Comparison of DNA extraction methods and environmental

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Comparison of DNA extraction methods and environmental Powered By Docstoc

 2   A comparison of methods for total community DNA preservation and extraction

 3                           from various thermal environments



 6                    Kendra R. Mitchell1 and Cristina D. Takacs-Vesbach1


 9             Department of Biology, University of New Mexico, Albuquerque, NM




13             Submitted to Journal of Industrial Microbiology & Biotechnology




17             Running Title: Nucleic acid preservation and extraction methods




21   *Corresponding author:, 505-277-3418, 505-277-3040 (Fax)


23   1. Abstract

24          The widespread use of molecular techniques in studying microbial communities

25   has greatly enhanced our understanding of microbial diversity and function in the natural

26   environment and contributed to an explosion of novel commercially viable enzymes.

27   Exemplified by the discovery of Taq polymerace, one of the most promising

28   environments for detecting novel processes, enzymes, and microbial diversity is hot

29   springs. This is the first study to specifically examine potential biases introduced by

30   DNA preservation and extraction methods in hot spring microbial samples. We compared

31   the quality, quantity, and diversity of environmental DNA samples preserved and

32   extracted by commonly used methods. We included samples from the whole spectrum of

33   environmental conditions that are found in Yellowstone National Park thermal features

34   and our results are applicable to future studies in the park and other extreme

35   environments. We found that samples preserved in a non-toxic sucrose lysis buffer,

36   along with a variation of a standard DNA extraction method using CTAB resulted in

37   higher quality and quantity DNA than the other preservation and extraction methods

38   tested here. Diversity determined using DGGE revealed that there was some variation

39   within replicates of a sample, but no statistical difference between the methods.

40   However, the sucrose lysis buffer preserved samples extracted by the CTAB method

41   were 15-43% more diverse than the other treatments.

42   Keywords: DGGE, DNA extraction, environmental microbiology, thermophile

43   2. Introduction

44          The impact of molecular studies on our knowledge of the extent of microbial

45   diversity can’t be over stated. This explosion of information has significantly changed

46   the whole field, from basic ecological research into the fundamental controls of microbial

47   communities to bioprospecting for novel and potentially commercially relevant enzymes.

48   Even with recent advances in culturing efforts [10, 22], the majority of microbes in the

49   environment still cannot be cultivated in the laboratory [18]. However, inability to

50   maintain an organism in culture is no longer a major impediment to accessing its genetic

51   diversity. Metagenomic studies similar to those that have been useful in exploring the

52   diversity of uncultivated organisms have also be used to mine for enzymatic diversity

53   [12]. The biotechnology applications that are currently targeting microbial metagenomic

54   studies range from the search for new antibiotics to environmentally sound biocatalysts

55   such as amyases [14, 19].

56          Thermal environments have been a particularly rich source of novel organisms [1,

57   7, 9, 11, 16, 26], processes [3], novel enzymes [10, 23], and on-going research in the

58   origin and diversity of microbes.   Fundamental to any of these studies is maximizing the

59   detectable diversity by maximizing the quality and quantity of DNA examined and

60   minimizing the biases of the methods. There are layers of potential bias in molecular

61   studies because of the sequential nature of the process, including those inherent in PCR

62   [27] which have been well studied and will not be addressed in this paper. In the

63   collection of an environmental sample for molecular study, the initial, and least

64   examined, possible source of biased detection of community diversity is the biases that

65   could be introduced by the preservation method. Another possible source of bias is DNA

66   extraction methods and post-extraction purification. The effect of DNA extraction

67   method on detectable diversity has been examined in soil [5, 6], marine sediments [15],

68   and compost [13, 28], but no study has examined the impact that temperature and pH

69   extremes may have on extraction efficiency.

70          We are concerned about the effects of sample preservation and DNA extraction

71   methods on the diversity of the community detected by examining molecular methods.

