CHARACTERIZATION OF NITROGEN-FIXING BACTERIAL RHIZOSPHERE COMMUNITIES USING RESTRICTION FRAGMENT LENGTH POLYMORPHISMS OF PCR- AMPLIFIED NIFH

Joe E. Lepo^a, Marisa K. Chelius^a, and David E. Weber^b

^aCenter for Environmental Diagnostics and Bioremediation, University of West Florida, Pensacola, Florida, 32514; and ^bNational Health Environmental Effects Research Laboratory, Gulf Ecology Division (formerly Environmental Research Laboratory), Environmental Protection Agency, Gulf Breeze, Florida 32651

DISCLAIMER: Mention of trade names in this publication does not constitute endorsement or recommendation for use by the U.S. EPA or by the Center for Environmental Diagnostics and Bioremediation, University of West Florida.

SUMMARY

Methods for studying the population diversity of N₂-fixing bacteria in soil usually require enrichments of culturable bacteria; and the acetylene reduction assay allows the assessment of functional contributions. These methods do not describe the structure of nonculturable N₂-fixing populations, and the acetylene reduction assay provides incomplete information on relative in situ contributions of diverse populations. We have developed a method to study the species diversity of N₂-fixing bacteria in the rhizosphere by examining the diversity of a segment of nifH, the conserved structural gene for dinitrogenase reductase. Total DNA was extracted from rhizosphere zones in natural and artificial sediments by bead-beating and purified by CsCl-EtBr gradient centrifugation. The average DNA yield was 5.5 µg g^-1 of soil and was of sufficient purity for PCR amplification of nifH. Label ([-³²P] dCTP) was incorporated into the PCR reaction and nifH PCR products were restriction digested. Restriction Fragment Length Polymorphism (RFLP) analysis of the amplified sequences revealed differences in the community structure of N₂-fixing rhizobacteria of the salt marsh plant, Spartina alterniflora, and a laboratory cultured Sesbania macrocarpa. Soil inoculation experiments were used to determine the efficiency of the methods, and amplified nifH DNA could be detected when 10⁴ cells each of Vibrio natriegens and Azotobacter vinelandii were added per gram of soil. Restriction patterns produced by each species were detected at 10⁶cells g^-1 soil. These results indicate that RFLP analysis of amplified nifH sequences from rhizosphere communities may provide information on species composition and reveal shifts in diversity.

Key words: Nitrogen fixation, PCR, rhizosphere, microbial diversity, nifH

INTRODUCTION

Studies on N₂-fixing bacteria in the environment are limited to those organisms that can be detected by existing methodologies. Molecular methods have made it possible to describe naturally occurring microbial communities that have been previously undetected using conventional methods. In this report, we describe a method to assess the community structure of N₂-fixing bacteria in a natural and synthetic wetland soil by direct DNA extraction and restriction analysis of amplified nifH sequences.

Nitrogen is usually the nutrient that limits plant production in wetlands (Buresh et al., 1990) and, under N-limited conditions, plant roots excrete compounds with high C/N ratios, favoring rhizosphere N₂fixation (Klein et al., 1990). Rates of N₂ fixation are usually higher in the root zone of flooded soil systems than in well-drained soils, and substantial N₂ fixation rates have been reported in the root zones of several non-nodulating marsh plants (Kana and Tjepkema, 1978), as well as in nodulating legumes of riparian systems (Moreira et al., 1992). In addition, occurrence and frequency of nodulation of legume species was higher in flooded soils than non-flooded sites (Moreira et al., 1992). The contribution of N₂-fixing bacteria to the N budget of salt marsh systems is significant. N₂ fixation accounted for the majority of the NH₃ entering a New England marsh (Valiela and Teal, 1979) and supplied about 50% of the annual plant N demand (Teal et al., 1979; White and Howes, 1994a). Additionally, reported N₂-fixation rates for decaying S. alterniflora are high (Newell et al., 1992) and may be a source of available N for new plant growth (White and Howes, 1994b).

