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Applied and Environmental Microbiology, April 2004, p. 2119-2128, Vol. 70, No. 4
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.4.2119-2128.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Determination of Nitrogen-Fixing Phylotypes in Lyngbya sp. and Microcoleus chthonoplastes Cyanobacterial Mats from Guerrero Negro, Baja California, Mexico

Enoma O. Omoregie,1 Lori L. Crumbliss,1 Brad M. Bebout,2 and Jonathan P. Zehr1*

Department of Ocean Sciences and Institute of Marine Sciences, University of California, Santa Cruz, California 95064,1 Exobiology Branch, NASA Ames Research Center, Moffett Field, California 940352

Received 25 August 2003/ Accepted 1 December 2003


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ABSTRACT
 
In many environments, biological nitrogen fixation can alleviate nitrogen limitation. The high rates of N2 fixation often observed in cyanobacterial mats suggest that N2 fixation may be an important source of N. In this study, organisms expressing nifH were identified in a Lyngbya sp.- and two Microcoleus chthonoplastes-dominated cyanobacterial mats. The pattern of nitrogenase activity was determined for the Lyngbya sp. mat and a Microcoleus chthonoplastes mat sampled directly in Guerrero Negro, Mexico. Their maximum rates were 23 and 15 µmol of C2H4 m–2 h–1, respectively. The second Microcoleus mat, which was maintained in a greenhouse facility, had a maximum rate of 40 µmol of C2H4 m–2 h–1. The overall diel pattern of nitrogenase activity in the three mats was similar, with the highest rates of activity occurring during the dark period. Analysis of nifH transcripts by reverse transcription-PCR revealed that several different organisms were expressing nifH during the dark period. nifH phylotypes recovered from these mats were similar to sequences from the unicellular cyanobacterial genera Halothece, Myxosarcina, and Synechocystis, the filamentous cyanobacterial genera Plectonema and Phormidium, and several bacterial nifH groups. The results of this study indicate that several different organisms, some of which were not previously known to fix nitrogen, are likely to be responsible for the observed dark-period nitrogenase activity in these cyanobacterial mats.


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INTRODUCTION
 
Microbial mats are benthic laminated microbial assemblages that are persistent biological features in many environments. They are found in many different aquatic habitats, including hydrothermal vents, hot springs, ponds, and lakes (20, 22, 24, 25). Cyanobacterial mats are microbial mats that contain cyanobacteria but typically host a large diversity of microorganisms that function in numerous microbially mediated processes. Microbial processes in mats include photosynthesis, sulfate reduction, methanogenesis, and nitrogen fixation (5, 7, 16). Thus, there are strong light and chemical gradients in mats that both control and result from the vertical stratification of microorganisms and biochemical activities in the mats.

In many environments, nitrogen availability can limit growth and ecosystem productivity (44). Biological nitrogen fixation, or diazotrophy, the fixation of atmospheric nitrogen gas (N2) into biologically available ammonia (NH3), is important in making nitrogen available in many ecosystems, including microbial mats (18, 27, 44). The ability to fix N2 is widely distributed among diverse members of the Bacteria and Archaea (46). The nitrogenase protein which catalyzes the conversion of N2 to NH3 is highly conserved among microorganisms and is likely to have evolved early (33, 46). Nitrogen fixation is inhibited by oxygen due to the sensitivity of nitrogenase to oxygen inactivation (17). Oxygen-evolving cyanobacteria have developed various strategies, such as temporal or spatial segregation of N2 fixation (into heterocysts) and oxygen evolution, to avoid oxygen inactivation (14). Cyanobacterial mats contain diverse types of microorganisms that could potentially fix N2 but also exhibit high concentrations of oxygen due to oxygenation from the atmosphere and cyanobacterial photosynthesis (2, 12). Thus, it is often unclear which microorganisms may be involved in N2 fixation and how N2 fixation in mats proceeds in the presence of oxygen-evolving phototrophs.

Cyanobacterial mats, which contain diverse cyanobacterial taxa, including heterocystous cyanobacteria, filamentous nonheterocystous cyanobacteria, and unicellular cyanobacteria, often exhibit high rates of nitrogen fixation (4, 5, 13, 28, 30). The temporal patterns of N2 fixation observed in cyanobacterial mats are often similar to those observed in cultures of cyanobacteria (35), suggesting that cyanobacteria may play a role in mat N2 fixation. However, N2 fixation may also be driven by the activities of anoxygenic photoautotrophs, including green and purple bacteria (31, 45). It is also possible that heterotrophic bacteria are involved in N2 fixation, fueled by photosynthate released from phototrophs (27, 29).

Most studies of the organisms involved in N2 fixation in cyanobacterial mats have focused on physiological and genetic characterizations (4, 26, 31, 36, 37, 48). Incubating microbial mats with the photosystem II inhibitor DCMU [3-(3,4-dichlorophenyl)-1,1-dimethylurea] and/or keeping mats in the dark during the daytime typically causes substantial reductions in nitrogenase activity during the subsequent night (4, 31). The decreased dark-period N2 fixation is most likely due to a reduction in photosynthetic products which would fuel N2 fixation.

Diversity studies in cyanobacterial mats have focused on amplifying and sequencing the nifH gene, one of the genes responsible for nitrogen fixation (26, 37, 48). These studies have shown that there is a diverse array of N2 fixers that could be involved in N2 fixation, including cyanobacteria, alpha-proteobacteria, gamma-proteobacteria, and delta-proteobacteria. One study attempted to identify which microorganisms actively participate in N2 fixation (36). However, there is relatively little information on how the diversity of organisms involved in N2 fixation differs between mats.