72   The objective of this study was to find the most effective sample preservation and DNA

73   extraction method that results in high molecular weight DNA, is relatively free from

74   contaminants, and maximizes the amount of diversity that we detect. We compared three

75   preservation methods and six nucleic acid extraction methods. The ideal preservation

76   and extraction method would work well with a variety of samples that included high and

77   low biomass, a wide range of pH (e.g. thermal features range from very acidic, <1, to

78   alkaline), as well as different sample types including biofilms, microbial mats, and

79   sediments. Additionally, this method should be quick to accommodate high throughput

80   of large sample numbers and facilitate sample collection from the remote backcountry

81   where liquid nitrogen, dry ice, and toxic preservatives are not practical. We included

82   samples from the whole spectrum of environmental conditions that are found in YNP

83   thermal features and our results are applicable to future studies in YNP, other extreme

84   environments, and microbial surveys in general.

85   3. Methods and Materials

86   Site description and sampling

87          Samples were collected from thermal features throughout Yellowstone National

88   Park, USA (YNP) during the summer of 2002 as part of our microbial inventory. A

 89   subset of fifteen samples was selected for this study that encompassed the full range of

 90   pH, temperature, and biomass types found in the park (Yellowstone sample n=15, see

 91   Table 1). The pH and temperature of the samples were split into three ranges, low (pH 0

 92   to 4, temperature 40 to 60 degrees Celsius), mid (pH 4.01 to 8, temperature 60.1 to 80),

 93   and high (pH above 8.01, temperature above 80.1). The samples were also categorized by

 94   the type of biomass collected: sediment, microbial mat, or filaments. Additionally, we

 95   collected samples from two neutral thermal springs in the Jemez Mountains, New

 96   Mexico. Approximately 1-2.5 ml of sample was collected at each site with either sterile

 97   forceps or a syringe.

 98   Sample Preservation

 99          Two replicate YNP samples were collected from each sampling site: one was

100   preserved in an equal volume of sucrose lysis buffer (20 mM EDTA, 200 mM NaCl, 0.75

101   M sucrose, 50 mM Tris-HCl, pH 9.0) [8] and the other replicate was preserved in an

102   equal volume of GIT (5 M guanidine isothiocyanate, 50 mM Tris pH 7.4, 25 mM EDTA

103   pH 8, 0.8% 2-mercaptoethanol)[4]. The YNP samples were held at ambient air

104   temperature (10 to 26 ºC) for up to five days before they were stored at –80 degrees

105   Celsius.

106          The two Jemez Springs, New Mexico samples were collected from neutral

107   thermal springs one from the Jemez Springs Bath House (BH) and the other from

108   Giggling Star Resort (GS). Three replicate samples were collected. Two of the replicates

109   were preserved in sucrose lysis buffer (SLB), one of these replicates was frozen

110   immediately in liquid nitrogen and the other was held at 20C for 7 days before being

111   frozen at –80C. The third replicate sample was collected and mixed with molten 2%

112   agarose while in the field for extraction with a high molecular weight agarose noodle

113   method (see below).

114   Extraction Methods

115          We used four extraction methods: lysis by pulse boil, a CTAB extraction, agarose

116   noodle which results in high molecular weight DNA [25], and the Mo Bio Soil DNA

117   Purification kit, which combined with the preservation methods results in six treatments.

118   With the pulse boil method, nucleic acids were extracted from 200 l of the YNP and

119   Jemez samples that were preserved in SLB (referred to here as “Boil”) [20]. Briefly: the

120   samples were boiled at 96C then cooled to 4C three times in a thermocycler, sodium

121   dodecyl sulphate (SDS) (to a final concentration of 2%) and proteinase K (final

122   concentration 250 g/ml) were added and the sample was incubated at 42C for 2.5 hrs,

123   then incubated at 60C for 30 minutes, extracted once with phenol/chloroform then twice

124   with chloroform, finally the DNA was precipitated and washed with ethanol.

125          We extracted nucleic acids from 200 l of the YNP and Jemez SLB preserved

126   samples using a variation of the CTAB [29] method (referred to as CTAB S). Briefly: 2

127   volumes of 1% CTAB buffer (1% CTAB, 0.75 M NaCl, 50 mM Tris pH 8, 10 mM

128   EDTA) and proteinase K (final concentration 100 g/ml) were added to the SLB

129   preserved samples; incubated for one hour at 60C, SDS (final concentration 2%) was

130   added and incubated one hour at 60C, extracted once with phenol/chloroform then twice

131   with chloroform, finally the DNA was precipitated with ethanol. The YNP samples that

132   were preserved in GIT (referred to as CTAB G) were also extracted using the CTAB

133   method with the modification of washing the sample three times with filter sterilized

134   water before the addition of CTAB to remove the GIT from the sample. GIT is a protein

135   denaturant and inhibits proteinase K activity if not removed.

