass desiccators. Alternatively, 569-0.2X was supplemented with 0.5% methanol or 0.15% formaldehyde or 0.25% methanol and 0.075% formaldehyde and incubation was carried out in an air atmosphere at the same temperature. The optimal concentrations of methanol and formaldehyde in the media were determined in preliminary experiments testing a range between 0.05 and 1%. Selection in the presence of ampicillin (10 μg ml⁻¹) was performed by incubating samples in 596-0.2X containing 0.5% methanol in an air atmosphere at 10°C. Solid 596-0.2X media containing 0.5% methanol or 0.15% formaldehyde were prepared by adding 0.8% agarose.

Analytical procedures.The physical-chemical parameters (temperature; pH; specific conductance; hardness; alkalinity; ammonia, nitrite, chloride, calcium, magnesium, and iron concentrations; salinity; and oxygen consumption) of the artesian well water were assayed by standard procedures (http://www.epa.gov/waterscience/methods/).

To evaluate methane uptake, almost-pure cultures of C. fusca (about 100 mg, dry weight) grown in 100 ml of 596-0.2X containing 0.15% formaldehyde were centrifuged, washed with 596-0.2X without added carbon sources, gently resuspended in 100 ml of the same medium, and incubated at 10°C in flasks under a methane (0.45%)/air atmosphere (headspace of 150 ml) at 10°C for 0 to 72 h. Flasks without bacteria were used as a control. Methane consumption was measured in headspace samples by using a Hewlett-Packard 5890 gas chromatograph equipped with a flame ionization detector and a Chrompack PLOT fused-silica capillary column (length, 50 m; inside diameter, 0.53 mm; film thickness, 10 μm) coated with Al₂O₃/KCl, according to standard methods (43).

Assimilation of [¹⁴C]methane into cellular compounds.Almost-pure C. fusca cultures (about 100 mg, dry weight) were incubated in 100 ml of 569-0.2X in the presence of 100 μCi [¹⁴C]methane (1 mCi mmol⁻¹) (American Radiolabeled Chemicals, Inc.) in a methane (about 1.5%)/air atmosphere (headspace of 150 ml) at 10°C. Incubation was stopped after 0, 24, and 72 h. Cells were collected by centrifugation, washed with 0.9% NaCl, resuspended in phosphate buffer (pH 7.4) and then homogenized in a glass Potter-Elvehjem homogenizer with a Teflon pestle attached by rubber tubing to a constant-speed stirring motor (19). For protein analysis, the homogenate was subjected to two cycles of freezing in liquid nitrogen and thawing, incubated in the presence of 3% sodium dodecyl sulfate for 30 min at 37°C, and then centrifuged at 10,000 × g for 10 min at 4°C. Supernatant was recovered and proteins were purified as described previously (44). The protein concentration was assayed according to Lowry's procedure (22). Total lipids were extracted according to the modified Bligh and Dyer method (2) as reported previously (46). Labeled neutral or polar lipids, were separated by thin-layer chromatography and stained with iodine vapors as described previously (33). Regions of thin-layer chromatography corresponding to the different classes of lipids were scraped with a scalpel and analyzed for radioactivity. The radioactivity in each fraction was measured on a scintillation analyzer (Packard Tri-carb 1900TR) using a scintillation cocktail for aqueous and nonaqueous samples (PerkinElmer). As a control, heat-killed C. fusca cells were used.

DNA procedures.To achieve maximum extraction of DNA from the total microbial community obstructing the filter or from enriched microbial fractions, samples (10 mg, dry weight) were subjected to two cycles of freezing in liquid nitrogen and thawing at 55°C. Homogenates were suspended in 1.7 ml SET buffer (75 mM NaCl, 25 mM EDTA, 20 mM Tris-Cl [pH 7.5]), processed with the Potter-Elvehjem system, and incubated in the presence of 0.5 mg/ml proteinase K and 1% sodium dodecyl sulfate for 2 h at 55°C. Genomic DNA was then purified as previously described (41).

