An improved electroelution method for ... - Wiley Online Library

RESEARCH ARTICLE

An improved electroelution method for separation of DNA from humic substances in marine sediment DNA extracts Jens Kallmeyer1,2 & David C. Smith1 1

¨ Potsdam, Graduate School of Oceanography, University of Rhode Island, Narragansett, RI, USA; and 2Institut fur ¨ Geowissenschaften, Universitat Potsdam, Germany

Correspondence: Jens Kallmeyer, Institut fur ¨ ¨ Potsdam, Geowissenschaften, Universitat Karl-Liebknecht Strasse 24, Haus 27, 14476 Potsdam, Germany. Tel.: 149 331 977 5694; fax: 149 331 977 5700; e-mail: [email protected] Received 26 June 2008; revised 19 March 2009; accepted 20 March 2009. Final version published online 1 May 2009. DOI:10.1111/j.1574-6941.2009.00684.x Editor: Gary King Keywords DNA extraction; electroelution; humic substances; PCR inhibition.

Abstract We present a method for the rapid and simple extraction of DNA from marine sediments using electroelution. It effectively separates DNA from compounds, including humic substances, that interfere with subsequent DNA quantification and amplification. After extraction of the DNA from the sediment into an aqueous solution, the crude sample is encased in 2% agarose gel and exposed to an electrical current, which draws the DNA out of the gel into a centrifugal filter vial. After electroelution, the sample is centrifuged to remove contaminants 100 000 Da. Recovery of DNA using this method is quantitative and does not discriminate on the basis of size, as determined using DNA standards and DNA extracts from environmental samples. Amplification of DNA is considerably improved due to removal of PCR inhibitors. For Archaea, only these purified extracts yielded PCR products. This method allows for the use of relatively large volumes of sediment and is particularly useful for sediments containing low biomass such as deeply buried marine sediments. It works with both organic-rich and -poor sediment, as well as with sediment where calcium carbonate is abundant and sediment where it is limited; consequently, adjustment of protocols is unnecessary for samples with very different organic and mineral contents.

Introduction The application of molecular methods that allow us to explore the diversity of microbial communities in a wide range of environments without reliance on cultivation has revolutionized our view of the microbial world, particularly in the marine environment (e.g. DeLong et al., 1994; Be´ ja´ et al., 2000). While the use of these methods has led to truly revolutionary findings, shortcomings have also been documented (Webster et al., 2003). Biases introduced at the stage of DNA extraction, purification, the choice of primers as well as the PCR amplification conditions can skew our view of the structure of the in situ microbial community. Extracting DNA from deeply buried marine sediments is particularly challenging, given the low biomass residing in a poorly characterized organic matrix and the large surface area of minerals present. Differences in the extraction procedures and analysis techniques have resulted in questions regarding the microbial diversity in deeply buried sediments at the highest taxonomic level. For instance, studies using samples from FEMS Microbiol Ecol 69 (2009) 125–131

the same expedition (Ocean Drilling Program, Leg 201, Site 1230; reviewed in Smith & D’Hondt, 2006) have concluded that the microbial community is dominated either by Bacteria (Mauclaire et al., 2004; Schippers et al., 2005; Schippers & Neretin, 2006) or by Archaea (Biddle et al., 2006; Sørensen & Teske, 2006). The natural variability within the samples may be masked by analytical problems resulting from biases introduced at the DNA extraction and purification stage. The detection of Archaea seems to be particularly problematic when using PCR. In some cases, their presence is indicated by geochemical evidence but not revealed by PCR (e.g. Kormas et al., 2003). This may be explained by primer mismatch (Teske & Sørensen, 2008), while studies by others have used whole-genome amplification (Lipp et al., 2008; Forschner et al., 2009) to reveal archaeal DNA in sediment samples. Several studies have evaluated extraction and purification protocols and generally conclude that the total yield of DNA and the observed diversity heavily depends on the techniques used (Martin-Laurent et al., 2001; Luna et al., 2006; 2009 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