The purpose of this study was to determine the types of microorganisms involved in N2 fixation in two microbial mats exhibiting very different rates of nitrogenase activity, the well-developed Lyngbya sp. and Microcoleus chthonoplastes mats, in Guerrero Negro, Baja California, Mexico. Previous work by Omoregie et al. (26) found numerous nifH sequences belonging to various members of the proteobacterial lineage as well as cyanobacteria and several unidentified bacteria. However, from that study it was unclear which of these organisms were active in N2 fixation. We examined the rates of nitrogenase activity by acetylene reduction in relation to the expression of specific nifH gene phylotypes by reverse transcription-PCR. This method has already been proven effective in several studies (23, 36, 51). Our results show that unicellular and filamentous cyanobacteria as well as heterotrophic bacteria were expressing the nitrogenase gene during the dark period.


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MATERIALS AND METHODS
 
Sampling.
A Lyngbya sp.-dominated microbial mat (LG), freshly collected (GNM), and greenhouse-maintained Microcoleus chthonoplastes (GHM)-dominated mats were studied. The mats used in this study were sampled from the Exportadora de Sal salt works in Guerrero Negro, Baja California, Mexico. The Exportadora de Sal salt works is located approximately 720 km south of the United States border with Mexico.

Microcoleus chthonoplastes mats were collected from a hypersaline pond (area 4; salinity, ca. 90 to 78{per thousand}) in June 2001 (GHM) and October 2001 (GNM). Core samples of GNM mats were obtained immediately upon collection of mats from area 4. The GHM mats were maintained in a greenhouse facility at NASA Ames Research Center in Mountain View, Calif. (3), for approximately 1 month prior to sampling for this study in July 2001. The GHM samples were obtained from the same cores used in the study by Omoregie et al. (26).

Lyngbya (LG) mats were collected from a desiccated intertidal flat in October 2001. Because the tidal flat from which the Lyngbya mats were collected is flooded at irregular intervals by water from the Ojo de Liebre lagoon, samples of the mat were collected in plastic trays (20 by 25 cm) and transported to a nearby laboratory for time course (diel) experiments. At the field laboratory, the trays were filled with lagoon water from the nearby Ojo De Liebre (salinity, ca. 40{per thousand}) and incubated at in situ irradiance and temperature. LG mats were sampled from these trays in all experiments. Circular cores approximately 15 mm by 15 mm in dimension were taken from all three mats. Cores were sectioned at approximately 3- to 5-mm intervals and placed in cryovials, which were immediately frozen in liquid nitrogen and transferred to a –80°C freezer. An additional set of replicate cores (n = 3) were placed in serum bottles at each time point and used for acetylene reduction rate measurements (incubation period, 3 to 4 h). The top 3- to 5-mm section of each mat core was used for this study.

Acetylene reduction assay.
Nitrogenase activity was measured by the acetylene reduction assay (38) following the procedure of Bebout et al. (4). Cores (ca. 15 mm by 5 mm) were placed in 38-ml serum bottles containing 20 ml of water (overlying water from each mat). Three replicate cores were taken from each mat during each sampling period. The bottles were sealed with rubber stoppers, and 2 ml of headspace was removed and replaced with 5 ml of acetylene. The bottles were returned to the flumes, trays, or ponds from which they were sampled to maintain the same light and temperature conditions as the mats from which they were sampled. After 3 to 4 h, incubations were terminated by shaking each bottle for 10 s. Two milliliters of headspace was injected into a 9.1-ml serum bottle filled with a 3.4 M solution of sodium chloride, displacing the salt solution through a vent needle, for short-term storage. Ethylene concentrations were quantified (usually within hours of sample collection) by injecting 0.1 ml of the headspace into a Shimadzu GC14A or GCMini2 gas chromatograph (Shimadzu, Kyoto, Japan).

Isolation of total RNA.
Total RNA was extracted from mat samples by modification of the Tillett and Neilan method (42). Preheated xanthogenate-sodium dodecyl sulfate buffers (1.2 ml) was added to 25 to 40 mg of mat in a 2-ml Bead Beat tube. The tubes were agitated in a Fast Prep machine (Bio 101, Carlsbad, Calif.) for 1.5 min at speed setting 6. Two hundred microliters of chloroform-isoamyl alcohol was added to each tube, vortexed briefly, and then centrifuged for 5 min. The aqueous phase was added to 500 µl of phenol-chloroform-isoamyl alcohol. Extractions were repeated until no visible material was left at the interface. The aqueous phase was removed to a microcentrifuge tube, to which 1 volume of isopropanol added, and then placed on ice for 15 min. The tubes were centrifuged for 10 min. The isopropanol was aspirated, and the tubes were washed twice in 70% ethanol. RNA pellets were then resuspended in 50 µl of nuclease-free water (Ambion). Residual DNA was removed by two rounds of DNase (Ambion) treatment, followed by purification with the RNEasy mini kit (Qiagen, Hilden, Germany) and eluted in 50 µl of RNA storage solution. To demonstrate that the RNA was not contaminated with genomic DNA, 1 µl of RNase (Ambion) was added to aliquots of extracted RNA prior to DNase treatment.

Reverse transcription and amplification.
Reverse transcription and amplification were carried out by a nested PCR approach with primers NIFH4 (5'-TTYTAYGGNAARGGNGG-3', positions 564 to 562 in A. vinelandii M11579) and NIFH3 (5'-ATRTTRTTNGCNGCRTA-3', positions 1102 to 1118 in A. vinelandii M11579) for the first round and NIFH1 (5'-TGYGAYCCNAARGCNGA-3', positions 639 to 655 in A. vinelandii M11579) and NIFH2 (5'-ADNGCCATCATYTCNCC-3' positions 984 to 1000 in A. vinelandii M11579) for the second round (N = A, G, C, or T; D = A, G, or T; Y = C or T; R = A or G) (50). Primers were synthesized and purified by polyacrylamide gel electrophoresis at New England Biolabs.