Primers and PCR conditions are listed in Table 1. All PCRs were performed as follows: initial denaturation at 95°C for 5 min followed by 30 cycles of denaturation at 95°C for 45 s, annealing for 30 s at the indicated temperature, and extension at 72°C for 1 to 2 min, depending on the amplicon length. They were carried out in a Perkin-Elmer Cetus DNA thermal cycler 2400.

To perform PCR single-strand conformational polymorphism (PCR-SSCP) experiments, PCR products were purified by using a High Pure PCR product purification kit (Boehringer Mannheim), denatured, and resolved on 10% polyacrylamide gel (acrylamide/N,N-methylenebisacrylamide, 49:1) in 0.8× TBE (72 mM Tris-borate, 1.6 mM EDTA) containing 5% glycerol (35). Bands identified after silver staining were excised with razorblades and single-strand DNAs were eluted from the gel by using a Qiaex II DNA purification kit (QIAGEN). The resulting DNAs were reamplified, cloned into pGEM-T Easy vector by using a TA cloning kit (Promega), and finally sequenced using primers M13F and M13R as a service of MWG Biotech.

For restriction fragment length polymorphism (RFLP) analysis, the PCR products generated by 16S-specific primers Com1-F and Com2-R were cloned into pGEM-T Easy vector to generate a library. Two hundred clones were screened by RFLP analysis with either EcoRI or HinfI on 6% polyacrylamide gel (35). The different clones were then grouped on the basis of their restriction profiles, and for each group, five representative clones were sequenced.

DNA similarity searches were carried out using BLAST at NCBI (http://www.ncbi.nlm.nih.gov/). Sequence alignments were performed with ClustalW at EBI (http://www.ebi.ac.uk/). Phylogenetic analyses were conducted using neighbor-joining with the MEGA 3.1 program (17). Bootstrap analysis (34) was used to estimate the reliability of phylogenetic reconstructions (1,000 replicates).

Light, fluorescence, and electron microscopy.For light and fluorescence microscopy, a Leica DMLB microscope and a Nikon Optiphot-2 fluorescence microscope were used, respectively. Fluorescence staining was performed with diamidino-2-phenylindole (DAPI) or the lipophilic dye FM4-64 (Molecular Probes, Invitrogen).

For transmission electron microscopy, samples were fixed with 2% glutaraldehyde and 1% formaldehyde in 0.04 M piperazine-N,N′-bis(2-ethanesulfonic acid) (PIPES) buffer at pH 7 for 2 h at room temperature. The samples were rinsed in 0.08 M PIPES buffer and twice in 0.08 M Na-cacodylate buffer and postfixed in 1% OsO₄ in 0.08 M Na-cacodylate buffer, pH 6.7, overnight at 4°C. Following dehydration in a step gradient of ethanol with three exchanges of anhydrous ethanol and one exchange of propylene oxide at 4°C, the samples were slowly infiltrated with Epon 912 resins at 4°C, transferred to polypropylene dishes and incubated at 70°C for 24 h. Thin sections were stained with 3% uranyl acetate in 50% methanol for 15 min and in Reynold's lead citrate for 10 min and, finally, observed with a Leo 912AB electron microscope.

Nucleotide sequence accession numbers.Nucleotide sequences determined in this study were deposited at GenBank under accession numbers DQ984190, partial 16S rRNA gene from C. fusca AW-b; DQ984191, partial 16S rRNA gene from Crenothrix sp. AW-a; DQ984192, partial pmoA gene from C. fusca AW-b; DQ984193, partial u-pmoA gene from Crenothrix sp. AW-a; DQ984194, partial pmoA gene from uncultured bacterium cAW-d; DQ984195, partial pmoA gene from uncultured bacterium cAW-e; DQ984196, partial pmoA gene from uncultured bacterium cAW-f; EF530351, partial mxaF gene from C. fusca AW-b; and EF530352, partial mxaF gene from uncultured bacterium cAW-h. The nucleotide sequences of partial or almost entire 16S rRNA, pmoA, and mxaF genes are reported in Tables S2 to S4 of the supplemental material.