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Lakay et al., 2007). While it has been shown that a higher DNA yield leads to a higher diversity of phylotypes detected in some cases (e.g. Martin-Laurent et al., 2001; Luna et al., 2006), other studies did not find such a relationship (e.g. Gabor et al., 2003). One of the most common problems when extracting DNA from marine sediments is coextraction of humic substances and other compounds that inhibit downstream applications including quantification or PCR (Zipper et al., 2003). Commercially available DNA extraction kits designed for use with soils/sediments have been used increasingly. However, many of these kits require modifications before they can be used for marine samples (Webster et al., 2003; Newberry et al., 2004), or they have significantly lower DNA yields (Martin-Laurent et al., 2001). Other approaches for extraction of DNA from marine sediments are more labor intensive and involve the use of hazardous chemicals including phenol and chloroform (Sørensen et al., 2006), or the uses of various resins to bind humic substances (Juniper et al., 2001). Chandler et al. (1997) introduced electroelution as a technique for purification of DNA from soil samples in a procedure that requires several cleaning steps, including a phenol/chloroform extraction, before electroelution. Regardless of the analyses one plans to conduct, starting with high-quality environmental DNA as a template is paramount. In order to extract DNA from sediments, the cells have to be lysed to release the DNA. There are different techniques to lyse cells (chemical, enzymatic, and mechanical), all of which lead to a suspension that contains cell remains, DNA, organic matter, and sediment particles. After centrifugation or filtration to remove particulate matter, the sample still contains not only the DNA but also a variety of other substances, including humic substances. The method presented here offers a simple solution by separating the DNA from unwanted compounds using electroelution, without labor-intensive steps or the use of hazardous chemicals. In order to evaluate the suitability of electroelution for separation of clean DNA from marine sediments, we quantified the following parameters: (1) recovery of DNA from environmental samples; (2) recovery of known amounts of dsDNA; (3) recovery efficiency as a function of size; and (4) PCR amplification for Bacteria and Archaea after electroelution.

Materials and methods Sediments Marine sediments were collected from the following locations: (1) Salt Pond Marina (SPM): The sediment was collected by hand at a water depth of c. 0.2 m in SPM in Wakefield, RI (41123 0 N, 71131 0 W). The sediment is dark brown to black, 2009 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

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J. Kallmeyer & D.C. Smith

clay-rich sand with a high organic content. Salinity in the marina is c. 29 psu. The sediment was stored in a plastic bag in the dark at 4 1C. (2) North East Atlantic, Station ‘Biotrans’ (BT): The sediment was collected during RV METEOR cruise M 21/6 in 1992 with a multicorer at a 4560 m water depth at 471N, 201W, and consists of beige carbonate ooze. The core was sliced into 1-cm sections, packed in plastic bags, and stored at 20 1C until analysis. (3) Kieler Bight, Friedrichsort (KB): The sediment was collected in September 2008 with a small box corer at 54122 0 N, 10110 0 E at 18 m water depth. It consists of an organic-rich sandy mud with high microbial activity. (4) Laptev Sea (LAP): The sediment was collected during Polarstern cruise ARK-IX/4 in 1993 with a multicorer in 3237 m water depth at 79114 0 N, 122151 0 E. It consists of light gray to greenish carbonate sands with clay. The core was sliced into 1-cm sections, packed in plastic bags, and stored at 20 1C until analysis.