Methods for studying the community structure of N₂-fixing bacteria in the rhizosphere usually require enrichments of culturable bacteria, and functional contributions are often assessed using the acetylene reduction assay (ARA) of cultured isolates or soil cores. These methods provide incomplete information because: i. only a small fraction of viable cells is represented by culture-based methods, ii. N₂-fixing bacteria that inhabit living root cells or tissues (McClung et al., 1983; You and Zhou, 1989) are not identified, iii. there are technical problems associated with the use of an indirect assay (ARA) for nitrogenase activity (Buresh et al., 1990; Howarth, et al., 1988; Smith, 1980), and iv. the relative functional in situ contributions of individuals in diverse populations are not assessed.

Methodology facilitating the characterization of the N₂-fixing bacterial community structure, when used with ARA and selective culturing methods, may provide a more thorough analysis of N₂-fixing bacterial population dynamics in the rhizosphere. The nucleic acids of N₂-fixing bacteria in the environment have been isolated and studied (Hilger and Myrold, 1991; Kirshtein et al., 1991; Ogram et al., 1987); however, the soil environment offers a particular challenge to the molecular ecologist due to the presence of organic contaminants that co-precipitate with DNA and inhibit many enzymes used with molecular techniques.

We describe here, a method for the isolation from soil of N₂-fixing rhizobacterial DNA of sufficient yield and purity for PCR amplification. Bacterial DNA was extracted by bead-beating and purified by CsCl-EtBr gradient centrifugation. The nifH gene, encoding dinitrogenase reductase, was PCR amplified (Saiki et al., 1988) and restriction patterns were analyzed. By this method, differences were detected in the rhizobacterial community structure of a laboratory-grown legume, Sesbania macrocarpa,and the salt marsh plant, Spartina alterniflora.

MATERIALS AND METHODS

Organisms. Azotobacter vinelandii ATCC 13705 and Azotobacter chroococcum ATCC 480 were grown in a N-free Burk (Burk, 1930) medium. Vibrio natriegens ATCC 14048 was grown in a minimal salts medium (Forsberg et al., 1970) and Pseudomonas aeruginosa ATCC 9027 on nutrient agar (Difco, Detroit, Mich.). For soil inoculation experiments, bacteria were grown at 30C to late exponential phase.

Plant material. Sesbania macrocarpa Muhl. were grown from seed and cultured in an artificial sediment that was contained in styrofoam pots of a size 7.5 cm x 5.5 cm (Weber et al., 1995). Seedlings were inoculated with 200 mg of Rhizobium spp. (Nitragen Corp., Milwaukee, Wis.) as they were planted in sediment and fertilized with a N-free nutrient solution (Hoagland and Arnold, 1950) three times per wk. Rhizospheres, operationally defined as roots and soil attached to roots after moderate shaking, were harvested after six weeks, and root nodules were removed prior to DNA extraction.

Spartina alterniflora Loisel. was collected from Bayou Texar (Pensacola, FL). Plants were removed from the marsh, and the roots and attached rhizosphere soil were processed for extraction of total DNA.

Direct DNA extraction. Plant roots and attached rhizosphere soil (10 g), an equal weight of washed zirconium beads (0.1 mm dia.), and 10 ml bead-beating buffer (10 mM Tris.HCl, pH 8.0; 150 mM NaCl; 100 mM EDTA; 4% sodium dodecyl sulfate [SDS]) were added to a bead-beater cup (Biospec Products, Bartlesville, Okla.). The mixture containing the soil was beaten on ice for 15 s with a 1 min interval between beating. This was repeated four additional times. After sample removal to a 50 ml Oakridge centrifuge tube, the bead-beater cup was washed with 3 ml of wash buffer (10 mM Tris.HCl, pH 8.0). This was added to the sample and centrifuged for 8 min at 8,000 x g and 4C. The resulting pellet was washed with 3 ml of wash buffer and centrifuged again. Two grams of acid-washed polyvinylpolypyrrolidone (PVPP; Sigma Chemical Co., St. Louis, MO) (Evans et al., 1972) was added to the combined fluid fractions and incubated on ice for 30 min, followed by centrifugation. Wash buffer (3 ml) was added to the pellet and centrifuged again. One volume of phenol, saturated with 10 mM Tris (pH 8.0) and 1 mM EDTA (TE), was added to the combined fluid fractions, vortexed briefly, and centrifuged. One volume of chloroform:isoamyl alcohol (24:1) was added to the resulting aqueous phase, vortexed briefly, and centrifuged again. Glycogen (20 µg ml^-1; Boehringer-Mannheim, Indianapolis, Ind.), ammonium acetate (2.5 M final concentration), and 2 volumes of ethanol were added to the supernatant. The DNA was precipitated overnight at -20C, followed by centrifugation for 20 min at 10,000 x g and 4C. The pellet was resuspended in 400 µl of TE.

