J. Phycol. 36, 251–258 (2000)
XANTHOGENATE NUCLEIC ACID ISOLATION FROM CULTURED
AND ENVIRONMENTAL CYANOBACTERIA1
Daniel Tillett and Brett A. Neilan2
School of Microbiology and Immunology, The University of New South Wales, Sydney 2052, NSW, Australia
The isolation of high-quality nucleic acids from
cyanobacterial strains, in particular environmental
isolates, has proven far from trivial. We present
novel techniques for the extraction of high molecular weight DNA and RNA from a range of cultured
and environmental cyanobacteria, including stains
belonging to the genera Microcystis, Lyngbya, Pseudanabaena, Aphanizomenon, Nodularia, Anabaena, and
Nostoc, based on the use of the nontoxic polysaccharide solubilizing compound xanthogenate. These
methods are rapid, require no enzymatic or mechanical cell disruption, and have been used to isolate
both DNA and RNA free of enzyme inhibitors or nucleases. In addition, these procedures have proven
critical in the molecular analysis of bloom-forming
and other environmental cyanobacterial isolates. Finally, these techniques are of general microbiological utility for a diverse range of noncyanobacterial
microorganisms, including Gram-positive and Gramnegative bacteria and the Archea.
lysis, but also interfere with most nucleic acid purification protocols (Porter 1988, Wilkins and Smart 1996).
Cyanobacterial cells are also rich in endo- and exonucleases and contain photosynthetic pigments, which
can inhibit enzymatic reactions, especially reverse
transcription (Cardellina et al. 1993, Lau et al. 1993)
and the PCR (Cohen et al. 1994, Giovannoni et al.
1990, Golden 1987, Neilan 1995a).
In consequence, the techniques developed to isolate cyanobacterial nucleic acids are often complex
and laborious, requiring either (1) mechanical cell
breakage (Golden 1987, Jackman and Mulligan 1995,
Leff et al. 1995, Luo and Stevens 1997, Reith et al.
1986, West and Adams 1997); (2) enzyme digestions
(Cohen et al. 1994, de Lorimer et al. 1984, Giovannoni et al. 1990, Golden 1987, Jackman and Mulligan
1995, Joset 1988, Lotti et al. 1996, Nishihara et al.
1997, Palenik 1994, Porter 1988, Smoker and Barnum
1988); (3) multiple organic solvent extractions (Cohen et al. 1994, Ferris et al. 1996, Giovannoni et al.
1990, Golden 1987, Jackman and Mulligan 1995, Leff
et al. 1995, Lotti et al. 1996, Mak and Ho 1991, Neilan
1995a, Nishihara et al. 1997, Palenik 1994, Porter
1988, Smoker and Barnum 1988, West and Adams
1997, Zehr and McReynolds 1989); (4) grinding under liquid nitrogen or dry ice (Kramer et al. 1996,
Luo and Stevens 1997, Van der Plas et al. 1989); (5)
hot phenol (Bovy et al. 1993, Kramer et al. 1996); (6)
or cesium chloride ultracentrifugation (de Lorimer et
al. 1984, Giovannoni et al. 1990, Golden 1987, Joset
1988, Leff et al. 1995, Reith et al. 1986). In addition,
many of these techniques fail to isolate nucleic acids
from all cyanobacterial strains or environmental isolates.
Jhingan (1992) introduced a novel method for the
extraction of DNA from plant matter based on the use
of metal xanthates. The formation of water-soluble
polysaccharide xanthates with potassium ethyl xanthogenate was used to disrupt plant cell walls. In the
presence of amine groups, these polysaccharide xanthates form insoluble complexes that are selectively
precipitated (Carr et al. 1975). In addition, the xanthates bind metal ions, thus potentially inhibiting the
activity of DNA degrading enzymes, as well as chelating ionic inhibitors of DNA amplification reactions.
This rapid xanthogenate-based method does not require the use of toxic organics, enzymatic digestions,
or cesium chloride ultracentrifugation. In addition, it
can be performed without mechanical tissue homogenizations. We reasoned, as the Cyanobacteria present
many of the same difficulties provided by plants, this
Key index words: cyanobacteria; DNA extraction; nucleic acids; PCR; xanthogenate
The Cyanobacteria are a diverse and cosmopolitan
bacterial phylum and possess a number of unique biological characteristics. All cyanobacteria perform oxygenic photosynthesis and contain chlorophyll a and
accessory photopigments, such as phycocyanin and
phycoerythrin (Castenholz and Waterbury 1989). Many
species are capable of fixing atmospheric nitrogen,
some of which differentiate specialized cell types for
this process and other functions (Castenholz and Waterbury 1989, Golden 1987). In addition, a number of cyanobacterial species can form symbiotic relationships
with a taxonomically wide range of organisms, including animals, plants, fungi, algae, nonphotosynthetic
protists, and heterotrophic bacteria (West and Adams
1997).
Progress in determining the molecular mechanisms underlining cyanobacterial form and function
has been hindered by the difficulty encountered with
by many cyanobacteria in isolating high-quality nucleic acids. Many cyanobacterial strains produce copious quantities of mucilaginous polysaccharides, which
not only make it difficult to achieve complete cellular
1
Received 29 April 1999. Accepted 27 October 1999.
