APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Apr. 1991, p. 1070-1074 0099-2240/91/041070-05$02.00/0 Copyright © 1991, American Society for Microbiology

Vol. 57, No. 4

Rapid Method for Direct Extraction of DNA from Soil and Sediments YU-LI TSAI

AND

BETTY H. OLSON*

Environmental Design and Analysis, Program in Social Ecology, University of California, Irvine, California 92717 Received 10 October 1990/Accepted 23 January 1991

A rapid method for the direct extraction of DNA from soil and sediments was developed. The indigenous microorganisms in the soil and sediments were lysed by using lysozyme and a freeze-thaw procedure. The lysate was extracted with sodium dodecyl sulfate and phenol-chloroform. In addition to a high recovery efficiency (>90%), the yields of DNA were high (38 and 12 ,ug/g [wet weight] from sediments and soil, respectively). This method generated minimal shearing of the extracted DNA. The crude DNA could be further purified with an Elutip-d column if necessary. An additional advantage of this method is that only 1 g of sample is required, which allows for the analysis of small samples and the processing of many samples in a relatively short (7 h) period.

Isolation of bacterial DNA from natural environments has become a useful tool with which to study the ecological functions of certain characterized genes that encode important metabolic pathways (25), allow tracking of genetically engineered organisms (8, 10), and reveal bacterial DNA diversity (21, 22) in microbial ecosystems. The application of DNA extraction methods to environmental samples can obviate the need for cell cultivation, since cell cultivation has the disadvantage of obtaining only a very small proportion of the total microbial community. DNA extraction methods can also help researchers to understand the occurrence of particular bacterial genes in situ through applications of nucleic acid technology. Several techniques have been described for direct detection and extraction of DNA from aquatic environments (3, 6, 13-15, 19), but there are only a few reports related to extraction of DNA from soil and sediments (8, 12, 23). Two current methods used to obtain microbial DNA from soils or sediments are (i) bacterial cell extraction followed by cell lysis and DNA recovery (8, 23) and (ii) direct extraction by alkaline lysis (12). The direct extraction method produces a better DNA yield than the cell extraction method (20). However, both approaches share three disadvantages, i.e., they (i) are lengthy and laborious, (ii) require a relatively large sample size (50 to 100 g), and (iii) produce low yields of DNA. In this study we have developed a rapid method for direct extraction of DNA from soil and sediments by applying a freeze-thaw approach. This method overcomes the drawbacks of the previous techniques. The high yield from a small sample size (1 g [wet weight]) means that the method described in this report can be used efficiently to study in situ gene occurrence and to detect the distribution of genetically engineered microorganisms in the environment.

inated with polynuclear aromatic hydrocarbons (6.5 lug/g [wet weight]) and mercury (0.07 ,ug/g), and the ORT sediments were contaminated with mercury (up to 80 ,ug/g) and other heavy metals (Global Geochemistry Corp., Canoga Park, Calif.; Oak Ridge National Laboratory, Oak Ridge, Tenn.). The heterotrophic bacterial population in both samples was enumerated on plate count agar (Difco, Detroit, Mich.). The indigenous mercury-resistant strains were recovered on plate count agar amended with 25 ,ug of Hg2+ as HgCI2 per ml. Naphthalene degraders were selectively cultivated on a minimal medium saturated with naphthalene vapor (18). Sterile soil and sediment samples were produced by autoclaving at 121°C for 30 min. All soil and sediment quantities are expressed as wet weight unless otherwise indicated. The percentage of water was determined to be 51% in the ORT sediments and 10% in the SC soil (2). Organisms. Two strains (V55 and VNM43) isolated from SC soil were used as seed organisms. They were identified as Pseudomonas luteola and Pseudomonas putida, respectively, by the API Rapid NFT? kits (Analytab Products, Plainview, N.Y.). In the remainder of this paper these strains are referred to as P. luteola V55 and P. putida VNM43. Both strains have the ability to degrade naphthalene and are resistant to mercuric ions. They were grown to late exponential phase in 50% plate count broth (Difco) with 25 pLg of Hg2+ per ml and were washed once with 10 mM phosphate buffer (pH 7.0) and resuspended in phosphate buffer before inoculation into the soils or sediments. The inoculated soils and sediments were shaken at 100 rpm (Orbit Shaker; Lab-Line Instrument, Inc., Melrose Park, Ill.) at 23°C for 30 min prior to DNA extraction. Direct extraction of DNA. SC soil or ORT sediment samples (1 g) were mixed with 2 ml of 120 mM sodium phosphate buffer (pH 8.0) by shaking at 150 rpm for 15 min. The slurry was pelleted by centrifugation at 6,000 x g for 10 min. The pellet was washed again with phosphate buffer, resuspended in 2 ml of lysis solution (0.15 M NaCl, 0.1 M Na2EDTA [pH 8.0]) containing 15 mg of lysozyme/ml, and incubated in a 37°C water bath for 2 h with agitation at 20- to 30-min intervals, and then 2 ml of 0.1 M NaCl-0.5 M Tris-HCl (pH 8.0)-10% sodium dodecyl sulfate was added. Three cycles of freezing in a -70°C dry ice-ethanol bath and thawing in a 65°C water bath were conducted to release DNA from the