Detection of C. polyspora and C. fusca by optical microscopy.The microbial community obstructing a filter of an artesian well water pumping system in a farm in Salento (Apulia, Italy) was studied over a period of 1 year (November 2005 to October 2006). The succession of two filamentous, sheathed microorganisms was observed by analyzing the microbial community by optical microscopy during this time. Tentative identification of these two microorganisms as C. polyspora and C. fusca was performed by macro- and micromorphological criteria according to Kolk (16) and Bergey's Manual (14), the physiological properties of these microorganisms being largely unknown.

C. polyspora exhibited its typical habitus, consisting of long, unbranched, sheathed clusters of cell filaments with brownish coloration at the cluster's basal area. The filaments were up to 1 cm long, about 2 to 4.0 μm wide, did not taper to the tip, often appeared attached to a mass substrate, and contained disk-shaped to cylindrical cells that reacted negatively to the Gram stain and exhibited variation in size (Fig. 1A). These cells were frequently divided by longitudinal or oblique septa leading to formation of microgonidia (small spherical cells, possibly spores) at the filament tips (Fig. 1A), a pattern typical of these kinds of bacteria. C. fusca formed filaments of about the same width and length as C. polyspora, attached or free, surrounded by a more or less distinct sheath that may be encrusted (possibly with iron or manganese oxides), giving a yellowish-brown color. At variance with those of C. polyspora, C. fusca filaments had the tendency to taper to the tip and contained gram-negative rod-shaped cells divided by cross-septation with considerable variation in size, up to 20 μm in length (Fig. 1A to C). Microgonidia were never observed, whereas massive “acropetal” production of macrogonidia (large spherical cells, possibly spores) was frequently detected at the tips of longer filaments as carefully described by Kolk (16) (Fig. 1C).

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FIG. 1.

Optical microscopy and culture-independent analysis of the microbial community obstructing the filter. (A to C) Gram stains of samples collected in December 2005, January 2006, and March 2006. The arrows indicate filamentous microorganisms resembling C. polyspora or C. fusca. The white bars represent 10 μm. (D to F) PCR-SSCP analysis of the microbial community. (D) PCR-SSCP profile of total sample (lanes 1 and 2) and filter-passing (lane 3) or filter-retained (lane 4) fractions with 16S rRNA gene-specific primers. The sample was collected in November 2005. (E and F) PCR-SSCP profile of filter-retained fractions with 16S rRNA gene-specific or pmoA-specific primers. The samples were collected in the period from November 2005 to March 2006 (lanes 1 to 5). In panels D to F, the arrows indicate specific bands (a, a′, b, b′, c, c′, d, d′, e, and e′) that were excised from the gel, eluted, and subjected to nucleotide sequence analysis. The lines indicate bands that were not characterized at the molecular level.

In our samples, C. polyspora was largely predominant during the first month (November 2005) (data not shown) and coexisted with C. fusca during the next three months (December 2005 to February 2006) (Fig. 1A and B), and then it almost disappeared, and C. fusca became prevalent (March to May 2006) (Fig. 1C). During the period from June to October 2006, the abundance of both microorganisms progressively decreased. In September to October 2006, luxuriant growth of C. polyspora causing severe obstruction of the filter was observed (data not shown). The available data of the standard water quality parameters suggested a possible correlation between the C. polyspora overgrowth and an increase of iron, calcium, magnesium, ammonia, and nitrite levels in the water, with a moderate decrease of pH values (see Table S1 in the supplemental material). At a depth of 54 m, where sampling was carried out, the water temperature, 18.5°C, was very close to the annual average temperature, slightly under 20°C.

