Strategy for Extracting DNA from Clay Soil and Detecting a Specific ...

APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Sept. 2009, p. 6017–6021 0099-2240/09/$08.00⫹0 doi:10.1128/AEM.00211-09 Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Vol. 75, No. 18

Strategy for Extracting DNA from Clay Soil and Detecting a Specific Target Sequence via Selective Enrichment and Real-Time (Quantitative) PCR Amplification䌤 Kweku K. Yankson† and Todd R. Steck* Department of Biology, The University of North Carolina at Charlotte, Charlotte, North Carolina 28223 Received 29 January 2009/Accepted 20 July 2009

We present a simple strategy for isolating and accurately enumerating target DNA from high-clay-content soils: desorption with buffers, an optional magnetic capture hybridization step, and quantitation via real-time PCR. With the developed technique, ␮g quantities of DNA were extracted from mg samples of pure kaolinite and a field clay soil. was determined from binding isotherms. The DNA used was a 1,040-bp region of the gfp gene (GC content, ⬃38%) PCR amplified from pGFPmut3.1 (3, 9, 14) by using primers specific for this gene (5⬘-ACTGGAAAGCGGGCAGTG and 5⬘-AAA CGCGCGAGACGAAAGGG). Kaolinite was sterilized by gamma irradiation by exposure to Cobalt-60 until 70 kGy was delivered (Radiation Science and Engineering Center, Penn State University, University Park, PA), a dose sufficient to kill all living microbes (25) and to directly and indirectly damage DNA (18, 30). By use of the PCR-generated gfp DNA fragment, adsorption data were fit to the Langmuir (33) and Freundlich (29) models in accordance with the method of Pietramellara et al. (27), with real-time quantitative PCR (qPCR) used to quantitate DNA. Adsorption data were also used to conduct a Scatchard plot analysis (39) of DNA binding to kaolinite particles. DNA was adsorbed to clay by using the protocol of Cai et al. (7), modified as follows. Twenty-two milligrams of kaolinite was brought up to 1 ml with distilled-deionized water and vortexed. To a 0.1-ml sample, a solution containing a known amount of the DNA fragment was added, and the resulting mixture was brought up to 1 ml with distilled, deionized water, with gentle shaking at 25°C over a 2-h period. The mixture was then centrifuged at 16,125 ⫻ g for 20 min at room temperature, the supernatant removed, and the pellet stored at ⫺20°C. The Scatchard Plot (q) indicates that there are 11.5 DNA binding sites per mg of kaolinite (source clay label [8], KGa1b), and the Kd value of 0.0786 reflects kaolinite’s strong affinity for DNA (R2, 0.943) (see reference 39 for explanation of parameters). The Freundlich coefficient (Kf) was 1.71 (1/n, 0.758; R2, 0.978) (see reference 29 for explanation of parameters). The Langmuir adsorption isotherm (⌫max) reveals that kaolinite can bind 81.3 ␮g DNA per mg kaolinite (L, 0.0014; R2, 0.846) (see reference 33 for explanation of parameters). These parameters indicate that kaolinite is a high-capacity clay (cation exchange capacity [4], 3 cmol/kg) that strongly binds DNA and is appropriate for use in establishing DNA extraction conditions. Field soil: removal of PCR inhibitors. Before desorption of DNA from field soil was examined, a procedure for removal of qPCR inhibitors was tested. Three humic acids, purchased from the International Humic Substance Society (St. Paul,