MATERIALS AND METHODS

Soil characterization. Subsurface soil samples from a manufactured gas site in Southern California (SC) and sediment samples from a settling pond in Oak Ridge, Tenn. (ORT), were used for DNA extraction. The SC soils were contam*

Corresponding author. 1070

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microbial cells in the soil or sediments. The efficiency of lysis was determined by direct cell counts with phasecontrast microscopy and by acridine orange direct cell counts (7) with fluorescence microscopy (model BH-2; Olympus, Tokyo, Japan) and was found to be greater than 95% for P. putida VNM 43 and P. luteola V55 as well as a Bacillus sp. After the freeze-thaw cycles, 2 ml of 0.1 M Tris-HCl (pH 8.0)-saturated phenol (17) was added and the sample was briefly vortexed to obtain an emulsion. The mixture was centrifuged at 6,000 x g for 10 min. A 3-ml sample of the top aqueous layer was collected and then mixed with 1.5 ml of phenol and 1.5 ml of chloroform mixture (chloroform/isoamyl alcohol ratio, 24:1). A 2.5-ml portion of the resulting extract was further extracted with an equal volume of chloroform mixture. Finally, nucleic acids in the extracted aqueous phase (2 ml) were precipitated with 2 ml of cold isopropanol at -20°C for 1 h or overnight. The pellet of crude nucleic acids was obtained by centrifugation at 10,000 x g for 10 min and then vacuum dried at 23°C. The nucleic acid pellets, which were brownish as a result of the humic materials in both ORT and SC samples, were resuspended in 100 ,ul of TE buffer (20 mM Tris-HCl, 1 mM EDTA [pH 8.0]). The RNA molecules in the crude extracts were removed by incubation with heat-treated pancreatic RNase A (final concentration, 0.2 jig/[ld) for 2 h at 37°C. The RNA-free DNA was then purified with an Elutip-d column (Schleicher & Schuell, Keene, N.H.) attached to a Schleicher & Schuell NA010/27 (0.45-,um [pore size] cellulose acetate) prefilter. DNA was recovered from the column as suggested by the manufacturer. The efficiency of recovery of total DNA from sediment and/or soil crude extracts was determined as the percentage of DNA extracted from a known density of P. putida VNM43 in seeded sterile samples compared with DNA recovered from the same density of pure culture controls. Cell numbers used in all seeding experiments resembled heterotrophic plate counts of the sediments and soils under investigation. The yield of DNA from the direct extraction was expressed in micrograms of crude DNA obtained per gram of unseeded nonsterile samples. Gel electrophoresis. Samples of extracted DNA were analyzed on a 0.7% agarose gel containing 1 p.g of ethidium bromide per ml. To determine the quality of extracted DNA, we analyzed undigested DNA extract and appropriate controls including DNase-digested (0.5 pig/p.l) and RNase-digested (0.2 p.g/p.l) extracts. The concentration of DNA in the crude extract was determined spectrophotometrically at 260 nm. Restriction digestion of DNA was conducted with EcoRI (2 U/p.l), BamHI (1 U/pul), and HindIll (1 U/pul) with incubation for 5 h at 37°C, followed by gel electrophoresis. Hybridization. A 1-kb fragment of the merA gene encoding mercuric reductase was labeled with [ot-32P]dCTP by random priming (specific activity, 108 dpm/pg of DNA) and used as a probe to detect the presence of mercury detoxification genes in the environmental samples. The merA fragment of plasmid pDU1358 was kindly provided by S. Silver. DNA was transferred from the agarose gel to membranes by vacuum blotting (11). The extracted DNA was also filtered onto membranes through a slot blot apparatus (Minifold II; Schleicher & Schuell). Nylon membranes (GeneScreen Plus; NEN Products, Boston, Mass.) and nitrocellulose membranes (BA85; pore size, 0.45 pum; Schleicher & Schuell) were used as solid supports. Hybridization was carried out at 42°C in the presence of 50% formamide, followed by high-stringency washes as previously described (24). The Radioanalytic Imaging System (AMBIS Systems, San Di-