Author for reprint requests; fax ⫹61 2 9385 1591; e-mail b.neilan
@unsw.edu.au.
2
251
252
DANIEL TILLETT AND BRETT A. NEILAN
technique may provide a safe, rapid, and efficient
means to extract DNA from cyanobacteria. Unfortunately, we, together with other investigators (Johns et
al. 1997, Ross 1995, Williams and Ronald 1994),
found the original protocol to be unreliable, and in
our hands most cyanobacterial strains yielded little or
no DNA.
In this paper we present new protocols, based on
the use of xanthogenate, for the rapid extraction of
high-quality DNA and RNA from a wide range of environmental and cultured cyanobacteria. In addition,
these protocols have proven of general utility for the
extraction of DNA and RNA from a diverse range of
microorganisms and environmental samples.
materials and methods
Bacterial and archeal strains. The strains used in this study are
listed in Table 1. Cyanobacterial strains with designations AWT,
NIES, or PCC were obtained from Australian Water Technology (Sydney, Australia), the National Institute for Environmental Studies (Tsukuba, Japan) (Natl. Inst. for Environmental
Studies 1991), and the Pasteur Culture Collection (Paris,
France) (Rippka and Herdman 1992), respectively. Microcystis
incerta HINDAK 1965/17 was obtained from the Institute of
Botany, Czech Academy of Science. Escherichia coli JM109 (Yanish-Perron et al. 1985) was obtained from Promega (Madison,
WI). Methanococcoides burtonii (Franzmann et al. 1992), Vibrio angustum S14, the Rhodococcus sp. (Paje et al. 1997), Helicobacter pylori SS1 (Lee et al. 1997), and the Acetobacter sp. (Bernardo et al.
1998) were kind gifts of T. Thomas, S. Srinivasan, C. Svenson,
B. Burns, and E. Bernardo respectively. The cyanobacterial
blooms samples were kindly supplied by P. Hawkins.
The cyanobacterial strains were maintained in either JM
(Natl. Inst. for Environmental Studies 1991) or BG-11 (Castenholz and Waterbury 1989) media at 25⬚ C with a light intensity
of approximately 20 ␮mol⭈m⫺2⭈s⫺1. E. coli JM109 was grown at
37⬚ C in LB media (Sambrook et al. 1989). M. burtonii was
grown at 22.5⬚ C in liquid methanogen growth media under
anaerobic conditions (Franzmann et al. 1992). V. angustum S14
was grown at 25⬚ C in LB media supplemented with 20 g⭈L
NaCl. The Rhodococcus strain was grown at 25⬚ C in PAS media
(Paje et al. 1997). H. pylori SS1 was grown at 37⬚ C under mi-
Table 1.
Bacterial and archeal strains used in this study.
Strain
Microcystis aeruginosu PCC 7806
Microcystis wesenbergii NIES 107
Microcystis viridis NIES 102
Microcystis incerta HINDAK1965/17
Lyngbya sp. AWT 211.
Pseudanabaena sp. AWT 210,
Aphanizomenon flos-aquae NIES 81
Nodularia spumigena PCC 73104
Anabaena circinalis AWT 006
Nostoc punctiforme PCC 73102
Escherichia coli JM109
Acetobacter sp.
Vibrio angustum S14
Helicobacter pylori SS1
Rhodococcus sp.
Methanococcoides burtonii
a
croaerophilic condition on CSA media (Lee et al. 1997). The
Acetobacter strain was grown at 30⬚ C in coconut water medium
(Bernardo et al. 1998).
DNA extraction. DNA was isolated using the methods of
Neilan et al. (1993, 1995), Porter (1988), Jhingan (1992), or
the following xanthogenate-SDS (XS) DNA extraction protocol. Briefly, 1 mL volumes of mid to late logarithmic growth
phase bacterial cell cultures were harvested by centrifugation
and the cell pellets resuspended in 50 ␮L of TER (10 mM TrisHCl, pH 7.4; 1 mM EDTA, pH 8; 100 ␮g⭈mL RNase A). Alternatively, small amounts of environmental samples, approximately
10–20 mg wet weight, were resuspended in 50 ␮L of TER. To
each cell suspension in a 1.5-mL microcentrifuge tube was
added 750 ␮L of freshly made XS buffer (1% potassium ethyl
xanthogenate [Fluka, Buchs, Switzerland]; 100 mM Tris-HCl,
pH 7.4; 20 mM EDTA, pH 8; 1% sodium dodecylsulfate; 800
mM ammonium acetate) and the tubes were inverted several
times to mix. The tubes were incubated at 70⬚ C for 10 to 120
min in a waterbath, with the time dependent on the microorganism under investigation. After incubation, the tubes were
vortexed for 10 s before being placed on ice for 30 min. Precipitated cell debris was removed by centrifugation at 14,000 rpm
for 10 min and the supernatants carefully transferred to fresh
eppendorf tubes containing 750 ␮L of isopropanol. Samples
were incubated at room temperature for 10 min and the precipitated DNA pelleted by centrifugation for 10 min at 12,000 g.
The DNA pellets were washed once with 70% ethanol, air-dried,
and finally resuspended in 100 ␮L of TE (10 mM Tris-HCl, pH
7.4; 1 mM EDTA, pH 8). To ensure that the DNA samples were
free of exo- and endonucleases, samples were incubated at 37⬚ C
for 2 h in 1⫻ HinfI restriction enzyme buffer before gel electrophoresis.