Isolating and characterizing DNA sequences for use in molecular methods are integral to evaluating microbial community diversity in soil (6, 21, 22, 24, 37). Any isolation protocol should maximize nucleic acid isolation while minimizing copurification of enzymatic inhibitors. Although several methods that focus on extraction of total community DNA from environmental soil and water samples have been published (7, 21, 26, 34), the lack of a standard nucleic acid isolation protocol (32) reflects the difficulty in accomplishing these goals, most likely due to the complex nature of the soil environment. DNA extraction is especially difficult for soils containing clay (3, 5), given the tight binding of DNA strands to clay soil particles (7, 10, 20). Additionally, extracellular DNA binds to and is copurified with soil humic substances (10), which inhibit the activity of enzymes such as restriction endonucleases and DNA polymerase (6, 13, 23). Although clay-bound DNA can be PCR amplified in the absence of inhibitors (1), it is often the case that inhibitors are present in the soil environment, among them bilirubin, bile salts, urobilinogens, and polysaccharides (40). Of these inhibitors, humic substances have been found to be the most recalcitrant (36). A promising technique for isolating specific target sequences from soil particles and enzymatic inhibitors is the magnetic capture hybridization-PCR technique (MCH-PCR) presented by Jacobsen (19) and used to obtain high detection sensitivities (11, 38).We have found no evidence in the published literature of the use of MCH-PCR on soils that have high clay contents and here present a three-step strategy for isolating specific DNA sequences from the most difficult soil environment—clay that contains humic substances—and enumerating a specific target sequence from the crude extract. Kaolinite clay: DNA binding. To determine whether kaolinite would be a suitable clay type for use in optimizing DNA desorption protocols, the nature of DNA binding to pure kaolinite (KGa-1b) (4) (Clay Minerals Society; Chantilly, VA)

* Corresponding author. Mailing address: Department of Biology, The University of North Carolina at Charlotte, Charlotte, NC 28223. Phone: (704) 687-8534. Fax: (704) 687-3128. E-mail: [email protected]. † Present address: Phase Forward, Clarix Products Group, Two Radnor Corporate Center, Radnor, PA 19807. 䌤 Published ahead of print on 24 July 2009. 6017

0.1 0.14 7.00 5.00 47 49 161 142 43 40 43 40 546 1032 28.0 28.0 62.0 62.0 12 12 0 0 5.7 5.9 b

a

Percentage of cation exchange capacity occupied by the basic cations Ca, Mg, and K. Measure of the active acidity in the soil solution. c Percentage of cation exchange capacity occupied by calcium. d Percentage of cation exchange capacity occupied by magnesium.

1.2 1.0 90.0 91.0 12.0 10.7 1.05 1.12 Mineral Mineral Field soil Kaolinite clay

0.22 0.32

Soluble-salt index S index Cu index Zn availability index Zn index Mn index % Mgd % Cac K index P index Exchangeable Soil acidity (meq/ pHb 100 cm3) % Base saturationa Wt/vol (g/cm3) % Humic matter Soil classification Source