DNA extraction All further sample handling was carried out inside a laminar flow hood (NuAire) and all solutions were autoclaved or 0.1-mm filtered before use. DNA was extracted using a combination of previously published methods (Stahl et al., 1988; Kallmeyer et al., 2008) that first dislodge the cells from the sediment particles before lysis with a combined chemical and mechanical treatment. One gram of carbonate-free sediment (SPM, KB) and 0.5 g of carbonate-rich sediment (BT, LAP) were each placed into sterile 15-mL centrifuge tubes and diluted with 2 mL 2.5% NaCl, 0.5 mL methanol, and 2 mL of an extraction solution containing 100 mM disodium EDTA dihydrate, 100 mM pyrophosphate decahydrate, 1% (v/v) Tween 80, and 300 mM sodium chloride, and 1% (w/v) sodium dodecyl sulfate. After 1 h of vortexing, the samples were placed in an icewater bath and treated with an ultrasonic probe (Fisher Scientific, Sonic Dismembrator 60) for 10 5 s with 10-s intermissions between treatments. The samples were then centrifuged for 20 min at 2000 g, followed by careful decantation of the supernatant into a sterile 50-mL centrifuge tube. The supernatants were brownish to almost black (SPM, KB) and colorless (BT, LAP), respectively, and were not turbid. DNA was precipitated by mixing the supernatants (c. 7 mL) with 18 mL of ice-cold 96% ethanol and stored at 20 1C overnight (DeFlaun et al., 1986). After precipitation, the samples were centrifuged (40 min at 2000 g) and the DNA formed a pellet with other substances in the sample (e.g. humics, reagents, calcium carbonate, etc.). The SPM pellet was small (c. 0.5 mL) and dark brown, whereas the BT and LAP pellets were much larger (c. 5 mL), white, and were composed mostly of FEMS Microbiol Ecol 69 (2009) 125–131

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An improved electroelution method for separation of DNA

calcium carbonate. After removal of the supernatant, the pellets formed in the BP and LAP samples were dissolved in 10 mL of a sterile, ice-cold carbonate dissolution mix (0.43 M glacial acetic acid, 0.43 M sodium acetate, and 0.17 M sodium chloride, buffered at pH 4.6). The pellets dissolved almost instantly and the DNA was precipitated again by adding 25 mL of ice-cold 96% ethanol, followed by overnight storage at 20 1C and centrifugation (40 min at 2000 g). The resulting pellets were white and of approximately the same size as the pellets from the carbonate-free sediments (0.5 mL). After removing the supernatant, all pellets were washed once in ice-cold 96% ethanol and air-dried in a laminar flow hood. After drying, the pellets were dissolved in steriledistilled water. These DNA extracts were used for electroelution without any further treatment.

The gel buffer [1 Tris-acetate-EDTA (TAE)] was degassed in a side-arm flask to prevent bubbles from becoming trapped beneath the membranes of the centrifuge filters. Electroelution was carried out at a constant voltage of 200 V for 2 h. The recovery of the DNA from the centrifugal filters was carried out according to the manufacturer’s recommendations. After electroelution, the volumes of the retentates varied, even between replicates of the same sample; therefore, they were brought up to a known volume (usually 50–500 mL) with sterile-distilled water.

DNA quantification DNA was quantified fluorometrically (Quant-IT HS dsDNA Assay Kit, Molecular Probes–Invitrogen) on a Wallac Victor 1420 Microplate Fluorophotometer, 485 and 535 nm, excitation and emission, respectively. All analyses were performed in triplicate.

Electroelution procedure Eighty microliters of agarose (2%, Promega) was cast into the electroelution vial (Millipore Centrilutor) to prevent the sample from leaking through the porous bottom, but to allow penetration of DNA. Between 50 and 200 mL of sample was placed in the plugged Centrilutor vial, and then 100 mL of agarose gel was added. After this second layer solidified, another 200 mL agarose gel plug was cast on top (Fig. 1). The electroelution vial was then inserted into a filter unit (Millipore Centricon); for separation of genomic DNA we used a filter with a molecular weight cut-off (MWCO) of 100 kDa. The electroelution apparatus (Millipore Centrilutor) consists of two acrylic chambers, filled with buffer solution. The two chambers are connected through the filter units, which have a tight fit at the bottom of the upper chamber. A schematic drawing and a detailed description is provided in Fig. 1. Further sample handling was performed according to the manufacturer’s recommendations.