Two microliters of sample was added to a 50-µl Access RT-PCR (Promega) reaction. Each reaction contained 1x reaction buffer, 1 mM MgSO4, 0.8 mM total deoxynucleoside triphosphates, 1 µM each NIFH3 and NIFH4, 5 U of Tfl DNA polymerase, and 5 U of avian myeloblastosis virus reverse transcriptase. Reaction master mixes containing all reagents excluding avian myeloblastosis virus reverse transcriptase were filtered through a 100-kDa filter (Millipore). After filtration, avian myeloblastosis virus reverse transcriptase was added to the master mix and aliquoted, and the template was added to each reaction mixture. Alternatively, reaction mixtures without avian myeloblastosis virus or Tfl reverse transcriptase were filtered through a 30-kDa filter (Millipore), and enzymes were added and then aliquoted. To demonstrate that reverse transcriptase was necessary for sample amplification and that contaminating genomic DNA was not present, control reactions without reverse transcriptase were also performed. Samples were reverse transcribed and amplified with the following cycling conditions: 48°C for 45 min and 94°C for 2 min, followed by 40 cycles of 94°C for 30 s, 57°C for 30 s, and 72°C for 1 min, followed by 72°C for 7 min.

A second round of nested amplification was performed with the NIFH1 and NIFH2 primers. The final concentration in each 50-µl reaction mixture was 4 mM MgCl2, 1x PCR buffer, 0.8 mM total deoxynucleoside triphosphates, 1 µM each forward primer and reverse primer, and 2.5 U of Taq (Promega). Core mixes were filtered as described above. The following cycling conditions were used for the second round: 94°C for 1 min, followed by 30 cycles of 94°C for 30 s, 57°C for 30 s, and 72°C for 1 min, and a final 72°C for 7 min.

Cloning and plasmid isolation.
Amplification products were ligated and transformed with the pGEM-T vector kit (Promega). Transformation reactions were plated on Luria-Bertani (LB)-agar plates containing 100 µg ml–1 ampicillin, 0.5 mM isopropylthiogalactopyranoside (IPTG), and 80 µg ml–1 5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside (X-Gal). Individual colonies were used to inoculate overnight cultures containing 3 ml of LB broth and 100 µg ml–1 ampicillin. Plasmids from cell cultures were purified with the QIAprep Spin miniprep kit (Qiagen, Hilden, Germany).

DNA sequencing and analysis.
Purified plasmids were sequenced in one direction with either the Sp6 or T7 primer (IDT) with the ABI Prism BigDye Terminator version 3.0 cycle sequencing kit (Applied Biosystems). Samples were sequenced on an ABI Prism 310 genetic analyzer (Applied Biosystems). Sequences were edited with the GCG program (Accelrys). Probabilistic alignments for all sequences and sequences from GenBank were generated from predicted amino acid sequences with the HMMR program in GCG. Percent identities were calculated with the distance program in GCG with Kimura correction; the calculated distances were then normalized to 100. Representative sequences were chosen and sequenced in the reverse direction with T7 or Sp6 primer. Phyloge-netic distances were calculated from the probabilistic alignments with the distance correction algorithm of Tajima-Nei, available in the program TREECON for Windows (39, 43). Phylogenetic reconstructions from distance approximations were done by the neighbor-joining method in TREECON (34). A total of 100 bootstrap replicates were performed for each tree. The cluster 1 tree was rooted with a nifH gene from Desulfovibrio vulgaris, and the cluster 2, 3, and 4 trees were rooted with a nifH-like gene from Plectonema boryanum.


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RESULTS
 
Nitrogenase activity.
Nitrogenase activity for both mats (Fig. 1 and 2) was maximal at night and minimal during the day. This pattern of activity was inversely correlated to light intensity. The maximum rates of acetylene reduction for the LG and the GNM mats were 23 and 15 µmol of C2H4 m–2 h–1, respectively.



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  		FIG. 1. Acetylene reduction (AR) rate measurements and light intensity for the October 2001 Microcoleus (GNM) mat. Solid squares ({blacksquare}) represent irradiance, and solid triangles ({blacktriangleup}) represent the rate of acetylene reduction. Horizontal bars indicate the length of incubation. Vertical bars represent ±1 standard deviation of the average for three replicates. The highest rates of N2 fixation (15 µmol of C2H4 m–2 h–1) were measured between 22:00 and 2:00 h during the dark period.



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  		FIG. 2. Acetylene reduction (AR) rate measurements and light intensity for the Lyngbya mat (LG). Measurements were made over a day and half of sampling (39 h). Solid squares ({blacksquare}) represent irradiance, and solid triangles ({blacktriangleup}) represent the rate of acetylene reduction. Horizontal bars indicate the length of incubation. Vertical bars represent ±1 standard deviation from the average for three replicates. The highest rates of N2 fixation (23 µmol of C2H4 m–2 h–1) were measured between 22:00 and 2:00 h (second day of measurements) during the dark period.

nifH gene transcription.
Since acetylene reduction rates were maximal at night, RNA samples collected at night were assayed for nifH expression. nifH transcripts were detected in all three mats during the dark. Figure 3 shows the product of an RT-PCR amplification of nifH mRNA from the LG mat. RT-PCR amplifications from the GHM and GNM mats (gels not shown) were similar to that of the LG mat. Samples taken from the LG and GHM mats at 22:00 and 2:00 h were positive for amplification of nifH. Additionally, samples taken from the GHM mat and the GNM mat at 18:00 h and from the GNM mat at 20:00 h were also positive for amplification of nifH. Samples to which RNase was added just prior to purification and samples in which reverse transcriptase was not included in the reaction mixture were all negative for amplification. In addition, no-template controls for all reactions were negative for amplification.



