Abstract
Little is known about Ceanothus-infective Frankia strains because no Frankia strains that can reinfect the host plants have been isolated from Ceonothus spp. Therefore, we studied the diversity of the Ceonothus-infective Frankia strains by using molecular techniques. Frankia strains inhabiting root nodules of nine Ceanothus species were characterized. The Ceanothus species used represent the taxonomic diversity and geographic range of the genus; therefore, the breadth of the diversity of Frankia strains that infect Ceanothus spp. was studied. DNA was amplified directly from nodular material by using the PCR. The amplified region included the 3′ end of the 16S rRNA gene, the intergenic spacer, and a large portion of the 23S rRNA gene. A series of restriction enzyme digestions of the PCR product allowed us to identify PCR-restriction fragment length polymorphism (RFLP) groups among the Ceanothus-infective Frankia strains tested. Twelve different enzymes were used, which resulted in four different PCR-RFLP groups. The groups did not follow the taxonomic lines of the Ceanothus host species. Instead, the Frankia strains present were related to the sample collection locales.
Frankia strains are nitrogen-fixing actinomycetes that form symbiotic relationships with 24 genera of woody dicotyledonous plants. The focus of this paper is the Ceanothus-Frankia symbiosis. Although this symbiotic relationship is of ecological and practical importance, very little is known about Ceanothus-infective Frankia strains. The main reason for this is the fact that no isolates that can reinfect the host after isolation have been recovered from Ceanothus spp. Consequently, we studied the diversity of the Ceanothus-infective Frankia strains by using molecular techniques. Prior studies in which the PCR and restriction fragment length polymorphism (RFLP) have been used have been useful in discriminating among closely related Frankia strains obtained from members of other host genera (8, 13, 15, 17). In this study, we used PCR-RFLP techniques to characterize the Frankia strains that infect nine Ceanothus species.
The Ceanothus species studied represent the taxonomic diversity and geographic distribution of the genus (4, 9). The genus Ceanothus, which is indigenous only to North America, contains 55 species. Two subgenera of the genus, subgenera Ceanothus and Cerastes, are recognized. The subgenus Ceanothus, the more ancient subgenus, includes evergreen species, such as C. cordulatus, C. velutinus, and C. thyrsiflorus, and deciduous species, such as C. integerrimus, C. sanguineus, and C. americanus. The subgenus Cerastes includes the evergreen species C. cuneatus, C. prostratus, and C. pumilus. Because we included the diversity of the host plants in our study, we expected that the breadth of the diversity of the Frankia strains that infect Ceanothus spp. would be represented.
In less comprehensive surveys, other workers have found a considerable amount of diversity in the Frankia-Ceanothus symbiosis (2, 14). Baker and Mullin (2) assessed the Frankia strains inhabiting C. americanus nodules at seven different locations in a 70-mile radius in eastern Tennessee by using restriction endonuclease digestion of genomic DNAs in combination with genetic probes. A total of 25 to 30 nodules were collected per site. These authors found differences in RFLP patterns between sites and among different plants collected from a single site. It is unclear whether they found more than one pattern per nodule.
Murry et al. (14) used repetitive extragenic palindromic (REP) PCR methods to characterize the diversity of the Frankia strains associated with six Ceanothus species located in the coastal region of southern California. The study area was an area that had a 10-mile radius and contained seven sites. These authors sampled two to five mature nodule clusters per species. In addition, they compared nodules collected from mature and immature plants. The results varied. Murry et al. found variation within and between sites for some species and found no variation within or between collection sites for other species. Also, they found more than one pattern per nodule cluster. No evidence of contaminating DNA was found, however. The conclusion of Murry et al. was that we need more collections of Ceanothus species to further elucidate the diversity of the Frankia strains that inhabit these plants.
Our study was more extensive than previous studies since we included a greater number of sites, host species, and environmental conditions. Therefore, we predicted that we would find even higher levels of diversity than were found previously.
MATERIALS AND METHODS
Sample collection.
