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Marine Geology 174 (2001) 241±247 www.elsevier.nl/locate/margeo Extraction and puri®cation of DNA from organic rich subsurface sediments (ODP Leg 169S) S.K. Juniper*, M.-A. Cambon, F. Lesongeur, G. Barbier Laboratoire de caracteÂrisation de microorganismes marins, IFREMER, Centre de Brest, B.P. 70, 29280 PlouzaneÂ, France Received 10 September 1999; accepted 13 December 1999 Abstract Molecular biology offers many new tools for the characterisation of mixed communities of microorganisms. Approaches that require the extraction and puri®cation of bulk community DNA from sediments and soils must contend with contaminants such as humic acids and heavy metals that can interfere with subsequent genetic analysis. This paper reports on the adaptation of DNA extraction and puri®cation techniques to samples of organic rich sediments collected during the Ocean Drilling Program Leg 169S in Saanich Inlet, British Columbia. In an extraction time series, DNA yield increased up to 48 h (at 378C), after which there were negligible increases in yield and signs of degradation. Resulting extracts, rich in humic substances, blocked the DNA polymerase enzyme even at high dilution. Standard puri®cation procedures (phenol/chloroform extraction followed by silicabased DNA binding or agarose gel electrophoresis) proved ineffective in removing PCR inhibitors. The inhibitory effect was eliminated by cesium chloride density gradient centrifugation with eukaryote DNA added as a carrier, permitting ampli®cation and cloning of SSU (small subunit) rRNA genes. A detailed extraction and puri®cation protocol is presented. These procedures, although time-consuming, may be applicable to other sediment types where microbial DNA is particularly dif®cult to extract or purify. q 2001 Elsevier Science B.V. All rights reserved. Keywords: Microbial; DNA; Subsurface sediments; ODP Leg 169S 1. Introduction Traditional microbiological culture techniques tend to be very selective and miss a huge amount (.99%) of information on prokaryote diversity in natural populations (Amann et al., 1995). For many years this limitation largely con®ned microbial ecologists to the quantitative study of bulk community processes, with little progress being made in understanding what * Corresponding author. Centre de recherche en GeÂochronologie et GeÂochimie Isotopique (GEOTOP) & DeÂpartement des sciences biologiques, Universite du QueÂbec aÁ MontreÂal, C.P. 8888, Succursale Centre-Ville, MontreÂal, QueÂbec, Canada H3C 3P8. Fax: 11-514-9873635. E-mail address: [email protected] (S.K. Juniper). organisms comprised these communities or how phylogenetic (and metabolic) composition and diversity were related to environmental factors. The availability of powerful, molecular techniques is now changing the way we approach both fundamental and applied environmental problems that involve microorganisms (Pace, 1997). Although still primarily used for community descriptions, molecular techniques have the potential to provide a completely new view of the inner workings of the microbial world. Recent successes of molecular techniques in environmental microbiology include the discovery a major new group of Archaea in Paci®c Ocean plankton (Fuhrmamn et al., 1992), novel groups of Eubacteria in Australian soils (Liesack and Stackebradt, 1992), 0025-3227/01/$ - see front matter q 2001 Elsevier Science B.V. All rights reserved. PII: S 0025-322 7(00) 00153-5 242 S.K. Juniper et al. / Marine Geology 174 (2001) 241±247 and a radical revision of our understanding of the phylogenetic diversity of Archaea from studies of hydrothermal pools and springs (Pace, 1997; Nealson, 1997). Information molecules like DNA and RNA can be used to globally characterise entire natural communities of microorganisms, through various forms of `molecular ®ngerprinting'. One may also identify and even quantify particular microbial species within samples by targeting speci®c DNA sequences with probes. The ®rst approach allows evaluation of microbial diversity in relation to environmental factors, and is particularly relevant to studies