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Applied and Environmental Microbiology, May 2002, p. 2236-2245, Vol. 68, No. 5
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.5.2236-2245.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Characterization of Trichodesmium spp. by Genetic Techniques

K. M. Orcutt,1* U. Rasmussen,1 E. A. Webb,2 J. B. Waterbury,2 K. Gundersen,1,{dagger} and B. Bergman1

Department of Botany, Stockholm University, S-106 91 Stockholm, Sweden,1 Department of Biology, Woods Hole Oceanographic Institute, Woods Hole, Massachusetts 025432

Received 11 September 2001/ Accepted 7 February 2002


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ABSTRACT
 
The genetic diversity of Trichodesmium spp. from natural populations (off Bermuda in the Sargasso Sea and off North Australia in the Arafura and Coral Seas) and of culture isolates from two regions (Sargasso Sea and Indian Ocean) was investigated. Three independent techniques were used, including a DNA fingerprinting method based on a highly iterated palindrome (HIP1), denaturing gradient gel electrophoresis of a hetR fragment, and sequencing of the internal transcribed spacer (ITS) of the 16S-23S rDNA region. Low genetic diversity was observed in natural populations of Trichodesmium spp. from the two hemispheres. Culture isolates of Trichodesmium thiebautii, Trichodesmium hildebrandtii, Trichodesmium tenue, and Katagnymene spiralis displayed remarkable similarity when these techniques were used, suggesting that K. spiralis is very closely related to the genus Trichodesmium. The largest genetic variation was found between Trichodesmium erythraeum and all other species of Trichodesmium, including a species of Katagnymene. Our data obtained with all three techniques suggest that there are two major clades of Trichodesmium spp. The HIP1 fingerprinting and ITS sequence analyses allowed the closely related species to be distinguished. This is the first report of the presence of HIP1 in marine cyanobacteria.


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INTRODUCTION
 
The genus Trichodesmium is a group of globally significant marine diazotrophic cyanobacteria that contribute up to 80 Tg of new N input into oligotrophic waters per year (7, 21). They are cosmopolitan in tropical and subtropical oceans and can form massive surface blooms (6, 20) large enough to be seen from the space shuttle (21). Trichodesmium thiebautii and Trichodesmium erythraeum are the most frequently reported species associated with blooms in the open ocean.

It has been observed that oceanic N2-fixing filamentous cyanobacteria have lower diversity than filamentous cyanobacteria found in coastal areas, freshwater, and terrestrial systems. With the notable exceptions of Nostoc, Anabaena, the endosymbiont Richelia intracellularis (8, 9, 37), and molecular evidence for heterocystous cyanobacteria at the Bermuda Atlantic Time-series Study (BATS) site (44), there are few heterocystous cyanobacteria in the open ocean relative to the numbers in brackish and freshwater systems. Additionally, there are few genera of nonheterocystous filamentous cyanobacteria in the open ocean, where the most notable such cyanobacteria are members of the genera Trichodesmium and Katagnymene (5).

Wille (40) described four species of Trichodesmium based on morphology (T. contortum, T. thiebautii, T. tenue, and T. erythraeum) and two species of Katagnymene (K. spiralis and K. pelagica) obtained on a cruise in the Atlantic Ocean. Later, Sournia (36) described one additional species, Trichodesmium hildebrandtii, and reluctantly placed T. tenue with T. thiebautii due to the large variation in filament size of the latter species (40). More recently, identification of Trichodesmium species based on ultrastructural and morphological characterization resulted in identification of five species of Trichodesmium, T. thiebautii, T. erythaeum, T. tenue, T. contortum, and T. hildebrandtii (19). In the latter study, species of Trichodesmium were found to be morphologically similar regardless of geographical origin.

Sequence analyses of nifH, hetR, and regions of the small-subunit rRNA (16S rRNA) have revealed very low genetic diversity among the species of Trichodesmium (4, 18, 42). The similarities of nifH sequences from other cyanobacterial species in a genus typically range from 92 to 95%, whereas T. thiebautii and T. erythraeum nifH sequences were found to be 98% identical (4, 42). Recently, Janson et al. (18) found Trichodesmium spp. to be very similar using 16S ribosomal DNA (rDNA) and hetR sequences and were able to resolve three clades containing (i) T. thiebautii and T. hildebrandtii, (ii) T. contortum and T. tenue, and (iii) T. erythraeum. In the latter study, the hetR sequence was more variable and gave better resolution between the species T. erythraeum and T. thiebautii than analysis of the 16S rDNA region gave.