Nodule, soil, and shoot samples were collected from each of the nine Ceanothus species examined (Table 1). Samples were obtained from 19 sites in Oregon (one site for each of the seven Oregon species and 12 sites for C. velutinus, which is more widespread than the other species assayed) and from one site in Tennessee. At least two plants were collected per site (Table 1). The samples consisted of at least five distinct nodule clusters per site (with the exception of the C. americanus samples [Table 1]). Each collection was made within the geographic range of the host plant species. All samples were transported to the laboratory on ice. Upon arrival, nodule samples were rinsed in deionized water, washed with Tween 80, surface sterilized with ethanol, and stored at −20°C until they were used.
TABLE 1.
Nodule collection information
| Site | Species | Location | No. of plants sampled | No. of nodule clusters collected |
|---|---|---|---|---|
| 1 | C. americanusa | Anderson County, Tennessee | 2 | 2 |
| 2 | C. cordulatus | Highway 138, Toketee, Oreg. | ≥5 | ≥10 |
| 3 | C. prostratus | Highway 138, Diamond Lake, Oreg. | ≥5 | ≥10 |
| 4 | C. sanguineus | H. J. Andrews Experimental Forest, Blue River, Oreg. | 3 | ≥10 |
| 5 | C. integerrimus | Highway 126, Blue River, Oreg. | 5 | ≥10 |
| 6 | C. cuneatus | Highway 99, Adair Village, Oreg. | ≥5 | ≥10 |
| 7 | C. thyrsiflorus | Bear Creek Recreation Site, Remote, Oreg. | 2 | 5 |
| 8 | C. pumilus | Siskiyou National Forest, Agness, Oreg. | ≥5 | ≥10 |
| A | C. velutinus | Mary’s Peak Road, Philomath, Oreg. | 3 | ≥5 |
| B | C. velutinus | Galice Road, Galice, Oreg. | 2 | ≥5 |
| C | C. velutinus | Highway 140, Fish Lake, Oreg. | 2 | ≥5 |
| D | C. velutinus | Highway 138, Falls Creek, Oreg. | 3 | ≥5 |
| E | C. velutinus | Metolius River, Camp Sherman, Oreg. | 2 | ≥5 |
| F | C. velutinus | Highway 26, Warm Springs, Oreg. | 2 | ≥5 |
| G | C. velutinus | Highway 395, Seneca, Oreg. | 2 | ≥5 |
| H | C. velutinus | Izee Road, Izee, Oreg. | 2 | ≥5 |
| I | C. velutinus | Cerro Gordo, Cottage Grove, Oreg. | 3 | ≥5 |
| J | C. velutinus | BLM Road 10-18, Roseburg, Oreg.b | 3 | ≥5 |
| K | C. velutinus | Buck Creek Road, Scottsburg, Oreg. | 2 | ≥5 |
| L | C. velutinus | Forest Service Road 25, Brickerville, Oreg. | 2 | ≥5 |
The C. americanus nodules were collected by Beth Mullin.
BLM, Bureau of Land Management.
DNA extraction.
Genomic DNA was prepared by using a protocol adapted from the protocol of Baker and Mullin (2). Individual lobe tips were excised with a scalpel and frozen in a dry ice-ethanol bath (−70°C). DNA was extracted from a minimum of 15 lobes per species (at least three replicates per nodule cluster). No composites were prepared to increase the chance of capturing the total genetic diversity of the Frankia strains that infect individual plants. Individual lobe tips were homogenized with a mortar and pestle by using 600 μl of cetyltrimethylammonium bromide extraction buffer (2% cetyltrimethylammonium bromide, 100 mM Tris [pH 8.0], 20 mM EDTA, 1.4 M NaCl) and were incubated at 65°C for 30 min. The DNA was extracted twice with an equal volume of chloroform-isoamyl alcohol (24:1) and was precipitated with 1 volume of ice-cold isopropanol. The precipitated DNA was resuspended in 50 μl of TE (10 mM Tris [pH 8.0], 0.1 mM EDTA) (10:0.1), reprecipitated by adding 0.25 volume of 10 M ammonium acetate and 1 volume of ice-cold isopropanol, and washed with 70% ethanol. DNA pellets were resuspended in 50 μl of TE (10:0.1). Samples were further purified by adding 1 volume of a polyethylene glycol-NaCl mixture (20% polyethylene glycol 8000, 2.5 M NaCl). Samples were kept at 37°C for 15 min and centrifuged. The DNA pellets were washed twice with 80% ethanol, dried, resuspended in 50 μl of TE (10:0.1), and stored at −20°C until they were used.