A

B

FIG. 1. Total DNA extracted from ORT sediments. (A) Ethidium bromide-stained agarose gel; (B) autoradiogram of DNA hybridization signals with merA after Southern blot of the gel in panel A. Aliquots (3 ,ul) of total extracts (100 ,ul) were loaded into each well. Lanes: 1, HindlIl-cut lambda bacteriophage molecular size marker (1 p.g); 2, unseeded; 3, unseeded and digested with RNase; 4, unseeded and digested with DNase; 5, seeded; 6, seeded and digested with RNase; 7, seeded and digested with DNase; 8, plasmid size marker from E. coli V517.

ego, Calif.) was used to quantify the radioactivity signals of hybridized DNA on the membranes.

RESULTS AND DISCUSSION

Figure 1A shows an ethidium bromide-stained agarose gel used to visualize the DNA extracted from ORT sediments after DNase and RNase treatments. The addition of 2.5 x 108 cells of P. luteola V55 to 1 g of nonsterile sediments resulted in a larger quantity of DNA than in the unseeded sample. The efficiency of recovery was calculated to be 92% when sterile ORT sediment was seeded with a known cell density of P. luteola V55. The sterile samples (autoclaved or gamma irradiated) did not contain indigenous DNA after extraction. The total DNA yield from nonsterile, unseeded ORT sediments was determined to be 38 ,ug/g (Table 1). The largest DNA was greater than 23 kb, and most DNA was in the size range of 6.5 to 23.1 kb. The DNA-shearing effect of the extraction procedure was less prominent than in the method described by Ogram et al. (12), as evidenced by smearing of smaller fragments. Because the DNA extracts before RNase digestion contained RNA contaminants (Fig. 1A), introduction of RNase during the extraction procedure was necessary. Impurities such as humic materials in the extracts did not affect RNase or DNase digestion, but may affect DNA hybridization efficiency. RNA was not observed in unseeded samples, indicating that RNA degradation or TABLE 1. Yields and efficiency of direct DNA extraction method Sample Sample

Heterotrophic (CFU/g) bacteria

ORT sediments (8.0 + 0.7) x 107 (4.1 +0.1) x 106 SC soil

Recovery(%) efficiency'

Yieldb [wet (pLg/g

92 ± 3 95 4

38 ± 5 12 4

wt])

aThe recovery efficiency was calculated from the total DNA obtained from a known cell density of sterile seeded samples compared with DNA extraction

from bacterial pure cultures. b The yield was measured from nonsterile unseeded samples.

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FIG. 2. RNase treatment of nucleic acids extracted from SC soil. (A) Ethidium bromide-stained agarose gel; (B) DNA hybridization signals with merA after Southern transfer. Aliquots (4 ,ul) of total extracts (100 pLI) were loaded into each well. Lanes: 1, HindIII-cut lambda phage molecular size marker (1 ,ug); 2, sterile unseeded; 3, sterile seeded; 4, nonsterile unseeded; 5, nonsterile seeded; 6 and 7, pure culture of P. putida VNM43; 8, 10 ng of merA 1-kb fragments.

less RNA synthesis was occurring in the nonsterile, uninoculated environmental samples. Southern blot analyses showed that ORT sediments contained merA genes in the indigenous microbial community (Fig. 1B). Small DNA fragments (less than 500 bp) resulting from DNase digestion of the extracted DNA were homologous to partial merA sequences (Fig. 1B, lane 7). The total heterotrophic bacterial population in the ORT sediments was (8.0 + 0.7) x 107 CFU/g, with 17% of the population exhibiting phenotypic mercury resistance. This determination supports the finding of merA sequences in the ORT sediments. Figure 2A illustrates the ethidium bromide-stained DNA extracted from unseeded nonsterile and sterile SC soils and from seeded (7.0 x 108 cells of P. putida VNM43 per g) nonsterile and sterile soils. No evidence of DNA was found in the sterile soil, but DNA was recovered from the nonsterile soil (lanes 2 and 4, respectively). The heterotrophic bacterial density in the nonsterile SC soil was (4.1 + 0.1) x 106 CFU/g. Of these bacteria, 9% were phenotypically mercury resistant and 15% were naphthalene degraders. Most of the DNA fragments extracted from the seeded soils were larger than 23.1 kb (Fig. 2A), indicating that the present protocol does not cause severe DNA shearing. In addition, the DNA eluted from agarose gels could easily be subjected to restriction analyses for further subcloning. The efficiency of extraction and the total recovery of DNA were 95% and 12 ,ug/g, respectively (Table 1). There was no linear correlation between the total yield of DNA and the density of heterotrophic bacteria based on dry weight when two different types of samples (sediment and soil) were used. This discrepancy could be due to the presence of eukaryotic organisms, chemolithotrophic bacteria, nonculturable bacteria, or other microorganisms which were not enumerated during the experiment. On the average, each Escherichia coli cell contains 9.0 x io-9 ,ug of DNA (9). Assuming that this holds true for average bacteria, the density of heterotrophic bacteria from ORT sediments and SC soil would represent 0.72 and 0.037 ,ug of DNA per g, which would amount to 1.9 and 0.3% of total extracted DNA, respectively. These results further