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  		FIG. 3. Agarose gel showing reverse transcription and amplification of nighttime nifH mRNA from the Lyngbya mat (LG). Samples were collected at 22:00 h during the first day of incubation and at 22:00 and 2:00 h during the second day of incubation. (A) mRNA samples treated with DNase, and DNased samples treated with RNase prior to purification of mRNA, as well as a no-template control. (B) DNase samples to which no reverse transcriptase was added.

nifH sequence analysis.
A total of 110 cDNA sequences were recovered by RT-PCR in this study (Table 1). There were 67 sequences from the GHM mat, eight sequences from the GNM mat, and 35 sequences from the LG mat. In general, there are four major phylogenetic clusters of nifH protein sequences (9). Clusters 1 and 3 contain conventional (Mo-containing) nitrogenases from various proteobacteria, cyanobacteria, and firmicutes (cluster 1) and from clostridia, sulfate reducers, methanogens, and green sulfur bacteria (cluster 3). Cluster 2 contains "second alternative" (non-Mo- and non-V-containing) nitrogenases, and cluster 4 contains nitrogenase homologs. The majority of predicted nifH protein sequences obtained from the mats were in cluster 1 or 3 (Fig. 4 and 5). The sequence type GN2084A04, which was recovered from the LG mat, was deeply divergent and fell between clusters 2 (alternative nitrogenases) and 4 (nifH homologs). The closest isolate sequence type to this sequence in amino acid identity was that of Desulfovibrio vulgaris (70% identity).


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  		TABLE 1. Summary of representative sequences recovered from the greenhouse Microcoleus (GHM), the October 2001 Lyngbya (LG), and the Microcoleus (GNM) matsa



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  		FIG.4. Phylogenetic tree of cluster 1 of nifH, showing the placement of representative mat sequences. Representative sequences were unique sequences from each mat or one sequence chosen to represent sequences from the same mat that showed 98% or greater amino acid sequence identity. Numbers in parentheses represent the number of sequences that were 98% or more identical to the representative sequence. Symbols indicate sequences recovered from genomic DNA (26) and mRNA from the Microcoleus (GHM) mat in July 2001({clubsuit}); genomic DNA from the Microcoleus mat (GNM) in June 2001 (26) and mRNA from October 2001 ({spadesuit}); and genomic DNA from the Lyngbya mat (LG) in June 2001 (26) and mRNA from October 2001 ({diamondsuit}). Bold type indicates sequences that were recovered from mRNA in this study. Bootstrap values of greater than 49 out of 100 replicates are shown at the nodes.



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  		FIG. 5. Phylogenetic tree of nifH clusters 2, 3, and 4, showing the placement of recovered representative sequences. Representative sequences were unique sequences from each mat or one sequence chosen to represent sequences from the same mat that showed 98% or greater amino acid sequence identity. Numbers in parentheses represent the number of sequences that were 98% or greater identical to the representative sequence. Symbols indicate sequences recovered from genomic DNA (26) and mRNA from the Microcoleus (GHM) mat in July 2001({clubsuit}); genomic DNA from the Microcoleus mat (GNM) in June 2001 (26) and mRNA from October 2001 ({spadesuit}); and genomic DNA from the Lyngbya mat (LG) in June 2001 (26) and mRNA from October 2001 ({diamondsuit}). Bold type indicates sequences that were recovered from mRNA in this study. Bootstrap values of greater than 49 out of 100 replicates are shown at the nodes.

A total of 91 sequences from this study were in cluster I of the nifH protein phylogeny. There were 63 amino acid sequences from all three microbial mats that ranged from 93 to 100% identity to the nifH protein from unicellular cyanobacteria, including Halothece sp. strain MPI96P605, Myxosarcina sp. strain ATCC 29377, and Synechocystis sp. strain WH8501. A total of 19 sequences from all three mats were 95 to 100% identical to the nifH protein from the filamentous cyanobacteria Plectonema boryanum and Phormidium sp. strain ATCC 29409. Five sequences from the GNM and LG mats were 95 to 97% identical to a second copy of nifH protein in the filamentous heterocystous cyanobacterium Anabaena variabilis ATCC 29413. Four sequences from the GHM mat were 86 to 88% identical to the nifH protein from the gamma-proteobacterium Azotobacter vinelandii.

nifH protein sequences in cluster 3 (Table 1 and Fig. 5) made up about 16% (18 sequences) of all the sequences recovered in this study. There were 3, 4, and 12 nifH protein sequences recovered from the GHM, GNM, and LG mats, respectively. This cluster of nifH sequences includes several lineages of anaerobic bacteria, such as clostridia, sulfate reducers, methanogens, and green sulfur bacteria. None of the nifH sequences in this cluster were closely related (88% or less identity) to sequences from bacterial isolates. Most of the cluster 3 sequences were more closely related (91 to 100%) to sequences recovered from other mats or ecosystems with anaerobic environments. The sequence types from cultivated isolates that were closest to any of the other mat cluster 3 sequences were those from delta-proteobacteria of the genus Desulfovibrio.

Of the 26 individual sequence types detected in this study, 12 were detected in DNA clone libraries (Table 1) in a previous study of these mats by Omoregie et al. (26). Two of the five sequence types recovered from the GHM mat were not detected by PCR in the same mat (26). Four of the seven RT-PCR sequence types from the GNM mat were detected by PCR (26). Furthermore, only one of these four sequence types from the GNM mat was recovered from the same mat (26). Five of the 13 sequence types from the LG mat were recovered by PCR of DNA (26). Only four of these sequence types were recovered from the same mat.


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DISCUSSION
 
The diel pattern of nitrogenase activity (Fig. 1 and 2) for the Lyngbya (LG) mat and the Microcoleus (GNM) mat was cyclic, with the highest rates during the night and the lowest rates during the day. The pattern of acetylene reduction for the GHM mat was similar to the observed pattern in the LG and GNM mats (26). This pattern is consistent with the temporal separation of photosynthesis and N2 fixation that has been observed in several filamentous nonheterocystous cyanobacterial mats and in cyanobacterial cultures (5, 6, 10, 21, 26). This pattern is typical for cyanobacteria which do not have heterocysts, the specialized cells in which nitrogen fixation is spatially separated from oxygenic photosynthesis in the vegetative cells (6). In filamentous nonheterocystous cyanobacteria, photosynthesis and nitrogen fixation are usually temporally separated, with photosynthesis occurring during the daytime and N2 fixation occurring during the nighttime (14). Nitrogenase activity at night was anticipated because filamentous nonheterocystous cyanobacterial species are abundant in these mats.