Recovery of DNA from sediment extracts Three different volumes (50, 100, and 150 mL) of the DNA extracts of BT and SPM sediment were electroeluted and reconstituted to their original volume. The DNA content was quantified in three volumes (5, 10, and 20 mL) of the crude (noneluted) DNA extract. In order to check whether sample size has any influence on recovery, the results were normalized for the different volumes used.

Quantitative recovery of dsDNA standard One microgram of l-dsDNA standard (Quant-IT BR dsDNA Assay Kit, Molecular Probes–Invitrogen) was diluted in 300 mL of sterile-distilled water, and divided into six 50-mL aliquots. Three aliquots were electroeluted and the retentate was reconstituted to 50 mL; the other three were quantified without any further treatment. For each of the six aliquots, the DNA content was measured fluorometrically.

Size discrimination assay

Fig. 1. Schematic of the loaded electroelution apparatus. The sample is solidified and encased between two layers of clean agarose gel and sits inside the Centrilutor vial, which has holes in the top and bottom to transmit the current. The upper and lower chambers are filled with buffer and the current runs from the upper chamber through the samples into the lower chamber. DNA is pulled out of the sample into the Centricons vial, where it is trapped on the filter membrane, while co-eluted humic substances pass through the filter due to their smaller size.

FEMS Microbiol Ecol 69 (2009) 125–131

To test whether the electroelution discriminates on the basis of DNA size, 10 mL of a quantitative DNA ladder (exACTGene 50-bp Mini DNA Ladder, 25–650 bp, Fisher Scientific) was electroeluted. The retentate of the electroeluted DNA ladder was brought up to 15 mL with sterile 2.5% NaCl solution. Then, two 3-mL aliquots of the retentate and two 2-mL samples of the untreated DNA ladder were separated into their different-size fractions on a 5% acrylamide gel in 0.5 TAE. Electrophoresis was carried out in a refrigerator at 4 1C in a Bio-Rad vertical gel box with 1 TAE buffer at a constant voltage of 120 V, for 35 min. The gel was stained for 15 min with 5 mL of SYBR Green 1 (Molecular Probes–Invitrogen) 2009 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

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at a 1 : 10 000 dilution. Band intensity was quantified using GENESNAP software.

PCR amplification Aliquots of the crude extracts of BT and SPM sediment were carried out using the electroelution procedure. PCR reactions using 16S rRNA gene primers for Bacteria and Archaea were performed on the electroeluted and nonelectroeluted extracts from both sediments. Using bacterial (B27F, 5 0 AGA GTT TGA TCC TGG CTC AG-3 0 ; B1392R, 5 0 -ACG GGC GGT GTG TRC-3 0 ) and archaeal (8F, 5 0 -TCC GGT TGA TCC TGC C-3 0 ; 1492R, 5 0 -GGC TAC CTT GTT ACG ACT T-3 0 ) primers (Integrated DNA Technologies, Coralville, IA), samples were amplified undiluted and with 10-, 100-, and 1000-fold dilutions. The total PCR reaction volume was 25 mL, made up from 1 Taq PCR Mastermix (Qiagen), 0.1 mM F primer and 0.1 mM R primer, 5 mL template, and molecular biology grade H2O. The PCR protocols were as follows: for Bacteria 16S rRNA gene primers – 94 1C for 3 min, 30 cycles of 94 1C for 45 s, 60 1C for 1 min, 72 1C for 1 min 45 s, and 72 1C for 10 min; and for Archaea 16S rRNA gene primers – 94 1C for 5 min, 30 cycles of 94 1C for 30 s, 52 1C for 1 min, 72 1C for 1 min 30 s, and 72 1C for 10 min. Amplifications were performed using an Eppendorf Mastercycler ep. PCR products (including a positive and negative control and a 1-kb DNA ladder) were visualized on a 1% agarose gel stained with ethidium bromide.