DNA purification. CsCl (4.50 g), 5 mg EtBr and 100 µl of soil DNA extract were added to an ultracentrifuge tube and brought to 5 ml with distilled water. The density gradient was achieved by ultracentrifugation at 80,000 rpm (NVT90 rotor, 4 h in a Beckman L 80 ultracentrifuge). The DNA band was recovered in approximately 2 ml and was added to an equal volume of sterile distilled water and 2 volumes of cold ethanol. The DNA was precipitated for 30 min at -20C, followed by centrifugation at 10,000 x g for 15 min. The DNA pellet was resuspended in 200 µl of TE. One volume of TE-saturated phenol was added to the DNA, mixed by inversion, and centrifuged. The supernatant was added to one volume of chloroform:isoamyl alcohol (24:1), mixed by inversion and centrifuged again. The DNA in the supernatant was combined with 20 µg

ml^-1 glycogen, ammonium acetate (2.5 M final concentration), and 2 volumes of ethanol, precipitated at -20C for several hours, centrifuged, and the pellet was resuspended in 50 µl of TE.

PCR amplification. The purified rhizosphere DNA extracts were diluted in distilled water, and 50 - 100 ng DNA (determined by A₂₆₀; Sambrook et al., 1989), was used as template in PCR. Whole cells (A.vinelandii, A. chroococcum, V. natriegens) were picked from agar media, boiled in water 5 min, and 2 µl was added directly to the PCR reaction. The primers (Zehr and McReynolds, 1989) are degenerate and amplify a 359-bp region of nifH. The reaction volume was 100 µl and consisted of 50 mM KCl, 10 mM Tris.HCl (pH 9.0), 1.0% Triton X-100, 6.7 mM MgCl₂, 0.2 mM each dATP, dCTP, dGTP, dTTP, 100 pmol each primer, and 2.5 Units of Taq DNA polymerase (Promega, Madison, WI). Deionized formamide (1%) was added to the whole-cell reactions to increase specificity (Sarkar et al., 1990). Thermal cycling conditions were an initial 3 min at 93C followed by 25 cycles of 1 min 12 sec at 93C, 1 min at 57C, and 1 min 30 sec at 70C, and a final extension for 3 min at 70C. The Taq DNA polymerase was added under "hot-start" conditions after 4 min of denaturation. PCR reactions containing radioisotope were as described above with the addition of 5 µCi of [- ³²P]dCTP per sample. PCR products (5 µl) were analyzed by electrophoresis on a 2% agarose gel containing 0.5 µg ml^-1 EtBr.

Restriction digests. Amplified nifH sequences (6-12 µl) were restriction-enzyme digested overnight with Cfo I, Hae III, or Dde I (Promega, Madison, Wis.). Digested samples (10 µl) were electrophoresed on an 8% polyacrylamide gel (17 x 15 cm) in Tris-acetate-EDTA (TAE) for 6 h at 100 V (Sambrook et al., 1989). After electrophoresis, the gel was dried and exposed on Kodak X-OMAT film.

Soil inoculation. To determine extraction efficiencies and detection limits, an artificial sediment, as described above, was used for soil inoculation experiments. Cell concentrations of A. vinelandii and V. natriegens were determined by direct counts of bacterial cells using a DNA-staining fluorochrome, 4'6-diamidino-2-phenylindole (DAPI) (Porter and Feig, 1980). The cells were diluted in 100 mM phosphate buffer (pH 6.9), 170 mM NaCl, and 2 µl of each dilution (5 x 10⁷ - 5 x 10²) was added to 10 g of wet soil. Inoculated soils were immediately frozen at -20C and maintained at this temperature until DNA extraction and purification as described above.

