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Extraction of high molecular weight DNA from microbial mats
Benjamin S. Bey, Erin B. Fichot, Gargi Dayama, Alan W. Decho, and R. Sean Norman
Department of Environmental Health Sciences, Arnold School of Public Health, University of South Carolina, Columbia, SC, USA
BioTechniques, Vol. 49, No. 3, September 2010, pp. 631–640
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Due to the presence of inhibitors such as extracellular polymeric substances (EPSs) and salts, most microbial mat studies have relied on harsh methods of direct DNA extraction that result in DNA fragments too small for large-insert vector cloning. High molecular weight (HMW) DNA is crucial in functional metagenomic studies, because large fragments present greater access to genes of interest. Here we report improved methodologies for extracting HMW DNA from EPS-rich hypersaline microbial mats. The protocol uses a combination of microbial cell separation with mechanical and chemical methods for DNA extraction and purification followed by precipitation with polyethylene glycol (PEG). The protocol yields >2 µg HMW DNA (>48 kb) per gram of mat sample, with A260:280 ratios >1.7. In addition, 16S rRNA gene analysis using denaturing gradient gel electrophoresis and pyrosequencing showed that this protocol extracts representative DNA from microbial mat communities and results in higher overall calculated diversity indices compared with three other standard methods of DNA extraction. Our results show the importance of validating the DNA extraction methods used in metagenomic studies to ensure optimal recovery of microbial richness.

Hypersaline microbial mats are layered, organosedimentary structures that provide unique systems to examine community dynamics and biogeochemical cycling (1-7). Given the high level of interactions occurring within microbial mats, they provide ideal model systems for metagenomic studies aimed at understanding complex microbial interactions. However, unlike soil, sediment, or water samples, microbial mats contain high levels of salts and extracellular polymeric substances (EPSs) (3), rendering the isolation of high-quality DNA extremely difficult. Currently, most molecular-based microbial studies use harsh methods of DNA extraction directly from samples, such as microbial mats (8) and soils/sediments (9-13). While these methods are effective, there are two major limitations that preclude a detailed understanding of microbial mat functioning: (i) it is not known how well the final DNA represents the microbial community and (ii) the DNA is sheared into small fragments that are sufficient for microbial diversity studies but too small for large-insert metagenomic studies.

In this study, we developed methods to extract microbial community DNA from hypersaline microbial mat samples and compared it with commonly used DNA extraction methods. Our goals were to extract DNA (i) with comparable efficiency and presumed minimal bias inherent to bead beating methods; (ii) with high molecular weight (HMW) by using gentle approaches that minimally shear DNA; and (iii) with high quality for downstream applications. Results suggest that removing bacterial and archaeal cells from sediment is an essential first step in DNA extraction from hypersaline mats. While this study focused on microbial mats as a model system, our results show the importance of validating DNA extraction methods used in metagenomic studies to ensure optimal microbial richness recovery. Furthermore, since microbial mats represent one of the most troublesome sample types for nucleic acid recovery, our approach can be utilized for other recalcitrant samples. Finally, this study highlights the importance of extensive DNA extraction quality control prior to any metagenomic studies based on clone library construction or pyrosequencing.

Materials and methods

Sample sites and collection

Microbial mats were collected from three hypersaline ponds in the Bahamas: Salt Pond (24°01′N, 74°27′W), French Pond (23°57′N, 74°31′W), and Darby Pond (23°50′N, 76°12′W). Samples were collected to a depth of 6 cm and frozen at −20°C for transport to the laboratory where they were maintained at −80°C until processing.

Microbial cell removal

Microbial cells were extracted from 30 g (wet weight) of homogenized mat samples to separate the cells from the background sediment matrix. First, samples were blended three times in 100 mL 1 M NaCl (BDH Chemicals, Dubai, UAE) using a Waring blender (Waring Laboratory Science, Torrington, CT, USA) at medium speed for 1 min with intermittent cooling for 1 min at −20°C to limit temperature-induced cell lysis (8,9). To enhance cell desorption from sediments, the final blended slurry was adjusted to 250 mL with 1 M NaCl and shaken (150 rpm) at room temperature for 30 min. It should be noted that this salinity is sample-specific and can be adjusted for different samples. The slurry was centrifuged at low speed (500× g for 15 min at 4°C) to separate particulates from microbial cells. The supernatant, containing microbial cells, was transferred into a 250-mL centrifuge bottle, and the sediment from the previous cell extraction was blended again, shaken, and centrifuged four additional times in 250 mL 1 M NaCl, as described earlier. Observation by epifluorescent microscopy of extracted cells and spent-sediment suggested that cyanobacteria were represented in the cell extracts. Supernatants from each of the five blending steps were combined, and microbial cells were pelleted by high-speed centrifugation (25,000× g for 15 min at 4°C). To reduce inhibitors of downstream applications, the cell pellet was resuspended in 250 mL 2% sodium hexametaphosphate (EMD Chemicals, Gibbstown, NJ, USA), a sequestrant/chelator of compounds such as humic acids, free DNA, and antibiotics (10,11). The resuspended cells were shaken at room temperature for 30 min at 150 rpm and centrifuged at 25,000× g for 15 min at 4°C. The cell pellet was washed with 250 mL Tris-EDTA (TE) buffer, pH 8.0, containing 50 mM EDTA, shaken at room temperature for 10 min, and centrifuged at 25,000× g for 15 min at 4°C. The final cell pellet was resuspended in 15 mL TE (10 mM EDTA), and 200-µL aliquots were stored at −80°C for subsequent DNA extractions.

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