Culture-independent analysis of the microbial community.The microbial community obstructing the filter was then analyzed by culture-independent methods. For this purpose, the DNA of the microorganisms was amplified using the bacterium-specific primers Com1-F and Com2-R (Table 1). These primers targeted a 409-nucleotide-long (in Escherichia coli) central region of the prokaryotic small-subunit rRNA gene (16S rRNA gene) and allowed high taxonomic resolution, often at the species level (18). The November 2005 sample was analyzed first. The SSCP profile of the amplified 16S rRNA gene pool revealed a relatively low degree of complexity of the microbial community, with a few species being predominant (Fig. 1D, lanes 1 and 2). The profile was further simplified when the starting sample was first passed through a 100-μm pore size fine-meshed sieve to retain the filamentous mass (Fig. 1D, lane 4) and eliminate microorganisms that were not arranged in long filaments (Fig. 1D, lane 3). The SSCP profile of the enriched sample was dominated by the presence of two major bands (a and a′) (Fig. 1D, lane 4) that were excised from the gel, eluted, cloned, and subjected to nucleotide sequence analysis. To avoid any bias due to the reamplification step, only PCR products showing the same SSCP profile as the original bands were further processed. The sequence corresponding to bands a/a′ exhibited the highest level of similarity (97.69% identity) with sequences published by Stoecker and coworkers (39) and attributed to C. polyspora Cohn (Table 2), suggesting that the filamentous microorganism in our sample might belong to the same genus as Cohn's Crenothrix.

Due to the inability to recover the other SSCP bands efficiently, the identities of the other microorganisms that did not form long filaments (Fig. 1D, lane 3) were determined by a different approach. The PCR products generated by the bacterium-specific primers targeting the total DNA from the filterable fraction were cloned to construct a library. Two hundred clones gave different RFLP profiles, and five clones representative of each profile were subjected to sequence analysis (Table 2). This approach led us to identify two microorganisms, AWR-3 and AWR-4, belonging to the genus Flavobacterium, closely allied to the species F. succinicans and F. columnare, respectively; an unknown microorganism, AWR-1, grouping with Niastella jeongjuensis of the Bacteroidetes/Chlorobi group; a β-proteobacterium, AWR-2, close to Methylophilus methylotrophus, a methylotrophic β-proteobacterium; and a Bacillus sp. strain isolate, AWR-5, close to B. cereus. Flavobacterium sp. strains AWR-3 and AWR-4 and Bacillus sp. strain AWR-5 were obtained in pure cultures.

The other samples, from December 2005 to March 2006, were then analyzed by PCR-SSCP. To recover the filamentous mass and to eliminate microorganisms that were not arranged in long filaments, the samples were filtered as above. The SSCP profiles of the amplified 16S rRNA gene pool showed a succession of the microbial community (Fig. 1E). During the last month (March 2006), the fingerprinting corresponding to Crenothrix (Fig. 1D, lane 4, and Fig. 1E, lane 1) disappeared, and two major bands (b and b′) became predominant (Fig. 1E, lane 5). These bands were subjected to nucleotide sequence analysis as described above. The results of this analysis suggested that the filamentous microorganism that was prevalent in this sample (March 2006), tentatively identified as C. fusca on the basis of its micromorphology (Fig. 1A to C), was a member of the type I methanotrophic γ-proteobacteria distinct from Crenothrix and from any other known member of this group of prokaryotes. Among the cultivated species, the psychrophilic methanotrophs Methylobacter tundripaludum SV96 and Methylobacter psychrophilus Z-0021 (3) exhibited the highest similarities, with sequence identities of 94.82% and 94.23%, respectively (Table 2). The levels of similarity were too low to assign our microorganism to any known genus of the methanotrophs.