TABLE 1. Elemental analyses of field soil and kaolinite clay

MN)—Summit Hill, Elloit Soil, and Peat—were taken to be representative of soil humic acids (15) and were used in Taq DNA polymerase inhibition assays. DNA fragment adsorption and soil sterilization were carried out as described for kaolinite; gamma irradiation has been shown to have the least effect on the physical and chemical properties of soil (35) and both directly and indirectly damages DNA (18, 30). Soil was collected from an unused field on the campus of the University of North Carolina at Charlotte (Charlotte, NC). Samples were isolated using an auger at depths of 53 cm and 91 cm below the surface and then combined for analysis and use. Particle size analysis of soil sample revealed 4.44% sand, 63.7% silt, and 31.8% clay. The results of elemental analyses for both clay and soil conducted by the North Carolina Department of Agriculture & Consumer Services, Agronomic Division, are given in Table 1. MCH, modified from the method of Crosby and Criddle (11), was used to remove enzymatic inhibitors. The desired volume of the postdesorption supernatants was heated at 95°C for 5 min and quenched on ice for 10 min, and 1 ␮l of a 10-pmol/␮l sample of the 98-bp gfp capture probe was added. Four hundred fifty microliters of DIG EasyHyb hybridization buffer (Roche Applied Science, Indianapolis, IN) was added, and the sample was rotated gently at 37°C for 2 h. Twenty minutes before the end of the hybridization period, magnetic beads (Promega, Madison, WI) were prepared by washing thrice with 2⫻ SSC buffer (1⫻ SSC is 0.15 M NaCl plus 0.015 M sodium citrate) via repipetting. Twenty microliters of a 1-mg/ml sample of beads was used per capture reaction (19). The supernatant from the third wash was removed just before the probe-containing sample was added. The target, probe, and beads were incubated at 37°C with gentle rotation for 20 min. Magnetic beads were separated from the solution with a magnetic separation stand (Promega, Madison, WI), and the supernatant was removed. The beads (with capture probe plus target sequence) were washed with 0.3 ml of 2⫻ SSC and then thrice with 0.3 ml of 1⫻ SSC. Each wash was incubated with gentle rotation at 35°C for 5 min. Captured DNA was eluted in 50 ␮l of nuclease-free water, 1 ␮l of which was used directly as a template in qPCR reactions. Target sequences were purified and detected from crude soil extracts containing humic acid at all concentrations tested: 1, 5, 15, 30, and 300 ␮g/ml. When the MCH purification step was omitted, however, qPCR was able to detect the DNA fragment in soil containing humic acid at concentrations of only 1 and 5 ␮g/ml. Detection sensitivity of qPCR and MCH-qPCR. Before qPCR was used on desorbed DNA, the detection limit/sensitivity (the smallest weight amount or copy number of DNA that could be captured by MCH and amplified by qPCR) and limit of accurate quantification (the smallest amount or copy number of DNA that could be captured by MCH and accurately quantified by qPCR) were determined. The primers used in qPCR (5⬘-AGCG TTCAACTAGCAGACCA and 5⬘-AAAGGGCAGATTGTGT GGAC) were designed with the Primer3 software program (31) to generate a 100-bp gfp product optimal for qPCR amplicon generation. The 98-bp MCH capture probe (5⬘-TTGTTAAAGT GTGTCCTTACAGCTATGACCATGATTACGCCAAGCTT GCATGCCTGCAGGTCGACTCTAGAGGATCC) was designed using the VectorNTi software program (Invitrogen; Carls-

APPL. ENVIRON. MICROBIOL. Na level (meq/100 cm3)

YANKSON AND STECK

Cation exchange capacity (meq/100 cm3)

6018

VOL. 75, 2009

METHOD FOR EXTRACTING DNA FROM CLAY SOIL

6019

FIG. 1. Contribution of each buffer to DNA desorption. (A) Kaolinite; (B) field soil. Data are averages of results from duplicate experiments. Error bars indicate the standard errors of the means.

bad, CA) and was synthesized and polyacrylamide gel electrophoresis purified by Bioneer (Alameda CA). Capture probe length was chosen based on published information regarding capture probe length (12, 19, 38). To obtain the amplicons, PCR products were gel purified using a QIAquick gel extraction kit (Qiagen. Inc., Valencia, CA). Standards and samples (postdesorption supernatants or MCH-purified DNA fragment) were coamplified to quantify the starting amount of DNA in the samples. qPCR was performed with a LightCycler 2.0 system (Roche Applied Science, Indianapolis, IN). Reaction mixtures were prepared using a QuantiTect SYBR green PCR kit (Qiagen, Valencia, CA) as follows: 1 ␮l DNA template, 3 ␮l nuclease free water, 5 ␮l master mix, and 0.5 ␮l each 5 ␮M primer. A final reaction volume of 10 ␮l was loaded into each microcapillary glass vessel. The template DNA was denatured at 95°C for 15 min, with a 20°C/s slope. Amplification occurred over 40 cycles of denaturation at 94°C for 15 s, primer annealing at 52°C for 30 s, and product extension at 72°C for 15 s (each step with a 20°C/s slope). An additional step was included for fluorescence data acquisition at 76°C (⬃3°C below the melting temperature of the product) for 5 s. Gels were run initially to confirm product identity. Melting curves were also plotted to determine the identity of the PCR products and to identify potential instances of nonspecific product formation (28). The reaction conditions for generating product melting curves consisted of a 94°C sample heating step for 0 s, followed by cooling at 50°C for 30 s. The sample was then slowly heated (0.2°C/s slope) to 99°C, while fluorescence was monitored. qPCR was performed both directly on desorbed DNA and after an additional MCH purification step. DNA at the lowest concentration tested, 3.46 ⫻ 10⫺12 ng/␮l, was detected using qPCR without MCH purification; addition of the MCH purification step decreased the detection limit to 7.67 ⫻ 10⫺10 ng/␮l. Regardless of MCH purification, qPCR had a limit of accurate quantification of 3.46 ⫻ 10⫺8 ng/␮l. Kaolinite clay: DNA desorption. Multiple buffers have been used to desorb DNA from soil. Here, five buffers were used sequentially according to the methods of Pietramellara et al. (27) and Cai et al. (7), modified as follows: 40 mg of powdered skim milk per gram soil was added to each of the Tris (10 mM Tris [pH 7]), NaCl (100 mM NaCl [pH 7]), and phosphate (100 mM NaPO3]6 [pH 7]) buffers prior to desorption. The pellet obtained from the adsorption procedure was washed (three times each) with 1 ml of Tris buffer, 1 ml of NaCl buffer, and 1 ml of phosphate buffer. Supernatants from each buffer wash were stored at 4°C. Desorbed DNA was stored for at most 1