PCR amplification.
Purified genomic DNA (5 to 10 ng) was used to amplify a 2,098-bp region of the 16S and 23S rRNA genes by the PCR. The amplified portion included the 3′ end of the 16S rRNA gene, the intergenic spacer (IGS), and a large portion of the 23S rRNA gene. All reactions were performed in 50-μl (final volume) reaction mixtures containing 1.5 μl of template DNA, 5 μl of GeneAmp 10× PCR buffer II (Perkin-Elmer Corp., Branchburg, N.J.), 1.5 mM MgCl2, each deoxynucleoside triphosphate (Perkin-Elmer) at a concentration of 0.2 mM, 0.2 μM forward primer 1649F (5′-GATTGGGACGAAGTCGT-3′) (20), 0.2 μM reverse primer 2309R (5′-ATCGCATGCCTACTACC-3′) (6), and 2 U of AmpliTaq DNA polymerase (Perkin-Elmer). An initial denaturation at 95°C for 2 min was followed by 35 cycles consisting of denaturation at 95°C for 45 s, annealing at 53°C for 45 s, and extension at 72°C for 1.5 min (Coy Thermocycler, Ann Arbor, Mich.). There was a final extension step consisting of 72°C for 5 min. The PCR products were loaded onto a 1% agarose gel (Sea Kem; FMC BioProducts, Rockland, Maine) and electrophoresed. After electrophoresis, bands were excised, purified with GeneClean II (BIO 101, Vista, Calif.), resuspended in 18 μl of TE (10:0.1), and stored at −20°C until they were used.
RFLP analysis.
Purified PCR products were digested with restriction endonucleases for at least 4 h by using the suppliers’ instructions. The following 12 restriction enzymes were used: HaeIII, HhaI, HinFI, TaqI, RsaI, NciI, NdeII, MvnI, AluI, MspI, BstUI, and Sau96I. The digested products were electrophoresed on chilled 3 to 5% Metaphor gels (FMC BioProducts). Gels containing higher percentages of Metaphor were used for smaller fragments. Each digestion was repeated at least twice to verify the patterns obtained.
Phylogenetic analysis.
The sizes and numbers of bands were determined manually, and the results were used to calculate similarity indices for each strain. Relationships were inferred from restriction patterns by using both parsimony (PAUP) (18) and distance-based (NT-SYS) methods.
Sequencing.
Amplification reactions were performed by using the primers used for the RFLP analysis. The PCR products were ethanol precipitated, resuspended in deionized water, and subjected to double-stranded sequencing. Primers 1649F and 23S12R (5′-TCCACCGTGTGCCCTTA-3′; synthesized for this study) were used to determine the sequence of the IGS region. Taq dye terminator chemistry was used to determine sequences with an ABI cycle sequencer (Center for Gene Research and Biotechnology, Central Services Laboratory, Oregon State University, Corvallis).
Sequence data analysis.
All of the sequences obtained were compared with sequences in the GenBank database by using BLAST (1). The sequences were aligned by using the ASSEMBLE CONTIGS option of the Genetic Data Environment (version 2.2) sequence analysis software (provided by Steven Smith, Millipore Corp., Marlborough, Mass.). The alignment was optimized by manual adjustment.
Nucleotide sequence accession numbers.
The nucleotide sequences determined in this study have been deposited in the GenBank database under accession no. AF050760 to AF050768.
RESULTS AND DISCUSSION
The data suggested that there were some geographic and taxonomic relationships between Ceanothus spp. and the microsymbiont genus Frankia. These relationships were not as clear as we anticipated, however.
PCR-RFLP analysis.
PCR amplification was successful with nodular microsymbionts of the nine Ceanothus species assayed. Only one 2,098-bp band was present after most amplification reactions. Twelve different restriction enzymes were utilized. HaeIII digestion resolved four distinct RFLP groups (Fig. 1). Digestion with HhaI, HinFI, TaqI, RsaI, NciI, NdeII, MvnI, AluI, MspI, BstUI, and Sau96I separated the microsymbionts into only two RFLP groups (data not shown). Only one RFLP group was identified for all of the individual lobe tips from each plant analyzed. Actually, except for sites A, D, and L, all of the lobe tips from a given site assayed produced the same RFLP pattern (Fig. 2).