To substantiate with further molecular data the idea that Roze's Clonothrix might be a type I methanotroph, we looked for the presence of the particulate methane mono-oxygenase (pMMO) α subunit-encoding gene pmoA in the filter-retained fraction of our samples by using PCR-SSCP (Fig. 1F). MMO is responsible for the initial transformation of methane into methanol. MMO exists in two forms, a cytoplasmic, soluble form and the membrane-bound form pMMO. Soluble MMO is found in only a subset of methanotrophs, whereas pMMO is a marker of most methanotrophs (7, 20). Using the primer pair pmof1/pmor, designed to amplify evolutionarily conserved regions of pmoA (4), we observed specific SSCP bands in all samples, from November 2005 to March 2006 (Fig. 1F, lanes 1 to 5). As seen with the 16S rRNA genes, the SSCP profile of the amplified pmoA sequences demonstrated a succession, with four major bands predominating in the November 2005 sample (d, d′, e, and e′) and two different bands (c and c′) predominating in the March 2006 sample. We deduced that these two bands might correspond to the two filaments encoded by the putative C. fusca pmoA gene, while the four bands in the sample of November 2005 might correspond to pmoA sequences of contaminating methanotrophs. Bands c and c′ were excised from the gel and subjected to nucleotide sequence analysis. The results confirmed that c and c′ represented the two cDNA strands of an unknown pmoA gene. Among characterized pmoA genes, that of Methylosarcina fibrata exhibited the highest similarity (91.59% identity at the level of deduced amino acid sequence). The other bands were similarly characterized. d and d′ and e and e′ corresponded, respectively, to the two strands of two pmoA genes from bacteria phylogenetically close to Methylomicrobium buryatense 5B (94.44% identity at the level of deduced amino acid sequence) and to Methylomonas methanica S1 (96.30% identity at the level of deduced amino acid sequence).

Enrichment media for C. fusca.The results of the 16S rRNA gene and pmoA sequence analyses prompted us to hypothesize that Roze's Clonothrix might be a methane oxidizer physiologically close to the psychrophilic type I methanotrophs. On the basis of this hypothesis, we attempted to enrich the starting sample (January 2006) in C. fusca by a three-step procedure. The three steps were (i) enrichment of the starting sample in a diluted medium for type I methanotrophs (596-0.2X) without added carbon sources, under a methane (10%)/air atmosphere at 10°C for 2 weeks in the dark; (ii) further selection in the presence of ampicillin in 596-0.2X medium containing methanol, in an air atmosphere for 24 h at the same temperature; and (iii) incubation in 596-0.2X containing methanol, formaldehyde, or both, in an air atmosphere at 10°C for 2 weeks in the dark. Methanol and, in some cases, formaldehyde, formate, and methylamines may be utilized as C₁ carbon (and energy) sources by methanotrophic microorganisms (3). However, as formaldehyde is a highly reactive molecule and a powerful poison for living systems (9), the optimal concentrations of formaldehyde and methanol (that generates formaldehyde) in the media were determined in preliminary experiments testing a range between 0.05 and 1%. The ampicillin treatment was effective in eliminating the rapid-growing nonfilamentous microorganisms in the starting sample.

In the media containing methanol, formaldehyde, or both, C. fusca was able to grow at extremely low rates, as deduced from the elongation and thickening of single inoculated filaments and from the increase in dry weight. In the medium containing formaldehyde, growth rates of about 0.01 h⁻¹ were estimated. In the media with either methanol or methanol and formaldehyde, growth rates could not be evaluated by dry weight, because rapid-growing bacteria had a selective advantage and outgrew C. fusca. In contrast, an almost pure culture of C. fusca could be obtained in 596-0.2X containing 0.15% formaldehyde, as determined by Gram staining (see Fig. S1A to D in the supplemental material), plating on complex organic media (nutrient agar) to exclude the presence of heterotrophic contaminants, and PCR-SSCP profiles (see Fig. S2 in the supplemental material). In the medium with formaldehyde, C. fusca filaments nucleating at the base and sporulating at the tip (see Fig. S1A to D in the supplemental material) were observed, in addition to spores and younger, short, thinner filaments often lacking a distinct sheath and exhibiting false-branching (data not shown), such as were carefully described by Kolk (16). The filaments were also observed by fluorescence microscopy after being stained with the membrane-labeling dye FM4-64 (see Fig. S1E in the supplemental material) or with DAPI (see Fig. S1F in the supplemental material). The DAPI staining results were suggestive of the presence of multiple nucleoids within each larger cell.

However, C. fusca did not grow on 596-0.2X containing agar with methane, methanol, or formaldehyde. The inability to grow on agar is a common feature of several methanotrophs (3). C. fusca also failed to grow in the medium 596-0.2X containing 0.5% glucose. This result was consistent with the taxonomic position of this microorganism. Indeed, all type I methanotrophs that have been isolated to date are able to utilize only reduced C₁ substrates (11, 13). In all tested media, Crenothrix was progressively lost.