day; in most cases, desorption, MCH, and qPCR were performed on the same day. After the third phosphate buffer wash, the pellet was washed with 0.1 ml of a saline desorption solution (SDS; 17 mM lactic acid, 3 mM KH2PO4, 27 mM Na2HPO4, 0.23 mM MgSO4, 11 mM NH4Cl, 19 ␮M CaCl2, 0.5 ␮M FeSO4, 86 mM sodium pyrophosphate, 57 mM EDTA). This particular wash step involved adding 0.1 ml of SDS to the clay pellet and vortexing the mixture at 1,200 ⫻ g for 1 h at room temperature. Because the kaolinite used in these experiments was pure and free of PCR inhibitors, DNA in the postdesorption supernatants was quantified directly by qPCR amplification, without a prior MCH purification step. An average, from two experiments, of 7.3% of DNA was recovered. The desorption yield after each wash step was then determined. Washing the clayDNA fragment sample with distilled water recovered 33.7% ⫾ 8.9% of the total recoverable DNA, suggesting that this fraction of DNA fragment was loosely bound to the soil particles (33). Of the remaining four buffers used to remove the tightly bound DNA fragment, phosphate removed more DNA (⬃45%) than the other three buffers combined (Fig. 1A). The use of water and phosphate buffer accounted for over 75% of the bound DNA fragment that was recovered. Desorbed DNA can rebind clay particles, reducing the net desorption efficiency. Addition of nonfat powdered milk increased DNA fragment recovery by approximately 50% when water, Tris, and NaCl buffers were used (Fig. 2A). That skimmed milk reduced the rebinding of desorbed DNA fragment is consistent with reports that skim milk competes with DNA for binding sites on clay minerals (16, 17). Phosphate and SDS buffers were not tested as in this experiment, because DNA was quantitated via qPCR, and agents in these two buffers inhibit PCR amplification. When each buffer is examined separately, the effect of milk on desorption is higher with Tris and NaCl than with water (Fig. 2B). Field soil: DNA binding and desorption. The desorption protocol described above was used with field soil having a clay content of 31.8%, with one additional step. MCH was included as a pre-PCR purification step due to the presence of PCR inhibitors in field soil. When all five buffers were sequentially used, similar amounts of DNA fragment were desorbed from the soil (783 ⫾ 280 ng) and from clay (622 ⫾ 14.6 ng). However, because more DNA fragment bound to soil (35.0 mg) than to clay (8.65 mg), the recovery efficiency from soil, 2.2%, was lower than the 7.3% recovered from clay. Although only approximately 7% of the total amount of

6020

YANKSON AND STECK

APPL. ENVIRON. MICROBIOL.

