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Marine Geology 174 (2001) 241±247
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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
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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.
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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.
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