Although the use of hetR in the study of Janson et al. (18) resulted in lower sequence similarity between T. erythraeum and T. thiebautii, the similarities between closely related species, such as T. thiebautii and T. hildebrandtii, were high (98%). In order to investigate the genetic diversity of the closely related species of Trichodesmium, genetic techniques that provide high resolution are required. In this study, we investigated the genetic diversity of Trichodesmium spp. by using three independent techniques that provide high resolution in order to resolve members of this closely related genus and to investigate the genetic diversity of natural populations. A PCR-based DNA fingerprinting method using base pair extended short oligonucleotide primers for HIP1 was used to distinguish Trichodesmium spp. and to investigate the genetic diversity within and between species. HIP1 is an octameric repetitive sequence found in many but not all cyanobacteria (33) and has been used in DNA fingerprinting to resolve cyanobacterial species and strains (35). The diversity of trichomes within single Trichodesmium colonies from natural populations and a comparison of culture isolates were examined by using denaturing gradient gel electrophoresis (DGGE) analysis of a fragment of the hetR gene. The sequence of the internal transcribed spacer (ITS) region between the 16S and 23S rRNA genes was determined to study inter- and intraspecific variability. In most bacteria the 5S, 16S, and 23S rRNA genes are separated by ITS regions. The spacer region between the 16S and 23S rRNA genes can encode zero, one, or two tRNA genes and can exhibit considerable length and sequence variation. Due to this variation analysis of the ITS region has been widely applied to many bacterial groups (2, 12, 17, 22) and cyanobacteria (24) in order to resolve closely related species.


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MATERIALS AND METHODS
 
Collection of Trichodesmium spp.
Natural populations of Trichodesmium colonies were collected in 1999 from the BATS site (31°50'N, 64°10'W) off Bermuda and during a research cruise off North Australia between Darwin and Townsville. Colonies were picked randomly and separated into T. thiebautii tufts (colonies with trichomes arranged in parallel), T. thiebautii puffs (colonies with trichomes arranged radially), and the easily recognized T. erythraeum colonies (colonies that are darker reddish brown and have trichomes arranged in parallel like a raft). The colonies were rinsed in filtered seawater, placed on 0.5-µm-pore-size Fluorophore FHLP filters (Millipore, Bedford, Mass.), and frozen immediately in sterile Eppendorf tubes. The samples were transported on dry ice and stored at -20°C until they were used in the PCR. Culture isolates of Trichodesmium spp. were collected from the Sargasso Sea off Bermuda (BATS) and from the West Indian Ocean off Zanzibar (East Africa). An isolate of K. spiralis (KAT) was also collected off Zanzibar. All isolates of Trichodesmium spp. and Katagnymene were identified by using the morphological descriptions of Janson et al. (19) and Geitler (15) and are listed in Table 1. A culture strain of T. erythraeum (BE) was isolated from the BATS site by placing several colonies into a 50:50 mixture of Whatman GF/F-filtered Sargasso seawater and YBC11 medium (11). Cultured Trichodesmium spp. used in the HIP1 fingerprinting and DGGE analyses were maintained at the Department of Botany, Stockholm University, Stockholm, Sweden, on YBC11 medium (11) in a climate chamber under the following conditions: daily cycle consisting of 12 h of light (52 microeinsteins m-2 s-1) and 12 h of darkness, 28°C, and 65% humidity. All other Trichodesmium spp. and K. spiralis strain KAT were isolated from single trichomes. These cultured isolates were maintained at the Department of Biology, Woods Hole Oceanographic Institution, Woods Hole, Mass., in a medium made of 0.75x Sargasso seawater filtered successively through 1.0- and 0.2-µm-pore-size Millipore membrane filters before it was Tyndalized by heating in a microwave oven to boiling in Teflon containers. The Sargasso seawater was diluted with steam-sterilized Millipore Q water. All chemicals added to the medium were tissue culture tested and purchased from Sigma. Concentrated stocks (1,000x) were sterilized by either filtration or steam and added to the 0.75x seawater base. The chemicals added included 1.5 µM EDTA, 8 µM phosphoric acid, 0.05 µM Fe (ferric citrate), 0.1 µM MnSO4, 0.01 µM ZnCl2, 0.01 µM NaMoO4, 0.0001 µM CoCl2, 0.0001 µM NiCl2, 0.0001 µM NaSeO3, and 1.5 µg of vitamin B12 per liter. All Trichodesmium cultures were grown in Nalgene polycarbonate flasks. The growth conditions were typically a cycle consisting of 14 h of light (30 to 300 microeinsteins m-2 s-1) and 10 h of darkness and temperatures ranging from 24 to 28°C. The small-volume cultures were gently shaken daily.


