RESEARCH ARTICLE
Soil pH regulates the abundance and diversity of Group1.1c
Crenarchaeota
Laura E. Lehtovirta, James I. Prosser & Graeme W. Nicol
Institute of Biological and Environmental Sciences, University of Aberdeen, Aberdeen, UK
Correspondence: Graeme W. Nicol, Institute
of Biological and Environmental Sciences,
University of Aberdeen, Cruickshank Building,
St Machar Drive, Aberdeen AB24 3UU, UK.
Tel.: 144 1224 272 258; fax: 144 1224 272
703; e-mail: [email protected]
Received 10 February 2009; revised 25 June
2009; accepted 29 June 2009.
Final version published online 1 September
2009.
DOI:10.1111/j.1574-6941.2009.00748.x
MICROBIOLOGY ECOLOGY
Editor: Christoph Tebbe
Keywords
Crenarchaeota; pH; 16S rRNA gene;
quantitative PCR; diversity; soil.
Abstract
Archaeal communities in many acidic forest soil systems are dominated by a
distinct crenarchaeal lineage Group 1.1c. In addition, they are found consistently
in other acidic soils including grassland pasture, moorland and alpine soils. To
determine whether soil pH is a major factor in determining their presence and
abundance, Group 1.1c community size and composition were investigated across
a pH gradient from 4.5 to 7.5 that has been maintained for > 40 years. The
abundances of Group 1.1c Crenarchaeota, total Crenarchaeota and total bacteria
were assessed by quantitative PCR (qPCR) targeting 16S rRNA genes and the
diversity of Group 1.1c crenarchaeal community was investigated by denaturing
gradient gel electrophoresis (DGGE) and phylogenetic analysis. The abundance of
Group 1.1c Crenarchaeota declined as the pH increased, whereas total Crenarchaeota and Bacteria showed no clear trend. Community diversity of Group 1.1c
Crenarchaeota was also influenced with different DGGE bands dominating at
different pH. Group 1.1c Crenarchaeota were also quantified in 13 other soils
representing a range of habitats, soil types and pH. These results exhibited the
same trend as that shown across the pH gradient with Group 1.1c Crenarchaeota
representing a greater proportion of total Crenarchaeota in the most acidic soils.
Introduction
Molecular techniques have revealed the abundance and
ubiquity of Archaea in all terrestrial and marine ecosystems
(Fuhrman et al., 1992; Buckley et al., 1998) despite earlier
indications that these organisms were restricted to extreme
environments. The presence of members of the two major
established archaeal kingdoms, Crenarchaeota and Euryarchaeota in nonthermophilic environments has been demonstrated by 16S rRNA gene surveys, with Crenarchaeota
often more common in soil environments (Nicol et al.,
2003). In particular, nonthermophilic ‘Group I’ Crenarchaeota have been reported in numerous environments including marine (Delong, 1992; Fuhrman et al., 1992) and
freshwater environments (Jurgens et al., 2000), sediments
(Hershberger et al., 1996; MacGregor et al., 1997; Schleper
et al., 1997), bulk soil (Bintrim et al., 1997; Jurgens et al.,
1997; Buckley et al., 1998; Sandaa et al., 1999; Ochsenreiter
et al., 2003) and the rhizosphere and mycorrhizosphere
(Simon et al., 2000; Bomberg et al., 2003). Estimates based
on 16S rRNA gene copy numbers suggest that Crenarchaeota
FEMS Microbiol Ecol 70 (2009) 367–376
constitute 1–5% of the total prokaryotic community in soils
(Buckley et al., 1998; Sandaa et al., 1999; Ochsenreiter et al.,
2003), with some estimates as high as 12–38% (Kemnitz
et al., 2007).