Methane uptake and assimilation of [¹⁴C]methane into cellular compounds.To obtain direct evidence of the ability of C. fusca to utilize methane, uptake and assimilation studies were carried out, utilizing cultures enriched in formaldehyde medium. After removing formaldehyde, methane uptake was evaluated in 596-0.2X medium under a methane (0.45%)/air atmosphere at 10°C. The measurements, made at various intervals (0, 24, 48, and 72 h) with triplicate samples, demonstrated that methane consumption occurred as a function of time (Fig. 2A). A decrease of about 27% in methane concentration was observed during the first 24 h, when methane uptake was maximal (about 5.4 μmol g⁻¹ [dry weight] h⁻¹).

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FIG. 2.

Methane uptake by C. fusca cultures and assimilation of [¹⁴C]methane or [¹⁴C]methanol into cellular compounds. (A) The methane uptake was estimated in triplicate samples. (B) ¹⁴C assimilation from [¹⁴C]methane into indicated lipids. The values are expressed as nmol g⁻¹ (dry weight). (C) ¹⁴C assimilation from [¹⁴C]methane into proteins. The values are expressed as nmol g⁻¹ (protein). In all graphs, the values represent means with standard deviations (bars).

The assimilation of [¹⁴C]methane into cellular compounds was then analyzed. In these experiments, C. fusca cultures were incubated in 596-0.2X under a [¹⁴C]methane (about 1.5%)/air atmosphere. ¹⁴C distribution into total lipids, phospholipids, neutral lipids (Fig. 2B), or proteins (Fig. 2C) was evaluated in triplicate samples at different incubation times (0, 24, and 72 h). The results demonstrated that incorporation of ¹⁴C in all macromolecules occurred as a function of time, and it was maximal during the first 24 h, consistent with the methane uptake data (Fig. 2A).

Ultrastructural analysis of C. fusca Roze.Transmission electron microscopy analysis of C. fusca from enrichment medium containing methane demonstrated multilayered sheaths surrounding cell filaments consisting of both granular and fibrillar material (Fig. 3A to D). The architecture of the cell wall was typical of that of gram-negative bacteria (Fig. 3B to D). The vegetative cells (Fig. 3A) exhibited a bacillar shape and were, in our microphotographs, about 1.0 μm wide and 2.5 μm long (without sheaths), although larger elements could be observed. They contained an elaborate membrane system in the cytoplasm, in the form of lamellar stacks arranged perpendicularly to the main cell axis and lining the periphery of the cytoplasm. At higher magnifications, the membrane system consisted of closely packed, swollen sacks (Fig. 3B and C). This membrane system resembles that of type I methanotrophs, and seems to be different from that of C. polyspora, which is characterized by flattened sacks (39, 42). Electron-dense storage granules resembling polyphosphate granules and large electron-transparent organelles delimited by membranes and resembling gas vesicles were also found (Fig. 3C). The role of these organelles is obscure; their close association with the lamellar stacks arranged perpendicularly to the membrane is noteworthy (Fig. 3C). Indeed, gas vesicles were also observed in M. psychrophilus, the psychrophilic methanotroph closely allied to C. fusca (27). Occasionally, slightly smaller cells arranged in unsheathed filaments with the above-mentioned features were observed (Fig. 3D to F). This finding may correspond to contaminant methanotrophs (for instance, to the uncultured bacterium cAW-f) (Fig. 4B and C, and see Fig. S2 in the supplemental material) or to younger, short, thinner filaments of C. fusca lacking a distinct sheath.

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FIG. 3.