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  		TABLE 1. Trichodesmium spp. and Katagnymene sp. isolates used in the genetic analysis

HIP1 fingerprinting.
The sequences of the oligonucleotide primers used for the PCR fingerprinting were based on the HIP1 consensus sequence 5'-GCGATCGC-3' with a 2-bp extension at the 3' end as described by Smith et al. (35). Extended HIP1 primers (HIP-AT, HIP-CA, HIP-GC, and HIP-TG) were used as either single primers or in combinations of two primers. The primers were synthesized by DNA Technology A/S (Aarhus, Denmark). The PCR samples were run in 25-µl mixtures containing a colony of field-collected Trichodesmium that was placed on a triangular piece of a 0.5-µm-pore-size Fluorophore FHLP filter (Millipore) (3). For the cultured Trichodesmium and Katagnymene strains, a 1-µl template was made from either one colony dispersed in sterile double-distilled freshwater or 2 to 5 ml of a dense culture filtered onto a 13-mm-diameter polycarbonate filter and dispersed in 20 to 50 µl of sterile double-distilled freshwater. Each reaction mixture contained a deoxynucleoside triphosphate mixture (Finnzymes OY, Espoo, Finland) (40 nM each), buffer (1/10 of the appropriate 10x buffer supplied by the manufacturer), 1.0 U of DyNAzyme DNA polymerase (Finnzymes OY), and primers (50 pmol each). All PCR samples were overlaid with sterile mineral oil. The PCR conditions were similar to those described by Smith et al. (35); reaction mixtures were cycled with a Perkin-Elmer Cetus DNA thermal cycler 480 using a temperature profile of 95°C for 5 min, 30 cycles of 95°C for 30 s, 30°C for 30 s, and 72°C for 60 s, and finally one cycle of 72°C for 5 min. Aliquots (12 µl) of each amplified PCR product were separated on a 1.5% agarose gel, visualized with ethidium bromide, and photographed under UV light using a Kodak DC 120 digital camera and Kodak Digital Science 1D Analysis software. A pGEM DNA marker (Promega, Madison, Wis.) was included on the agarose gel. At least 20 individual colonies of each species (T. thiebautii and T. erythraeum) and morphology (T. thiebautii puffs and tufts) were analyzed from both cruises (off Bermuda and off North Australia), and the cultured isolates were each examined with the PCR at least five times.

The specificity of the HIP1 primers was tested by using DNA templates from strains known to not contain HIP1 repeats (Prochlorococcus marinus strain MED4, Synechococcus sp. strain 8103, and Synechococcus sp. strain 7803) as a negative control. No PCR products were generated from these strains.

PCR amplification of hetR.
The forward primer used in the PCR was hetR1 (5'-AARTGYGCNATHTAYATGAC-3') described by Janson et al. (18) with an additional 39-bp GC clamp at the 5' end. The reverse primer (5'-GCATCAGGCATARTTGAAGGA-3') was designed to generate a 272-bp fragment by alignment of hetR genes from Trichodesmium, using the GenBank accession numbers reported by Janson et al. (18). Each reaction mixture contained a 2-µl template treated as described above for the HIP1 PCR, a deoxynucleoside triphosphate mixture (Finnzymes OY) (20 nM each), buffer (1/10 of the appropriate 10x buffer supplied by the manufacturer), 1.0 U of DyNAzyme DNA polymerase (Finnzymes OY), and primers (100 pmol each) and was overlaid with sterile mineral oil. The PCR conditions were those described by Janson et al. (18); reactions were cycled with a Perkin-Elmer Cetus DNA thermal cycler 480 using a temperature profile of 95°C for 6 min, 30 cycles of 93°C for 60 s, 52°C for 60 s, and 70°C for 60 s, and finally one cycle of 70°C for 10 min.