RNA extraction. RNA was extracted using the following xanthogenate-SDS-phenol (XSP) protocol. Briefly, cells were pelleted and resuspended in 50 ␮L of culture medium or TE. The
concentrated cells were added to 1.3 mL of preheated (65⬚ C)
XSP buffer (1:1 volumes of XS buffer and phenol). The tubes
were incubated at 65⬚ C for 5 min. During this incubation, the
tubes were inverted several times to facilitate the mixing of the
organic and aqueous phases, important in ensuring efficient
cell lysis. The tubes were vortexed for 10 s, and 200 ␮L of chloroform:isoamyl alcohol (24:1) was added to each tube. The
phenol and aqueous phases were separated by centrifugation at
14,000 rpm for 5 min. The aqueous phases were carefully transferred to fresh eppendorf tubes containing 500 ␮L of phenol:chloroform:isoamyl alcohol (25:24:1). The tubes were incu-
Phyluma
Kingdom
DNAb
RNAc
Cyanobacteria (Chroococcales)
Cyanobacteria (Chroococcales)
Cyanobacteria (Chroococcales)
Cyanobacteria (Chroococcales)
Cyanobacteria (Oscillatoriales)
Cyanobacteria (Oscillatoriales)
Cyanobacteria (Nostocales)
Cyanobacteria (Nostacales)
Cyanobacteria (Nostacales)
Cyanobacteria (Nostacales)
Purple Bacteria (Gamma)
Purple Bacteria (Alpha)
Purple Bacteria (Gamma)
Purple Bacteria (Epsilon)
Gram positive (High G⫹C)
Methanogen (Group V)
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Bacteria
Archea
21
16
25
9
11
8
14
8
10
5
⫹
⫹
⫹
⫹
⫹
⫹
42
44
50
14
ND
10
ND
15
11
ND
⫹
ND
ND
⫹
ND
ND
Phylum (order) of strain as based on 16S rRNA sequence.
Plus indicates DNA isolated using XS protocol (␮g⭈mg dry cell weight). Noncyanobacterial strains were not quantified during this
study.
c Plus indicates total RNA isolated using XSP protocol (␮g⭈mg dry cell weight). ND, not determined. Noncyanobacterial strains were
not quantified during this study.
b
253
XANTHOGENATE DNA EXTRACTION FROM CYANOBACTERIA
bated at 65⬚ C for 2 min, vortexed briefly, then centrifuged for
4 min at 14,000 rpm. The aqueous phases were once again carefully transferred to fresh tubes. The phenol–chloroform extractions were repeated until no opaque interface was visible between the two phases; in most cases two extractions were
sufficient. After the final phenol–chloroform extraction, the
aqueous phases were transferred to fresh tubes and 1 volume of
isopropanol was added. The tubes were inverted several times
and the RNAs allowed to precipitate on ice for 15 min. Total
RNA was isolated by centrifugation at 14,000 rpm for 10 min.
The RNA pellets were washed once with 70% ethanol, air-dried,
and finally resuspended in 50 ␮L of DEPC-treated water (Sambrook et al. 1989). To ensure that the RNA samples were free of
ribonucleases, samples were incubated at 42⬚ C for 16 h before
gel electrophoresis.
PCR amplification. Amplification of the phycocyanin intergenic spacer region (PC-IGS) and 16S rRNA gene were performed as described previously (Neilan et al. 1995a, 1997).
Briefly, the PC-IGS PCR contained 2 ␮L of 10⫻ PCR buffer
(Biotech International, Perth, Australia), 2 ␮L of 25 mM
MgCl2, 0.5 ␮L of 10 mM of each deoxynucleotide triphosphate,
5 pmol of each of the two PC-IGS primers (Table 2),] 10 ng of
genomic DNA, 1 unit of Taq polymerase (Biotech International, Perth, Australia), and water to 20 ␮L. The PC-IGS PCRs
were subjected to 30 cycles of 94⬚ C for 10 s, 50⬚ C for 20 s, and
72⬚ C for 40 s in a Perkin-Elmer 2400 PCR thermocycler.
The cyanobacterial 16S rDNA gene PCR amplification was
performed as described previously, except that primers 27F and
408R (Table 2) were used (Neilan et al. 1995a). The cyanobacterial 16S rDNA PCR thermal cycling conditions consisted of 30
cycles of 94⬚ C for 10 s, 60⬚ C for 20 s, and 72⬚ C for 40 s.
Restriction enzyme digestion. Approximately 200 ng of XS isolated DNA was digested with 5 units of the restriction enzyme
HinfI in 15 ␮L of 1⫻ buffer (Boehringer, Mannheim, Germany). Reaction mixtures were incubated overnight at 37⬚ C
prior to analysis by agarose gel electrophoresis.
results
XS DNA isolation from cultured cyanobacteria. The XS
DNA extraction protocol was tested on a range of cultured cyanobacteria from the orders Chroococcales,
Oscillatoriales, and Nostocales (Table 1). High-quality
DNA was isolated from all strains tested, including Microcystis aeruginosa PCC 7806, M. wesenbergii NIES 107,
M. viridis NIES 102, M. incerta HINDAK 1965/17, Lyngbya sp. AWT 211, Pseudanabaena sp. AWT 210, Aphanizomenon flos-aquae NIES 81, Nodularia spumigena PCC
73104, Anabaena circinalis AWT 006, and Nostoc punctiforme PCC 73102 (Fig. 1A). The DNA yield varied between 5 ␮g⭈mg⫺1 and 25 ␮g⭈mg⫺1 of dry cell weight,
depending on the strain studied.