Results For both SPM and BT sediment extracts, the recovery of DNA standards after electroelution is independent of the sample volume (Fig. 2, open symbols). The SD averaged over the three sample volumes is 9.5% and 12.5% for BT and SPM, respectively. It was not possible to determine the DNA content of the crude (noneluted) DNA extract (Fig. 2, closed symbols). For both samples, the signal from the crude extract decreases with increasing volume of sample and is generally much weaker than the signal from the eluted sample. For the dark brown SPM pellet, this result was not surprising. The crude BT extract, however, showed only a faint white coloration, but apparently still contained compounds that interfere with the quantification. The recovery of electroeluted l-dsDNA standard is 101 7.5% as compared with the untreated standard, independent of the sample volume used for electroelution (data not shown). Experiments with the quantitative DNA ladder showed that there is no discrimination against specific size fractions 4 100 bp. The overall recovery of the DNA fragments after electroelution is 95.3 5.35%, with no visible trend between larger and smaller fragments (Table 1). Run times of up to 4 h did not result in higher recoveries. The recovery of both the l-dsDNA standard and the 2009 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

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Fig. 2. DNA concentration of the BT and SPM sediment extracts. The DNA concentrations are normalized to the weight of the sediment and the volume of extract used. Open symbols: the DNA extracts were exposed to electroelution and brought back to their original volume after the treatment. Solid symbols: crude (noneluted) DNA extracts were measured directly. For the untreated samples, the calculated concentration of DNA is much lower, indicating that the contaminants are interfering with the detection of the fluorescent signal.

Table 1. Recovery of a quantitative DNA ladder Size (bp)

DNA ladder added (ng)

DNA recovered (ng)

Yield (%)

650 600 550 500 450 400 350 300 250 200 150 100 Average

12.4 0.8 8.8 0.5 5.8 0.4 5.8 0.6 7.2 1.1 17.2 2.9 5.6 0.7 5.2 0.2 3 0.4 11 1.4 6.4 0.4 6.4 0.8

12.9 0.5 8.8 0.2 6.0 0.1 5.7 0.2 6.9 0.5 16.1 0.4 5.0 0.2 4.6 0.2 2.7 0.1 9.8 0.0 6.2 0.2 5.9 0.2

103.7 3.7 99.6 2.7 103.7 0.8 98.4 4 95.8 6.2 93.8 2.3 88.8 3 89.2 4 90.6 3 89.3 0.1 97.1 3.6 93 3.4 95.3 5.35

Overall recovery is 95 5.8% with no apparent trend with fragment size.

quantitative DNA ladder is very close to the 98% recovery for radiolabeled DNA from agarose gels given by the manufacturer of the electroelution apparatus. FEMS Microbiol Ecol 69 (2009) 125–131

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Fig. 3. (a) Negative image of ethidium bromide-stained gel with the PCR products with 16S bacterial primers. (b) Negative image of ethidium bromide-stained gel with the PCR products with 16S archaeal primers.

The PCR results clearly demonstrate the improvement as a result of cleaning the DNA extracts with electroelution. For Bacteria, there were faint bands in the crude BT extract in the 1 : 100 and the 1 : 1000 dilutions; no bands were visible in the crude extracts of the other sediments (SPM, KB, and LAP; Fig. 3a). No amplification of archaeal DNA from the crude extracts of any sediment at any dilution was evident (Fig. 3b). The electroeluted extracts, however, exhibited bands at various dilutions and at different intensities. BT shows bands at 1 : 10 and 1 : 100 dilutions; there was only one band for the SPM sediment in the undiluted sample. Bands were present in the LAP 1 : 10 to 1 : 1000 dilutions with increasing intensity with higher dilutions; KB exhibited the opposite pattern, but with bands in all dilutions including the undiluted sample. Separation of DNA from a sample using centrifuge filters without electroelution did not yield clean extracts. Some of the organic matter adhered onto the filters and coeluted with the DNA. Because the retentate of the SPM sediment was brownish and appeared almost identical to the dissolved pellet, we did not pursue this approach any further. In those samples that contained visible amounts of contaminants, it could be observed that during electroelution some of the dark material moved toward the cathode along with the DNA, while another fraction moved toward the anode. After electroelution, some faint coloration still remained in the agarose gel FEMS Microbiol Ecol 69 (2009) 125–131

plug. This coloration could not be removed by longer run times of the electroelution, indicating that these compounds either have no net charge or moved too slowly to be observed due to their large size. The filter membranes did not show any coloration after the eluted sample had passed through, suggesting that humic substances and other organic matter that coeluted with the DNA passed the filter membrane.