FIG. 1.
PCR fingerprints generated after HaeIII digestion of amplified sequences from uncultured microsymbionts of Ceanothus spp. Lanes 1 and 13, 100-bp ladder; lanes 3 to 12, symbiotic Frankia strains obtained from C. velutinus (site J), C. velutinus (site E), C. thyrsiflorus, C. sanguineus, C. pumilus, C. prostratus, C. integerrimus, C. cuneatus, C. cordulatus, and C. americanus nodule DNA, respectively. Lane 2 contained the marker pBR322:MspI. There are four different band patterns. The pattern in lane 12 is the RFLP group I pattern; the pattern in lane 10 is the RFLP group II pattern; the pattern in lanes 3 and 5 is the RFLP group III pattern; and the pattern in lanes 4, 6 to 9, and 11 is the RFLP group IV pattern. All band patterns were verified by repeating each digestion experiment a minimum of five times.
FIG. 2.
Map of Ceanothus sites as identified by RFLP group. RFLP group I Frankia strains were collected in Tennessee (site 1). Site 6 is the only place where RFLP group II Frankia strains were collected. The shaded region in the Willamette Valley of Oregon (sites AII, DI, I, J, K, LII, and 7) contained Ceanothus species exhibiting the RFLP group III Frankia pattern. RFLP group IV Frankia strains were collected from Ceanothus species growing in the unshaded region of Oregon.
Data obtained with only 7 of the 12 enzymes (HaeIII, HinFI, TaqI, NciI, NdeII, AluI, and Sau96I) were used for analysis. We used data only from digestions that generated clear, coherent band patterns, which enabled us to quantify the number of bands and determine the differences between samples. A total of 57 bands were generated with the seven enzymes used, and each strain was scored either positive or negative for each band. The resulting information was used to create a similarity index (Table 2) and to perform a cluster analysis (Fig. 3). The dendrogram in Fig. 3 is consistent with the results of the parsimony analysis (data not shown).
TABLE 2.
Similarities of Ceanothus microsymbionts
| Source of micro-symbiont | Similarity to microsymbiont ofa:
|
|||||||||
|---|---|---|---|---|---|---|---|---|---|---|
| Cameb | Ccor | Ccun | Cint | Cpro | Cpum | Csan | Cthy | Cvelb | Cvelj | |
| Came | 1.000 | |||||||||
| Ccor | 0.526 | 1.000 | ||||||||
| Ccun | 0.474 | 0.912 | 1.000 | |||||||
| Cint | 0.526 | 1.000 | 0.912 | 1.000 | ||||||
| Cpro | 0.526 | 1.000 | 0.912 | 1.000 | 1.000 | |||||
| Cpum | 0.526 | 1.000 | 0.912 | 1.000 | 1.000 | 1.000 | ||||
| Csan | 0.526 | 1.000 | 0.912 | 1.000 | 1.000 | 1.000 | 1.000 | |||
| Cthy | 0.509 | 0.947 | 0.965 | 0.947 | 0.947 | 0.947 | 0.947 | 1.000 | ||
| Cvelb | 0.526 | 1.000 | 0.912 | 1.000 | 1.000 | 1.000 | 1.000 | 1.000 | 1.000 | |
| Cvelj | 0.509 | 0.947 | 0.965 | 0.947 | 0.947 | 0.947 | 0.947 | 0.947 | 0.947 | 1.000 |
The values are levels of similarity based on the percentage of shared fragments. A total of 57 fragments were included in the study.
Abbreviations: Came, C. americanus; Ccor, C. cordulatus; Ccun, C. cuneatus; Cint, C. integerrimus; Cpro, C. prostratus; Cpum, C. pumilus; Csan, C. sanguineus; Cthy, C. thyrsiflorus; Cvelb, C. velutinus (site B); Cvelj, C. velutinus (site J).
FIG. 3.
Dendrogram (as determined by the NT-SYS method) showing the relationships of amplified 16S ribosomal DNA-23S ribosomal DNA sequences based on the results of a PCR-RFLP analysis performed with seven restriction endonucleases (HaeIII, HinFI, TaqI, NciI, NdeII, AluI, and Sau96I). Unless otherwise noted, the Ceanothus species are members of the subgenus Ceanothus.