The quality of the extracted DNA for successive enzymatic reactions was assessed by restriction enzyme
digestion and PCR (Fig. 1B and 1C). All DNAs were
found to be free of active DNases and were efficiently
cleaved by the restriction enzyme HinfI. Because
many cultures of filamentous cyanobacterial strains
Table 2.
contain contaminating heterotrophic bacteria, the cyanobacterial phycocyanin intergenic spacer PCR was
performed to ensure that cyanobacterial DNA had
been extracted (Fig. 1C). PCR amplicons were obtained from all cyanobacterial strains tested, indicating that not only was cyanobacterial DNA isolated, but
the isolated DNA was free from PCR inhibitors.
XS DNA isolation from environmental cyanobacterial
blooms. A large toxic cyanobacterial bloom event occurred in the Botany Ponds, Sydney, Australia, over
the late summer and autumn months of 1996. This
mixed bloom was dominated by species of the genera
Microcystis and Anabaena, and underwent a number of
complex population successions as assessed by microscopy and toxin data (Baker et al. 1998). Bloom samples had been collected on a weekly to monthly basis
and frozen stocks were stored. We undertook to study
the genetic population structure by analyzing the phycocyanin intergenic spacer and 16S rDNA region
(Neilan et al. 1995a, 1997).
Initial attempts to obtain DNA free of PCR inhibitors from the Botany Ponds bloom samples proved
difficult. A range of techniques were tried, including
the methods of (Neilan et al. 1993, 1995a, Porter
1988). Many of the Botany Ponds bloom samples were
highly pigmented and the DNA extracted from these
samples failed to allow PCR amplification. This difficulty in isolating enzyme inhibitor-free DNA has been
commonly observed with environmental samples
(Giovannoni et al. 1990, Golden 1987, Neilan 1995a).
We investigated the ability of the XS DNA extraction protocol to isolate PCR inhibitor-free DNA from
the Botany Ponds bloom samples. This technique provided DNA free of PCR inhibitors with PCR amplicons
of the PC-IGS and cyanobacterial 16S rDNA regions
obtained from all eight examined bloom samples
(Fig. 2). These included samples that were originally
highly pigmented and had not provided amplifiable
template when other DNA extraction methods were
used (Neilan 1995a, Neilan et al. 1993, Porter 1988).
XS DNA isolation from other microorganisms. The XS
DNA extraction protocol was tested on a range of representative noncyanobacterial strains, to assess its general microbiological utility. A range of Gram-positive
and Gram-negative bacteria and an archea were examined, including E. coli JM109, an Acetobacter sp., V.
angustum S14, a Rhodococcus sp., H. pylori SS1, and
Methanococcoides burtonii (Table 1). High-quality DNA
was isolated from all Gram-negative and Gram-positive bacteria and the archea examined (Fig. 3), including strains that have previously proven recalci-
Oligonucleotide primers used in this study.
Primer
Sequence
Reference
27F
408R
PC␤F
PC␣R
AGAGTTTGATCCTGGCTCAG
TTACAA(C/T)CCAA(G/A)(G/A)(G/A)CCTTCCTCCC
GGCTGCTTGTTTACGCGACA
CCAGTACCACCAGCAACTAA
Neilan et al. 1997
Neilan et al. 1997
Neilan et al. 1995
Neilan et al. 1995
254
DANIEL TILLETT AND BRETT A. NEILAN
Fig. 1. DNA isolated from cultured
cyanobacteria using the XS DNA extraction method. (A) 2 ␮L of genomic DNA
electrophoresed on a 0.65% agarose gel in
1⫻ Tris–Acetate–EDTA (TAE) buffer with
100 ng of lambda HindIII DNA marker
(lane 12). (B) HinfI restriction enzyme digest of XS isolated genomic DNA electrophoresed on a 1.5% agarose gel in 1⫻
TAE with 50 ng of lambda HindIII DNA
marker (lane 12). (C) Amplification products from the cyanobacterial phycocyanin
intergenic spacer PCR electrophoresed on
a 3% agarose gel in 1⫻ TAE with 100 ng of
Phi-X 174 HaeIII DNA marker (lane 12).
All gels were stained with ethidium bromide and photographed under UV transillumination.
trant with other methods (Bernardo et al. 1998, Paje
et al. 1997).
The DNA quality and yield obtained from noncyanobacteria was assessed by restriction enzyme digestion (Fig. 3). All DNA samples were digested with the
restriction enzyme HinfI and were observed to be free
of extraneous nucleases, including the H. pylori strain,
which had been previously found to contain high levels of exonucleases (de Ungria et al. 1998).