that extend across environmental gradients such as oxic±anoxic interfaces or, in the present case, sections through subsurface sediments. While the molecular tools for diversity studies are well developed, their application to sediments and soils is problematic, and requires the extraction and puri®cation of nucleic acids, usually DNA, from a complex organo-mineral matrix. DNA extraction from sediment need not be absolutely quantitative since information rather than mass is being analysed. But it must be as representative (i.e. non-selective) as possible of the total microbial community present in the sample, and the extraction process must provide enough DNA for subsequent ampli®cation using the polymerase chain reaction (PCR). In typical Ocean Drilling Program (ODP) legs that sample deep subsurface sediments, sample volume available to all investigators is very limited, so that microbiological studies must be able to optimise DNA extraction from small amounts of core material. Following extraction from sediments, DNA must be puri®ed. The PCR ampli®cation reaction is very sensitive to various substances that co-extract with DNA from sediments and soils, such as heavy metals, pigments and humic compounds (Holben et al., 1988; Rochelle et al., 1992). Several authors have developed DNA extraction and puri®cation for soils and sediments (Holben et al., 1988; Tsai and Olson, 1991; Liesack and Stackebradt, 1992; Rochelle et al. 1992; Erb and Wagner-DoÈbler 1993; Kuske et al., 1998) and comparative evaluations have been performed by Zhou et al. (1996) and more extensively by Miller et al. (1999). However, comparative studies are still few, and careful testing and adaptation remain necessary, given the complexity of sediment types and multiplicity of factors affecting the performance of extraction and puri®cation procedures. After unsuccessfully attempting to apply several standard DNA extraction± puri®cation protocols to sediments sampled during ODP Leg 169S in Saanich Inlet, British Columbia, we undertook to devise a protocol that was adapted to the organic- and clay-rich subsurface sediments found in productive coastal environments such as Saanich Inlet. This paper describes a method for optimal extraction and puri®cation of DNA from these sediment samples in preparation for ampli®cation by PCR and subsequent cloning. 2. Methods and results 2.1. Sample collection and storage All sediments used for this study were obtained during ODP Leg 169S by recovering material in the core catcher of the hydraulic piston corer, at a depth of 45 m below sea¯oor in Hole 1033-B. Material was removed from the core catcher with a sterile spatula, packed into sterile 50 ml centrifuge tubes and immediately frozen by immersion in liquid nitrogen. Samples were stored at 2808C until laboratory experiments. 2.2. DNA extraction Prior to all manipulations, samples were thawed overnight at 48C following which 5 g lots of sediment were placed in 50 ml centrifuge tubes. Our ®rst objective was to optimise the period of extraction, using a standard chemical/enzymatic lysis extraction procedure, modi®ed to include treatments with polyvinylpolypyrrolidone (PVPP) resin to remove humic substances (Holben et al., 1988) and Chelex (Sigma) resin to remove heavy metals. Both humic substances and heavy metals are known to interfere with the PCR ampli®cation of DNA. No physical disruption techniques were included in the extraction protocol, in order to avoid shearing of DNA molecules (Leff et al., 1991), although Miller et al. (1999) have recently demonstrated that DNA shearing can be minimised in carefully controlled bead mill homogenisation. The thawed sediments were resuspended in 6 ml of TE-Na buffer (100 mM Tris with 50 mM EDTA and 100 mM NaCl, pH 8.0) with 1.5 g PVPP prepared by S.K. Juniper et al. / Marine Geology 174 (2001) 241±247 243 Fig. 1. Results of time series tests of extraction yield for microbial DNA in core catcher sediments from Hole 1033-B, core 3. Extracts from each time interval were migrated on agarose gel for visualisation of total yield (band density) and DNA quality (band smearing indicates