136          SLB (Mo Bio S) and GIT (Mo Bio G) preserved YNP samples were extracted

137   using the Mo Bio Soil DNA purification kit following manufacture’s suggested protocol

138   (Mo Bio Laboratories, Inc., Solana Beach, CA).

139          Nucleic acids were extracted from YNP SLB preserved samples and all Jemez

140   samples using the high molecular weight noodle extraction method (Noodle). Briefly:

141   the sample was mixed with molten 2% agarose and cooled in 1 mL syringes forming

142   agarose noodles, the noodles were incubated for 3 hours at 37C in a lysis buffer (10 mM

143   Tris, 50 mM NaCl, 50 mM EDTA, 0.2% SDS, 1% Sarkosyl, 1mg/ml lysozyme) then

144   incubated at 37C in ESP buffer (1% Sarkosyl, 0.1 M EDTA, 1 mg/ml proteinase K), the

145   ESP buffer was changed once a day for a total of four days, the noodles were then stored

146   at 4C in TE storage buffer (10 mM Tris and 50 mM EDTA). The purified nucleic acids

147   were extracted from the agarose noodles by incubating at 60C to melt the agarose then

148   adding agarase and incubating overnight at 37C. The nucleic acids were purified from

149   the agarose/agarase slurry by extracting once with phenol/chloroform then twice with

150   chloroform. The nucleic acids were precipitated with ethanol. [25]

151   Measuring extraction success

152          Environmental DNAs were electrophoresed on an ethidium bromide stained 1.2%

153   agarose gel. The size and quality (evidence of shearing etc.) of the DNA was evaluated

154   using a size standard DNA ladder (EZ Load 1Kb, BioRad).

155          The purity of the extracted DNA was quantified by calculating the ratio of the

156   absorbance at 260 nm and 280 nm (A260/A280). Nucleic acids extracted from

157   environmental samples are often contaminated with humic organic carbons, metals, and

158   other compounds that makes using the 260 nm absorbance to calculate DNA

159   concentration unreliable. To circumvent this problem, we also quantified the DNA using

160   PicoGreen dsDNA Quantitation Reagent (Molecular Probes, Inc., Eugene, OR, USA).

161   Extracted DNA (5 l) was mixed with a 1:200 solution of PicoGreen in 1X tris-acetate

162   buffer (TAE) and absorbance was read in a fluorometer.

163          The third parameter that was qualified was the ability to amplify 16S rDNA from

164   the environmental genome of each extraction replicate. Dilutions of the genomic DNA

165   were used as template DNA. The reaction included (1X Promega buffer with 1.5 mM

166   MgCl2, bovine serum albumin (0.04 % final), 2.5 U Taq DNA polymerase (Promega

167   U.S.) 2.5% Igepal CA-630 (Sigma-Aldrich), 10 M each dATP, dGTP, dCTP, dTTP

168   (BioLine USA, Inc.), 20 M Bacterial specific primers 338FGC


170   AGCAG) and 519R (ATTACCGCGGCTGCTGG)). The PCR reaction (50 l) was

171   incubated in a thermocycler (ABI GeneAmp 2700) for 5 minutes at 94.0C then for 30

172   cycles of 30 seconds at 94.0C, 30 seconds at 50.0C and 30 seconds at 72.0C. The

173   reaction was incubated at 72.0C for 7 minutes for final extension.

