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Applied and Environmental Microbiology, January 2009, p. 252-256, Vol. 75, No. 1
0099-2240/09/$08.00+0     doi:10.1128/AEM.01630-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Selective Extraction of Bacterial DNA from the Surfaces of Macroalgae

Catherine Burke, Staffan Kjelleberg,^* and Torsten Thomas

School of Biotechnology and Biomolecular Sciences and Centre for Marine Bio-Innovation, The University of New South Wales, Sydney 2052, Australia

Received 15 July 2008/ Accepted 27 October 2008

Top ABSTRACT INTRODUCTION REFERENCES

 
A novel method has been developed for the selective extraction of DNA from surface-associated bacterial communities from the two model marine benthic algae Ulva australis and Delisea pulchra. The extracted DNA had no detectable contamination with host DNA, was recovered in high yield and quality, and was representative of the bacterial community on the algal surfaces. The DNA is suitable for a variety of subsequent applications, including the construction of large-insert clone libraries and metagenomic sequencing.

Top ABSTRACT INTRODUCTION REFERENCES

 
Marine living surfaces are often inhabited by diverse and complex communities of bacteria (2, 13, 14, 17, 18, 35, 36) and hence harbor biodiversity, ecological processes, and biotechnological potential distinct from those of surrounding planktonic communities (7, 19-21, 24, 39, 41). Bacterial epiphytic communities on algae have been studied in terms of nutrient acquisition (5, 10), carbon exchange (4), resistance to biofouling (6, 37), the production of bioactive compounds (1, 14), and, perhaps most notably, the lack of the host's normal morphological development in the absence of a bacterial community (23, 25, 30, 32).

Most studies of marine epibiotic microbial communities have been carried out using cultured isolates; however, it is commonly accepted that the majority of bacteria which exist in the environment are not readily cultured. As such, comprehensive studies of bacterial communities are increasingly based on the analysis of total environmental DNA, which is required in high yield and on molecular quality, and should ideally be free of eukaryotic or host DNA. For some environments, such as the planktonic water phase, the separation of bacterial cells from eukaryotic cells prior to the extraction of DNA is relatively straightforward (i.e., serial filtration) (33, 40). However, many bacterial communities, and in particular the ones associated with surfaces, exist in biofilms, which prevent the straightforward removal of bacterial cells from eukaryotic cells or host surfaces.

Currently available DNA extraction techniques typically result in the coextraction of the DNA of the host (e.g., alga) along with that of the associated bacterial community. This can lead to substantial problems in the subsequent processing of the sample, as illustrated by frequent contamination of 16S rRNA gene clone libraries with mitochondrial and chloroplast sequences derived from the host (21, 38). Coextraction of the host and community DNA is also prohibitive for metagenomic applications where total DNA is cloned and subsequently sequenced or screened. Here, large proportions of eukaryotic host DNA in combination with the typically larger genome size of the host would result in metagenomic libraries largely biased toward the host rather than the desired bacterial surface community sequences.

In this study, we have developed an efficient method for the extraction of DNA from the epiphytic bacterial community of the green alga Ulva australis and, further, applied the method to the red alga Delisea pulchra. We demonstrate representative and selective extraction from the bacterial community with a high-quality DNA yield, which is suitable for subsequent applications.

Several plants of Ulva australis (approximately 30 g total) were collected from three different rock pools (replicates), and Delisea pulchra (approximately 20 g total) was sampled offshore at a depth of 10 m from Bare Island (Sydney, NSW, Australia) during October 2006 and February 2007. Plants were transferred into sterile plastic bags containing seawater from the same sampling location and immediately transported on ice to the laboratory. Plants were washed three times in sterile seawater to remove loosely associated bacterial cells. Holdfasts were removed, U. australis was cut into sections of approximately 2 cm², and D. pulchra was cut into sections of approximately 5-cm length. Ten grams wet weight of each sample of U. australis was removed and freeze-dried, and the remaining material was subjected to DNA extraction as described below.