XSP RNA extraction from cyanobacteria and other microorganisms. In an effort to study the transcriptional regulation of the microcystin synthetase gene in Microcystis aeruginosa PCC 7806 (Dittmann et al. 1997 and
Tillett and Neilan unpublished data) we required a
robust RNA extraction protocol suitable for cyanobacteria. Previous investigators have also noted difficulties in RNA isolation from cyanobacteria and other
polysaccharide-rich samples (Bugos et al. 1995, Luo
XANTHOGENATE DNA EXTRACTION FROM CYANOBACTERIA
Fig. 2. Isolation of PCR inhibitor-free DNA from environmental cyanobacterial blooms using the XS DNA extraction
method. Phycocyanin intergenic spacer pyc PCR and cyanobacterial 16S rDNA (16S) were performed on XS-extracted DNA
from the Botany Ponds cyanobacterial bloom samples collected
in 1996 on the dates indicated. Both PCR products from each
sample were pooled, and a total of 4 ␮L run on a 2% agarose
gel in 1⫻ TAE with 100 ng of Phi-X 174 HaeIII DNA marker.
The gel was stained with ethidium bromide and photographed
under UV transillumination.
and Stevens 1997, Wilkins and Smart 1996). We reasoned that a combination of the XS buffer with the
hot phenol method (Giovannoni et al. 1990, Kramer
et al. 1996, Sambrook et al. 1989, Wilkins and Smart
1996) might enable high-quality RNA to be isolated
rapidly from cyanobacteria in general and from M.
aeruginosa PCC 7806 in particular.
The XSP RNA extraction protocol was tested on a
range of cyanobacterial and noncyanobacterial strains,
255
including M. aeruginosa PCC 7806, M. wesenbergii NIES
107, M. viridis NIES 102, M. incerta HINDAK 1965/17,
Nodularia spumigena PCC 73104, Pseudanabaena sp.
AWT 210, Anabaena circinalis AWT 006, E. coli JM109,
and H. pylori SS1 (Table 1). Total RNA yield varied
among the strains examined, ranging from 10 ␮g⭈mg⫺1
to 50 ␮g⭈mg⫺1 of dry cell weight (Fig. 4). We regularly
obtained ⬎2 ␮g of total RNA per milliliter of M. aeruginosa PCC 7806 cell culture. This compares favorably
with the method of Luo (Luo and Stevens 1997), who
obtained 82 ␮g of total RNA from 2 L of cyanobacterial culture. All XSP-extracted RNA samples were observed to be free of extraneous RNases, with no degradation visible after incubation at 42⬚ C for 16 h (Fig.
4). In addition, RNA isolated from M. aeruginosa PCC
7806 and H. pylori SS1 using the XSP method has
been used successfully for primer extension and nuclease protection analysis (Tillett and Neilan 1999a).
discussion
The advent of cyanobacterial molecular investigations has created the need for rapid and efficient
techniques for the extraction of nucleic acids from
both cultured and environmental isolates. This paper
presents novel methods for the extraction of highquality nucleic acids from a wide range of environmental and cultured cyanobacteria, as well as from
other bacterial and archeal microorganisms.
XS DNA isolation. The XS DNA extraction method
rapidly provided high-quality DNA of high molecular
weight from a diverse range of cyanobacteria and
other microorganisms. The DNA was free of enzymatic inhibitors and was suitable for restriction enzyme digestion, cloning, and PCR (Tillett and Neilan
1999b). The XS buffer contains no hazardous chemicals and is of low cost. In addition, this technique re-
Fig. 3. DNA isolated from noncyanobacterial microorganisms using the
XS DNA method. Lane 1: 50 ng of
lambda HindIII DNA marker. Lanes 2,
4, 6, 8 and 10: 2 ␮L of genomic DNA
isolated using the XS protocol. Lanes
3, 5, 7, 9, 11: 7.5 ␮L of HinfI-digested,
XS-isolated genomic DNA. Lane 12:
100 ng Phi-X 174 HaeIII DNA marker.
DNA samples were electrophoresed on
a 1.5% agarose gel in 1⫻ TAE, stained
with ethidium bromide, and photographed under UV transillumination.
256
DANIEL TILLETT AND BRETT A. NEILAN
quires minimal handling of the microbial sample and
does not require mechanical homogenization. This
not only reduces the risk of sample contamination,
but minimizes the production of aerosols, of particular importance if the sample contains possible human
pathogens (Ross 1995). This technique is routinely
used in our laboratory and, to date, the XS protocol
has been used successfully on ⬎200 cyanobacteria
strains and other microorganisms (Tillett and Neilan
unpublished data). While we have found there is generally a good correlation between cyanobacterial culture density and DNA yield, it should be noted that
the DNA yield is generally greater and of better quality from cells extracted in mid-log phase. Finally, the
XS DNA extraction method provided PCR inhibitorfree microbial DNA from a range of environmental
samples, including cyanobacterial blooms, river sediment, worm castings, and marine biofilms (Thompson 1997 and Tillett and Neilan unpublished data).
Important steps in the XS DNA extraction procedure should be noted. First, the XS buffer should be
freshly prepared. We routinely make the XS buffer
from stored premade stock solutions and potassium
ethyl xanthogenate powder. Second, the XS buffer
should not be overloaded with biomass by attempting
to extract too much sample, as this results in DNA of
poor quality. We therefore recommend using 750 ␮L
of XS buffer per 1–2 mL of log-phase cell culture.
Third, the cell pellet should be fully resuspended in
TER buffer before the addition of the XS buffer.