fragmentation of high molecular weight DNA due to shearing or other factors). Data from UV-spectrophotometric determination of DNA concentration are superimposed on gel image. Error bars represent standard deviation of duplicate measurements. A 500 bp ladder of standard DNA is shown for comparison. The single bands at 24 and 48 h greater exceed the molecular weight of the top (8000 bp) marker band. Composite image from larger, single gel. acid washing as described by Evans et al. (1972) and 250 mg Chelex. This was followed by successive additions of 1.0 ml Sarkosyl (10%), 1.0 ml SDS (10%) and 1.0 ml proteinase K (20 mg/ml) to initiate lysis and extraction of nucleic acids. Samples were then brie¯y vortexed and incubated in duplicate at 378C for 3, 24, 48, 72 and 96 h while slowly mixing on an angled turntable. Following incubation, an equal volume of buffered (pH8.0) PCI (Phenol/Chloroform/Isoamyl Alcohol: 25/24/1) was added to sample tubes and gently mixed to begin separation of DNA from other cell components. Samples were then centrifuged at 6000g for 30 min at 58C, and the upper aqueous phase containing nucleic acids was collected by pipetting. Removal of RNA from this phase was accomplished by addition of 5 mg/ml RNAse and incubating for 60 min at 608C. Samples were then cooled, extracted again with an equal volume of PCI and centrifuged. The aqueous phase was extracted with an equal volume of pure chloroform and centrifuged. The ®nal aqueous phase was removed and gently mixed with two volumes of ice-cold EtOH and stored overnight at 2208C to precipitate DNA. After centrifugation at 12,000g for 30 min the supernatant was gently poured off and the DNA pellet was air dried. The pellets were then redissolved in 0.5 ml sterile demineralised water. DNA in all extracts was quanti®ed using a GeneQuant UV/Visible spectrophotometer, and visualised following electrophoresis on agarose gel. Optimal extraction time under the conditions used was 48 h (Fig. 1). Shorter periods yielded less DNA, while for longer incubations more variable results and smearing in gels (Fig. 1) indicated DNA degradation was beginning to occur. 2.3. DNA puri®cation Even after PVPP washing all extracts were dark tan coloured, likely due to the presence of humic compounds with similar solubility properties to 244 S.K. Juniper et al. / Marine Geology 174 (2001) 241±247 Fig. 2. Results of tests of inhibition of PCR by soluble contaminants in DNA extracts from 48 h extraction core catcher sediments from Hole 1033-B, core 3. Agarose gel image shows PCR product from ampli®cation using universal primers for the bacterial SSU rRNA gene. Left two lanes represent nucleic acid extracts from IFREMER bacterial strain 721 without addition of sediment extract. For extract additions, 0.1, 1.0 and 4.0 ml of extract replaced equal quantities of distilled water in the 50 ml PCR mix. A 500 bp ladder of standard DNA is shown for comparison. Composite image from larger, single gel. DNA. Initial attempts to amplify DNA from these extracts, using universal bacterial and archaeal primers for the small subunit (SSU) rRNA gene, were unsuccessful. Treatment of extracts on mini silica-based DNA binding columns or by agarose gel electrophoresis followed by excision and gel extraction did not relieve the inhibitory effect. To test the inhibitory effect of soluble impurities on the PCR reaction, DNA from a laboratory strain of bacteria (strain 721) was ampli®ed by PCR in the presence of small amounts of sediment extract. Results (Fig. 2) showed that addition of as little as 0.1 ml to the 50 ml PCR mix resulted in noticeable inhibition, with complete inhibition occurring with a 1.0 ml addition of extract, despite the presence of nonspeci®c binding protein (BSA) in the reaction mixture which has been shown to decrease inhibition (Kreader, 1996). A further puri®cation step was performed using cesium chloride (CsCl) density gradient centrifugation. S.K. Juniper et al. / Marine Geology 174 (2001) 241±247 This procedure usually requires a minimum of around 200 mg of DNA in order to visualise the DNA layer in the density gradient. Most of