Twenty grams of U. australis was placed into 100 ml of calcium- and magnesium-free artificial seawater (CMFSW) containing 0.45 M NaCl, 10 mM KCl, 7 mM Na₂SO₄, and 0.5 mM NaHCO₃ and supplemented with 10 mM EDTA and 1 ml filter-sterilized rapid multienzyme cleaner (3M, Sydney, NSW, Australia). Samples were incubated for 2 hours at room temperature and 80 rpm and then vortexed for 2 minutes. Plant material was removed and the remaining liquid centrifuged at 300 x g for 15 min to remove any remaining algal material. The supernatant was transferred to new tubes, and DNA was extracted by adding an equal volume of phenol, chloroform, and isoamyl alcohol (25:24:1 ratio, respectively) (Fluka, Seelze, Germany) to each sample. Tubes were mixed by inversion and centrifuged at 10,000 x g for 10 min, and the aqueous phase was removed to new tubes. DNA was precipitated with 0.3 M sodium acetate (pH 5.2) and 3 volumes of ethanol at –20°C overnight and then centrifuged at 20,000 x g and 4°C for 30 min. Pelleted DNA was washed once with 70% ethanol, air dried, and resuspended in a total of 8 ml sterile deionized water. The DNA was again precipitated, samples were centrifuged as described above, and DNA pellets were resuspended in a total of 1.4 ml deionized water. DNA was treated with RNase A (0.2 mg/ml) at 4°C overnight.

D. pulchra was subjected to the same procedure as described above; however, only 10 g of material was processed per 50 ml of solution, and after the 2-hour incubation period, the liquid was filtered through first an 11-µm filter and then a 3-µm filter to remove diatoms and very small plant fragments. DNA was extracted from this filtered solution as described above. For comparison, DNA was also extracted from 200 mg of each freeze-dried U. australis algal sample by using the FastDNA Spin kit for soil (Q-Biogene, Carlsbad, CA) according to the manufacturer's instructions and as previously described (21). This method involves bead beating of freeze-dried samples to break open the cells prior to DNA extraction and will be referred to as the bead-beating method.

Twenty random segments of U. australis (10 from before and 10 from after enzyme treatment) were stained with 5 µM SYTO9 nucleic acid stain (Invitrogen, Carlsberg, CA) and examined under a fluorescence confocal microscope (Leica, Wetzlar, Germany) to assess cell removal from the algal surface. The bacterial community of D. pulchra could not be examined with a fluorescent stain, due to a high level of background fluorescence. U. australis exhibits a morphologically diverse bacterial surface community consisting of multilayer microcolonies of rods and coccoid bacterial cells in addition to long filamentous chains (Fig. 1a to d). Initial trials to remove this biofilm community by various enzymatic treatments and physical methods such as sonication were not successful. However, the incubation of the algal samples with a buffer combining CMFSW, 10 mM EDTA, and rapid multienzyme cleaner showed reproducible and almost complete removal of the surface community (Fig. 1). Very few bacterial cells remained, and importantly, the U. australis and D. pulchra tissues were intact without any visible lesions as assessed by light microscopy (data not shown). The concentration of the rapid multienzyme cleaner was important, as lower concentrations often did not completely remove the cells and biofilms, while higher concentrations resulted in damage to the algal surface. The addition of EDTA was also critical, as the DNA was of a lesser quality (i.e., lower molecular weight) and lower yield when extracted in the absence of EDTA.

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FIG. 1. SYTO9 staining of the surface community of U. australis from four random sections before treatment (a to d) and after treatment (e to h) with CMFSW, EDTA, and rapid multienzyme cleaner. Cells are stained in green, and all pictures were taken at a magnification of x640.

This DNA extraction yielded an average of 2 and 0.8 µg DNA per gram of U. australis and D. pulchra wet materials, respectively. The yield for D. pulchra being lower than that for U. australis is consistent with the report of a relatively low bacterial biomass on the surface of the red alga (26). Pulsed-field gel electrophoresis analysis showed high-quality DNA fragments in the range of 5 to >80 kb, the majority being between 10 and 50 kb for U. australis and 5 and 30 kb for D. pulchra (data not shown). For comparison, DNA extracted from U. australis by the bead-beating method (see above) yielded approximately 2 µg DNA per g wet weight and appeared sheared on a standard electrophoresis gel, with the largest fragments at around 5 kb (data not shown). A clear advantage of the new protocol is the size distribution of DNA fragments obtained, making the DNA amenable for cloning into a variety of vectors, including the popular cosmid and fosmid vectors which are routinely used in environmental metagenomic studies (8, 11, 15, 29, 31). In fact, the DNA obtained from U. australis in this study has been successfully used to generate several fosmid libraries of between 20,000 and 50,000 clones (C. Burke, T. Thomas, S. Kjelleberg et al., unpublished data).

