INTRODUCTION

Top Abstract Introduction Materials and Methods Results Discussion References

Most known bacterial polycyclic aromatic hydrocarbon (PAH) degraders have been isolated from heavily contaminated terrestrial environments (1, 4, 11, 33, 36). Various strains of Pseudomonas, Comamonas, Acinetobacter, and Sphingomonas have been obtained from such sources. However, PAH pollution is not constrained to sites impacted by fuel spills or locally high levels of fossil fuel use. PAHs occur ubiquitously, even in apparently pristine environments with relatively little fossil fuel use. Although there has been relatively little emphasis on such systems, the impact of long-term, chronic PAH exposure may be determined by assessing the presence and activity of nutritionally versatile bacteria that are also capable of degrading PAHs.

As is the case for terrestrial systems, marine environments include both heavily impacted sites and far more numerous sites exposed to low levels of PAH input. A variety of PAH degraders (mostly Pseudomonas and Vibrio spp.) have been isolated from polluted systems, including Boston Harbor, the Chesapeake Bay, and hydrocarbon-contaminated salt marshes (for examples, see references 6, 10, and 40). Several novel marine PAH degraders, e.g., Cycloclasticus spp. and Neptunomonas naphthovorans, have also been isolated from contaminated sediments (12, 15, 19). Some of these isolates occur in presumably pristine systems (14). The potentially ubiquitous distribution of marine PAH degraders suggests that the capacity for PAH degradation in polluted systems depends on the diversity and characteristics of naturally occurring populations and their responses to environmental conditions, rather than on the introduction of new taxa or selective modification of existing ones.

Although efficient and rapid PAH degradation generally depends on molecular oxygen availability (for examples, see references 8, 9, and 29), oxygen is often limited to the top several millimeters of surface sediment in marine systems (for an example, see reference 31), which severely constrains the potential for PAH degradation. However, the activities of benthic macrofauna, which physically mix sediments and introduce oxygen to subsurface sediments by burrow ventilation (2), may create unique sites for enhanced PAH degradation. This proposal is supported by patterns of PAH degradation in slurries of burrow sediment from different macrofaunas. When incubated under oxic conditions, these slurries exhibited enhanced PAH degradation potentials, relative to those of slurries of nonburrow sediment, for a number of compounds, including naphthalene, phenanthrene, acenaphthene, and dibenzothiophene (9). Similarly, the rhizosphere of marine plants enhances oxygen availability in sediments (20) and supports populations of aerobic PAH degraders (10).

Accordingly, we report here results of efforts to enrich, isolate, and characterize PAH degraders from macrofaunal burrow sediments. We describe the isolation of two aerobic, obligately marine PAH-degrading bacteria from burrows of a polychaete (Nereis virens) and a mollusc (Mya arenaria) in the intertidal mudflat of Lowes Cove, Maine. Both strains use naphthalene and phenanthrene aerobically as their sole carbon sources. 16S ribosomal DNA (rDNA) phylogenetic analysis and phenotypic characterization suggest that strain LC8 belongs to a new genus and species within the Sphinogmonadaceae that is provisionally designated Lutibacterium anuloederans LC8. Likewise, strain M4-6 is a new Cycloclasticus species, provisionally designated Cycloclasticus spirillensus M4-6. However, only LC8 appeared to maintain a significant PAH degradation potential after inoculation into contaminated and uncontaminated sediment slurries incubated with or without added phenanthrene.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results Discussion References

Sampling and slurry preparation. Animal burrow sediments were collected from the intertidal zone of Lowes Cove during the summer of 1999 with a sterile spatula during low tide. This site and the burrow sediment collection method have been described in detail previously (3, 17, 18, 23, 24). Based on PAH assays, the site is not known to be contaminated (PAH concentrations, <50 ng g [dry weight] of sediment¹). Samples collected were transferred to the laboratory within 1 h and processed immediately. Surface sediment (the top 1 to 3 mm) and bulk sediment (from a depth of 10 to 15 cm) were also collected and processed similarly. M. arenaria burrow sediments were collected after being exposed with a clam fork. Only the oxidized, light-brown layer (1 to 2 mm thick) was scraped off with a spatula. Burrow sediments of N. virens were collected similarly. In all cases, burrow identification was based on the presence of animals and distinctive burrow morphologies. Burrow sediment slurries were prepared with artificial seawater (37). Unless stated otherwise, 10% (wt/vol) slurries were used in the following experiments. For abiological controls, bulk sediments were autoclaved at 121°C and then cooled; this cycle was repeated three times over a period of 72 h.

