APPLIED AND ENVIRONMENTAL MICROBIOLOGY, July 2009, p. 4620–4623
0099-2240/09/$08.00⫹0 doi:10.1128/AEM.00582-09
Copyright © 2009, American Society for Microbiology. All Rights Reserved.
Vol. 75, No. 13
[FeFe] Hydrogenase Genetic Diversity Provides Insight into Molecular
Adaptation in a Saline Microbial Mat Community䌤†
Eric S. Boyd,1 John R. Spear,2 and John W. Peters1*
Department of Chemistry and Biochemistry and Astrobiology Biogeocatalysis Research Center, Montana State University, Bozeman,
Montana 59717,1 and Division of Environmental Science and Engineering, Colorado School of Mines, Golden, Colorado 804012
Received 10 March 2009/Accepted 29 April 2009
Degenerate primers for the [FeFe] hydrogenase (hydA) were developed and used in PCRs to examine hydA
in microbial mats that inhabit saltern evaporative ponds in Guerrero Negro (GN), Mexico. A diversity of
deduced HydA was discovered that revealed unique variants, which may reflect adaptation to the environmental conditions present in GN.
427 (TGGVMEAA) in the Clostridium pasteurianum protein
sequence, were designed for use in specific gene amplifications.
Mat core samples (1 by 6 cm) were collected at 2:00 pm from
pond 4 (35°C, pH 8.0, and 80 ppt salt) near pond 5 at the
Exportadora de Sal saltworks, GN, on 13 February 2005. Mat
cores were sectioned in 1-mm increments on site and immediately flash frozen in liquid nitrogen as previously described
(16). Genomic DNA was extracted by bead beating in the
presence of phenol-chloroform-isoamyl alcohol and sodium
dodecyl sulfate as previously described (7) and was quantified
by using the High DNA Mass Ladder (Invitrogen, Carlsbad,
CA). Approximately 500-bp fragments of hydA were amplified
in triplicate using primers FeFe-272F (5⬘-GCHGAYMTBAC
HATWATGGARGA-3⬘, where H ⫽ A, C, T; Y ⫽ C, T; M ⫽
A, C; B ⫽ C, G, T; W ⫽ A, T; R ⫽ A, G; 432-fold degeneracy)
and FeFe-427R (5⬘-GCNGCYTCCATDACDCCDCCNGT-3⬘,
where N ⫽ A, C, T, G; Y ⫽ C, T; D ⫽ A, G, T; 864-fold
degeneracy) and 35 cycles of PCR as previously described (4).
An empirically determined annealing temperature of 56.5°C, a
MgCl2 concentration of 1.5 mM, and a primer concentration of
1 ␮M for each forward and reverse primer were utilized in
50-␮l PCR mixtures containing 10 ng of genomic DNA as the
template. Equal 40-␮l volumes of three replicate PCR products were pooled, purified using the Promega Wizard purification kit (Madison, WI), quantified using the Low Mass DNA
Ladder (Invitrogen), cloned using the pGEM Easy Vector
System (Promega), and sequenced by using the M13F-M13R
primer pair as previously described (5).
Deduced amino acid sequences were screened for the presence of the L1 and L2 HydA sequence motifs as described
above. ClustalX (version 2.0.8) (15) was employed to align
inferred amino acid sequences and to create distance matrices
for use in identifying and clustering operational taxonomic
units (OTUs) with DOTUR (26). Calculations of Shannon
diversity and Chao1 and Ace1 deduced amino acid sequence
richness were completed with DOTUR by using the furthestneighbor algorithm and a precision of 0.01. The phylogenetic
position of putative HydA was assessed with MRBAYES (10,
25). Tree topologies were sampled every 500 generations for
2,000,000 generations (burnin ⫽ 1,000,000) by using the WAG
evolutionary model with fixed amino acid frequencies and
gamma-shaped rate variation with a proportion of invariable
Hydrogen (H2) is an important intermediate in the decomposition of organic matter in anaerobic environments and is
the basis for many syntrophic interactions that occur in the
food chain such as those between acetogens and methanogens
and/or sulfate reducers. Little is known concerning the potential role of fermentative bacteria and/or H2 cycling in phototrophic microbial mat systems, such as the 6-cm-thick and extremely diverse microbial mats inhabiting the saline ponds at
the Exportadora de Sal SA saltern in Guerrero Negro (GN),
Baja California Sur, Mexico. Recent 16S rRNA gene surveys in
GN revealed an abundance of bacteria in the upper 2 mm of
the mat that, based on their phylogenetic affiliation, are
thought to harbor fermentative metabolisms (6, 8, 13, 16, 20,
28). Similarly, previous studies have shown that the flux of H2
from the surface of the phototrophic mats present in GN is
⬃150-fold higher during the night than during the day (9).