Analysis of PCR products by DGGE.
DGGE was performed by using the Dcode universal mutation detection system as described in the manufacturer's manual (Bio-Rad, Hercules, Calif.). Aliquots (15 µl) of each PCR product were combined with 15 µl of loading buffer (0.05% bromophenol blue, 0.05% xylene cyanol, 70% glycerol) and applied directly onto 6% (wt/vol) polyacrylamide gels in 0.5x TAE buffer (0.02 M Tris base, 0.01 M acetic acid, 0.5 mM EDTA; pH 8.0) with a linear 20 to 45% denaturant gradient (100% denaturant was defined as 7 M urea and 40% [vol/vol] formamide). Gel electrophoresis of cultured strains (T. tenue Z-1, T. hildebrandtii #11 and II-4, and K. spiralis KAT) and natural samples (T. thiebautii puffs and tufts) was performed at 60°C (constant temperature) for 6.5 h at 150 V; the T. erythraeum strains (IMS 101, BE, and Z-9) and natural samples of T. erythraeum were electrophoresed under the same conditions for 24 h at 150 V. For quality control in the DGGE analysis, Nostoc sp. strains PCC 9229 and PCC 9231 were included to ensure that denaturation had occurred (32). After electrophoresis the gel was stained for 10 min with ethidium bromide and photographed as described above.

Sequencing of the ITS region.
A high-fidelity PCR protocol with the Pfu polymerase (Promega) was used to amplify the ITS region from the following culture isolates: T. erythraeum strains IMS 101 and Z-9, T. tenue strain Z-1, T. hildebrandtii strains II-4 and #11, T. thiebautii strain II-3, and K. spiralis strain KAT (Table 1). The DNA sequence of the ITS was obtained by using some modifications of the method described by Rocap et al. (34). The cultured cells (3 x 106 cells ml-1) were filtered onto 5-µm-pore-size polycarbonate membranes to dryness, resuspended in 100 µl of sterile double-distilled freshwater, heated at 95°C for 10 min, and centrifuged in an Eppendorf microfuge at 14,000 rpm for 5 min. The supernatant fraction was used as the template for high-fidelity PCR with Pfu polymerase and primers tri16S-1247F (5'-CGTACTACAATGGTTGGG-3') and 23S-241R (5'-TTCGCTCGCCRCTACT-3'). Primer tri16S-1247F was designed to be specific for filamentous nonheterocystous cyanobacteria (Trichodesmium and Oscillatoria) by using the Ribosomal Database Project check probe functionality (26). The reverse primer 23S-241 sequence was taken from the study of Rocap et al. (34), and only one band was obtained for each strain. Amplified products from two independent reaction mixtures were purified from the agarose gel by using a Qiaquick gel extraction kit (Qiagen, Valencia, Calif.) and were sequenced entirely on both strands at the University of Maine DNA Sequencing Center. Two additional primers, AlaF (5'-TWTAGCTCAGTTGGTAGAG-3') and AlaR (5'-CTCTACCAACTGAGCTAWA-3'), were used in order to get complete coverage of the ITS region (34).

Analysis of ITS sequences.
The DNA sequences were aligned by using Macvector (Oxford Molecular, Oxford, United Kingdom) and ClustalX (39) software. The evolutionary relationships of the cultures were determined with PAUP (38) by using optimality settings of distance, maximum likelihood, and maximum parsimony. In these analyses the insertions and deletions were treated as missing data. The HKY85+gamma model was used for the likelihood analyses, while LogDet was used for neighbor joining. The gamma parameter and nucleotide frequency were determined empirically. Bootstrap values were generated for all three methods by using 1,000 replicates. Similar relationships were inferred for the remaining Trichodesmium species when the three T. erythraeum strains were omitted.

Nucleotide sequence accession numbers.
The GenBank accession numbers for the sequences determined in this study are as follows: Trichodesmium strain Z-9, AF399646; Trichodesmium strain IMS 101, AF399647; Trichodesmium strain II-3, AF399648; Trichodesmium strain #11, AF399649; Trichodesmium strain II-4, AF399650; Trichodesmium strain Z-1, AF399651; and Katagnymene strain KAT, AF399652.