Enrichment and isolation of phenanthrene-degrading bacteria. Two hundred fifty milliliters of 10% sediment slurry was prepared in a 1-liter flask. A 2% phenanthrene stock solution was prepared in acetone and added to the slurry at an initial concentration of 10 ppm. Additional phenanthrene was added when the concentration of phenanthrene remaining in the slurry dipped below 0.5 ppm. The phenanthrene concentration in the slurry was slowly increased to about 100 ppm over a period of 2 weeks and was maintained at this level for the duration of the enrichment. After 4 weeks of enrichment, subsamples from the flasks were diluted serially with mineral salts medium ONR7a (12). Spread plates were prepared with ONR7a that had been solidified with 1.5% agar and sprayed with a 2% phenanthrene solution in acetone. The plates were incubated at room temperature (about 23°C) for up to 3 weeks. Colonies showing a clearing zone on the crystalline phenanthrene layer were picked and streaked on new minimal agar plates. A rapidly growing, visually distinct colony and a separate, morphologically unique isolate were selected for further analysis and purified by repeated plating.

Isolate characterization. Cell motility was examined with motility agar and by phase-contrast microscopy at ×1,000 magnification. Routine microbiological tests, including those for Gram stain reaction, nitrate reduction, oxidase, catalase, gelatinase, lipase, phosphatase, and glucose fermentation, were performed according to standard methods (35). Sodium ion requirements were tested by replacing sodium salts in the medium with the corresponding potassium salts. Salinities were established by adjusting inorganic salt concentrations to final values of 3.5, 10, 35, and 70 ppt. Isolate growth was tested at temperatures between 4 and 42°C. Poly--hydroxybutyrate (PHB) inclusions were visualized with Sudan black. For bacteriochlorophyll detection, strain LC8 was grown to late log phase on pyruvate. The culture was centrifuged, washed twice with ONR7a, and collected by centrifugation at about 6,500 × g. The cell pellet was extracted by vortexing with 5 ml of an acetone-dimethyl sulfoxide mixture (9:1) for 2 min. The absorbance at 300 to 800 nm of the organic layer was measured with a Beckman model DU-600 spectrophotometer. The following compounds were tested at 15 to 20 mM as the sole carbon sources for growth of the isolates in ONR7a or ONR7a plus 0.05% yeast extract (ONR7y): arabinose, fructose, galactose, glucose, lactose, mannose, ribose, sucrose, alanine, aspartate, glycine, phenylalanine, proline, serine, valine, sodium citrate, sodium fumarate, gluconic acid, glucuronic acid, sodium glycolate, malic acid, sodium malonate, sodium acetate, sodium propionate, sodium pyruvate, sodium succinate, sodium tartarate, betaine, ethanol, glycerol, mannitol, and hexadecane. The following aromatics were also tested as the sole carbon sources for growth: phthalate, salicylate, biphenyl, naphthalene, 1,4-dimethyl naphthalene, 1,8-dimethyl naphthalene, acenaphthene, fluorene, anthracene, phenanthrene, dibenzothiophene, benz[a]anthracene, pyrene, chrysene, and fluoranthene. Isolate growth was scored as positive or negative by comparing the turbidity of the medium after 72 h of incubation at room temperature with shaking (120 rpm) to that of a control receiving no carbon source. Whole-cell fatty acid analyses were performed by Microbial ID, Inc. (Newark, Del.).