Replicate DNA samples from both U. australis and D. pulchra were analyzed for 16S rRNA and 18S rRNA genes by PCR to estimate the proportion of bacterial versus eukaryotic DNA in the samples. Standard PCRs were carried out with primers 27F and 1492R (16) for the 16S rRNA gene and primers EK1F (5'-CTGGTTGATCCTGCCAG-3') (22) and 18Sr-b (5'-GATCCTTCYGCAGGTTCACCTA-3'), modified on the basis of a study by Medlin et al. (28), for the 18S rRNA gene on serial dilutions of DNA (500 pg to 0.05 pg per reaction). For all samples of U. australis extracted with the enzyme-based procedure, there appears to be a weak band for the 18S rRNA gene at 500 pg but none in further dilutions of the DNA sample (Fig. 2a; data from only one replicate shown), while the 16S rRNA gene PCR yielded products at DNA concentrations as low as 0.5 pg (i.e., 3 orders of magnitude lower). For DNA extracted with the bead-beating method, 18S rRNA gene products were obtained at concentrations as low as 0.5 pg (weak band) and 16S rRNA gene products at 0.5 pg (Fig. 2b, data from same replicate shown). For the D. pulchra DNA extracted using the enzyme treatment, strong 16S rRNA gene bands were obtained at a concentration of 5 pg, while no 18S rRNA gene band was detected (Fig. 2c).

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FIG. 2. 16S (lanes 2 to 6) and 18S (lanes 7 to 11) rRNA gene PCR products for one replicate (2006) with the enzyme-based DNA extraction from U. australis (a), the bead-beating extraction from the same sample (b), and the enzyme-based method for D. pulchra (c). Lanes 1 and 12 contain 200 ng EcoRI plus HindIII marker; lanes 2 and 7 are 16S and 18S rRNA gene internal controls (i.e., DNA was spiked with 100 ng Pseudoalteromonas tunicata [marine epiphytic bacterium] or Saccharomyces cerevisiae genomic DNA, respectively). Lanes 3 and 8 are the products starting from 500 pg template DNA; lanes 4 and 9, from 50 pg; lanes 5 and 10, from 5 pg; and lanes 6 and 11, from 0.5 pg.

To determine the source of 18S rRNA gene DNA from the bead-beating method, an 18S rRNA gene clone library was created from the PCR products obtained from two replicates in the 500-pg template DNA PCR. Clone libraries were created using the TA cloning kit for sequencing (Invitrogen), as per the manufacturer's instructions, and screened by colony PCR. Sequencing and BLASTn analysis of 25 clones from the two libraries revealed that the 18S rRNA gene sequences aligned closely (85% to 99% identity over 1,000 to 1,700 bp) to small eukaryotes, including those of the ciliate Ephelota sp. (13 clones), the ribbon worm Oerstedia dorsalis (two clones), the copepods Bradya sp. (three clones) and Ecbathyrion prolixicauda (one clone), the yellow-green alga Tribonema marinum (one clone), the diatom Asterionellopsis glacialis (one clone), and the fungus Tremiscus helvelloides (three clones).

An additional 18S rRNA gene PCR was carried out on U. australis DNA samples by using alga-specific primers AB1F (5'-GGAGGATTAGGGTCCGATTCC-3') and TW4R (5'-CTTCCGTCAATTCCTTTAAG-3') (12). A band was obtained from 500 and 50 pg of bead-beating-extracted DNA, while no bands could be visualized for the DNA extracted with the new enzyme-based method. Sequencing of the PCR product obtained from the bead-beating DNA extraction method and BLASTn analysis revealed that the sequence had its highest identity with Ulva rigida (99% identity over 731 bp). It should be noted that the U. australis 18S rRNA gene sequence is not in the NCBI database.