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RESULTS
 
HIP1 fingerprinting.
Each of the four HIP1 extended primers was tested for its ability to distinguish between cyanobacterial cultures of the different species. The DNA of Trichodesmium spp. contain large numbers of HIP1 sequences that produced smearing banding patterns when nonextended HIP1 primers were used (data not shown). Therefore, both single extended primers and two extended primers in combination were tested (Fig. 1). The single HIP-TG extended primer gave distinct fingerprints for T. erythraeum strain BE and T. thiebautii strain II-3 but with few products that were similar in size in both species (Fig. 1). In order to optimize the fingerprint resolution by producing more complex banding patterns, double-primer combinations were used (Fig. 1). The combination HIP-GC and HIP-CA gave the highest resolution and was able to distinguish between species in culture (Fig. 2). Cultured isolates of T. thiebautii strain II-3, T. hildebrandtii strains II-4 and #11 , T. tenue strain Z-1, and K. spiralis strain KAT displayed fingerprints that were similar but had banding patterns that were distinguishable from each other. T. erythraeum strains IMS 101 and Z-9 displayed a pattern that was distinctly different from the other patterns (Fig. 2). However, within the T. erythraeum species group it was possible to distinguish the HIP1 patterns of strains IMS 101 and Z-9 (Fig. 2). In contrast, the two strains of T. hildebrandtii (#11 and II-4) collected 4 years apart from the Sargasso Sea had virtually identical fingerprint patterns (Fig. 2).



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  		FIG. 1. HIP1 fingerprints obtained with different extended primers, including a single primer (HIP-TG [TG]) and combinations of two primers (HIP-AT and HIP-CA [AT/CA], HIP-TG and HIP-CA [TG/CA]), for culture isolates of T. erythraeum strain BE (lanes 1) and T. thiebautii strain II-3 (lanes 2). Lane M contained DNA molecular weight standards.



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  		FIG. 2. Fingerprint patterns of culture isolates of Trichodesmium spp. and Katagnymene sp. obtained with a combination of primers HIP-GC and HIP-CA. The results for two replicate samples of each strain are shown, and lanes M contained DNA molecular weight standards.

Natural populations of T. thiebautii puffs and tufts collected off Bermuda at the BATS site and off North Australia gave nearly identical fingerprint patterns in the 222- to 460-bp region (Fig. 3). Morphologically different T. thiebautii colonies had similar fingerprints, even though there were differences in trichome width within a colony (range, 4 to 10 µm) or in the arrangement of trichomes as puffs or tufts (Fig. 3). In addition, the cultured T. thiebautii strain II-3 puffs gave a fingerprint identical to that of the natural populations of T. thiebautii puffs and tufts (Fig. 3). Natural populations of T. erythraeum from both hemispheres had similar banding patterns in the 176- to 679-bp region (Fig. 4). However, cultured isolates of T. erythraeum strains IMS 101, BE, and Z-5 showed small genetic differences compared to each other (Fig. 4).



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  		FIG. 3. Fingerprint patterns obtained with a combination of primers HIP-GC and HIP-CA for T. thiebautii puffs and tufts collected at the BATS site near Bermuda (Bda) and in the Arafura Sea off North Australia (Aus). For each group the results for 3 of a minimum of 20 samples analyzed are shown. A culture of T. thiebauttii strain II-3 was used for comparison. The arrows show the range of the primary bands used for pattern identification, and lanes M contained DNA molecular weight standards.



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  		FIG. 4. Fingerprint patterns obtained with a combination of primers HIP-GC and HIP-CA for natural populations of T. erythraeum collected at the BATS site (Bda) and off North Australia (Aus). Culture isolates of T. erythraeum strain IMS 101 (1) and T. erythraeum strain BE (2), both from the North Atlantic, as well as T. erythraeum strain Z-5 (3) collected off Zanzibar, were included for comparison. For each station the results for 6 of a minimum of 20 samples analyzed are shown. The arrows show the range of the primary bands used for pattern identification, and lanes M contained DNA molecular weight standards.