We expected to find a taxonomic relationship between Frankia strains and the host species that they infect; however, this was not the case. There are two subgenera in the genus Ceanothus, the subgenera Ceanothus and Cerastes. We studied the Frankia strains associated with nine species, six species belonging to subgenus Ceanothus (C. americanus, C. cordulatus, C. integerrimus, C. sanguineus, C. thyrsiflorus, and C. velutinus) and three species belonging to subgenus Cerastes (C. cuneatus, C. prostratus, and C. pumilus). The C. cuneatus-infective Frankia strains belonged to a single group (RFLP group II); however, the Frankia strains associated with the two other members of subgenus Cerastes were members of RFLP group IV, which includes Frankia strains obtained from nodules of members of subgenus Ceanothus. Thus, there does not appear to be a strong taxonomic relationship between the genus Ceanothus and the genus Frankia at this level of resolution.
The four RFLP groups did generally follow a geographic pattern (Fig. 2). RFLP group I contained Frankia strains obtained from C. americanus, the only plant species assayed outside Oregon. The C. cuneatus-infective Frankia strains (RFLP group II) and all members of RFLP group III were collected in the Willamette Valley. RFLP group IV was the largest and most wide-ranging group. The strains from six of the nine plant species studied are members of this group. Many of the Ceanothus-infective Frankia strains in RFLP group IV were collected in the Cascades. The Frankia strains obtained from C. velutinus nodules collected in the mountainous areas of eastern Oregon, in the Coast Range, and in the Siskiyous were also members of RFLP group IV.
The influence of geography was most evident when we evaluated the C. velutinus-infective Frankia strains. Interestingly, C. velutinus-infective Frankia strains were members of both RFLP group III and RFLP group IV. With the exception of strains obtained from three sites (sites A, D, and L), all of the C. velutinus-infective Frankia strains collected in the Willamette Valley were members of RFLP group III. The Frankia strains associated with the plants collected at sites A, D, and L were not the same; some of these strains were members of RFLP group III, and others were members of RFLP group IV. These sites are on the boundary between the valley floor and the adjacent foothills. All of the other C. velutinus-infective Frankia strains were members of RFLP group IV.
A number of environmental properties were characterized (Table 3). In a previous study, pH played a significant role in determining what kind of Frankia strains were present in Elaeagnus nodules (8). There did not appear to be a relationship between pH and the Frankia type present in this study, however. We did observe a trend in RFLP type with elevation.
TABLE 3.
Characteristics of collection sites
| Site | Species | Elevation (ft) | Soil type | Classification | Parent material |
|---|---|---|---|---|---|
| 1 | C. americanus | NAa | NA | ||
| 2 | C. cordulatus | 2,650 | Gravelly sandy loam | ||
| 6 | C. cuneatus | 300 | Witzel very cobbly loam | Lithic Ultic Haploxerolls | Igneous rock |
| 5 | C. integerrimus | 1,200 | Klickitat stony loam | Typic Haplumbrepts | Basalt, intrusive rock |
| 3 | C. prostratus | 4,500 | Loamy sand with pumice | Pumice | |
| 8 | C. pumilus | 1,800 | Snowcamp | Serpentinite | |
| 4 | C. sanguineus | 2,600 | NA | ||
| 7 | C. thyrsiflorus | 900 | Digger gravelly loam | Dystric Eutrochrepts | Sandstone, siltstone |
| A | C. velutinus | 2,300 | Trask gravelly loam | Umbric Dystrochrepts | Sandstone |
| B | C. velutinus | 1,200 | Abegg gravelly loam | Ultic Haploxeralfs | Sedimentary and intrusive igneous rock |
| C | C. velutinus | 4,200 | NA | ||
| D | C. velutinus | 1,200 | Shotty loam, gravelly loam | ||
| E | C. velutinus | 1,200 | NA | ||
| F | C. velutinus | 1,500 | NA | ||
| G | C. velutinus | 4,800 | NA | ||
| H | C. velutinus | 4,000 | Laycock-Logdell complex very shaly loam | Ultic Haploxerolls | Shale, sandstone, graywacke |
| I | C. velutinus | 1,300 | Nekia silty clay loam | Xeric Haplohumults | Basalt, tuff sedimentary rock |
| J | C. velutinus | 1,800 | Jory silty clay loam | Xeric Haplohumults | Volcanic and sedimentary rock |
| K | C. velutinus | 1,200 | Preacher clay loam | Typic Haplumbrepts | Sedimentary rock |
| L | C. velutinus | 1,300 | Blachly silty clay loam | Umbric Dystrochrepts | Sandstone |
NA, information not available.