Together, these results suggest the potential for fermentative
metabolisms in H2 and carbon cycling in the GN saline mat
environment.
The [FeFe] hydrogenase has a central role in primary and
secondary substrate fermentations, catalyzing the oxidation of
excess reducing equivalents coupled to the reduction of protons, yielding H2 (2, 17, 29, 30). The [FeFe] hydrogenase is
only known to occur in anaerobic bacteria and a small number
of green algae (23, 29, 30), making this a useful biomarker for
examining the diversity and distribution of this functional class
of organism. Eighty-two putative [FeFe] hydrogenase largesubunit protein sequences (HydA) present in the GenBank
database that represent the known diversity of putative HydA
were screened for the L1 ([FLI]TSC[C/S]P[GAS]W[VIQH])
and L2 ([IVLF]MPCx[ASRD]K[KQ]xE) (conserved residues
are in bold and underlined, and bracketed positions indicate
“semiconserved” residues at that position) signature sequence
motifs (17, 30). Degenerate PCR primers, corresponding to
positions 272 to 279 (AD[M/L]TIMEE) and positions 420 to
* Corresponding author. Mailing address: Department of Chemistry
and Biochemistry, 103 Chemistry Research Building, Montana State
University, Bozeman, MT 59717. Phone: (406) 994-7211. Fax: (406)
994-5407. E-mail: John.peters@chemistry.montana.edu.
† Supplemental material for this article may be found at http://aem
.asm.org/.
䌤
Published ahead of print on 8 May 2009.
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VOL. 75, 2009
[FeFe] HYDROGENASE DIVERSITY IN A SALINE MAT
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FIG. 1. (A) Phylogenetic tree based on deduced putative HydA amino acid sequences from GN and reference HydA sequences. Nodes with
posterior probabilities of ⬍50% were collapsed in this analysis (posterior probabilities of 100 are denoted by asterisks). Bar, one substitution per
10 sites. (B) Partial-length deduced amino acid sequences of environmental and reference deduced HydA sequences illustrating the phylogenetic
coherence of novel substitutions and insertions in and upstream of L1 sequence motif. Cluster designations (C1 to C7) correspond to those
presented in Table 1.
sites as recommended by ProtTest (1). The Saccharomyces
cerevisiae Narf protein, a homolog of HydA, was used as the
outgroup. The phylogenetic tree was projected using TreeView
(version 1.6.6) (21).
An unexpectedly diverse assemblage of putative HydA variants was present in the top 1 mm of the GN microbial mat,
corresponding to the photic zone. At a sequence identity
threshold (SIT) of 100%, the predicted Shannon diversity index was 3.84 and the mean Chao1 HydA richness was 187
unique OTUs (see Fig. S1a in the supplemental material).
When a 99% SIT was applied, the predicted Chao1 richness
estimate decreased 56% to 82 unique OTUs and the Shannon
4622
BOYD ET AL.
APPL. ENVIRON. MICROBIOL.
TABLE 1. Phylogeny of partial-length putative HydA sequence clusters
Sequence
clustera
No. of clones
in cluster
Maximal intracluster
sequence divergence
(%)b
C1
C2
C3
C4
C5
C6
C7
22
16
6
9
3
7
2
49.4
41.6
28.8
38.6
38.6
51.8
32.9
Most closely related sequencec
Phylumd
Mean % sequence identity
(range)e
Moorella thermoacetica ATCC 39073
Opitutus terrae PB90-1
Bacteroides thetaiotaomicron VPI-5482
Alkaliphilus oremlandii OhILAs
Moorella thermoacetica ATCC 39073
Heliobacillus mobilis
Thermoanaerobacterium saccharolyticum
JW/SL-YS485
Firmicutes
Verrucomicrobia
Bacteroidetes
Firmicutes
Firmicutes
Firmicutes
Firmicutes
61.8 (56.4–67.3)
64.3 (56.3–71.1)
72.1 (69.8–73.2)
65.1 (61.4–66.4)
61.4 (NA)
50.3 (47.5–54.6)
61.2 (58.5–64.2)
a
Sequence clusters correspond to those presented in Fig. 1.