174   Relative species Richness

175          Differences in relative species richness among the preservation and extraction

176   methods were determined using denaturing gradient gel electrophoresis (DGGE). Ten l

177   of PCR product (approximately 500 ng) was run on a 6% (wt/vol) acrylamide gel with

178   1X TAE (40 mM Tris, 20 mM acetate, and 1mM EDTA) with a denaturing gradient of

179   20% (8% (vol/vol) formamide and 8.4% (wt/vol) urea) to 60% (24% formamide and

180   25.2% urea). The gels were run in a Bio-Rad DCode Universal Mutation Detection

181   System (Cat# 170-9080) at 180V for 3.5 hours. The gels were stained in 1X TAE

182   containing SybrGreen (100 l/L) for 30 minutes then destained in 1X TAE for 15

183   minutes. The gels were photographed under UV light and DGGE bands were identified

184   and analyzed using Kodak 1D software. For our analyses, only bands with a minimum

185   intensity of 72% were recognized (program default). Bands were distinguished based on

186   migration distance within each gel, as determined by the software using the DGGE

187   product from Escherichia coli as a standard. . Although re-amplification and sequencing

188   of individual DGGE bands can be particularly important in resolving heteroduplex

189   fragments, we did not sequence the bands we detected in this study because we were

190   interested in detecting the greatest number of bands possible. A relative species richness

191   index was computed for each case by dividing the number of bands detected within a

192   treatment by the total number of unique bands found within a sample. Relative species

193   indices were computed for samples where at least 2 of the treatments amplified (YNP

194   samples n=12).

195   Statistical Analysis

196          Statistical analyses were performed using Minitab software version 13 and SPSS

197   11 (for Mac OS X). The purity of the DNA among treatments was compared using the

198   absolute value that resulted from subtracting the A260/A280 from 1.8, the A260/A280 of pure

199   DNA [21]. Univariate ANOVA was used to detect statistical differences in the quantity

200   and purity of the DNA and relative species richness (dependent variables) among the

201   treatments and by pH, temperature, and biomass type (fixed factors). For analysis of the

202   DGGE bands, only samples where at least 2 of the treatments amplified were included,

203   otherwise a statistical comparison would not be possible. The Bonferroni test (which is

204   more powerful for small sample sizes than Tukey’s test) was performed to identify

205   treatments that were significantly different.



208   Sample Preservation

209          Sucrose lysis buffer preserved the DNA of our extreme environment samples

210   better than the GIT solution. The extracted DNAs were quantified using the PicoGreen

211   reagent. Using our protocol, the PicoGreen assay has a dsDNA detection limit of 500

212   pg/ml. We were able to extract more DNA from the samples preserved in SLB than

213   from the GIT replicates (paired t-test p=0.01). We also compared the amount of DNA

214   recovered from samples that were frozen or mixed with agarose in the field with those

215   that were held at ambient temperature, there was no statistical difference between the two

216   replicates (paired t-test =0.05). Even when samples cannot be frozen immediately, the

217   DNA is adequately preserved in sucrose lysis buffer. Although this study did not assess

218   the relative effectiveness of the different methods in extracting RNA, we have

219   successfully produced cDNA from samples preserved in SLB using reverse transcriptase

220   PCR (unpublished data).

221          There was no significant difference between the preservation methods in the

222   diversity we detected (paired t-test =0.05). However, the 16S rDNA from the samples

223   that were preserved in SLB were amplified by PCR more successfully; 80.5% of SLB

224   preserved samples PCR amplified compared to just 50% of the GIT preserved samples.

225   DNA extraction

226          The CTAB extraction on SLB preserved samples resulted in the greatest yield of

227   DNA. The results of the quantification are displayed in Fig. 1a. ANOVA showed a

228   statistically significant difference in DNA yield between the extraction methods

229   (p<0.001). Post-hoc multiple comparison analysis indicates that the CTAB extraction on

230   SLB preserved samples resulted in the highest quantity of DNA.

231          DNA purity, as measured by A260:A280 ratio, was greatest in samples extracted

232   using the commercial Mo Bio extraction kits, however the kit results in less DNA and

233   smaller fragments (presumably from shearing and the selecting column used in the kit)

234   (Fig. 1b). There was no significant difference between the purity of the DNA extracted

235   with the Boil, CTAB S, or CTAB G methods (ANOVA =0.05). The quantity of DNA

236   extracted by the Noodle method was not sufficient to obtain reliable absorbance data,

237   therefore the purity of the DNA extracted with that method could not be compared to the

238   other methods. Most of the amplification success we observed was in sample extracted

239   with either the CTAB method (89% amplified) or the Mo Bio kit (95% amplified)

240   (ANOVA and Bonferroni test p<0.001).