Transmission electron microscopy of C. fusca filaments. (A to C) Longitudinal sections of C. fusca filaments at different magnifications. The bars represent 1 μm in panel A and 0.5 μm in panels B and C. Note in panel B (i) the thick, multilayered sheath; (ii) the gram-negative type of cell wall; and (iii) the membrane system in the cytoplasm, consisting of closely packed, swollen sacks arranged perpendicularly to the main axis of the cell. In the filament shown in panel C, the lamellar stacks are arranged perpendicularly to the membrane of a large, electron-transparent organelle associated with electron-dense granules. (D) Cross section of C. fusca filaments. The bar represents 0.5 μm. Note the circular shape and thickness of the sheath. (E to G) Longitudinal sections at different magnifications of unsheathed microorganisms arranged in chains. The bars represent 2 μm in panel E, 0.5 μm in panel F, and 0.2 μm in panel G.

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FIG. 4.

Taxonomic position of C. fusca with respect to C. polyspora and other methanotrophic α- and γ-proteobacteria. (A) Phylogenetic analysis of 16S rRNA gene sequences. Bootstrap values ≥70 are reported at the branch points. (B) Phylogenetic analysis based on amino acid sequences deduced from pmoA, u-pmoA, and amoA. (C) Phylogenetic analysis based on amino acid sequences deduced from mxaF. In panels B and C, bootstrap values ≥50 are reported at the branch points. In all panels, the arrows indicate the positions of isolates from the microbial community obstructing the filter.

Phylogeny of C. fusca Roze.Using C. fusca cultures in formaldehyde medium as a source of DNA, almost the entire 16S rRNA-encoding gene was amplified by using the proteobacteria-specific primers 16SEB20-42-F and 16SEB1488-R (Table 1) and was cloned into a multicopy plasmid. Three of these clones were subjected to nucleotide sequencing that gave identical results. Phylogenetic analysis demonstrated the taxonomic position of our isolate in a subgroup of type I methanotrophs including C. polyspora and the psychrophilic methanotrophs M. tundripaludum and M. psychrophilus (Fig. 4A). Almost the entire 16S rRNA gene sequence of the Crenothrix sp. which was predominant during the first month (November 2005) was then determined by using the sample fraction enriched in filaments as a source of DNA (Fig. 1D, lane 4). The results confirmed the taxonomic position of our Crenothrix in a clade including the other C. polyspora Cohn clone sequences from Stoecker and coworkers' study (39) (Fig. 4A). However, if we consider the phylogenetic distances, we may hypothesize that our and Stoecker and coworkers' microorganisms may belong to the same genus but to different species.

The DNA from C. fusca cultures in formaldehyde medium was then used to amplify pmoA by using the above-mentioned primer pair (4). We could observe a specific PCR product of the expected size that was subjected to direct DNA sequencing or to DNA cloning and sequencing. Both approaches gave identical sequences, corresponding to those of the c and c′ PCR-SSCP bands (Fig. 1F). Phylogenetic analysis confirmed the presence in C. fusca of a pmoA gene typical of type I methanotrophs (Fig. 4B). The phylogenetic analysis also included the pmoA sequences from bands d/d′ (uncultured bacterium cAW-d) and e/e′ (uncultured bacterium cAW-e) of Fig. 1F.

Stoecker and coworkers' study (39) described the presence in C. polyspora Cohn-enriched material of a very unusual pMMO gene (u-pmoA). Thus, we looked for the presence of this gene in C. fusca cultures by PCR using a u-pmoA-specific primer pair. However, we did not obtain any specific PCR product. In contrast, we were able to amplify u-pmoA-specific DNA sequences when the starting sample (January 2006) was used as the DNA template. u-pmoA-specific PCR products were cloned and subjected to nucleotide sequencing. This analysis confirmed the presence of the u-pmoA gene in the unenriched sample, exhibiting high similarity (95.23% identity at the level of the nucleotide sequence; 97.14% identity at the level of the deduced amino acid sequence) with that of C. polyspora (Fig. 4B).

The presence of mxaF, a marker gene of both methanotrophs and methylotrophs, encoding the α subunit of the methanol dehydrogenase (24), was also investigated in C. fusca cultures. The primer pair mxaFf and mxaFr (Table 1) amplified a specific PCR product that was subjected to sequencing. Phylogenetic analysis demonstrated that the C. fusca methanol dehydrogenase was typical of that of type I methanotrophs (Fig. 4C).