RESULTS AND DISCUSSION

Extraction, purification and radiolabelling of DNA. Cell lysis by bead-beating was simple and rapid. More et al. (1994) reported that bead-beating was more efficient than a freeze-thaw method for separating DNA from other cell components (Tsai and Olson, 1991). Other studies found that contamination by humic material was less when cells were lysed by bead-beating than with an SDS/heat treatment (Marsh and Wellington, 1994), and when crude DNA was extracted with cold rather than hot phenol (Smalla et al., 1993). Because of soil organic and other contaminants, we were unable to PCR amplify the extracted DNA without purification by CsCl-EtBr gradient centrifugation. We found that the DNA recovery from the CsCl-EtBr gradient was optimum when samples were centrifuged for 80,000 rpm for at least 4 h, rather than at a lower speed and longer time. A low-melt agarose purification method (Herrrick et al., 1993) also yielded amplifiable DNA in about the same amount of time as did CsCl-EtBr gradient centrifugation. However, the yield of amplified sequences, as determined by band intensity on an EtBr-stained gel, was consistently higher after purification by CsCl-EtBr gradient centrifugation.

The majority of purified DNA was from 2-9 kb in length (Figure 1). The average yield of purified DNA was about 5.5 µg per gram (fresh weight) of root and attached soil. When we eliminated the phenol and chloroform extractions after gradient centrifugation, the yield increased about 5-fold, as determined by comparisons to DNA standards on an EtBr-stained gel. However, without the organic extractions, an additional dilution of template DNA in water was necessary for successful PCR amplification.

PCR products were detected by EtBr staining (Figure 2), but unambiguous detection of restriction fragments from environmental DNA was possible only with the use of radioisotopes. Hybridization of the restriction fragments to a labeled probe required several additional steps, and some fragments were not detected after transfer of DNA by electroblotting from a polyacrylamide gel to a hybridization membrane (data not shown). Detection of restriction fragments by [-³²P]dCTP incorporation into the PCR product was rapid and simple.

RFLP analysis of amplified nifH from pure cultures. To determine the suitability of various restriction enzymes for RFLP analysis, PCR-amplified nifH from pure cultures of Azotobacter spp. and from V. natriegens was restriction digested with four- and five-base cutting enzymes (Figure 3). Hae III (4-base cutter) cut all three species tested and the patterns obtained were unique to each organism, whereas Cfo I (4-base cutter) and Dde I (5-base cutter) did not cut the amplified nifH sequence of V. natriegens(Figure 3). Restricted fragments smaller than 118 bp were not detectable; therefore, some fragment lengths from individual species did not total the expected 359 bp (Figure 3). Restricted fragments from combined species totaled more than 359 bp; this is explained by the heterogeneity of nifH sequences from the mixed population. These results indicate that the complexity of the restriction patterns may reflect the complexity of the natural population of nifH-bearing bacteria.

Soil inoculation. We assessed the efficiency of the methods by determining the limit of detection of the amplified nifH sequences in an inoculated synthetic soil. Previous soil inoculation experiments using natural autoclaved soils did not allow detection of the introduced target DNA because of a high background of detectable nifH sequences. The synthetic soil was useful because it lacked interfering, indigenous N₂-fixing bacterial DNA, as demonstrated by the absence of PCR product in the uninoculated control. The detection limit for PCR-amplified nifH DNA was 10⁴ cells each of V. natriegens and A. vinelandii per gram of soil (Figure 4A). HaeIII cuts nifH from both species, whereas CfoI will cut only the A. vinelandii nifH sequence (Figure 3). Both species were detected at a concentration each of 106 cells g-1 of soil, as evidenced by the restriction fragments produced with Cfo I and Hae III (Figure 4B). These are the same patterns as those observed in Figure 3.

RFLP analysis of nifH amplified from rhizospheres. S. macrocarpa, a legume found in flooded and wet soils, forms N₂-fixing nodules on the roots following infection of the root hairs by rhizobia. The rhizosphere of the inoculated S. macrocarpa should be enriched for this group of diazotrophs, owing to the lack of native N₂-fixing bacteria in the synthetic soil. The S. alterniflora rhizosphere may support a more diverse population of N₂-fixing bacteria, such as Vibrio spp. (Payne et al., 1961; Urdacai et al., 1988), cyanobacteria (Jones, 1974; Valiela and Teal, 1979), and the sulfate-reducing bacteria (Herbert, 1975, Postgate and Kent, 1985). The results in Figure 5 support these assumptions. The restriction patterns produced by Cfo I and Hae III from the S. macrocarpa-amplified nifH sequences implied the presence of a single nifH product. Conversely, the complex restriction patterns from the amplified S. alterniflorasequences suggest a more diverse population of N₂-fixing rhizobacteria.