16S rDNA sequencing and phylogenetic analysis. Total genomic DNA was extracted from 3 ml of late-log-phase phenanthrene-grown cells by a standard sodium dodecyl sulfate-proteinase K lysis procedure (32). The 16S rDNA gene was amplified by PCR with published primers (27f and 1492r) (41) in a 100-µl volume in accordance with standard protocols (28, 32). PCR products were separated in 0.8% agarose gels. DNA bands (1.5 kb) from the gel were excised and purified by using a Qiagen QIAquick gel extraction kit according to the manufacturer's instructions. Both strands of the purified DNA were then sequenced by an automatic DNA sequencer (ABI) with published primers (the PCR primers mentioned above and internal primers 926f, 907r, 357f, and 519) (41).

Phenanthrene uptake potential in slurries inoculated with strain LC8 or strain M4-6. Slurries of Lowes Cove sediment were prepared as described above. Additional slurries were prepared with sediment from Little Mystic Channel (LMC), a heavily contaminated site in Boston Harbor that contains substantial concentrations of a variety of PAHs (34, 39). LMC sediment was collected at low tide with acrylic cores (7 cm [diameter] by 32 cm) that were transported to the Darling Marine Center within 8 h of collection. The cores were stored under flowing seawater until use (about 7 days), at which time the top 15-cm section was removed and diluted with artificial seawater (1:10, vol/vol), as for Lowes Cove slurries. Abiological controls for Lowes Cove and LMC sediments were prepared from slurries autoclaved three times at intervals over 72 h.

3

1

3

PAH and PAH metabolite analysis. Loss of PAHs from culture media and slurries was measured by gas chromatography-mass selective detection after organic solvent extraction. In brief, slurries or cultures were acidified to a pH of <2.0 with 5 N HCl and were mixed with one-half volume of n-hexane in a Teflon-lined screw-cap tube. The tubes were shaken for 2 min, and aqueous and organic phases were separated by centrifugation at ~2,400 × g. The organic layer was concentrated under a stream of nitrogen, and PAHs were reconstituted in n-hexane. Samples were assayed with a Hewlett-Packard model 6890 gas chromatograph together with a model 5972A mass selective detector equipped with an HP-5MS capillary column (30 m by 0.25 mm by 0.25 µm [film thickness]) and helium as a carrier gas.

Nucleotide sequence accession number. The 16S rDNA sequence of strains LC8 and M4-6 have been deposited in the GenBank database under the accession numbers AY026916 and AY026915, respectively.

	RESULTS

Top Abstract Introduction Materials and Methods Results Discussion References

Enrichment culture and strain isolation. Macrofaunal burrow sediment slurries degraded substantial amounts of phenanthrene after a 2-week incubation (Fig. 1). Phenanthrene degradation rates of 15 µg ml¹ day¹ after 2 weeks were typical for an enrichment flask with N. virens sediment. Degradation was preceded by a lag period of about 8 days. Comparable results were obtained for slurries of M. arenaria burrow sediments (data not shown).

View larger version (22K): [in this window] [in a new window]   FIG. 1.   Time course of phenanthrene loss in N. virens burrow sediment slurries () and bulk surface sediment slurries (). Arrows indicate the addition of phenanthrene. All data are means of results for triplicate determinations; error bars represent ±1 standard error.

Isolate characteristics. Both strain LC8 and strain M4-6 were aerobic and obligately marine, requiring sodium chloride for growth. Both isolates tolerated salinities of up to 70 ppt. Phenotypic assays established that strain LC8 was a gram-negative, catalase-positive, oxidase-positive, nonmotile rod of ~0.5 by 1.5 to 2.0 µm (Fig. 2A) that formed pale-yellow colonies on minimal agar plates with phenanthrene as a carbon source. It also reduced nitrate and exhibited phosphatase activity but did not hydrolyze gelatin, show lipase activity, ferment glucose, or accumulate PHB. Strain LC8 grew between 15 and 30°C. Acetone-dimethyl sulfoxide (9:1) extracts of strain LC8 revealed none of the absorption maxima between 600 and 800 nm that are characteristic of bacteriochlorophylls. Strain LC8 used the following compounds as its sole carbon sources in ONR7y: arabinose, fructose, galactose, glucose, mannose, ribose, sucrose, citrate, fumurate, glucuronic acid, gluconic acid, acetate, pyruvate, tartarate, glycerol, mannitol, and hexadecane. Strain LC8 also utilized naphthalene and phenanthrene as its sole carbon and energy sources, transformed dibenzothiophene to dibenzothiophene sulfoxide, and cometabolized pyrene in the presence of phenanthrene (data not shown). Strain LC8 produced 1-hydroxynaphthoic acid transiently from phenanthrene, produced dibenzothiophene sulfoxide from dibenzothiophene, and produced pyrene-diol and pyrene dicarboxylic acid as pyrene metabolites (data not shown).