Discussion Our results show that electroelution separates DNA from interfering compounds even in very organic-rich samples that are otherwise not suitable for PCR amplification. Although, there is a technique to improve the performance of SYBR Green assays for quantifying DNA in the presence of humic substances (Zipper et al., 2003), it does not improve DNA amplification in these difficult samples. The ratio of Bacteria and Archaea in marine sediments is still under debate and the results may be biased by methodological limitations. Different techniques have yielded significantly different results so far, even on the very same sediments (Mauclaire et al., 2004; Schippers et al., 2005; Biddle et al., 2006); currently, no definitive answer can be provided (Jørgensen & Boetius, 2007). The results of our study show that amplification of DNA is considerably improved by purification through electroelution. 2009 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

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For the sediments used in this study, the improvement in the PCR results due to electroelution appeared to be much greater for Archaea than for Bacteria as only the electroeluted samples allowed amplification of archaeal DNA. This may, however, also be caused by a generally lower abundance of archaeal DNA in the samples. The 100-kDa MWCO filters should not retain any dsDNA c. o 150 bp (assuming 650 Da bp–1). As there was a clear band for the 100-bp fragment and still a faint but visible band for the 50-bp fragment (data not shown), the precision of the MWCO of the filters is apparently not very sharp and should be used with some caution. It is not surprising that the 25-bp fragments from the ladder could not be detected in the gel as it should have passed through the filter. Electroelution of the quantitative DNA ladder shows that there is apparently no discrimination as a function of size above the MWCO of the filter. As with DNA, humic substances carry a negative charge (Elfarissi & Pefferkorn, 2000). Humic substances will move through the gel faster than genomic DNA because of their much lower molecular weight. Chandler et al. (1997) used relatively short run times of up to 20 min at 100 V to remove humic compounds from the gel while retaining most of the DNA. The end of the treatment was determined visually, when brown components no longer eluted. Still, depending on the run time, between 20% and 65% of the DNA was lost. For applications such as PCR, this may not be a problem, as the increase in sensitivity is of several orders of magnitude. The loss of material may, however, prevent the exact quantification of DNA. Our approach differs from that of Chandler et al. (1997) as we use longer run times (2 h) and a higher voltage (200 V) to remove both DNA and negatively charged humics from the gel. While the DNA becomes trapped on the filter, humic substances will pass through because of their smaller size. Other contaminating compounds with a positive charge will move in the opposite direction; those that are either too large to move through the gel during the run time (e.g. small organic particles) or have no net charge will remain in the agarose plug. Apparently, this separation of the contaminants according to their charge and size is a key factor for obtaining clean DNA, as separation of DNA by just filtering the crude extract was not successful.

Conclusions Electroelution is a simple and rapid technique for retrieving clean DNA from marine sediment samples with very different organic and mineral contents (e.g. high vs. low calcium carbonate content). It eliminates the need for labor-intensive steps and the use of hazardous chemicals or expensive kits. By selectively pulling out the DNA with an electrical current, it is possible to obtain very clean DNA extracts, suitable for a wide 2009 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved

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variety of downstream applications. The suitability of the samples for PCR is considerably improved, in some cases enabling the use of PCR of otherwise unusable samples. Using different buffers and filters with different MWCOs, this technique offers a wide range of possible applications.

Acknowledgements The authors thank Steve D’Hondt for fruitful discussions and a review of the unpublished manuscript. Antje Boetius and Tina Treude provided the samples, and Tatiana Rynearson provided help and equipment for the DNA quantification. Roberta Sheffer helped with the PCR. This work was funded by the NASA Astrobiology Institute and NOAA Oceans and Human Health Initiative.

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