The GNM mat had the lowest rates of acetylene reduction of the three mats studied. Low acetylene reduction rates are characteristic of this particular cyanobacterial mat (5; B. M. Bebout, unpublished data). This mat seems to be at or near steady state with respect to growth and decomposition (12) and therefore probably has a low external N requirement (5). The GHM mat had a maximum rate of 40 µmol of C2H4 m–2 h–1. The reason for the increase in rates in the GHM mat (26) is unclear but may reflect seasonal changes of N dynamics within the mat.

The LG mat had the second highest rates of acetylene reduction. These rates of acetylene reduction, although low relative to rates measured previously in this mat (5; B. M. Bebout, unpublished data), suggest that this mat has a high requirement for fixed N. The LG mat is from an intertidal region, where it is periodically flooded and desiccated. This pattern of alternating wet and dry, as well as displacement of the mat caused by tides, forces the LG mat to be in a state of perpetual growth, in which its demand for fixed N is high (5).

RT-PCR of all three mat samples showed that nifH transcripts were present during the night (Fig. 3; data from GNM and GHM mats not shown). Results from negative controls (RNase-treated samples and PCRs without reverse transcriptase) showed that the RT-PCR amplification product was derived from mRNA in the sample and not from contaminating genomic DNA.

nifH gene expression was assayed (Fig. 3) at night, when nitrogenase activity was maximum (18:00 to 2:00 h). Nighttime, or dark-phase, N2 fixation is typical for most non-heterocyst-forming cyanobacteria (10, 21), such as those found in these mats. Dark-period N2 fixation could also be a strategy employed by other bacteria to escape inactivation of their nitrogen-fixing apparatus due to high oxygen concentrations. In any event, the nifH genes expressed at night are likely to be responsible for the observed acetylene reduction activity. This activity could be regulated by a circadian rhythm, at least for the cyanobacteria, which has previously been demonstrated in several nonheterocystous cyanobacterial species (8, 11, 19).

Cyanobacterial transcripts were the most numerous nifH phylotypes in RT-PCR clone libraries in all three mats at night (Table 1 and Fig. 4 and 5). N2 fixation is an energetically expensive process, as it requires 16 ATP molecules as well as eight reducing equivalents to reduce one molecule of N2 (32). Cyanobacteria are typically (nondesiccated conditions) not limited by energy in mat environments and thus can probably easily supply the energy and reductant necessary to drive N2 fixation. Experiments have previously shown that nighttime N2 fixation is substantially reduced when the activities of photoautotrophs are inhibited during the day by a lack of light or addition of the photosystem II inhibitor DCMU (4, 31). These results suggested the involvement of cyanobacteria in N2 fixation and indirectly hinted at the type of cyanobacteria involved in this process.

Cyanobacterial nifH sequences 97 to 100% identical to sequences from the unicellular halotolerant cyanobacterium Halothece sp. strain MPI96P605 were found in the GHM and GNM mats (Table 1 and Fig. 4). The genus Halothece is part of a cluster (16S rRNA) of extremely halotolerant cyanobacteria, which are ubiquitous in hypersaline microbial communities (15, 25). Sequences 93 to 97% identical to sequences from other species of unicellular cyanobacteria, such as Myxosarcina and Synechocystis were found only in the LG mat. It is interesting that unicellular cyanobacteria have been implicated in N2 fixation in these mats, as they have been detected in numerous environments where N2 fixation has been suspected (37, 47, 49). However, they have not received attention as major diazotrophs in mats. Recently, unicellular cyanobacteria have been found to be potentially important diazotrophs in the open ocean (51). The results of this study implicate unicellular cyanobacteria in N2 fixation in mat environments as well.

Phylotypes 95 to 97% identical to that of the second nitrogenase of Anabaena variabilis strain ATCC 29413 (41) were also detected in the GNM and LG mats. The second Anabaena variabilis nitrogenase is one of three possessed by this organism and is only expressed under anaerobic conditions in vegetative cells (40, 41). Anabaena variabilis is a heterocystous cyanobacterium that can fix N2 during the day in heterocysts (aerobic) or at night in vegetative cells (anaerobic). It is unclear how widely distributed the vegetative nitrogenase is among genera and morphotypes of cyanobacteria and whether it might be evolutionarily closely related to nifH genes in nonheterocystous cyanobacteria. Thus, the phylotypes found in the mats could be from heterocystous cyanobacteria very closely related to A variabilis or to uncharacterized nonheterocystous cyanobacteria that contain a nitrogenase that is of the second Anabaena variabilis type. Heterocysts, which are very visually distinctive, have not been observed directly at the microscope level. We also detected Phormidium sp. phylotypes (98 to 100% identical) in the LG and GNM mats, as well as Plectonema boryanum phylotypes (98% identical) in the LG and GHM mats by RT-PCR. These genera of cyanobacteria are filamentous nonheterocystous, and the finding of these sequences is consistent with the observation of filamentous nonheterocystous species in the mats.

In addition to cyanobacterial nifH phylotypes, phylotypes relating to gamma- and beta-proteobacteria (nifH does not adequately distinguish between gamma and beta lineages), and alpha-proteobacteria were found in the GHM and GNM mats. Phylotypes 86 to 88% identical to the Azotobacter vinelandii nifH sequence were detected in the GHM mats. This degree of identity is not high enough to assign those sequences to the gamma-proteobacterial lineage. However, these sequences group strongly within the gamma- and beta-proteobacterial clade when data sets of nifH sequences larger than those used in the analysis presented in Fig. 4 are used, indicating that these sequences are likely to be from gamma- or beta-proteobacteria.