View larger version (133K): [in this window] [in a new window]   FIG. 2.   Transmission electron micrographs of strain LC8 (A) and strain M4-6 (B) at ×15,500 magnification. Scale bars indicate 1 µm.

18:17c

16:17c

17:17c

16:17c

Phylogenetic analysis. PCR amplification of 16S rDNA from strains LC8 and M4-6 resulted in sequences of 1,452 and 1,496 bp, respectively. Based on a BLASTN search of GenBank, the closest matches to strain LC8 were Erythrobacter spp. (nucleotide identity, 96.7%), Erythromicrobium spp. (nucleotide identity, 94.8%), Porphyrobacter spp. (nucleotide identity, 94.9%), and Sphingomonas spp. (nucleotide identity, 92.0%). These and other related sequences were used to construct a consensus tree based on maximum likelihood phylogenetic analysis (100 bootstrap samples). The results indicated that strain LC8 was phylogenetically closest to Erythrobacter with an Erythrobacter-LC8 branch (supported at a level of 95% by bootstrap analysis) that is distinct from those of other sphingomonads (Fig. 3). Porphyrobacter and Erythromicrobium formed another clade distinct from the Erythrobacter-LC8 branch and those of the rest of the sphingomonads. The trees generated from the maximum parsimony and neighbor-joining methods shared the overall topography of the maximum likelihood tree and presented the same relationship among strain LC8, Erythrobacter spp., and the sphingomonads (data not shown).

View larger version (17K): [in this window] [in a new window]   FIG. 3.   Phylogenetic tree based on maximum likelihood analysis of 16S rDNA gene sequences of strain LC8 and selected representatives of the Sphingomonadaceae and -Proteobacteria. Numbers indicate the present bootstrap support for 100 resamplings of the data set.

View larger version (12K): [in this window] [in a new window]   FIG. 4.   Phylogenetic tree based on maximum likelihood analysis of 16S rDNA gene sequences of strain M4-6 and selected representatives of Cycloclasticus and the -Proteobacteria. Numbers indicate the percentages of bootstrap support for 100 resamplings of the data set.

Phenanthrene uptake in slurries. Phenanthrene uptake by slurry samples amended with strain LC8 was readily detected by using time course assays with added phenanthrene (Fig. 5). Uptake rates in autoclaved slurries and uninoculated slurries were negligible compared to rates for inoculated slurries incubated continuously with phenanthrene (Fig. 6). Essentially equivalent results were obtained for uninoculated and autoclaved slurries from Lowes Cove and LMC (data not shown). However, uptake rates for inoculated slurries varied over a 35-day interval depending on the sediment source and treatment (Fig. 6A and 7).

View larger version (12K): [in this window] [in a new window]   FIG. 5.   Time course of phenanthrene concentrations in slurries of autoclaved Lowes Cove sediment (), slurries containing no added strain LC8 (), and slurries incubated routinely with phenanthrene and inoculated with strain LC8 (). All data are means of results for triplicate determinations; error bars represent ±1 standard error.