our samples contained a few hundred micrograms of DNA. To assure visibility after centrifugation, all samples were supplemented with 200 mg of eukaryote DNA derived from herring sperm. Prior tests showed that the herring sperm DNA contained no prokaryote contaminants that were ampli®ed by bacterial or archaeal PCR primers. The puri®cation procedure began with the redissolution of the dried DNA pellets in 1.5 ml TE buffer (100 mM Tris with 50 mM EDTA, pH 8.0) containing 1.075 g/ml CsCl and transferred by syringe to bell-top Quick-Seal centrifuge tubes (Beckman). To this were added 200 mg herring sperm DNA solution (10 ml of 20 mg/ml) and 100 ml of 10 mg/ml ethidium bromide (EtBr). Tubes were ®lled to the shoulder and balanced with TE buffer, sealed and centrifuged at 400,000g for 12 h at 158C. The CsCl gradients were then viewed under UV illumination and the rose-coloured DNA layer was removed by piercing the side of the tube with a 2 ml syringe. Samples were placed in 1.5 ml centrifuge tubes and washed and centrifuged seven times until all EtBr colour had disappeared. This was followed by overnight dialysis at 58C against TE buffer, using Cellu SEP H1 membranes with a nominal molecular weight cut-off (MWCO) of 25,000 Da. DNA concentration in extracts was measured with a GeneQuant spectrophotometer and adjusted to 100 ng/ml prior to PCR. 2.4. PCR and cloning The universal bacterial primers used were Sa dir: 3 0 AGA GTT TGA TCA TGG CTC AG 5 0 and S17 rev: 3 0 GTT ACC TTG TTA CGA CTT5 0 . The PCR reaction mix contained 2 ml dTNP's, 5 ml Taq buffer, 0.5 ml Taq polymerase plus primers, made up to 50 ml with sterile distilled water. The PCR parameters were 3 min initial denaturation at 948C followed by 30 cycles of 1 min denaturation at 948C, 2 min annealing at 508C, and 4 min extension at 728C, ®nishing with 6 min at 728C before holding at 68C. After veri®cation on agarose gel, PCR product was puri®ed for cloning using a QIAquick PCR puri®cation kit; eluting DNA from columns with two successive 30 ml additions of EB buffer. 245 The puri®ed SSU rRNA gene sequences were successfully cloned into the E. coli SURE strain using a pGEM-T Easy Vector plasmid kit (Promega Corp.). Obtained clones were then compared by Restriction Fragment Length Polymorphism and distinct clones were identi®ed for further study by molecular sequencing. 3. Discussion Published techniques for the direct extraction and puri®cation of DNA from sediments and soils vary in complexity and time requirements. Zhou et al. (1996) and Miller et al. (1999) have tested and optimised procedures for a variety of sediment and soil types although both studies conclude that no single method will be appropriate to all situations and research goals. For most work involving SSU rDNA, a ®rst goal of the extraction±puri®cation process is to optimise for yield of high molecular weight genomic DNA (HMW gDNA), in order to reduce the potential for formation of chimeric PCR ampli®cation products. This restricts the use of physical disruption of cells during extraction as resulting shear forces can fragment HMW gDNA (Leff et al. 1993, Miller et al. 1999). The 48 h, gentle agitation extraction procedure developed here met this requirement but was time consuming and produced a dark extract that inhibited the PCR reaction. We were somewhat surprised by the increase in DNA yield provided by lengthy extraction periods, although similar results have been obtained in optimisation of DNA extraction from archival museum material (Jackson et al., 1991). With careful testing, our long extraction procedure could possibly be replaced by a rapid bead mill treatment as per Miller et al. (1999), and still avoid shear-fragmentation of high molecular weight DNA. However, bead-mill homogenisation is also known to recover contaminating humic acids (Ogram et al., 1987; Smalla et al., 1993; Leff et al., 1995) so that DNA puri®cation prior to PCR is likely to be unavoidable with sediments such as those sampled in ODP Leg 169S. These sediments were much richer in organic matter and terrigenous clays and humic substances than the oceanic sediments used by Rochelle et al. (1992, 1994) to develop DNA extraction and sample handling procedures for deep subsurface microorganisms. 