Cluster 3 nifH sequences (Table 1 and Fig. 5) were obtained by RT-PCR from all three cyanobacterial mats. This cluster of nifH represents several lineages of bacteria, including many anaerobes such as clostridia, sulfate reducers, methanogens, and green sulfur bacteria. None of the recovered sequences were very similar (83 to 89% identity) to any previously recovered isolates. The sequences were most identical (91 to 100% identity) to sequences obtained from identical microbial mats (26) or other habitats that contain anaerobic environments, such as sea grass bed communities (1). The identity of the organisms containing these nifH sequences is unclear. The sequences from isolates that were closest to any of the recovered sequences were from sulfate-reducing bacteria. Sequences very similar to those of sulfate-reducing bacteria have also been recovered from cyanobacterial mats in previous studies (26, 37). Recent work by Steppe et al. (36) suggested that nifH genes are expressed in sulfate-reducing bacteria and that N2 fixation in cyanobacterial mats does involve sulfate-reducing bacteria. However, it is also possible that the cluster 3 sequences recovered in this study correspond to multiple and as yet uncharacterized groups of bacteria that are not sulfate reducers.

The sequence GN2084A04 represents a novel and at present uncharacterized nifH. Since this sequence could not be ascribed to any particular nitrogenase cluster, it is unclear whether it even functions in N2 fixation. nifH sequences similar to this sequence (91%) have been recovered from similar microbial mats (26). It seems that this sequence does have a function in the environment, since it is actively expressed. This sequence includes a transcription stop codon in its cDNA sequence. Whether or not this stop codon is a result of nucleotide misincorporation during RT-PCR or a natural occurrence is unclear.

Over half of the sequence types recovered in this study were not found in nifH PCR clone libraries generated from genomic DNA on similar and identical mats from Guerrero Negro (26). Furthermore, in that study the most abundant sequences recovered were those belonging to cluster 3 organisms (63%), followed by cyanobacteria, which were almost a third less abundant (23%). In this study, cyanobacterial sequences were by far the most abundant sequences (78%), followed by those in cluster 3 (16%). These results suggest that less abundant members of the diazotroph community may have larger roles in N2 fixation than can be inferred from the abundance of their genes in a DNA-based clone library. However, these differences could also be a result of sampling, environmental, or extraction efficiency differences (26). This was most notable in the LG and GNM mats when these mats were not sampled for DNA and RNA simultaneously. Regardless of the reason for these differences, this study highlights the need to examine expression to determine which organisms are active in N2 fixation.

The patterns of acetylene reduction and the expression of nighttime nifH transcripts indicate that N2 fixation in these three cyanobacterial mats occurs on a diurnal pattern. Furthermore, the results of this study demonstrate that several groups of cyanobacteria, including unicellular and filamentous types, heterotrophic bacteria, and as yet uncharacterized groups of bacteria express nifH in cyanobacterial mats. The nifH phylotypes recovered in this study were very similar among the three mats studied, indicating that the same or similar organisms may be involved in N2 fixation in all three mats. In addition, several of the phylotypes recovered from this study clustered with phylotypes from different environments, such as Mono Lake, a North Carolina estuary (the Neuse River), the Pacific Ocean, and other microbial mats, which suggests that similar organisms may be involved in N2 fixation in other environments as well. The nifH phylotypes identified will provide targets for studies aimed at quantifying the role of diazotrophs in mats and other environments by approaches such as quantitative PCR and RT-PCR as well as DNA and cDNA arrays.


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ACKNOWLEDGMENTS
 
We thank Mary Hogan and Steven Carpenter for assistance with sampling. We also thank Dave Des Marais, Mykell Discipulo, and Moira Doty for logistical support. We are also grateful for continued access to the field site and for the logistical support provided by Exportadora de Sal, S.A. de C.V.

This work was funded by the NASA Exobiology Program and the NASA Astrobiology Institute.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Ocean Sciences and Institute of Marine Sciences, Earth and Marine Science Bldg., University of California, Santa Cruz, 1156 High St., Santa Cruz, CA 95064. Phone: (831) 459-4009. Fax: (831) 459-4882. E-mail: zehrj{at}ucsc.edu. Back