RFLP group IV Frankia strains were obtained from nodules that were collected at a higher average elevation than the elevation at which RFLP group III strains were collected (P < 0.1, as determined by Student’s t test) (Fig. 4). The elevation of the site with the RFLP group II strains was significantly lower than the elevations of the sites with RFLP group III or IV strains; however, there was only one RFLP group II site. More samples containing RFLP group II strains are needed to adequately evaluate the potential relationship between elevation and RFLP group II Frankia strains.
FIG. 4.
Relationship between elevations of collection sites and RFLP groups. Data were compiled for each of the 19 sites (RFLP group II, 1 site; RFLP group III, 7 sites; RFLP group IV, 14 sites); 3 sites had representatives belonging to two groups. The data for samples from each RFLP group were averaged, and standard errors were calculated.
In two previous studies performed with rhizobia the workers also found a correlation between elevation and nodule occupancy (11, 19). Bacterial strains are presumably affected by ecological factors determined by elevation, such as temperature and precipitation.
Other environmental features, such as soil type, soil classification, and parent material, were assessed (Table 3). Qualitatively, there did not appear to be a connection between any of these characteristics and the infective Frankia strains present.
The diversity of the Oregon Ceanothus-infective strains appeared to be related to sample collection location rather than to host taxonomy or to any of the environmental properties measured. The differences that did exist among Frankia strains assayed in Oregon were not great. These strains shared >91% of all of the bands identified (Table 2). The differences that were present were based only on HaeIII digestion results (the results obtained with all of the other enzymes indicated that these strains were identical) and consisted of the results for either one or two restriction sites; there was one difference between the restriction sites of RFLP groups III and IV, there was one difference between the restriction sites of RFLP groups II and III, and there were two differences between the restriction sites of RFLP groups II and IV. There was, however, a marked decrease in similarity (to 47 to 52%) when the samples were compared with the microsymbionts associated with C. americanus, a species found in the eastern United States. The microsymbionts associated with C. americanus are very different from those that nodulate the Oregon species.
Prior to using the IGS region used in this study, we tried to specifically amplify Frankia sequences located in the IGS between nifD and nifK (8). Unfortunately, we were able to amplify the sequences of only a subset of our samples. The sequences that we were able to amplify (the sequences of the C. americanus, C. thyrsiflorus, C. prostratus, C. cordulatus, and C. integerrimus microsymbionts) were digested with two restriction endonucleases, HhaI and HaeIII (data not shown). The results of this limited survey were the same as the results obtained with the IGS region between 16S rRNA and 23S rRNA genes. It appeared that the same Frankia strain infected C. prostratus, C. cordulatus, and C. integerrimus, whereas a different strain infected C. thyrsiflorus. Again, the Frankia strains associated with C. americanus were markedly different than any strain found in Oregon.
Sequencing.
Because RFLP group IV contains Frankia strains obtained from six of the nine Ceanothus species sampled, we sequenced the IGS between the 16S rRNA and 23S rRNA genes to confirm the homogeneity of this group. Almost all of the members of RFLP group IV had identical sequences; the only exceptions were the C. pumilus-infective Frankia strains. The C. pumilus-infective Frankia strains differed from the rest of the strains in the group at 2 of 450 bases (level of similarity, 99.6%). The exception of the C. pumilus-infective Frankia strains is not surprising since C. pumilus grows only in a chemically unique environment, serpentine soils.
The sequences of the C. cuneatus-infective Frankia strains and the RFLP group IV strains differed by three mismatches and two indels (level of similarity, 98.9%). There was a difference of 22 nucleotides and 18 indels between the sequences of the C. americanus-infective strains and the RFLP group IV Frankia strains (level of similarity, 91.1%). These differences between the sequences of the RFLP group IV strains and the group RFLP II and I strains are consistent with the findings of the PCR-RFLP analysis. Therefore, the sequencing results verified the PCR-RFLP analysis results, suggesting that RFLP group IV is quite homogeneous.