241          The most important parameter to consider when choosing a preservation and

242   extraction method is the effect a particular method has on the detectable diversity. The

243   number of bands or sequence diversity detected using DGGE varied within the replicates

244   of the samples (Fig. 2). The only statistically significant difference we detected was

245   between the CTAB S and the noodle method (p = 0.014). However, CTAB S resulted in

246   15% to 43% more bands than any of the other treatments (Fig, 3). There were no

247   differences at the fixed effects level (pH, temperature, and biomass type), nor were any

248   interactions (e.g., pH X method) among the factors detected.


250          It is standard practice that investigators informally evaluate methods to determine

251   which are appropriate for their study. However, few of these studies have been published

252   [6] and in many cases, statistical analysis are not used to determine the most appropriate

253   methods [5]. This study presents a framework for the systematic comparison and

254   evaluation of methods by statistical analysis. The current interest in the organisms that

255   inhabit extreme environments makes this study, which used samples from a wide variety

256   of thermal environments, of importance to thermal biology researchers, and applicable to

257   other microbial studies in general.

258   Sample Preservation

259          Sucrose lysis buffer preserved our extreme environment samples better than the

260   GIT solution. This was likely due to more effective lysing, as the SLB initiates the lysis

261   process while the sample is being stored. The alkaline SLB raised the pH within the

262   sample vials, potentially slowing the degradation of the DNA in low pH samples. Even

263   when samples could not be frozen immediately, the DNA is adequately preserved in

264   SLB. This result is especially important because of restrictions on carrying and shipping

265   dry ice and for samples that are collected from remote areas where it is logistically very

266   difficult to freeze samples immediately and maintain them frozen. The non-toxic nature

267   of sucrose lysis buffer may be an additional attraction for field microbiologists, and for

268   those shipping samples either commercially or hand-carrying on airplanes.

269   DNA extraction

270          When considering preservation alone, among the techniques we tested, SLB with

271   or without freezing preserved environmental DNA best. However, we also were

272   interested in the effect of extraction method on the amount and quality of extracted DNA

273   and detectable diversity.

274          CTAB extraction combined with SLB preserved samples resulted in the most

275   DNA. The greater efficiency of the CTAB extraction method has previously been found

276   in several environments, marine sediments [15] and by one of us in caterpillar gut and

277   filtered water (Takacs-Vesbach, unpublished data) indicating that this method may be

278   effective on a wide range of sample types. An extraction comparison on compost

279   microbial communities found no difference in the amount of DNA recovered by the

280   CTAB method, but it resulted in the highest percentage of cells lysed [28]. It is likely

281   that the combination of proteinase K and hot SDS lyses more cells, including cell types

282   more resistant to lysis (i.e. gram-positive bacteria)[17]. There were two exceptions

283   where larger quantities of DNA were not recovered by the CTAB method. The PicoGreen

284   analysis on two samples, 066-MV and 139-LS, indicates that more DNA was extracted

285   using the Mo Bio DNA extraction kit than was extracted with the Boil and CTAB

286   methods. The Boil and CTAB extracted DNA from these two samples was dark brown,

287   which were most likely photosynthetic pigments that were co-extracted with the DNA.

288   The Mo Bio kit, which produced clear DNA extracts, was effective at removing these

289   organics. We suspect that the co-extracted pigments interfered with PicoGreen

290   fluorescence, but both of these samples were successfully amplified with all of the

291   extraction methods, and the co-extracted pigments from the Boil and CTAB methods do

292   not appear to inhibit the PCR reaction.

293   DNA Purity

294          Extracting pure DNA from environmental samples is nearly as important as

295   extraction efficiency and is one of the most difficult technical problems in using

296   molecular techniques on environmental samples. Most DNA extraction procedures co-

297   extract humic organic carbon, pigments, heavy metals, and other contaminants. These

298   contaminants play havoc with PCR reactions and can degrade the DNA during storage.