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REFERENCES
 
    1
  1. Bagwell, C. E., J. R. La Rocque, G. W. Smith, S. W. Polson, M. J. Friez, J. W. Longshore, and C. R. Lovell. 2002. Molecular diversity of diazotrophs in oligotrophic tropical seagrass bed communities. FEMS Microbiol. Ecol. 39:113-119.[CrossRef]
    2. 2
  2. Bebout, B., and F. Garcia-Pichel. 1995. UV B-induced vertical migrations of cyanobacteria in a microbial mat. Appl. Environ. Microbiol. 61:4215-4222.[Abstract]
    3. 3
  3. Bebout, B. M., S. P. Carpenter, D. J. Des Marais, M. K. Discipulo, T. Embaye, F. Garcia-Pichel, T. M. Hoehler, M. Hogan, L. L. Janke, R. M. Keller, S. R. Miller, L. E. Prufert, C. Raleigh, M. Rothrock, and K. Turk. 2002. Long term manipulations of intact microbial mat communities in a greenhouse collaboratory: Simulating earth's present and past field environments. Astrobiology 2:383-402.[CrossRef][Medline]
    4. 4
  4. Bebout, B. M., M. W. Fitzpatrick, and H. W. Paerl. 1993. Identification of the sources of energy for nitrogen fixation and physiological characterization of nitrogen-fixing members of a marine microbial mat community. Appl. Environ. Microbiol. 59:1495-1503.[Abstract/Free Full Text]
    5. 5
  5. Bebout, B. M., H. W. Paerl, J. E. Bauer, D. E. Canfield, and D. J. Des Marais. 1994. Nitrogen cycling in microbial mat communities: The quantitative importance of N-fixation and other sources of N for primary productivity, p. 265-278. In L. Stal and P. Caumette (ed.), Microbial mats. Springer-Verlag, Berlin, Germany.
    6. 6
  6. Bergman, B. B., J. R. Gallon, A. N. Rai, and L. J. Stal. 1997. N2 fixation by non-heterocystous cyanobacteria. FEMS Microbiol. Rev. 19:139-185.[CrossRef]
    7. 7
  7. 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.
    8. 8
  8. Chen, Y. B., B. Dominic, M. T. Mellon, and J. P. Zehr. 1998. Circadian rhythm of nitrogenase gene expression in the diazotrophic filamentous nonheterocystous cyanobacterium Trichodesmium sp. strain IMS 101. J. Bacteriol. 180:3598-3605.[Abstract/Free Full Text]
    9. 9
  9. Chien, Y.-T., and S. H. Zinder. 1996. Cloning, functional organization, transcript studies, and phylogenetic analysis of the complete nitrogenase structural genes (nifHDK2) and associated genes in the archaeon Methanosarcina barkeri 227. J. Bacteriol. 178:143-148.[Abstract/Free Full Text]
    10. 10
  10. Colon-Lopez, M. S., D. M. Sherman, and L. A. Sherman. 1997. Transcriptional and translational regulation of nitrogenase in light-dark- and continuous-light-grown cultures of the unicellular cyanobacterium Cyanothece sp. strain ATCC 51142. J. Bacteriol. 179:4319-4327.[Abstract/Free Full Text]
    11. 11
  11. Colon-Lopez, M. S., H. Tang, D. L. Tucker, and L. A. Sherman. 1999. Analysis of the nifHDK operon and structure of the nifH protein from the unicellular, diazotrophic cyanobacterium, Cyanothece strain sp. ATCC 51142. Biochim. Biophys. Acta 1473:363-375.
    12. 12
  12. Des Marais, D. J. 1995. The biogeochemistry of hypersaline microbial mats. Adv. Microb. Ecol. 14:251-274.
    13. 13
  13. Fernandez-Valiente, E., A. Quesada, C. Howard-Williams, and I. Hawes. 2001. N2-fixation in cyanobacterial mats from ponds on the McMurdo ice shelf, Antarctica. Microb. Ecol. 42:338-349.[CrossRef][Medline]
    14. 14
  14. Gallon, J. R. 1992. Tansley review no. 44/reconciling the incompatible: N2 fixation and O2. New Phytol. 122:571-609.[CrossRef]
    15. 15
  15. Garcia-Pichel, F., U. Nubel, and G. Muyzer. 1998. The phylogeny of unicellular, extremely halotolerant cyanobacteria. Arch. Microbiol. 169:469-482.[CrossRef][Medline]
    16. 16
  16. 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.[CrossRef][Medline]
    17. 17
  17. Howard, J. B., and D. C. Rees. 1996. Structural basis of biological nitrogen fixation. Chem. Rev. 96:2965-2982.[CrossRef][Medline]
    18. 18
  18. Howarth, R. W., and R. Marino. 1988. Nitrogen fixation in freshwater, estuarine, and marine ecosystems. 2. Biogeochemical controls. Limnol. Oceanogr. 32:688-701.
    19. 19
  19. Huang, T. C., R. F. Lin, M. K. Chu, and H. M. Chen. 1999. Organization and expression of nitrogen-fixation genes in the aerobic nitrogen-fixing unicellular cyanobacterium Synechococcus sp. strain RF-1. Microbiology 145:743-753.
    20. 20
  20. Jeanthon, C. 2000. Molecular ecology of hydrothermal vent microbial communities. Antonie van Leeuwenhoek 77:117-133.[CrossRef][Medline]
    21. 21
  21. Misra, H. S., and R. Tuli. 2000. Differential expression of photosynthesis and nitrogen fixation genes in the cyanobacterium Plectonema boryanum. Plant Physiol. 122:731-736.[Abstract/Free Full Text]
    22. 22
  22. Moorhead, D. L., C. F. Wolf, and R. A. Wharton, Jr. 1997. Impact of light regimes on productivity patterns of benthic microbial mats in an antarctic lake: a modeling study. Limnol. Oceanogr. 42:1561-1569.
    23. 23
  23. Noda, S., M. Ohkuma, R. Usami, K. Horikoshi, and T. Kudo. 1999. Culture-independent characterization of a gene responsible for nitrogen fixation in the symbiotic microbial community in the gut of the termite Neotermes koshunensis. Appl. Environ. Microbiol. 65:4935-4942.[Abstract/Free Full Text]
    24. 24
  24. Nold, S. C., and D. M. Ward. 1995. Diverse Thermus species inhabit a single hot spring microbial mat. Syst. Appl. Microbiol. 18:274-278.[Medline]
    25. 25
  25. Nubel, U., F. Garcia-Pichel, E. Clavero, and G. Muyzer. 2000. Matching molecular diversity and ecophysiology of benthic cyanobacteria and diatoms in communities along a salinity gradient. Environ. Microbiol. 2:217-226.[CrossRef][Medline]