Analysis of the hetR fragment by DGGE.
DGGE analysis of the PCR-amplified 272-bp hetR fragment was performed for both natural populations and cultured isolates. The hetR fragments from the heterocystous cyanobacteria Nostoc strains PCC 9229 and PCC 9231 were included as controls in the DGGE analysis to ensure that denaturation occurred. The amplified hetR fragments from the T. thiebautii tufts collected from natural populations off Bermuda at the BATS site and off North Australia were all identical to each other (Fig. 5A), as were the amplified hetR fragments from the morphologically distinctive T. thiebautii puffs and tufts from the BATS site (Fig. 5B). All of the DGGE analyses of colonies from natural populations showed that individual colonies contained trichomes that were similar, forming one distinct band on the gel (Fig. 5A and B). The cultures of Trichodesmium isolates, including T. thiebautii strain II-3, T. hildebrandtii strains #11 and II-4, T. tenue strain Z-1, and K. spiralis strain KAT, exhibited small differences in the hetR fragment compared to the fragment of the T. erythraeum strains, which showed the largest difference in mobility (Fig. 5C to E). The hetR fragments of cultured T. thiebautii strain II-3 and T. hildebrandtii strains #11 and II-4 had migration patterns similar to each other and could not be distinguished easily (Fig. 5C). Similarly, the hetR fragments of T. tenue strain Z-1 and K. spiralis strain KAT migrated close to each other on the DGGE gel (Fig. 5D). The largest difference observed between cultured species was that separating T. erythraeum strains IMS 101, BE, and Z-9 from the group encompassing T. thiebautii strain II-3, T. hildebrandtii strains #11 and II-4, T. tenue strain Z-1, and K. spiralis strain KAT. The hetR fragment of T. erythraeum denatured higher in the gel on a lower gradient compared to the hetR fragment of the T. thiebautii group and required a melting time as long as 24 h to resolve a single band in the DGGE analysis (Fig. 5E). The hetR fragments from natural populations of T. erythraeum obtained from the BATS site and off North Australia comigrated (data not shown), and the hetR fragments from T. erythraeum strains IMS 101, BE, and Z-9 from different geographical areas (Table 1) were found to have similar mobilities (Fig. 5E).



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  		FIG. 5. DGGE analysis of the hetR fragment from natural populations of T. thiebautii tufts collected at the BATS site (Bda) and off North Australia (Aus) (A); natural populations of T. thiebautii puffs and tufts collected at the BATS site and Nostoc strains PCC 9229 and PCC 9231 (B); culture isolates of T. hildebrandtii strains II-4 and #11, K. spiralis strain KAT, and T. thiebautii tuft strain II-3 (C); cultured isolates of K. spiralis strain KAT, T. tenue strain Z-1, and T. hildebandtii strain II-4 (D); and cultured isolates of T. erythraeum strains IMS 101, BE, and Z-9 (E).

Analysis of the ITS region.
Six ITS sequences were determined from cultured isolates of Trichodesmium spp., and one ITS sequence was determined from Katagnymene sp. strain KAT (Fig. 6). These sequences did not contain the duplications seen in cultured strain NIBB 1067 analyzed by Wilmotte et al. (41). All of the cultured Trichodesmium strains in this study and Katagnymene sp. strain KAT showed high sequence similarity (Fig. 6). Strains of T. erythraeum, including isolates from the Atlantic (IMS 101), Pacific (NIBB 1067), and Indian (Z-9) oceans, were readily separated from T. thiebautii strain II-3, T. hildebrandtii strains #11 and II-4, T. tenue strain Z-1, and K. spiralis strain KAT (Fig. 7). The members of the group containing T. thiebautii strain II-3, T. hildebrandtii strains #11 and II-4, T. tenue strain Z-1, and K. spiralis strain KAT were all very similar. K. spiralis strain KAT was most similar to T. tenue strain Z-1. However, all isolates sequenced in this study could be distinguished from each other (Fig. 6 and 7).



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  		FIG. 6. Alignment of ITS sequences (16S-23S rDNA). The sequences were aligned by using ClustalW in the MacVector program (Oxford Molecular). Conserved bases are shaded, and tRNAIle and tRNAAla are depicted as boxes below the sequences (shaded and cross-hatched, respectively). NIBB 1067 (41), Z-9, and IMS 101 are all strains of T. erythraeum, while #11 and II-4 are strains of T. hildebrandtii. Strains Z-1, KAT, and II-3 are strains of T. tenue, K. spiralis, and T. thiebautii, respectively.



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  		FIG. 7. Evolutionary relationships inferred from whole Trichodesmium ITS sequences. The consensus tree shown was constructed by using maximum-likelihood analysis and neighbor joining. Similar relationships were obtained from distance and maximum-parsimony analyses. Bootstrap values were generated from 1,000 replicates, and only values greater than 50 are shown at the nodes.


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DISCUSSION
 
Our results strongly support the results of other studies that found Trichodesmium spp. to be closely related (4, 18, 42). Genetic characterization of Trichodesmium spp. based on HIP1 fingerprinting, DGGE analysis of the hetR fragment, and ITS sequencing showed that Trichodesmium spp. cluster into two major groups. This suggests that two distinct clades are present in the oceans, one including the closely related species T. thiebautii, T. tenue, T. hildebrandtii, and K. spiralis and the other containing only T. erythraeum (Fig. 7).