299   Not surprisingly, the purest DNA was extracted using the commercial Mo Bio extraction

300   kit, which was the only method that incorporated a post-extraction purification, however

301   the kit results in less DNA and smaller fragments. The Mo Bio kit contains a proprietary

302   inhibitor removal solution and a column purification step that were designed to remove

303   humic acid contaminants [2]. There was no significant difference in the purity of the

304   DNA extracted with the Boil, CTAB S, or CTAB G methods (ANOVA =0.05). The

305   concentration of the DNA extracted with the Noodle method was not sufficient to obtain

306   reliable absorbance data, therefore the purity of the DNA extracted with that method

307   could not be compared to the other methods.

308   Diversity

309          PCR amplification of environmental DNA is not always a straightforward process.

310   The co-extraction of humics, metals, and other potential inhibitors figure significantly in

311   the analysis of microbial communities. In our experience, all but a very small proportion

312   of the samples that we have examined can be amplified using PCR. Some samples

313   require more effort, troubleshooting, and processing time, but in the end most can be

314   amplified. However, that being said, the samples preserved in SLB and extracted with

315   either the CTAB method or the Mo Bio kit were more successfully amplified relative to

316   the other cases. It should be noted that some of the samples with no detectable DNA, as

317   determined using the PicoGreen fluorometer quantification, still had enough DNA to

318   amplify with PCR (theoretical detection limit for PCR amplification < 1 pg/ml)[24].

319          Although the CTAB S treatments consistently had more bands than any of the

320   other methods (15-43%), the only statistical difference detected was between the CTAB

321   S and the noodle extraction method. We also tested for fixed effects such as pH,

322   temperature, and biomass type, but did not detect any statistical differences. However, as

323   stated earlier, there were three samples that we were unable to amplify using two or more

324   of the extraction treatments. It is interesting to note that all three of these samples were

325   sediments from low pH systems (<pH 3.5), indicating that there may be a sample type or

326   geochemistry effect on extraction efficiency or amplification success. This may be a

327   result of low biomass or inhibition by some unknown factor associated with sediments.

328          The widespread use of molecular techniques utilizing 16S rDNA to describe the

329   diversity of a microbial community, such as clone libraries, DGGE, and T-RFLP

330   (terminal restriction fragment length polymorphisms), have greatly increased our

331   understanding of microbial ecology and revealed novel enzymes. We are interested in

332   not only expanding our knowledge of novel species and the environments they inhabit,

333   but also in developing a baseline inventory of microbial diversity. Because of this

334   emphasis on a baseline inventory, it is very important that we maximize our ability to

335   detect all possible diversity. DGGE is an attractive method for microbial ecology studies

336   because of the relatively rapid ease with which a community profile can be generated.

337   Many researchers prefer this method to others, such as T-RFLPS, because individual

338   bands can be sequenced. This can be particularly important in distinguishing truly unique

339   bands. DGGE can resolve minor base differences among samples; therefore, distinct

340   bands may represent sequences that differ by less than 1%. Depending on the nature of

341   the study being performed, 1% may or may not be significant to the research question.

342   Because our objective was to identify the method that resulted in the greatest detectable

343   diversity, we did not sequence the bands we observed in the study. Conversely, we are

344   aware that individual bands can be comprised of several different sequences, however,

345   because our methods were standardized across samples, this effect would not figure in

346   our conclusions. We repeated the PCR amplification and DGGE for selected samples to

347   ensure reproducibility in the number of bands detected.

348          The main goal of this study was to determine the sample preservation and DNA

349   extraction method that would maximize the detectable diversity in molecular analysis of

350   environmental communities. For the purposes of preserving and extracting very high

351   molecular weight DNA, many researchers prefer the noodle method. Based on the results

352   from this study, this preservation method does not seem to work well with the samples

353   we collect which often have low biomass in a dense extracellular matrix. In many

354   samples, no DNA was visible on the ethidium bromide stained gels from the noodle

355   treatments. Additionally, we have found that this method is particularly difficult to use in

356   the remote backcountry. Originally, the noodle method was used to recover large

357   fragments of DNA for metagenomic analysis (25), but the development of new

358   sequencing technologies and metagenomic approaches has made the quality of DNA

359   from routine DNA extraction procedures such as CTAB S acceptable. While the Mo Bio

360   kit resulted in the cleanest DNA of the methods examined, the quantity of DNA obtained

361   with the kit was much lower than the concentration recovered with the Boil, CTAB G,

362   and especially the CTAB S methods. More high molecular weight DNA was extracted

363   using the Boil or CTAB method than was using the Mo Bio kit. The impurities in the

364   DNA extracted with the CTAB S method did not seem to adversely impact our ability to

365   PCR amplify the target sequence. Furthermore, CTAB S resulted in more species

366   richness than of the other treatments. Therefore, the CTAB DNA extraction method on

367   samples preserved in SLB seems to be the most appropriate for our purposes and should

368   be of interest to other projects with similar objectives.