These PCR analyses show that the new method effectively avoids the coextraction of eukaryotic DNA (Fig. 2). The bead-beating method coextracts not only the host DNA but also the DNA of other eukaryotic organisms, such as copepods and ciliates, that are often found in association with algal surfaces (9, 27). In contrast, we were not able to detect DNA from these organisms or the algal host by using the enzyme-based approach. Additionally, sequencing of metagenomic DNA obtained from this study has not revealed any eukaryotic DNA sequences (Burke et al., unpublished).

While our extraction procedure efficiently removes bacterial cells from the surface of U. australis (see above), it is also important to show that it captures a representative DNA sample of this surface community. The bead-beating DNA extraction method has previously been used to generate 16S rRNA gene clone libraries to describe the surface community of U. australis and D. pulchra (21) and, as such, was considered a good standard against which to test our new method. We therefore used 16S rRNA gene-based denaturing gradient gel electrophoresis (DGGE) fingerprinting to compare the profiles of DNA extracted with the new enzyme-based procedure and the bead-beating method, using DNA extracted from seawater (S. Longford et al., unpublished) as an outgroup. Genes were amplified with primers GM5F(GC) and 907RC (34), and products were separated on 6% acrylamide gels with a 35 to 55% denaturing gradient of 7 M urea plus 40% (vol/vol) formamide for 16 h at 60°C and 75 V. The DGGE banding patterns between samples taken from three different rock pools in 2006 and extracted with the two extraction protocols can be seen in Fig. 3a.

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FIG. 3. (a) DGGE gel image comparing 16S rRNA gene profiles from DNA extracted from triplicate samples of U. australis with two different methods, the enzyme-based DNA extraction (lanes B1, B2, and B3) and bead beating (lanes labeled B1BB, B2BB, and B3BB), from 2006 samples. Lanes marked M are rpoB markers from a random bacterial assemblage. Profiles from triplicate seawater samples (outgroup) are marked SW1, SW2, and SW3. The dendrogram generated from this community profile is shown in panel b.

Bands on gels were scored manually into a presence/absence matrix and subjected to multidimensional scaling analysis with Bray-Curtis similarity. These results are presented as a dendrogram (Fig. 3b) and show that the U. australis community profiles cluster closely together and are separate from the seawater outgroup. These observations were also made for the 2007 samples (data not shown). The matrices from the different DNA extraction methods and samples were also subjected to analyses of similarities by using PRIMER v5.2.2 (PRIMER-E, Plymouth, United Kingdom) (3), which gave global R values of –0.074 and 0.037 for the 2006 and 2007 samples, respectively. Together, these results indicate that there is no significant difference (R < 0.25) between the community profiles generated from the two different DNA extraction techniques, giving confidence that the bacterial community is adequately represented in DNA extracted with our new method.

In conclusion, we have developed a simple and efficient method for the selective extraction of representative bacterial community DNA from the surface of the marine green alga U. australis and have also applied the technique to the red alga D. pulchra. The extracted DNA is largely devoid of DNA from the host background and other associated small eukaryotes and is of sufficient quality and quantity for a number of applications (e.g., metagenomic analysis). We believe that the method presented here will have a wider application to other macroalgae and surfaces and may allow for a more thorough and comprehensive analysis of a range of surface-associated bacterial communities, allowing for the detailed study of their composition, function, and ecology.

 
This work was funded by the Australian Research Council, the Hermon Slade Foundation, and the Centre for Marine Bio-Innovation at the University of New South Wales, Australia.

 
* Corresponding author. Mailing address: School of Biotechnology and Biomolecular Sciences and Centre for Marine Bio-Innovation, The University of New South Wales, Sydney 2052, Australia. Phone: (61 2) 93852102. Fax: (61 2) 93851779. E-mail: S.Kjelleberg{at}unsw.edu.au

Published ahead of print on 31 October 2008.

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Applied and Environmental Microbiology, January 2009, p. 252-256, Vol. 75, No. 1
0099-2240/09/$08.00+0     doi:10.1128/AEM.01630-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

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