FIG. 2. Effect of skim milk on desorption of bound DNA. (A) Effect on combined efficiency of desorption with water, Tris, and NaCl buffers. (B) Effect on individual buffer desorption efficiencies. Data are averages of results from duplicate experiments. Error bars indicate the standard errors of the means.

clay-bound DNA fragment was released via the desorption procedure, additional DNA fragment should be obtained by scaling up this procedure. The 7% recovery of bound DNA fragment is significantly less than the 25-to-93% recovery reported by Cai et al. (7), who used multiple buffers. Other authors have also reported desorption efficiencies greater than 7%, using desorption buffers (6). These recovery differences could be due to the method of DNA quantitation used in each study. Quantification of DNA in crude extracts by use of absorbance at only 260 nm is susceptible to errors if non-DNA UV absorbing materials, such as phosphate, are present (Giacomo Pietramellara, University of Florence, personal communication). In control experiments in which suspended kaolinite particles were present in the postdesorption supernatants, we noticed an overestimation of the amount of DNA in test samples when they were quantified using UV spectrophotometry. Micron-size clay particles can reduce the transmission of light through a sample. The use of UV absorbance in quantifying DNA recovered from soil can be validated by examining absorbance at 320 nm. Using MCH as a pre-PCR enrichment step, Tsai et al. (38) report a 10,000-fold increase in detection sensitivity over conventional PCR. Crosby and Criddle (11) have also reported a detection of minority populations as low as 0.01% of the total microbial community population via MCH enrichment and subsequent PCR amplification. We found that MCH functions well as a pre-PCR purification step for samples with humic acids at concentrations that inhibit PCR amplification, which were above 5 ␮g/ml in our experiments. For crude extracts in which inhibitor concentrations do not affect qPCR amplification, MCH can be omitted, reducing the total sequence detection time by up to half. It is possible that MCH may also be susceptible to the presence of complex metal groups in crude extracts that can adversely affect the ability of the streptavidin

coat on the magnetic beads to bond with the biotin moiety on the capture probe. If true, this susceptibility was not found when MCH was conducted on crude extracts from a field soil, suggesting that the technique can be used on a range of environmental soil types. Overall, the developed method was found to be effective at extracting and purifying DNA of sufficient quantity and purity for qPCR detection and quantification from high-clay-content soil types. We thank Nury Steuerwald (Carolinas Medical Center) for invaluable assistance with qPCR troubleshooting; North Carolina Department of Agriculture & Consumer Services, Agronomic Services, for soil and clay analysis; Martha Epps (Department of Geography and Earth Sciences, UNC—Charlotte) for soil particle size analysis; Craig Allan (Department of Geography and Earth Sciences, UNC—Charlotte) for assistance with experimental design; and Candace Davison (Radiation Science & Engineering Center, Pennsylvania State University) for gamma sterilization of the soil samples. This work was supported, in part, by funds provided by the University of North Carolina at Charlotte. REFERENCES 1. Alvarez, A. J., M. Khanna, G. A. Toranzos, and G. Stotzky. 1998. Amplification of DNA bound on clay minerals. Mol. Ecol. 7:775–778. 2. Reference deleted. 3. Andersen, J. B., C. Sternberg, L. K. Poulsen, S. P. Bjorn, M. Givskov, and S. Molin. 1998. New unstable variants of green fluorescent protein for studies of transient gene expression in bacteria. Appl. Environ. Microbiol. 64:2240– 2246. 4. Borden, D., and R. F. Giese. 2001. Baseline studies of the clay mineral society source clays: cation exchange capacity measurements by the ammonia-electrode method. Clays Clay Miner. 49:444–445. 5. Braid, M. D., L. M. Daniels, and C. L. Kitts. 2003. Removal of PCR inhibitors from soil DNA by chemical flocculation. J. Microbiol. Methods 52:389–393. 6. Buergmann, H., M. Pesaro, F. Widmer, and J. Zeyer. 2001. A strategy for optimizing quality and quantity of DNA extracted from soil. J. Microbiol. Methods 45:7–20. 7. Cai, P., Q. Huang, X. Zhang, and H. Chen. 2006. Adsorption of DNA on clay

VOL. 75, 2009

8.