    26. 26
  26. Omoregie, E. O., L. L. Crumbliss, B. M. Bebout, and J. P. Zehr. 2004. Comparison of diazotroph community structure in Lyngbya sp. and Microcoleus chthonoplastes dominated microbial mats from Guerrero Negro, Baja, Mexico. FEMS Microbiol. Ecol. 47:305-318.
    27. 27
  27. Paerl, H. W. 1990. Physiological ecology and regulation of N2 fixation in natural waters. Adv. Microb. Ecol. 8:305-344.
    28. 28
  28. Paerl, H. W., B. M. Bebout, C. A. Currin, M. Fitzpatrick, and J. Pinckney. 1994. Nitrogen fixation dynamics in microbial mats, p. 323-337. In P. Caumette (ed.), Microbial mats. Springer-Verlag, Berlin, Germany.
    29. 29
  29. Paerl, H. W., K. M. Crocker, and L. E. Prufert. 1987. Limitation of N2 fixation in coastal marine waters: Relative importance of molybdenum, iron, phosphorus, and organic matter availability. Limnol. Oceanogr. 31:525-536.
    30. 30
  30. Paerl, H. W., M. Fitzpatrick, and B. M. Bebout. 1996. Seasonal nitrogen fixation dynamics in a marine microbial mat: Potential roles of cyanobacteria and microheterotrophs. Limnol. Oceanogr. 41:419-427.
    31. 31
  31. Pinckney, J., and H. Paerl. 1997. Anoxygenic photosynthesis and nitrogen fixation by a microbial mat community in a Bahamian hypersaline lagoon. Appl. Environ. Microbiol. 63:420-426.[Abstract]
    32. 32
  32. Postgate, J. 1998. Nitrogen fixation, 3rd ed. University Press, Cambridge, England.
    33. 33
  33. Postgate, J. R., and R. R. Eady. 1988. The evolution of biological nitrogen fixation, p. 31-40. In H. Bothe, F. J. de Bruijn, and W. E. Newton (ed.), Nitrogen fixation: hundred years after. Gustav Fischer, Stuttgart, Germany.
    34. 34
  34. Saitou, N., and M. Nei. 1987. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol. Biol. E vol. 4:406-425.
    35. 35
  35. Stal, L. J. 1995. Physiological ecology of cyanobacteria in microbial mats and other communities. New Phytol. 131:1-31.
    36. 36
  36. Steppe, T. F., and H. W. Paerl. 2002. Potential N2-fixation by sulfate-reducing bacteria in a marine intertidal microbial mat. Aquat. Microb. Ecol. 27:1-12.
    37. 37
  37. Steppe, T. F., J. L. Pinckney, J. Dyble, and H. W. Paerl. 2001. Diazotrophy in modern marine Bahamian stromatolites. Microb. Ecol. 40:35-44.
    38. 38
  38. Stewart, W. D. P., G. P. Fitzgerald, and R. H. Burris. 1967. In situ studies of N2 fixation using the acetylene reduction technique. Proc. Natl. Acad. Sci. USA 58:2071-2078.[Free Full Text]
    39. 39
  39. Tajima, F., and M. Nei. 1984. Estimation of evolutionary distance between nucleotide sequences. Mol. Biol. E vol. 1:269-285.
    40. 40
  40. Thiel, T. 1993. Characterization of genes for an alternative nitrogenase in the cyanobacterium Anabaena variabilis. J. Bacteriol. 175:6276-6286.[Abstract/Free Full Text]
    41. 41
  41. Thiel, T., E. M. Lyons, J. C. Erker, and A. Ernst. 1995. A second nitrogenase in vegetative cells of a heterocyst-forming cyanobacterium. Proc. Natl. Acad. Sci. USA 92:9358-9362.[Abstract/Free Full Text]
    42. 42
  42. Tillett, D., and B. A. Neilan. 2000. Xanthogenate nucleic acid isolation from cultured and environmental cyanobacteria. J. Phycol. 36:251-258.[CrossRef]
    43. 43
  43. Van de Peer, Y., and R. De Wachter. 1994. TREECON for windows: A software package for the construction and drawing of evolutionary trees for the Microsoft Windows environment. Comput. Appl. Biosci. 10:569-570.[Free Full Text]
    44. 44
  44. Vitousek, P. M., and R. W. Howarth. 1991. Nitrogen limitation on land and in the sea: How can it occur? Biogeochemistry 13:87-115.
    45. 45
  45. Wahlund, T. M., and M. T. Madigan. 1993. Nitrogen fixation by the thermophilic green sulfur bacterium Chlorobium tepidum. J. Bacteriol. 175:474-478.[Abstract/Free Full Text]
    46. 46
  46. Young, J. P. W. 1992. Phylogenetic classification of nitrogen-fixing organisms, p. 43-86. In G. Stacey, H. J. Evans, and R. H. Burris (ed.), Biological nitrogen fixation. Chapman and Hall, New York, N.Y.
    47. 47
  47. Zani, S., M. T. Mellon, J. L. Collier, and J. P. Zehr. 2000. Expression of nifH genes in natural microbial assemblages in Lake George, N.Y., detected with RT-PCR. Appl. Environ. Microbiol. 66:3119-3124.[Abstract/Free Full Text]
    48. 48
  48. Zehr, J. P., M. Mellon, S. Braun, W. Litaker, T. Steppe, and H. W. Paerl. 1995. Diversity of heterotrophic nitrogen fixation genes in a marine cyanobacterial mat. Appl. Environ. Microbiol. 61:2527-2532.[Abstract]
    49. 49
  49. Zehr, J. P., M. T. Mellon, and S. Zani. 1998. New nitrogen-fixing microorganisms detected in oligotrophic oceans by the amplification of nitrogenase (nifH) genes. Appl. Environ. Microbiol. 64:3444-3450.[Abstract/Free Full Text]
    50. 50
  50. Zehr, J. P., and P. J. Turner. 2001. Nitrogen fixation: Nitrogenase genes and gene expression, p. 271-286. In J. H. Paul (ed.), Methods in marine microbiology. Academic Press, San Diego, Calif.
    51. 51
  51. Zehr, J. P., J. B. Waterbury, P. J. Turner, J. P. Montoya, E. Omoregie, G. F. Steward, A. Hansen, and D. M. Karl. 2001. Unicellular cyanobacteria fix N2 in the subtropical North Pacific Ocean. Nature 412:635-638.[CrossRef][Medline]

Applied and Environmental Microbiology, April 2004, p. 2119-2128, Vol. 70, No. 4
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.4.2119-2128.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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