FIG. 3. Autoradiogram of hybridization signals of extracted crude DNA with merA. (A) RNase-treated DNA from nonsterile unseeded (slot 1) and sterile seeded (slots 3 to 8) SC soils. The sterile SC soils were inoculated with (cells of P. putida VNM43 per g [wet weight]): no inoculum (slot 2), 1.1 x 108 (slot 3), 1.1 x 107 (slot 4), 1.1 x 106 (slot 5), 1.1 x 105 (slot 6), 1.1 X 104 (slot 7), and 1.1 X 103 (slot 8). Slots 1, 3, 4, 5, and 6 each contained 1/10 and slots 7 and 8 each contained 1/2 of total DNA extracts. (B) RNase-treated DNA from nonsterile unseeded ORT sediments (slots al to a4), seeded ORT sediments (slots aS to a9), nonsterile seeded SC soils (slots bl to b5), and pure culture of P. putida VNM43 (slots b6 to b9). The nonsterile ORT sediments and SC soil were seeded with 2.5 x 108 p. luteola V55 and 7.0 x 108 P. putida VNM43 per g (wet weight), respectively. Slots: aS, 10 Rg; al, 5 jig; a2 and a6, 1 ,ug; a3 and a7, 0.1 ,ug; a4 and a8, 0.01 ,ug; a9, 0.001 ,ug; bl, 10 ,ug; b2 and b6, 1 ,ug; b3 and b7, 0.1 ,ug; b4 and b8, 0.01 pug, b5 and b9, 0.001 jig of DNA.

support observations made by other investigators, who have reported that only a very small portion of the natural bacterial community is culturable (1, 4, 5, 16). Because the ORT sediment contains more water (51%) than the SC soil (10%), the yields based on dry weight would be 77 and 13 ,ug of DNA per g, respectively. Southern hybridization analyses did not indicate the occurrence of merA gene sequences in the nonsterile unseeded

A

B

FIG. 4. Restriction enzyme digestion patterns of DNA isolated from SC soil. (A) Ethidium bromide-stained agarose gel. (B) Autoradiogram of merA hybridization signals from Southern blot. Lanes 1 to 3 and 4 to 6 each contained 1/10 of DNA extracts from P. putida VNM43 and seeded soil samples, respectively. Lanes: 1' Hindlllcut lambda phage molecular size marker (1.25 p.g); 1 and 4, uncut; 2 and 5, EcoRI cut; 3 and 6, Hindlll cut; 7, 20 ng of merA fragment.

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FIG. 5. Restriction enzyme digestion patterns of DNA extracted from ORT sediments. (A) Ethidium bromide-stained agarose gel. (B) Autoradiogram of merA hybridization signals from Southern blot of gel in panel A. Lanes 2 to 4 and 5 to 7 each contained 1/30 of DNA extracts from unseeded and seeded samples, respectively. Lanes 8 to 10 and 11 to 13 contained 1/20 of Elutip-d column-purified DNA from unseeded and seeded samples, respectively. Lanes: 1, HindIll-cut lambda phage molecular weight marker with ladder of 23.1, 9.4, 6.5, 4.4, 2.3, and 2.0 kb; 2 and 8, unseeded and uncut; 3 and 9, unseeded and EcoRI cut; 4 and 10, unseeded and BamHI cut; 5 and 11, seeded and uncut; 6 and 12, seeded and EcoRI cut; 7 and 13, seeded and BamHI cut.