9. 10.

11.

12.

13.

14.

15. 16.

17.

18.

19.

20.

21.

22.

23.

24.

minerals and various colloidal particles from an alfisol. Soil Biol. Biochem. 38:471–476. Chipera, S. J., and L. B. David. 2001. Baseline studies of the Clay Mineral Society source slays: powder X-ray diffraction analysis. Clays Clay Miner. 49:398–409. Cormack, B. P., R. H. Valdivia, and S. Falkow. 1996. FACS-optimized mutants of the green fluorescent protein (GFP). Gene 173:33–38. Crecchio, C., and G. Stotzky. 1998. Binding of DNA on humic acids: effect on transformation of Bacillus subtilis and resistance to DNase. Soil Biol. Biochem. 30:1061–1067. Crosby, L. D., and C. S. Criddle. 2007. Gene capture and random amplification for quantitative recovery of homologous genes. Mol. Cell. Probes 21:140–147. de Moraes, R. R., J. E. Maruniak, and J. E. Funderburk. 1999. Methods for detection of Anticarsia gemmatalis nucleopolyhedrovirus DNA in soil. Appl. Environ. Microbiol. 65:2307–2311. Dong, D., A. Yan, H. Liu, X. Zhang, and Y. Xu. 2006. Removal of humic substances from soil DNA using aluminium sulfate. J. Microbiol. Methods 66:217–222. Fire, A., S. W. Harrison, and D. Dixon. 1990. A modular set of lazZ fusion vectors for studying gene expression in Caenorhabditis elegans. Gene 93:189– 198. Hengpraprom, S., C. M. Lee, and J. T. Coates. 2006. Sorption of humic acids and ␣-endosulfan by clay minerals. Environ. Toxicol. Chem. 25:11–17. Hoshino, Y. T., and N. Matsumoto. 2004. An improved DNA extraction method using skim milk from soils that strongly adsorb DNA. Microbes Environ. 19:13–19. Hoshino, Y. T., and N. Matsumoto. 2007. DNA- versus RNA-based denaturing gradient gel electrophoresis profiles of a bacterial community during replacement after soil fumigation. Soil Biol. Biochem. 39:434–444. Jackson, N. E., J. C. Corey, L. R. Frederick, and J. C. Picken, Jr. 1967. Gamma irradiation and the microbial population of soils at two water contents. Soil Sci. Soc. Am. 31:491–494. Jacobsen, C. S. 1995. Microscale detection of specific bacterial DNA in soil with a magnetic capture-hybridization and PCR amplification assay. Appl. Environ. Microbiol. 61:3347–3352. Khanna, M., and G. Stotzky. 1992. Transformation of Bacillus subtilis by DNA bound on montmorillonite and effect of DNase on the transforming ability of bound DNA. Appl. Environ. Microbiol. 58:1930–1939. Krsek, M., and E. M. H. Wellington. 1999. Comparison of different methods for the isolation and purification of total community DNA from soil. J. Microbiol. Methods 39:1–6. LaMontagne, M. G., F. C. Michel, P. A. Holden, and C. A. Reddy. 2002. Evaluation of extraction and purification methods for obtaining PCR-amplifiable DNA from compost for microbial community analysis. J. Microbiol. Methods 49:255–264. Mangiapan, G., M. Vokurka, L. Schouls, J. Cadranel, D. Lecossier, J. van Embden, and A. J. Hance. 1996. Sequence capture-PCR improves detection of mycobacterial DNA in clinical samples. J. Clin. Microbiol. 34:1209–1215. Martin-Laurent, F., L. Philippot, S. Hallet, R. Chaussod, J. C. Germon, G. Soulas, and G. Catroux. 2001. DNA extraction from soils: old bias for new

METHOD FOR EXTRACTING DNA FROM CLAY SOIL

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37. 38.