View larger version (23K): [in this window] [in a new window]   FIG. 6.   (A) Potential phenanthrene uptake rates for slurries from Lowes Cove (circles) and LMC (squares) inoculated with strain LC8 and incubated with (open symbols) or without (closed symbols) routine phenanthrene additions. All data are means of results for triplicate determinations; error bars represent ±1 standard error. (B) Potential phenanthrene uptake rates for uninoculated slurries from Lowes Cove incubated with () or without () routine phenanthrene additions and for autoclaved slurries (). All data are means of results for triplicate determinations; error bars represent ±1 standard error.

View larger version (16K): [in this window] [in a new window]   FIG. 7.   Potential phenanthrene uptake rates for slurries from Lowes Cove (circles) and LMC (squares) inoculated with strain M4-6 and incubated with (open symbols) or without (closed symbols) routine phenanthrene additions. All data are means of results for triplicate determinations; error bars represent ±1 standard error.

View larger version (15K): [in this window] [in a new window]   FIG. 8.   Residual phenanthrene remaining in LMC sediment slurries incubated without routine phenanthrene additions and inoculated with strain LC8 () or strain M4-6 (). Results for autoclaved control slurries () are shown, as well. All data are means of results for triplicate determinations; error bars represent ±1 standard error.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

Macrofaunal burrows enhance a number of microbial activities. For example, elevated levels of nitrification (26, 27) and sulfate reduction (18) have been documented for macrofaunal burrow sediments. Burrows also support higher levels of microbial biomass than do bulk sediments (17, 23). In addition, previous studies have shown that N. virens and M. arenaria burrow sediments harbor PAH-degrading bacteria and that degradation rates for added PAHs are greater in burrow sediments than in nonburrow sediments (9). Since Lowes Cove has not been significantly contaminated with PAHs, it appears that burrows provide an environment suitable for maintenance of PAH-degrading organisms of certain taxa. Accordingly, burrows in uncontaminated systems may provide a source of organisms that can respond to chronic or acute PAH pollution and perhaps serve as a resource for neighboring heavily contaminated systems.

Two distinct PAH degraders were isolated from Lowes Cove burrow sediments. Morphologically, phylogenetically, and phenotypically, strain LC8 appears to be a sphingomonad (Fig. 2 and 3). Many Sphingomonas spp. have been reported to degrade a variety of PAHs, which is not surprising, since most of these isolates have been obtained from contaminated terrestrial systems (for examples, see references 1, 5, 11, 22, and 36); a marine Sphingomonas sp. capable of degrading PAH has likewise been isolated from a contaminated site (16).

Nonetheless, 16S rDNA sequence analysis indicates that strain LC8 is most similar to the genus Erythrobacter (nucleotide sequence identity, 96.7%), with Erythromicrobium (sequence identity, 94.8%) and Porphyrobacter (sequence identity, 94.9%) being the next closest relatives. In addition, maximum likelihood analysis places strain LC8 and Erythrobacter on the same phylogenetic branch, distinct from the Erythromicrobium-Porphyrobacter branch as well as from that of the sphingomonads. However, the possibility that strain LC8 belongs to the genus Erythrobacter, Erythromicrobium, or Porphyrobacter is remote, since strain LC8 does not appear to contain photosynthetic pigments, while all Erythrobacter, Erythromicrobium, and Porphyrobacter spp. do (13, 42, 43). Thus, strain LC8 appears to be a new genus within the family Sphingomonadaceae (25), for which we provisionally assign the name Lutibacterium anuloederans LC8.

BLASTN search results and phylogenetic analysis of strain M4-6 16S rDNA sequence data conclusively establish it as a member of the genus Cycloclasticus. Maximum likelihood analysis shows that strain M4-6 forms a very tight cluster with all of the Cycloclasticus species analyzed. However, none of the previously reported Cycloclasticus species has a spirillum morphology, which suggests that strain M4-6 is a novel Cycloclasticus species. Strain M4-6 is further differentiated from extant Cycloclasticus species by a number of phenotypic traits (Table 1). For example, Dyksterhouse et al. (12) describe several Cycloclasticus strains from Puget Sound, all of which were short rods. Cycloclasticus pugetii reduces nitrate to nitrite, has lipase activity, and lacks PHB accumulation, while strain M4-6 shows no nitrate reduction or lipase activity but accumulates PHB (Table 1). Thus, we propose the new species, C. spirillensus M4-6.