SC soil (Fig. 2B, lane 4), but the direct application of DNA to membranes by the slot blot hybridization technique showed that crude DNA extracted from the same soil did contain merA genes (Fig. 3A, slot 1). The discrepancy between these two observations was due to incomplete transfer of DNA from the gel to the filters during Southern

blotting. Therefore, the existence of mercury-resistant bacterial strains such as P. putida VNM43 and P. luteola V55 in the polynuclear aromatic hydrocarbon-contaminated SC soils was further supported by the gene probe data. The DNA extracts from seeded sterile (Fig. 2B, lane 3) and nonsterile (lane 5) soils revealed smaller DNA fragments (between 1 and 20 kb) containing homologous merA sequences. This shearing is probably due to the grinding effect of the sand particles in SC soil during the extraction procedure because pure-culture controls (lane 6 and 7) did not exhibit this effect. The two soils differ in cation exchange capacity and particle size distribution, with SC soils having low cation exchange capacity (9 mEq/100 g) and high sand concentration (18.4%), and ORT sediments having high cation exchange capacity and low sand concentration (Uni-

versity of California, Riverside, Department of Soil and Environmental Sciences; Oak Ridge National Laboratory). Figure 3A shows the merA hybridization signals from the crude DNA extracted from sterile SC soil seeded with different quantities of P. putida VNM43. Signals were detected in a 50% aliquot of total DNA extract when as few as 1.1 x 104 cells per g (wet weight) were inoculated into soil, indicating that the sensitivity was 5.5 x 103 cells per g. A comparative sensitivity of 4.3 x 104 cells per g (dry weight) was reported for the cell extraction method (8). Figure 3B illustrates hybridization signals of putative merA genes extracted from nonsterile unseeded and seeded ORT sediments and nonsterile seeded SC soils. As the extracted DNA was normalized at 1 ,ug, DNA from seeded samples (Fig. 3B, slots a6 and b2) demonstrated higher merA hybridization signals (7,049 and 24,477 cpm) than that from unseeded samples (800 cpm; slot a2). As would be expected, the signal for the SC soils inoculated with a higher cell density (slot b2) had a higher-intensity signal of merA sequences than did ORT sediments inoculated with a lower cell density (slot a6). The pure-culture DNA control showed the highest signal

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(28,648 cpm) in 1 ,ug of DNA extract. The radioactivity counts for hybridized DNA were lower from soil and sediment samples than from pure cultures, indicating that DNA had absorbed into the soil particles or that more nonhomologous merA sequences were present in the former samples. The detection limit of homologous merA DNA was 10 ng of total extracted DNA from both P. putida VNM43 pure culture and P. putida VNM43-seeded SC soils (Fig. 3B). The merA hybridization signals were found in 1 ,ug of total DNA extracted from nonsterile unseeded ORT sediments. In comparison, Ogram et al. (12) required 1.2 ,ug of total DNA to detect nif nitrogen fixation genes from sediments other than those used in the present study. In the present study there is no indication that humic materials in the crude extract affect DNA-DNA hybridization (Fig. 3). Figure 4 shows the restriction patterns of DNA extracts from a pure bacterial culture and from a seeded SC soil. The unpurified DNA extract was clearly digested by Hindlll but only slowly digested by EcoRI. The application of restriction enzyme analyses to directly extracted DNA may allow more precise studies of in situ gene rearrangements or amplification. The impurities in ORT sediments did affect the restriction enzyme digestion (Fig. 5). EcoRI and BamHI did not digest DNA extracts from either unseeded or seeded samples within the 5-h incubation period. The DNA yield suffered a 40% loss during the Elutip-d column purification step, and the purified DNA (A260/A280 = 1.8) was cut by EcoRI but not by BamHI. Therefore, for studying in situ gene transfer the purification step might be needed depending on the soil type, but it is not necessary for determination of gene occurrence. In conclusion, this rapid method for the direct extraction of DNA needs only 1 g of soil or sediments and can detect the presence of a target gene, such as merA, from a minimum of 5,000 bacterial cells. It is simple and can be completed in 7 h. In addition, because of the small sample size (1 g), a large number of soil or sediment samples can be analyzed at one time. Finally, the high yield in combination with the high quality of DNA will enable microbial ecologists to study DNA from environmental samples in a more detailed fashion by molecular biology techniques.

6.

7. 8.

9.

10. 11. 12. 13.

14.

15.

16. 17. 18.

19. ACKNOWLEDGMENTS

This study was funded by grant 8000-25 provided by the Electric Power Research Institute. We are grateful to Marie Park for developing the autoradiograms. We also thank 0. A. Ogunseitan, P. A. Rochelle, and C. C. Tebbe for valuable discussions and suggestions and R. Turner for provision and analysis of ORT sediments. 1. 2.

3. 4. 5.

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