39.

40.

6021

microbial diversity analysis methods. Appl. Environ. Microbiol. 67:2354– 2359. McNamara, N. P., H. I. J. Black, N. A. Beresford, and N. R. Parekh. 2003. Effects of acute gamma irradiation on chemical, physical and biological properties of soils. Appl. Soil Ecol. 24:117–132. Picard, C., C. Ponsonnet, E. Paget, X. Nesme, and P. Simonet. 1992. Detection and enumeration of bacteria in soil by direct DNA extraction and polymerase chain reaction. Appl. Environ. Microbiol. 58:2717–2722. Pietramellara, G., J. Ascher, M. T. Ceccherini, P. Nannipieri, and D. Wenderoth. 2007. Adsorption of pure and dirty bacterial DNA on clay minerals and their transformation frequency. Biol. Fertil. Soils 43:731–739. Ririe, K. M., R. P. Rasmussen, and C. T. Wittwer. 1997. Product differentiation by analysis of DNA melting curves during the polymerase chain reaction. Anal. Biochem. 245:154–160. Rochette, E. A., and W. C. Koskinen. 1996. Supercritical carbon dioxide for determining atrazine sorption by field-moist soils. Soil Sci. Soc. Am. J. 60:453–460. Romanovskaya, V. A., P. V. Rokitko, Y. R. Malashenko, T. P. Krishtab, and N. A. Chernaya. 1999. Sensitivity of soil bacteria isolated from the alienated zone around Chernobyl nuclear power plant to various stress factors. Microbiology 68:465–469. Rozen, S., and H. J. Skaletzky. 2000. Primer3 on the WWW for general users and for biologist programmers, p. 365–386. In S. Krawetz and S. Misener (ed.), Bioinformatics methods and protocols: methods in molecular biology. Humana Press, Totowa, NJ. Schneegurt, M. A., S. Y. Dore, and C. F. Kulpa. 2003. Direct extraction of DNA from soils for studies in microbial ecology. Curr. Issues Mol. Biol. 5:1–8. Schulthes, C. P., and D. K. Dey. 1996. Estimation of Langmuir constants using linear and nonlinear least squares regression analysis. Soil Sci. Soc. Am. J. 60:433–442. Smalla, K., N. Cresswell, L. C. Mendoca-Hagler, A. Wolters, and J. D. van Elsas. 1993. Rapid DNA extraction protocol from soil for polymerase chain reaction-mediated amplification. J. Appl. Bacteriol. 74:78–85. Stroetmann, I., P. Kaempfer, and W. Dott. 1994. Untersuchungen zur Effizienz von Sterilisationsverfahren an unterschiedlichen Boeden. Zentralbl. Hyg. Umweltmed. 195:111–120. Tebbe, C. C., and W. Vahjen. 1993. Interference of humic acids and DNA extracted directly from soil in detection and transformation of recombinant DNA from bacteria and a yeast. Appl. Environ. Microbiol. 59:2657–2665. Torsvik, V., J. Goksoyr, and F. L. Daae. 1990. High diversity in DNA of soil bacteria. Appl. Environ. Microbiol. 56:782–787. Tsai, Y. L., J. Y. Le, and B. M. Olson. 2003. Magnetic bead hybridization to detect enterotoxigenic Escherichia coli strains associated with cattle in environmental water samples. Can. J. Microbiol. 49:391–398. Vettori, C., L. Calamai, M. Yoder, G. Stotzky, and E. Gallori. 1999. Adsorption and binding of AmpliTaq DNA polymerase on the clay minerals, montmorillonite and kaolinite. Soil Biol. Biochem. 31:587–593. Wilson, I. G. 1997. Inhibition and facilitation of nucleic acid amplification. Appl. Environ. Microbiol. 63:3741–3751.