Fourth, the 70⬚ C incubation time required varies
Fig. 4. Total RNA isolated from cyanobacteria and noncyanobacteria using
the XSP method. Two ␮L of XSP isolated
RNA samples were electrophoresed on a
nondenaturing 2% agarose gel in 1⫻ TAE
with 100 ng of Phi-X 174 HaeIII DNA
marker. The gel was stained with ethidium
bromide and photographed under UV
transillumination.
from bacterial strain to strain, and it is advisable to determine the ideal time empirically. We have found
that Gram-negative bacteria and archea require 15
min or less, unicellular cyanobacteria 30 to 60 min,
and filamentous cyanobacteria and environmental
samples up to 2 h. Fifth, it is recommended the samples be vortexed for at least 10 s after the 70⬚ C incubation step. Without vortexing, chromosomal DNA
remains associated with other cellular components
and is lost during the debris precipitation step. Sixth,
the samples should be left on ice for a minimum of 15
min, or until the XS buffer becomes opaque and viscous. This ensures that proteins and other cellular debris have aggregated before centrifugation.
XSP RNA isolation. We found the combination of
hot phenol and XS buffer (XSP) to be a rapid and effective method to isolate total RNA from cyanobacteria and other microorganisms. This protocol allows
the isolation of greater than 100 ␮g of nuclease-free
total RNA in ⬍2 h. RNA isolated using the XSP protocol has been used for reverse transcriptions, nuclease
protection assays, primer extensions, and Northern
blotting (Tillett and Neilan 1999a). This technique
does not require expensive chemicals such as guanadinium salts, grinding under liquid nitrogen, or cesium chloride ultracentrifugation. In addition, it has
proven to be of general utility with a range of noncyanobacterial microorganisms.
The important steps in the XSP RNA extraction
procedure should also be noted: First, the XSP buffer
should be preheated to ensure the immediate inacti-
XANTHOGENATE DNA EXTRACTION FROM CYANOBACTERIA
vation of RNases upon cell lysis. Second, inversion
and vortexing of the tubes is important in ensure efficient cell lysis and separation of the RNA from other
cellular components. Third, heating the sample with
phenol–chloroform. The failure of cold phenol–chloroform to remove RNases is enigmatic, however, the
use of hot phenol–chloroform proved critical for the
removal of endogenous RNases from certain cyanobacterial strains. Interestingly, no cautions were taken
to exclude exogenous RNases beyond the wearing of
gloves and DEPC treatment of the RNA resuspension
water. This suggests that the major problems encountered with RNA isolations are from endogenous and
not exogenous sources. Denaturation of genomic
DNA by the hot phenol in the XSP extraction technique results in a relatively low level of DNA contamination, from ⬍1 ng to 20 ng of DNA per microgram
of total RNA (Fig. 4). Depending on the organism,
this method enriches the RNA fraction of total nucleic acids by 20- to 100-fold (Table 1). This level of
DNA contamination should not prove problematic
for many applications (e.g. Northern blots). If required, however, the DNA can be easily removed by
enzymatic DNase treatment (Sambrook et al. 1989).
We have successfully used a post-XSP DNase treatment step to remove contaminating DNA in the development of an RT-PCR-based transcript mapping protocol (Tillett and Neilan 1999a). Finally, the XSP
extraction technique has proven of general utility in
the extraction of DNA from nuclease-rich samples. To
date, it has been used to extract DNA from DNaserich Vibrio vulnificus cultures (Weichart et al. 1997),
plasmids from clinical Helicobacter isolates (de Ungria
et al. 1998), as well as many cyanobacterial strains
(Tillett and Neilan unpublished data).
The work described here presents novel nucleic
acid extraction protocols for cultured and environmental microorganisms. In addition, these protocols
may be of utility to other fields, as they have also
proven of use in the extraction of nucleic acids from a
diverse range of sources, including rocks, fungi, blood,
mouse gastrointestinal tissue, and plant matter.
This work was supported by the Australian Research Council
and Australian Water Technologies. We would like to thank
R. A. Bass for her expert technical assistance and Carolina Beltran for her support and encouragement.
Baker, J., McKay, D., Choice, M., Chandrasena, N. & Hawkins P. R.
1998. Bloom and bust: toxin production in cyanobacteria. In
Proc. 4th Int. Conf. Conference on Toxic Cyanobacteria, Beaufort,
NC, p. 17.
Bernardo, E. B., Neilan, B. A. & Couperwhite, I. 1998. Characterization, differentiation and identification of two wild-type bacterial cellulose-synthesizing Acetobacter strains involved in nata
production. Syst. Appl. Microbiol. 21:599–608.
Bovy, A., de Kruif, J., de Vrieze, G., Borrias, M. & Weisbeek, P. 1993.
Iron-dependent protection of the Synechococcus ferredoxin I
transcript against nucleolytic degradation requires cis-regulatory sequence in the 5⬘ part of the messenger RNA. Plant Mol.
Biol. 22:1047–65.
Bugos, R. C., Chiang, V. L., Zhang, X.-H., Campbell, E. R., Podila,
257
G. K. & Campbell, W. H. 1995. RNA isolation from plant tissue
recalcitrant to extraction in guanidine. BioTechniques 19:734–7.