369   Acknowledgements

370          We are grateful for the comments and input by Robert Sinsabaugh and Diana

371   Northup who kindly reviewed an earlier version of the manuscript. This work was

372   supported by NSF Biodiversity Surveys and Inventories grant 02-06773 to CTV and NPS

373   funding to Ann Rodman.


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462   Figure Legend

463   Fig. 1 a) Mean DNA concentration as measured using PicoGreen assay for each

464   preservation and extraction method with 95% confidence intervals, ourtliers are filled

465   circles. b) Mean DNA purity with 95% confidence interval and outliers. Pure DNA has

466   a A260/A280 ratio of 1.8, showen as the gray bar. The concentrations of the DNAs from

467   the Noodle extractions were not high enough concentration to be detected by the

468   florometer.


470   Fig. 2 Example DGGE of PCR products, primers 338FGC and 519R, on 6% acrylamide

471   gel with 20%-60% urea/formamide denaturing gradient, stained with SybrGreen

472   [Treatments: a) Boil; b) CTAB S; c) CTAB G; d) Mo Bio S; e) Mo Bio G; f) Noodle].


474   Fig. 3 a) Rarefaction curves of DGGE bands detected with each extraction method. The

475   95% confidance interval for the diversity of all samples and extraction methods

476   combinded is in gray to demonstrate that there was no significant difference between

477   diversity detected by each method. b) Mean diversity detected as number of DGGE

478   bands with 95% confidence intervals, outliers are filled circles.

479   Table 1 Sample site, description and environmental parameters.

      Sample ID        Sample Area1        pH T (C) Sample description         Northing2        Easting2

      007-L         YNP Lower GB          7.54   79.9 black powder             4932385.831 517103.977

      010-L         YNP Lower GB          3.55   90.6 gray clay-like sediment 4933099.227 515316.123

      022-L         YNP Lower GB          6.89   85.9 black powder             4933820.851 513269.383

      045-L         YNP Lower GB          2.68   42.4 brown foam and water     4934500.042 513403.550

      048-L         YNP Lower GB          3.39   48.8 grey mud sediment        4934417.810 513990.150

      058-L         YNP Lower GB          2.96   61.4 yellow and tan powder 4953191.966 522871.118
                                                      orange mat and black
      066-MV        YNP Mud Volcano       6.41   67.4 filaments             4939784.796 544533.185

      072-CH        YNP Crater Hills      5.47   55.6 yellow powder            4952741.659 523490.278

      088-L         YNP Lower GB          8.46   52.8 orange and green mat     4935485.818 515973.327

      126-MM        YNP Mary Mountain 6.58       79.8 grey powder              4940597.962 532861.547

      131-LS        YNP Lone Star GB      4.24   43.9 yellow filaments         4916381.813 514106.247
                                                      green mat and gray
      139-LS        YNP Lone Star GB      2.49   54.8 sediment                 4919322.475 515169.854

      171-S         YNP Shoshone GB       8.63   68.7 orange and green mat     4911109.135 515791.782

      184-S         YNP Shoshone GB       8.92   77.3 tan filament             4911522.596 515932.872

      190-S         YNP Shoshone GB       9.08   44.5 layered orange mat       4911267.100 515925.080

      BH            Jemez Bath House      7.15   76.0 yellow filaments         3959977.588 347240.017
                    Jemez Giggling Star
      GS            Resort                6.45   53.4 green mat                3959468.000 347235.000
481       Geyser Basin is abbreviated GB. 2Northing and Easting are given in UTM, grid 12N for

482   YNP, 13N for Jemez Springs, NM, datum NAD83


487   Mitchell et. al. Fig. 1

493   Mitchell et. al. Fig. 2


498   Mitchell et. al. Fig. 3


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