246 S.K. Juniper et al. / Marine Geology 174 (2001) 241±247 Potential PCR inhibitors are probably numerous and omnipresent in sediments and soils. Most publications do not usually identify inhibitory substances beyond the level of general groups such as colloids, humic substances and heavy metals (Fisk et al., 2000; Holben et al., 1988; Rochelle et al., 1992). More study of the nature or effect of these contaminants is clearly necessary as is the further development of DNA puri®cation procedures for environmental samples. Rochelle et al. (1992) were able eliminate PCR inhibition in some samples by diluting extracted DNA to reduce contaminant concentrations. We diluted some samples by 2±25 £ (data not shown) prior to PCR but were unable to eliminate inhibitory effects. Our experiments with adding sediment extract to positive controls showed that inhibitory effects remained at dilutions of up to 50 £ . Dilution of sample beyond 10:1 is probably not a solution to the contamination problem since more dilute DNA will yield less and less PCR product (Rochelle et al., 1992a,b), to point where subsequent manipulation of sample DNA becomes impracticable. The CsCl density gradient centrifugation used here was effective but rather time consuming, particularly because of the repeated washings required to remove EtBr. Careful attention to optimising DNA extraction yield now allows us to eliminate the addition of eukaryote DNA (herring sperm). The amount of EtBr added to the CsCl preparations can also be adjusted down, on a sample by sample basis, thereby reducing the number of subsequent washing steps. Zhou et al. (1996) and Miller et al. (1999) have demonstrated that high-throughput DNA puri®cation procedures that use minicolumns or gel extraction kits are suf®cient for many soil and sediment types. Although mini-column and gel extraction puri®cation did not eliminate PCR inhibitors from our Leg 169S sediment extracts (data not shown), we still recommend testing of these simpler procedures prior to resorting to CsCl density gradients. Since initial descriptions of intact, active microbial cells in deep subsurface sediments in ODP Leg 112 (Cragg et al., 1990; Parkes et al., 1994), it has become apparent that the in¯uence of microorganisms on the diagenesis and evolution of buried organic matter extends far below the sediment±water interface. Besides an obvious interest for microbiologists, understanding how and what microorganisms modify the sedimentary record will be important to paleooceanographic studies. Molecular tools such as SSU rRNA gene analysis can reveal how the composition of microbial communities responds to changing environmental conditions during burial in marine sediments. Organic matter properties, sediment porosity and temperature undergo fundamental changes down through the sediment column. Are groups of microorganisms simply eliminated or inactivated, as conditions become more severe? Or does the microbial community adapt to exploit new opportunities for growth; such as has been suggested from observations of effects of deep subsurface temperature increases on acetate availability for methanogenic organisms (Wellsbury et al., 1997)? Molecular techniques provide a promising approach to these questions. Further application and methodological adaptation are clearly desirable. Acknowledgements This research was funded by NSERC Canada, through a collaborative Special Projects grant to M. Whiticar and a Research Grant to S.K. Juniper, and by IFREMER, France. Travel support for S.K.J. was provided through the Canadian Association of Universities and Colleges ªGoing Global Ð Science and Technology with European Partnersº program. We thank Yves Prairie for collection of sediment samples aboard the JOIDES Resolution, and the Ocean Drilling Program for technical support during Leg 169S. Reviewer comments and suggestions enabled substantial improvement of this manuscript. References Amann, R.I., Ludwig, W., Scheifer, K.