Maximum intracluster sequence divergence among sequences within a cluster as calculated by the ClustalX sequence identity matrix following alignment using the
Gonnet protein weight matrix (gap opening penalty of 13, gap extension penalty of 0.05).
c
The most closely related sequence is defined as the taxon with the HydA sequence most closely related to that of members within the cluster as presented
in Fig. 1.
d
Phylum designation of most closely related HydA sequence as defined in Fig. 1.
e
Sequence identity means and ranges were calculated by using the ClustalX sequence identity matrix as described in footnote b. NA, not available.
b
diversity index decreased slightly to 3.55 (data not shown). A
further decrease in the SIT from 99 to 85% resulted in a slight
decrease (23%) in the Chao1 predicted richness and a modest
decrease in the Shannon diversity index from 3.55 to 3.28,
suggesting that the majority of the putative HydA OTUs
present in the top 1 mm of the mat is a result of recent
evolution or speciation.
The putative HydA variants from GN resolved into 42 OTUs
which clustered into one of seven distinct phylogenetic clusters
(Fig. 1A), although clusters 5 and 7 could not be adequately
resolved. Intriguingly, the mean identity scores determined
from sequence alignment matrices for members of each of the
seven sequence clusters recovered from GN, when compared
to sequences in the GenBank database, were very low (47.5 to
73.2%) (Table 1), suggesting that the putative HydA sequences
recovered from GN represent novel sequence space.
The highest proportion of putative HydA sequences recovered from the top 1-mm transect of the GN microbial mat were
related to acetogens most closely affiliated with the Firmicutes
(66.2% of the clones) and the Verrucomicrobia (24.6%), with a
lower proportion related to the Bacteroidetes (9.2%). These
observations support those of previous studies that identified
the highest abundance of 16S rRNA genes related to fermentative/acetogenic organisms within the phyla Firmicutes, Verrucomicrobia, and Bacteroidetes in the upper few millimeters of
microbial mat (16), coinciding with the mat transect where
nighttime H2 concentrations were also elevated (9). The recovery of putative HydA sequences in the photic zone of the
GN microbial mat is consistent with the results of previous
studies conducted in the phototrophic mats in Yellowstone
National Park, WY, which documented the potential for the
secondary fermentation of cyanobacterial primary fermentation products in the top photic layers of the mat at night
(3). Importantly, cyanobacterial fermentation may also contribute to the production of H2 in the mats during periods of
anoxia (9).
Deduced amino acid sequences from GN had significantly
lower (P ⫽ ⬍0.01 for both GRAVY and aliphatic indices)
hydropathy indices than nonhalotolerant HydA sequences
from GenBank (see Table S1 in the supplemental material), a
feature which may reflect molecular adaptation to salinity.
These observations are consistent with the results of previous
investigations of deduced amino acid sequences from a variety
of halophilic organisms that indicate hydrophobicity indices
lower than those of their nonhalophilic counterparts, which is
hypothesized to reduce the effects of salt-driven protein misfolding and/or aggregation (11, 12, 14, 24, 27, 31). In addition,
many of the putative HydA sequences recovered from GN
exhibited previously unobserved substitutions in the L1 motif
and novel insertion domains upstream from the L1 motif (Fig.
1B). Residues within L1 are involved in the coordination of the
oxygen-labile [4Fe-4S] subcluster of the H cluster of HydA (18,
19, 22). Substitutions in this region may have implications for
the redox properties of this [4Fe-4S] cluster and thus may be
important in conferring stability in the presence of oxygen (23).