Cardellina, J. H., Munro, M. H. G., Fuller, R. W., Manfriedi, K. P.,
McKee, T. C., Tischler, M., Bokesch, H. R., Gustafson, K. R.,
Beutler, J. A. & Boyd, M. R. 1993. A chemical screening strategy for the dereplication and prioritization of HIV-inhibitory
aqueous natural products extracts. J. Nat. Prod. 56:1123–9.
Carr, M. E., Hofreiter, B. T. & Russel, R. C. 1975. Starch xanthatepolyethylenimine reaction mechanisms. J. Polym. Sci. Polym.
Chem. 13:1441–56.
Castenholz, R. W. & Waterbury, J. B. 1989. Section 19. Oxygenic
photosynthetic bacteria, group 1, cyanobacteria. In Staley, J. T.,
Bryant, M. P., Pfennig, N. & Holt, J. G. [Eds.] Bergey’s Manual of
Systematic Bacteriology. Williams & Wilkins, Baltimore, pp. 1710–99.
Cohen, M. F., Wallis, J. G., Campbell, E. L. & Meeks, J. C. 1994.
Transposon mutagenesis of Nostoc sp. strain ATCC 29133, a filamentous cyanobacterium with multiple cellular differentiation alternatives. Microbiology 140:3233–40.
de Lorimer, R., Bryant, D. A., Porter, R. D., Liu, W.-Y., Jay, E. &
Stevens, S. E., Jr. 1984. Genes for the ␣ and ␤ subunits of phycocyanin. Proc. Natl. Acad. Sci. USA 81:7946–50.
de Ungria, M. C. A., Tillett, D., Neilan, B. A., Cox, P. & Lee, A.
1998. A novel method of extracting plasmid DNA from Helicobacter spp. Helicobacter 3:269–77.
Dittmann, E., Neilan, B. A., Erhard, M., von Dohren, H. & Borner,
T. 1997. Insertional mutagenesis of a peptide synthetase gene
that is responsible for hepatotoxin production in the cyanobacterium Microcystis aeruginosa PCC 7806. Mol. Microbiol.
26:779–87.
Ferris, M. J., Muyzer, G. & Ward, D. M. 1996. Denaturing gradient
gel electrophoresis profiles of 16S rRNA-defined populations
inhabiting a hot spring microbial mat community. Appl. Environ. Microbiol. 62:340–6.
Franzmann, P. D., Springer, N., Ludwig, W., De, M. E. C. & Rohde,
M. A. 1992. A methanogenic archaeon from Ace Lake, Antarctica: Methanococcoides burtonii, new species. Syst. Appl. Microbiol.
15:573–81.
Giovannoni, S. J., DeLong, E. F., Schmidt, T. M. & Pace, N. R. 1990.
Tangential flow filtration and preliminary phylogenetic analysis of marine picoplankton. Appl. Environ. Microbiol. 56:2572–5.
Golden, S. S. 1987. Genetic engineering of the cyanobacterial chromosome. Methods Enzymol. 153:215–31.
Jackman, D. M. & Mulligan, M. E. 1995. Characterization of a nitrogen-fixation (nif) gene cluster from Anabaena azollae 1a shows
that closely related cyanobacteria have highly variable but
structured intergenic regions. Microbiology 141:2235–44.
Jhingan, A. K. 1992. A novel technology for DNA isolation. Methods
Mol. Cell Biol. 3:15–22.
Johns, M. A., Skrock, P. W., Nienhuis, J., Hinrichsen, P., Bascur, G.
& Munoz-Schick, C. 1997. Gene pool classification of common
bean landraces from Chile based on RAPD and morphological
data. Crop Sci. 37:605–13.
Joset, F. 1988. Transformation in Synechocystis PCC 6714 and 6803:
Preparation of chromosomal DNA. Methods Enzymol. 167:712–4.
Kramer, J. G., Wyman, M., Zehr, J. P. & Capone, D. G. 1996. Diel
variability in transcription of the structural gene for glutamine
synthetase (glnA) in natural populations of the marine diazotrophic cyanobacterium Trichodesmium thiebautii. FEMS Microbiol. Ecol. 21:187–96.
Lau, A. F., Siedlecki, J., Anleitner, J., Patterson, G. M., Caplan, F. R. &
Moore, R. E. 1993. Inhibition of reverse transcription activity by
extracts of cultured blue-green algae. Planta Med. 59:148–51.
Lee, A., O’Rourke, J., De Ungria, M. C., Robertson, B., Daskalopoulos, G. & Dixon, M. F. 1997. A standardized mouse model of
Helicobacter pylori infection: introducing the Sydney strain. Gastroenterology 112:1386–97.
Leff, L. G., Dana, J. R., McArthur, J. V. & Shimkets, L. J. 1995. Comparison of methods of DNA extraction from stream sediment.
Appl. Environ. Microbiol. 61:1141–3.
Lotti, F., Giovannetti, L., Margheri, M. C., Ventura, S. & Materassi,
R. 1996. Diversity of DNA methylation pattern and total DNA
restriction pattern in symbiotic Nostoc. World J. Microbiol. Biotechnol. 12:38–42.
258
DANIEL TILLETT AND BRETT A. NEILAN
Luo, X.-Z. J. & Stevens, S. E., Jr. 1997. Isolation of full-length RNA
from a thermophilic cyanobacterium. BioTechniques 23:904–9.