These results demonstrate that the methods used in this study can be used to characterize and compare the community structure of N₂-fixing bacteria from different soil sources. The DNA extraction and purification procedures could be applied to studies on the soil species composition of other groups of naturally occurring microbial communities. The methods may also prove useful for studying population dynamics in response to a stressor or for tracking particular groups of N₂-fixing bacteria in the soil.

ACKNOWLEDGEMENTS

We are grateful to Stephen Francesconi, Wade Jeffrey, Stephanie Willis, and Yujing Yang for useful advice and to Richard Devereux for helpful discussions and critical review of the manuscript. We thank Matthew A. MacGregor for the laboratory-culture of Sesbania macrocarpa and for assistance with materials that comprised the artificial sediments.

This work was supported by a cooperative agreement (CR-822236-01-0) between the EPA Environmental Research Laboratory, Gulf Breeze, FL. and the University of West Florida.

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Figure 1. Total DNA extracted from synthetic and natural soil. Lane 1, soil inoculated with 5 x 10⁶ cells g^-1 each of V. natriegens and A. vinelandii; lane 2, no cells; lanes 3 and 4, rhizosphere samples of S. macrocarpa and S. alterniflora, respectively; lane 5, 500 ng of Hind III digest of lane 1 DNA. Lanes 1,2 and 3 are synthetic soil and lane 4 is Bayou Texar salt marsh soil. Samples are aliquots (20% of the total volume) of purified DNA.

Figure 2. Agarose gel of amplified nifH sequences. Lanes 1 and 2, S. macrocarpa and S. alterniflora rhizosphere, respectively; lane 3, A. vinelandii. Samples are aliquots (10 µl) of each PCR reaction. Molecular size standards (base pairs) are indicated on the left.

Figure 3. Restriction patterns of amplified nifH sequences. Restriction enzymes are (lane groups): (1), Dde I; (2),Cfo I; (3), Hae III; (4), no enzyme used. Within lane groups, whole cell PCR of: lane a, A. vinelandii; lane b, A. chroococcum; lane c, V. natriegens; lane d, one PCR reaction using whole cells of all 3 species; lane m, X174 Hae III molecular size marker. Samples are aliquots (17 µl) of each PCR reaction digested in 30 µl total volume and EtBr-stained following electrophoresis on an 8% polyacrylamide gel.

Figure 4. Detection of amplified nifH sequences in inoculated soil. Sequences were labeled by the addition of [-³²P] dCTP to PCR reactions. Panel A is an autoradiogram of amplified nifH sequences. Soil was inoculated with V. natriegens and A. vinelandii in equal concentrations each of (lanes): (1), 5 x 10⁶; (2), 5 x 10⁵; (3), 5 x 10⁴; (4), 5 x 10³, cells g^-1 of soil. The 8% poloyacrylamide gel in lane 1 was exposed on X-Omat for 24 h, and lanes 2-4 were exposed for 72 h. Panel B is an autoradiogram of restriction patterns of amplified nifH sequences. Lane groups (1) and (2) were from soils inoculated with 5 x 10⁶ and 5 x 10⁵ cells, respectively, of each microorganism as in Panel A. Lanes a and b are amplified nifH sequences digested with Hae III and Cfo I, respectively. The molecular size standards (base pairs) are indicated on the left.

Figure 5. Autoradiogram of restriction patterns of amplified soil nifH sequences labeled during PCR with [-³²P] dCTP. Restriction enzymes are indicated over the lanes: (1), Dde I; (2), Cfo I; (3), Hae III; (4) Cfo I and Hae I; (5), no enzyme used. Lanes a and b are rhizosphere samples of S. macrocarpa and S. alterniflora, respectively; lane c, whole cell PCR of P. aeruginosa. Molecular size standards (base pairs) are indicated on the right.