When the results of the PCR-RFLP analysis and the results of the sequencing analysis were combined, the Ceanothus-infective Frankia strains could be separated into five groups. This is far less diversity than previous workers have observed in Ceanothus-infective Frankia strains (2, 14) and other host-infective groups (7, 8, 17).
Rouvier et al. (17) used PCR-RFLP analysis to assess the diversity of Casuarina and Allocasuarina microsymbionts. They also examined the IGS regions in the ribosomal and nif operons and found host specificity and high levels of diversity among their symbionts. The method which they used allowed them to assess strain level variations. In addition, the IGS between the 16S rRNA and 23S rRNA genes has been used successfully to characterize strains of other microbes, such as Escherichia coli and Nitrobacter spp. (5, 15). Therefore, it is reasonable to assume that if there were high levels of diversity among Oregon Ceanothus microsymbionts, we would have been able to detect them with this method.
There are a few reasons why previous studies may have found more diversity than we did. One reason may be that we examined only limited regions of the genome (the ribosomal operon and nifDK with limited success). Murry et al. (14) assessed the entire genome by using REP-PCR methods, and it is possible that we would have found greater diversity if we had assessed the entire genome. In subsequent work in our laboratory, workers have used REP-PCR methods to assess the population level diversity of Frankia strains in C. prostratus-C. velutinus, C. integerrimus-C. sanguineus, and C. velutinus-C. integerrimus copopulations (10). These workers found eight different patterns in 10 nodules; however, many of the patterns were quite similar. The differences observed among different sites were often greater than the differences between infected plant species at a given site, which is consistent with our finding that some factor besides host plant taxonomy dictates which Frankia type is present.
A second potential reason for the discrepancy between diversity levels is the possibility that the Ceanothus microsymbionts assayed by Murry et al. (14) may have been much different than the microsymbionts which we studied. We characterized some of the Ceanothus microsymbionts used in this study by using full-length 16S rRNA sequences (16) and found that our Ceanothus-infective Frankia strains were much different than the strains studied by Murry et al. (14) and were more similar to the strains studied by Benson et al. (3).
Third, Murry et al. (14) assessed the diversity of Ceanothus symbionts in southern California, which harbors a greater diversity of Ceanothus spp. More than 40 species of Ceanothus are endemic to California, which is believed to be the center of distribution of the genus (12). Thus, the greater diversity of the Ceanothus microsymbionts characterized by Murry et al. (14) may be related to the greater diversity of actinorhizal hosts in southern California.
The results obtained with two different methods (PCR-RFLP analysis and sequencing) and two different locations on the genome (16S rRNA-23S rRNA and nifDK) lead us to conclude that there is little genetic diversity in the Frankia strains that infect Oregon Ceanothus species and that C. americanus microsymbionts are considerably different than Oregon microsymbionts. The limited diversity that is present is not related to plant taxonomy or any known environmental condition. More samples and potentially more discriminatory methods may be needed to further clarify the diversity of Frankia strains associated with Ceanothus species.
ACKNOWLEDGMENTS
We thank Katharine Field, Ena Urbach, and Soon-Chun Jeong for their advice and criticisms; Beth Mullin for providing nodules; and the staff at the Oregon State University Herbarium and Central Services Laboratory for their assistance.
This research was supported by USDA-NRICGP grant 93-60017860-A4.
Footnotes
Oregon Agricultural Experiment Station technical paper 11473.