The genus Cycloclasticus has been isolated from a variety of uncontaminated and PAH-polluted sites, and its ability to degrade PAHs has been well documented in cultures (12, 14, 15). However, C. spirillensus M4-6 did not substantially enhance or sustain phenanthrene uptake when it was added to slurries from Lowes Cove or Boston Harbor (LMC) that were prepared with or without exogenous phenanthrene (Fig. 6 and 7). Qualitative observations of the distinctive spirillum morphology in slurries with added C. spirillensus M4-6, but not in unamended slurries, indicate that the rapid decrease in phenanthrene uptake for C. spirillensus M4-6 treatments is likely not due to cell loss alone. Loss of uptake over time in Lowes Cove slurries with added phenanthrene and in LMC slurries with elevated background PAH levels also indicates that the continuous presence or absence of PAHs likely had little impact on C. spirillensus M4-6. At present, the factors that control phenanthrene uptake potential by C. spirillensus M4-6 remain uncertain. Nonetheless, results from this study suggest that C. spirillensus M4-6 may not be a particularly effective PAH degrader in situ.

Relative to uninoculated slurries, inoculation of Lowes Cove and LMC slurries with L. anuloederans LC8 significantly enhances and sustains the phenanthrene uptake of treatments both with and without periodic phenanthrene addition (Fig. 6 and 8). Lower uptake rates over time for Lowes Cove slurries without periodic phenanthrene addition may reflect decreased expression of the dioxygenase(s) that initiates phenanthrene degradation, since uptake rates were higher and more stable for slurries with periodic phenanthrene addition (Fig. 6). In contrast, lower uptake rates over time in LMC slurries with periodic phenanthrene additions may reflect inhibition due to desorption of toxic PAHs or other organics by phenanthrene.

In summary, burrows of marine macrofauna in uncontaminated sediments exhibit enhanced PAH degradation rates (9) and harbor novel PAH-degrading bacteria. Based on responses of burrow isolates to reinoculation in contaminated and uncontaminated sediments, L. anuloederans LC8 (from a polychaete burrow) appears somewhat better suited for PAH degradation than does C. spirillensus M4-6 (isolated from a mollusc burrow).

Description of Lutibacterium anuloederans LC8 gen. nov., sp. nov. Lutibacterium anuloederans LC8 gen. nov., sp. nov. (lu.ti.bac' teri.um. L. n. lutum, mud; M. L. Lutibacterium, the mud bacterium; an' ulo.eder.ans. L. n. anulus, ring; L. v. ederans, to eat; M.L. anuloederans, ring-eating). Gram-negative, nonmotile, short rod with a single polar flagellum. Aerobic. Requires sodium ion for growth. Catalase and oxidase positive. PAHs, including naphthalene and phenanthrene, are used as sole or principal carbon sources for growth. In addition, strains utilize selected organic acids and amino acids, including citrate, acetate, pyruvate, and phenylalanine. Nitrate is reduced to nitrite. Colonies are small, round, opaque, and entire, with yellow pigmentation. The principal fatty acids in whole cells are C_18:17c, C_16:7c, and C_16:0.

Description of Cycloclasticus spirillensus M4-6, sp. nov. Cycloclasticus spirillensus M4-6, sp. nov. (spi.ril.len' sus. L. adj. spirillum, of or pertaining to a spiral). Gram-negative, motile spirillum with a single polar flagellum. 1.5 to 2.0 by 0.5 µm. Aerobic. Requires sodium ion for growth. Catalase and oxidase positive. PAHs, including naphthalene and phenanthrene, are used as sole or principal carbon sources for growth. In addition, strains utilize selected organic acids and amino acids, including citrate, acetate, pyruvate, alanine, and proline. Nitrate is not reduced to nitrite. Colonies are small, round, translucent, and entire, with no pigmentation. The principal fatty acids in whole cells are C_15:0 iso, C_15:0 anteiso, and C_16:7c.