-H., 1995. Phylogentic identi®cation and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59, 143±169. Cragg, B.A., Parkes, R.J., Fry, J.C., Herbert, R.A., Wimpenny, J.W.T., Getliff, J.M., 1990. Bacterial biomass and activity pro®les within deep sediment layers. In: Suess, E., von Huene, R. et al. (Eds.), Proceedings of the Ocean Drilling Program, Scienti®c Results, vol. 112, pp. 607±619. Erb, R.W., Wagner-DoÈbler, I., 1993. Detection of polychlorinated biphenyl genes in polluted sediments by direct DNA extraction and polymerase chain reaction. Appl. Environ. Microbiol. 59, 4065±4073. Evans, H.J., Koch, B., Klucas, R., 1972. Preparation of nitrogenase S.K. Juniper et al. / Marine Geology 174 (2001) 241±247 from nodules and separation into components. Methods Enzymol. 24, 470±476. Fisk, M.R., Thorseth, I.H., Urback, E., Giovannoni, S.J., 2000. Investigation of microorganisms and DNA from thermal waters of Site 1026. Proceedings of Leg 168, Ocean Drilling Program, pp. 167±174. Fuhrmamn, J.A., McCallum, K., Davis, A.A., 1992. Novel major archaebacterial group from marine plankton. Nature 356, 148±149. Holben, W.E., Jansson, J.K., Chelm, B.K., Tiedje, J.M., 1988. DNA probe method for the detection of speci®c microorganisms in the soil bacterial community. Appl. Environ. Microbiol. 54, 703±711. Jackson, D.J., Hayden, J.D., Quirke, P., 1991. Extraction of DNA from archival material. In: McPherson, N.G., Quirke, P., Taylor, G.R. (Eds.), PCR: A Practical Approach. Oxford University Press, New York, pp. 33±40. Kreader, C.A., 1996. Relief of ampli®cation inhibition in PCR with bovine serum albumin or T4 gene 32 protein. Appl. Environ. Microbiol. 62, 1102±1106. Kuske, C.R., Banton, K.L., Adorada, D.L., Stark, P.C., Hill, K.K., Jackson, P.J., 1998. Small-scale DNA sample preparation method for ®eld PCR detection of microbial cells and spores in soil. Appl. Environ. Microbiol. 64, 2463±2472. Leff, L.G., Dana, J.R., McAarthur, J.V., Shimkets, L.J., 1995. Comparison of methods of DNA extraction from stream sediments. Appl. Environ. Microbiol. 61, 1141±1143. Liesack, W., Stackebradt, E., 1992. Occurrence of novel groups of the domain Bacteria as revealed by analysis of genetic material isolated from an Australian terrestrial environment. J. Bacteriol. 174, 5072±5078. Miller, D.N., Bryant, J.E., Madsen, E.L., Ghiorse, W.C., 1999. Evaluation and optimization of DNA extraction and puri®cation procedures for soil and sediment samples. Appl. Environ. Microbiol. 65, 4715±4724. 247 Nealson, K.H., 1997. Sediment bacteria: who's there, what are they doing, and what's new? Annu. Rev. Earth Planet. Sci. 25, 403±434. Ogram, A., Sayler, G.S., Barkay, T., 1987. The extraction and puri®cation of microbial DNA from sediments. J. Microb. Methods 7, 57±66. Pace, N.R., 1997. A molecular view of microbial diversity and the biosphere. Science 276, 734±740. Parkes, R.J., Cragg, B.A., Bale, S.J., Getliff, J.M., Goodman, K., Rochelle, P.A., Fry, J.C., Weightman, A.J., Harvey, S.M., 1994. Deep bacterial biosphere in Paci®c Ocean sediments. Nature 371, 410±413. Rochelle, P.A., Fry, J.C., Parkes, R.J., Weightman, A.J., 1992. DNA extraction for 16S gene analysis to determine genetic diversity in deep sediment communities. FEMS Microbiol. Lett. 100, 59± 66. Rochelle, P.A., Cragg, B.A., Fry, J.C., Parkes, R.J., Weightman, A.J., 1994. Effect of sample handling on estimation of bacterial diversity in marine sediments by 16S rRNA gene sequence analysis. FEMS Microbiol. Ecol. 15, 215±226. Smalla, K., Cresswell, N., Mendonca-Hagler, L.C., Wolters, A., van Elsa, J.D., Rapid, D.N.A., 1993. extraction protocol from soil for polymerase chain reaction-mediated ampli®cation. J. Appl. Bacteriol. 74, 78±85. Tsai, Y., Olson, B.H., 1991. Rapid method for direct extraction of DNA from soil and sediments. Appl. Environ. Microbiol. 57, 1070±1074. Wellsbury, P., Goodman, K., Barth, T., Cragg, B.A., Barnes, S.P., Parkes, R.J., 1997. Deep marine biosphere fuelled by increasing organic matter availability during burial and heating. Nature 388, 573±576. Zhou, J., Bruns, M.A., Tiedje, J.M., 1996. DNA recovery from soils of diverse composition. Appl. Environ. Microbiol. 62, 316±322.