REFERENCES
- 1.Altschul S F, Gish W, Miller W, Myers E W, Lipman D J. Basic local alignment search tool. J Mol Biol. 1990;215:403–410. doi: 10.1016/S0022-2836(05)80360-2. [DOI] [PubMed] [Google Scholar]
- 2.Baker D D, Mullin B C. Diversity of Frankia nodule endophytes of the actinorhizal shrub Ceanothus as assessed by RFLP patterns from single nodule lobes. Soil Biol Biochem. 1994;26:547–552. [Google Scholar]
- 3.Benson D R, Stephens D W, Clawson M L, Silvester W B. Amplification of 16S rRNA genes from Frankia strains in root nodules of Ceanothus griseus, Coriaria arborea, Coriaria plumosa, Discaria toumatou, and Purshia tridentata. Appl Environ Microbiol. 1996;62:2904–2909. doi: 10.1128/aem.62.8.2904-2909.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4.Conrad S G, Jaramillo A E, Cromack K, Jr, Rose S. The role of the genus Ceanothus in western forest ecosystems. USDA Forest Service General Technical Report PNW-182. Portland, Oreg: Pacific Northwest Forest and Range Experiment Station; 1985. [Google Scholar]
- 5.García-Martínez J, Martínez-Murcia A, Antón A I, Rodríguez-Valera F. Comparison of the small 16S to 23S intergenic spacer region of the rRNA operons of some Escherichia coli strains of the ECOR collection and Escherichia coli K-12. J Bacteriol. 1996;178:6374–6377. doi: 10.1128/jb.178.21.6374-6377.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6.Hönerlage W, Hahn D, Zepp K, Zeyer J, Normand P. A hypervariable 23S rRNA region provides a discriminating target for specific characterization of uncultured and cultured Frankia. Syst Appl Microbiol. 1994;17:433–443. [Google Scholar]
- 7.Jamann S, Fernendez M P, Moiroud A. Genetic diversity of Elaeagnaceae-infective Frankia strains isolated from various soils. Acta Œcol. 1992;13:395–406. [Google Scholar]
- 8.Jamann S, Fernendez M P, Normand P. Typing method for N2-fixing bacteria based on PCR-RFLP analysis—application to the characterization of Frankia strains. Mol Ecol. 1993;2:17–26. doi: 10.1111/j.1365-294x.1993.tb00095.x. [DOI] [PubMed] [Google Scholar]
- 9.Jeong S-C, Liston A, Myrold D D. Molecular phylogeny of the genus Ceanothus using ndhF and rbcL sequences. Theor Appl Genet. 1997;94:852–857. [Google Scholar]
- 10.Jeong, S.-C., A. Liston, and D. D. Myrold. Specific genomic fingerprinting of microsymbiont Frankia from Ceanothus copopulations using repetitive sequences and PCR. Submitted for publication.
- 11.Mahler R L, Bezdicek D F. Distribution of Rhizobium leguminosarium in peas in the Palouse of eastern Washington. Soil Sci Soc Am J. 1980;44:292–295. [Google Scholar]
- 12.McMinn H E. Ceanothus vol. II: a systematic study of the genus Ceanothus. Santa Barbara, Calif: Santa Barbara Botanical Garden; 1942. [Google Scholar]
- 13.Mirza M S. Characterization of uncultured Frankia strains by 16S rRNA sequence analysis. Ph.D. thesis. Wageningen, The Netherlands: Wageningen Agricultural University; 1993. [Google Scholar]
- 14.Murry M A, Konopka A S, Pratt S D, Vandergon T L. The use of PCR-based typing methods to assess the diversity of Frankia nodule endophytes of the actinorhizal shrub Ceanothus. Physiol Plant. 1997;99:714–721. [Google Scholar]
- 15.Navarro E, Simonet P, Normand P, Bardin R. Characterization of natural populations of Nitrobacter spp. using PCR/RFLP analysis of the ribosomal intergenic spacer. Arch Microbiol. 1992;157:107–115. doi: 10.1007/BF00245277. [DOI] [PubMed] [Google Scholar]
- 16.Ritchie, N. J., and David D. Myrold. Phylogenetic placement of uncultured Ceanothus microsymbionts using 16S rRNA gene sequences. Can. J. Bot., in press.
- 17.Rouvier C, Prin Y, Reddell P, Normand P, Simonet P. Genetic diversity among Frankia strains nodulating members of the family Casuarinaceae in Australia revealed by PCR and restriction fragment length polymorphism analysis with crushed nodules. Appl Environ Microbiol. 1996;62:979–985. doi: 10.1128/aem.62.3.979-985.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Swofford D L. PAUP: phylogenetic analysis using parsimony, version 3.0. Champaign: Illinois National History Survey; 1991. [Google Scholar]
- 19.Woomer P, Singleton P W, Bohlool B B. Ecological indicators of native rhizobia in tropical soils. Appl Environ Microbiol. 1988;54:1112–1116. doi: 10.1128/aem.54.5.1112-1116.1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20.Yaeger, C., and D. D. Myrold. 1996. Unpublished data.