The discovery of a rich assemblage of putative HydA variants in the top 1 mm of the GN microbial mat underscores the
utility of these primers in examining the natural diversity of
putative HydA and fermentative organisms in complex microbial ecosystems. The composition of putative HydA sequences
from the top 1 mm of the microbial mat suggests adaption to
the conditions present in the evaporative salterns in GN.
Nucleotide sequence accession numbers. A total of 65 putative hydA gene sequences determined in the present study
have been deposited in the GenBank, DDBJ, and EMBL databases under accession numbers FJ623894 to FJ623958.
We gratefully thank the Exportadora de Sal, GN, Baja California
Sur, Mexico, for access to their saltworks, as well as the NASA Ames
GN researchers who facilitated the permitting process and gave advice.
We are also grateful to Laura Beer for helpful comments during the
preparation of the manuscript.
This work was supported by Air Force Office of Scientific Research
Multidisciplinary University Research Initiative Award FA9550-05-010365 and the NASA Astrobiology Institute Funded Astrobiology Biogeocatalysis Research Center (NNA08C-N85A to J.W.P). E.S.B. was
supported by a NASA Postdoctoral Fellowship awarded by the NASA
Astrobiology Institute. J.R.S. was supported by a National Science
Foundation Microbial Biology Postdoctoral start-up award (0624789)
and a U.S. Air Force Office of Scientific Research award (R-8196-G1).
REFERENCES
1. Abascal, F., R. Zardoya, and D. Posada. 2005. ProtTest: selection of best-fit
models of protein evolution. Bioinformatics 21:2104–2105.
VOL. 75, 2009
2. Adams, M. W. W. 1990. The structure and mechanism of iron hydrogenases.
Biochim. Biophys. Acta 1020:115–145.
3. Anderson, K. L., T. A. Tayne, and D. M. Ward. 1987. Formation and fate of
fermentation products in hot spring cyanobacterial mats. Appl. Environ.
Microbiol. 53:2343–2352.
4. Boyd, E. S., D. E. Cummings, and G. G. Geesey. 2007. Mineralogy influences
structure and diversity of bacterial communities associated with geological
substrata in a pristine aquifer. Microb. Ecol. 54:170–182.
5. Boyd, E. S., S. King, J. K. Tomberlin, D. K. Nordstrom, D. P. Krabbenhof,
T. Barkay, and G. G. Geesey. 2009. Methylmercury enters an aquatic food
web through acidophilic microbial mats in Yellowstone National Park, Wyoming. Environ. Microbiol. 11:950–959.
6. Canfield, D. E., and D. J. Des Marais. 1993. Biogeochemical cycles of
carbon, sulfur, and free oxygen in a microbial mat. Geochim. Cosmochim.
Acta 57:3971–3984.
7. Dojka, M. A., P. Hugenholtz, S. K. Haack, and N. R. Pace. 1998. Microbial
diversity in a hydrocarbon- and chlorinated-solvent-contaminated aquifer
undergoing intrinsic bioremediation. Appl. Environ. Microbiol. 64:3869–
3877.
8. Feazel, L. M., J. R. Spear, A. B. Berger, J. K. Harris, D. N. Frank, R. E. Ley,
and N. R. Pace. 2008. Eucaryotic diversity in a hypersaline microbial mat.
Appl. Environ. Microbiol. 74:329–332.
9. Hoehler, T. M., B. M. Bebout, and D. J. Des Marais. 2001. The role of
microbial mats in the production of reduced gases on the early Earth. Nature
412:324–327.
10. Huelsenbeck, J. P., and R. Ronquist. 2001. MRBAYES: Bayesian inference
of phylogeny. Bioinformatics 17:754–755.
11. Hutcheon, G. W., N. Vasisht, and A. Bolhuis. 2005. Characterization of a
highly stable ␣-amylase from the halophilic archaeon Haloarcula hispanica.
Extremophiles 9:487–495.
12. Joo, W. A., and C. W. Kim. 2005. Proteomics of halophilic archaea. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 815:237–250.
13. Kunin, V., J. Raes, J. K. Harris, J. R. Spear, J. J. Walker, N. Ivanova, C. von
Mering, B. M. Bebout, N. R. Pace, P. Bork, and P. Hugenholtz. 2008.