In particular, our ITS data indicate that the genus Trichodesmium is composed of a closely related group of species (Fig. 6 and 7). The conservation of length and sequence variation within the ITS of the strains examined so far is quite remarkable, but nonetheless differences appear to be sufficient to be able to delineate species clusters. In contrast to the hetR sequence analysis described by Janson et al. (18), it appears that the ITS sequences are able to distinguish T. thiebautii strain II-3 from T. hildebrandtii strains #11 and II-4.

We have demonstrated that DNA fingerprinting using the repetitive sequence of HIP1 combined with ITS sequence analysis provides sufficient resolution to identify closely related species in the genus Trichodesmium. However, DGGE analysis of the hetR fragments could not resolve the small differences between closely related species, such as T. thiebautii (strain II-3) and T. hildebrandtii (strains #11 and II-4). Therefore, the DGGE technique could only determine that trichomes within an individual colony were not a mixture of members of the two clades of Trichodesmium spp. in natural populations.

The use of HIP1 fingerprinting provided sufficient resolution in an analysis of the genetic diversity of natural populations of Trichodesmium from two distant geographical locations (Fig. 3 and 4). Although whole colonies from the natural environment can be used directly as templates in the PCR, variation in template size (variable colony size) can affect the amplification efficiency of some PCR products. For example, we experienced incomplete amplification of large PCR products when small colonies were used (Fig. 3 and 4). Smaller T. erythraeum colonies were found off Bermuda at the BATS site than in the waters off North Australia, and this may have resulted in a lower efficiency of amplification of large PCR products from the BATS samples (Fig. 4). The presence of polysaccharides that are common in some cyanobacteria may also cause inhibition in the PCR (35). The Fluorophore filters were useful for collecting and transporting Trichodesmium colonies from the field to the laboratory and can be used directly in the PCR (3). Finally, due to the large number of products obtained with HIP1 primers, any folding of the DNA template prevents primers from annealing and results in faint bands or a loss of bands in the DNA fingerprints. Therefore, multiple samples are recommended for any investigation of natural populations.

The fact that HIP1 is present in the Trichodesmium genome is an interesting result in itself. HIP1 sequences are involved in adaptive response and were first identified in cells of Cd-tolerant Synechococcus strain PCC 6301, where they traversed an excised region of the metallothionein locus (16). HIP1 sequences are unique among repetitive sequences in that they are abundant in protein-encoding regions of the genome (33). No HIP1 sequences have been identified in the genomes of marine Synechococcus sp. strain WH8102 and Prochlorococcus sp. strain MED4 (27), and it was suggested that marine cyanobacteria might not contain HIP1 sequences due to the homeostatic environment of the open ocean (10). To our knowledge, this is the first report of the presence of HIP1 sequences in a marine cyanobacterium. Therefore, the possible role of HIP1 sequences in the adaptive response and genome plasticity of Trichodesmium spp. will be of great interest in future studies.

Genetic comparisons of bacterioplankton from the Atlantic and Pacific oceans have shown that certain microbial species are widely distributed in subtropical oceans (29). For example, high- and low-light-adapted Prochlorococcus isolates from two different regions of the North Atlantic were found to have high sequence similarity in their 16S rRNA genes (28). Molecular evidence also suggests that transequatorial mixing of Arctic and Antarctic populations of foraminiferans occurs (13). Our data based on HIP1 fingerprinting indicate that members of the genus Trichodesmium have low genetic diversity in natural populations and that individual species may have a global distribution.

The cultured isolates of T. erythraeum strain BE from the BATS site, T. erythraeum strains Z-5 and Z-9 from the West Indian Ocean, and T. erythraeum strain IMS 101 from Gulf Stream waters off the Carolinas exhibited similar primary banding patterns but also slight differences (Fig. 2 and 4). These results could indicate that there are several subspecies within T. erythraeum. However, the HIP1 fingerprints of natural populations of T. erythraeum from the two hemispheres showed little genetic diversity within this species (Fig. 4). The changes in banding patterns observed in culture isolates compared to natural populations may be indicative of gene rearrangements that can occur as a result of laboratory maintenance in culture (33). This phenomenon appears to be unique to T. erythraeum as isolates of T. hildebrandtii strains #11 and II-4 had virtually identical fingerprints even though they were isolated 4 years apart (Fig. 2). Similarly, T. thiebautii strain II-3 isolated from the BATS site in 1994 had the same fingerprint as the natural samples collected in 1999 (Fig. 3).