Mak, Y. M. & Ho, K. K. 1991. An improved method for the isolation
of chromosomal DNA from various bacteria and cyanobacteria. Nucleic Acids Res. 20:4101–2.
National Institute for Environmental Studies 1991. Microbial culture
collection list of strains. Environmental agency, Tsukuba, Japan.
Neilan, B. A. 1995. Identification and phylogenetic analysis of toxigenic cyanobacteria using a RAPD PCR. Appl. Environ. Microbiol. 61:2286–91.
Neilan, B. A., Gurvitz, A., Leigh, D. A., Lai, L. Y. C. & McDonald, B.
1993. Rapid preparation of limited biological samples for
small-volume PCR. PCR Methods Appl. 2:261–2.
Neilan, B. A., Jacobs, D., Del Dot, T., Blackall, L. L., Hawkins, P. R.,
Cox, P. T. & Goodman, A. E. 1997. rRNA sequences and evolutionary relationships among toxic and nontoxic cyanobacteria
of the genus Microcystis. Int. J. Syst. Bacteriol. 47:693–7.
Neilan, B. A., Jacobs, D. & Goodman, A. E. 1995. Genetic diversity
and phylogeny of toxic cyanobacteria determined by DNA
polymorphisms within the phycocyanin locus. Appl. Environ.
Microbiol. 61:3875–83.
Nishihara, H., Miwa, H., Watanabe, M., Nagashima, M., Yagi, O. &
Takamura, Y. 1997. Random amplified polymorphic DNA
(RAPD) analyses for discriminating genotypes of Microcystis cyanobacteria. Biosci. Biotechnol. Biochem. 61:1067–72.
Paje, M. L., Neilan, B. A. & Couperwhite, I. 1997. A Rhodococcus that
thrives on a medium saturated with liquid benzene. Microbiology 143:2975–81.
Palenik, B. 1994. Cyanobacterial community structure as seen from
RNA polymerase gene sequence analysis. Appl. Environ. Microbiol. 60:3212–9.
Porter, R. D. 1988. DNA transformation. Methods Enzymol. 167:
703–12.
Reith, M. E., Laudenbach, D. E. & Straus, N. A. 1986. Isolation and
nucleotide sequence analysis of the ferredoxin I gene from the
cyanobacterium Anacystis nidulans R2. J. Bacteriol. 168:1319–24.
Rippka, R. & Herdman, M. 1992. Pasteur Culture Collection (PCC) of
Cyanobacterial Strains in Axenic Culture. Vol. 1. Catalogue of
Strains. Institut Pasteur, Paris, France.
Ross, I. K. 1995. Non-grinding method of DNA isolation from human pathogenic filamentous fungi using xanthogenates. BioTechniques 18:828–30.
Sambrook, J., Fritsch, E. F. & Maniatis, T. 1989. Molecular Cloning, A
Laboratory Manual. 2nd ed. Cold Spring Harbor Laboratory
Press, Cold Spring Harbor, New York.
Smoker, J. A. & Barnum, S. R. 1988. Rapid small-scale DNA isolation
from filamentous cyanobacteria. FEMS Microbiol. Lett. 56:119–22.
Thompson, L. 1997. Honours thesis. The University of New South
Wales, Sydney.
Tillett, D. & Neilan, B. A. 1999. Enzyme-free cloning: a rapid
method to clone PCR products independent of vector restriction enzyme sites. Nucleic Acids Res. 27:e26.
Tillett, D. & Neilan, B. A. 2000. Optimized rapid amplification of
cDNA ends (RACE) for mapping bacterial mRNA transcripts.
BioTechniques (in press).
Van der Plas, J., Bovy, A., Kruyt, F., de Vrieze, G., Dassen, E., Klien,
B. & Weisbeek, P. 1989. The gene for the precursor of plastocyanin from the cyanobacterium Anabaena sp. PCC 7937: isolation, sequence and regulation. Mol. Microbiol. 3:275–84.
Weichart, D., McDougald, D., Jacobs, D. & Kjelleberg, S. 1997. In
situ analysis of nucleic acids in cold-induced non-culturable
Vibrio vulnificus. Appl. Environ. Microbiol. 63:2754–8.
West, N. Y. & Adams, D. G. 1997. Phenotypic and genotypic comparison of symbiotic and free-living cyanobacteria from a single field site. Appl. Environ. Microbiol. 63:4479–84.
Wilkins, T. & Smart, L. B. 1996. Isolation of RNA from plant tissue.
In Krieg, P. A. [Ed.] A Laboratory Guide to RNA: Isolation, Analysis, and Synthesis. Wiley–Liss, New York, pp. 21–42.
Williams, C. E. & Ronald, P. C. 1994. PCR template-DNA isolated
quickly from monocot and dicot leaves without tissue homogenization. Nucleic Acids Res. 22:1917–8.
Yanish-Perron, C., Vieria, J. & Messing, J. 1985. Improved M13 phage cloning vectors and host strains: nucleotide sequence of
the M13mp18 and pUC19 vectors. Gene 33:103–19.
Zehr, J. P. & McReynolds, L. A. 1989. Use of degenerate oligonucleotides for amplification of the nifH gene from the marine cyanobacterium Tricodesmium thiebautii. Appl. Environ. Microbiol.
55:2522–6.