Millimeter-scale genetic gradients and community-level molecular convergence in a hypersaline microbial mat. Mol. Syst. Biol. 4:198.
14. Lanyi, J. K. 1974. Salt-dependent properties of proteins from extremely
halophilic bacteria. Bacteriol. Rev. 38:272–290.
15. Larkin, M. A., G. Blackshields, N. P. Brown, R. Chenna, P. A. McGettigan,
H. McWilliam, F. Valentin, I. M. Wallace, A. Wilm, R. Lopez, J. D. Thompson, T. J. Gibson, and D. G. Higgins. 2007. Clustal W and Clustal X version
2.0. Bioinformatics 23:2947–2948.
[FeFe] HYDROGENASE DIVERSITY IN A SALINE MAT
4623
16. Ley, R. E., J. K. Harris, J. Wilcox, J. R. Spear, S. R. Miller, B. M. Bebout,
J. A. Maresca, D. A. Bryant, M. L. Sogin, and N. R. Pace. 2006. Unexpected
diversity and complexity of the Guerrero Negro hypersaline microbial mat.
Appl. Environ. Microbiol. 72:3685–3695.
17. Meyer, J. 2007. [FeFe] hydrogenases and their evolution: a genomic perspective. Cell. Mol. Life Sci. 64:1063–1084.
18. Nicolet, Y., B. J. Lemon, J. C. Fontecilla-Camps, and J. W. Peters. 2000. A
novel FeS cluster in Fe-only hydrogenases. Trends Biochem. Sci. 25:138–143.
19. Nicolet, Y., C. Piras, P. Legrand, C. E. Hatchikian, and J. C. FontecillaCamps. 1999. Desulfovibrio desulfuricans iron hydrogenase: the structure
shows unusual coordination to an active site Fe binuclear center. Structure
7:13–23.
20. Nübel, U., F. Garcia-Pichel, M. Kuhl, and G. Muyzer. 1999. Quantifying
microbial diversity: morphotypes, 16S rRNA genes, and carotenoids of oxygenic phototrophs in microbial mats. Appl. Environ. Microbiol. 65:422–430.
21. Page, R. D. M. 1996. TREEVIEW: an application to display phylogenetic
trees on personal computers. Comput. Appl. Biosci. 12:357–358.
22. Peters, J. W., W. N. Lanzilotta, B. J. Lemon, and L. C. Seefeldt. 1998. X-ray
crystal structure of the Fe-only hydrogenase (CpI) from Clostridium pasteurianum to 1.8 angstrom resolution. Science 282:1853–1858.
23. Posewitz, M. C., D. W. Mulder, and J. W. Peters. 2008. New frontiers in
hydrogenase structure and biosynthesis. Curr. Chem. Biol. 2:178–199.
24. Reistad, R. 1970. On the composition and nature of the bulk protein of the
extremely halophilic bacteria. Arch. Mikrobiol. 71:353–360.
25. Ronquist, F., and J. P. Huelsenbeck. 2003. MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19:1572–1574.
26. Schloss, P. D., and J. Handelsman. 2005. Introducing DOTUR, a computer
program for defining operational taxonomic units and estimating species
richness. Appl. Environ. Microbiol. 71:1501–1506.
27. Spahr, P. F. 1962. Amino acid composition of ribosomes from Escherichia
coli. J. Mol. Biol. 4:395–406.
28. Spear, J. R., R. E. Ley, A. Berger, and N. R. Pace. 2003. Complexity in
natural microbial ecosystems: the Guerrero Negro experience. Biol. Bull.
204:168–173.
29. Vignais, P. M., and B. Billoud. 2007. Occurrence, classification, and biological function of hydrogenases: an overview. Chem. Rev. 107:4206–4272.
30. Vignais, P. M., B. Billoud, and J. Meyer. 2001. Classification and phylogeny
of hydrogenases. FEMS Microbiol. Rev. 25:455–501.
31. Visentin, L. P., C. Chow, A. T. Matheson, M. Yaguchi, and F. Rollin. 1972.
Halobacterium cutirubrum ribosomes. Properties of the ribosomal proteins
and ribonucleic acid. Biochem. J. 130:103–110.