Colonies of T. thiebautii puffs and tufts were found to be identical as determined by HIP1 fingerprinting and DGGE analysis of the hetR fragment (Fig. 5B). These results are similar to those of Ben-Porath et al. (4), who reported that a nifH DNA sequence of a T. thiebautii puff colony was identical to that of a T. thiebautii tuft (42, 43). The DGGE analysis produced only one band for T. thiebautii puffs and tufts as well as for T. erythraeum, indicating that the colonies were comprised of trichomes with similar genetic compositions and not a mixture of the two species. This may imply that there is a sensing mechanism among trichomes for self-identification if the colonies assemble from free trichomes. On the other hand, the colonies could also be clonal, resulting from growth and division originating from a single trichome. Although in this study T. thiebautii puffs and tufts were found to be genetically similar, an earlier study (14) showed that protein extracts obtained from the two morphologies produced slightly different banding patterns when they were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis. These observations suggest that the differences in morphology of T. thiebautii puffs and tufts are reflected at the protein level. The different morphologies of T. thiebautii puffs and tufts may be adaptations to the physical environment (turbulence) of the open ocean. For example, T. thiebautii puff colonies are the dominant form during winter conditions at the BATS site (30).

The trichome widths within T. thiebautii puff and tuft colonies displayed variation that overlapped the trichome widths of previously described species (19). Barker et al. (1) also found no correlation among trichome width, degree of coiling or gas vesicle strength, and genotype of Baltic Sea Nodularia sp. and emphasized the importance of genetic studies for species identification. In the past, morphological characteristics have been used for species identification of Trichodesmium isolates. However, due to overlapping characteristics (e.g., trichome width), a polyphasic approach using genetic tools coupled with morphology is probably more appropriate. For example, in one report of T. contortum a trichome width of 30 to 40 µm was described (19), while in another (36) the spiraled and flexed trichomes were described as >16 µm wide and as wide as 52 µm. This overlap has perhaps caused some confusion in the distinction between the genera Katagnymene and Trichodesmium. Our results based on ITS sequence, DGGE, and HIP1 fingerprinting analyses suggest that K. spiralis should be incorporated in the genus Trichodesmium. Until recently (25), Katagnymene was poorly characterized, as is the species T. contortum. K. spiralis is perhaps not a new species of Trichodesmium but is similar to the spiral trichomes described as T. contortum by Sournia (36) and the screw-like trichomes identified as T. contortum in the North Pacific Ocean by Letelier and Karl (23). Our finding that T. tenue and K. spiralis (or T. contortum) were closely related was also reflected in the study of Janson et al. (18), who found a close relationship between T. tenue and T. contortum.

The use of HIP1 fingerprinting will be useful for further diversity studies of the population dynamics of this important group in oceanic new production and biogeochemistry. HIP1 analysis can be used as a screening method to resolve differences in a population prior to sequencing, which will save time and expense. Although DGGE analysis of the hetR fragment could not distinguish between closely related strains, the HIP1 PCR and ITS sequencing were able to resolve the small differences. The functional role of HIP1 as an adaptive response in these marine diazotrophs may also in future studies reveal important information regarding the unique occupation of a very specific niche in the global ocean.


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ACKNOWLEDGMENTS
 
This work was funded by the Swedish Foundation for International Co-operation in Research and Higher Education (STINT) (B.B.), the Swedish Natural Science Research Council (B.B.), and the Swedish Institute (K.M.O.).

The captains and crews of R/V Weatherbird II and R/V Maurice Ewing are acknowledged for their assistance at sea. Ken Halanych helped us create the phylogenetic tree, and Alfonso Mateo assisted us with French translation. We thank Anthony H. Knap and the BATS staff for their hospitality and use of their lab at Bermuda Biological Station. Doug Capone and Ed Carpenter are thanked for sharing cruise time off North Australia. We thank Jon Zehr and two anonymous reviewers for valuable comments on an early version of the manuscript.


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FOOTNOTES
 
* Corresponding author. Present address: Darling Marine Center, University of Maine, 193 Clark's Cove Rd., Walpole, ME 04573. Phone: (207) 563-3146. Fax: (207) 563-3119. E-mail: karen.orcutt{at}maine.edu. Back

{dagger} Present address: Bigelow Laboratory for Ocean Sciences, West Boothbay Harbor, ME 04575. Back


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Applied and Environmental Microbiology, May 2002, p. 2236-2245, Vol. 68, No. 5
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.5.2236-2245.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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