Evidence from both molecular and cultivation-based
studies suggests that nonthermophilic Crenarchaeota play a
role in ammonia oxidation (Treusch et al., 2005; Francis
et al., 2007; Prosser & Nicol, 2008). Much of this evidence is
based on analysis of the functional amoA gene, encoding a
subunit of ammonia monooxygenase. Comparison of amoA
gene abundance and transcript abundance suggests that
ammonia-oxidizing archaea may be more active in the soil
than ammonia-oxidizing bacteria (Leininger et al., 2006). In
addition, all Group I Crenarchaeota cultivated so far are
ammonia oxidizers. The only nonthermophilic Group 1.1a
crenarchaeon isolated, Nitrosopumilus maritimus, grows
autotrophically with ammonia as the sole energy source
(Könneke et al., 2005). In addition, two highly enriched
cultures obtained from thermophilic environments, Nitrosocaldus yellowstoneii (de la Torre et al., 2008) and Nitrososphaera gargensis (Hatzenpichler et al., 2008), placed within
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368
two distinct lineages of Group I Crenarchaeota, are also
autotrophic ammonia oxidizers.
Several nonthermophilic crenarchaeal lineages have been
identified within Group I. Most aquatic sequences form a
distinct phylogenetic cluster termed 1.1a, while most soil
sequences fall within Group 1.1b (Ochsenreiter et al., 2003).
However, in acidic forest soils a different lineage dominates,
originally described in boreal Finnish forest soil (Jurgens
et al., 1997) and later described as Group 1.1c Crenarchaeota
(DeLong, 1998). This lineage has been estimated to account
for up to 85% of the total archaeal 16S rRNA gene sequences
in two soil cores of acidic forest soil (Kemnitz et al., 2007).
Group 1.1c Crenarchaeota share 80–85% of 16S rRNA gene
sequence identity with the two major crenarchaeal lineages
dominating the soil and marine environments. In a study
over a range of habitats, crenarchaeal Group 1.1c organisms
were most abundant in the montane forest soil that had the
lowest pH of all the sites studied (Oline et al., 2006), and
Group 1.1c Crenarchaeota in alpine soils were only detected
in the mature soil with lowest pH and highest nutrient levels
(Nicol et al., 2005). Group 1.1c organisms have been found
to have a specific association with boreal forest mycorrhizosphere (Bomberg & Timonen, 2007) and the occurrence of
Group 1.1c Crenarchaeota is also influenced by soil horizon
(Pesaro & Widmer, 2002), with apparent preference for the B
and unsaturated C soil horizons with lower pH (Hansel et al.,
2008). In addition, Group 1.1c Crenarchaeota have been
found in freshwater and sediments (Ochsenreiter et al., 2003)
as well as peat (Juottonen et al., 2008). The high abundance
of sequences of these organisms suggests that they may play
an important ecological role in these specific habitats. This
group has no cultivated representative and its potential
involvement in ammonia oxidation remains unknown (e.g.
as a lineage adapted to ammonia oxidation at low pH).
Soil pH has been demonstrated to affect microbial community structure (Bååth & Anderson, 2003; Nicol et al.,
2004; Fierer & Jackson, 2006) and the balance between
bacterial and crenarchaeal ammonia oxidizers is influenced
by soil pH, as indicated by analysis of abundances of amoA
genes and gene transcripts (Nicol et al., 2008). There is
currently no evidence that Group 1.1c Crenarchaeota are
ammonia oxidizers, but previous studies indicate that pH
may be an important determinant of their community
composition. No study has been carried out to verify this
and this investigation was performed to test the hypothesis
that soil pH is a major driver of 1.1c crenarchaeal abundance. This was achieved using soil plots in which pH has
been maintained at different values for over 40 years, using
16S rRNA gene techniques to determine the effects of soil
pH on the abundance and community structure of Group
1.1c Crenarchaeota. In addition, we investigated their abundance in 13 other soils with varying pH and representative of
a range of soil types and vegetation.
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L.E. Lehtovirta et al.
Materials and methods
Site description, soil sampling and nucleic acid
extraction
Soil samples were obtained from pH-manipulated plots at
the Scottish Agricultural College on Craibstone Estate,
NE Scotland (Grid ref. NJ 867112) in August 2006. The
soil plots have been maintained at 0.5 pH unit intervals in
the range 4.5–7.5 for the past 40 years, by the addition of
lime or aluminium sulphate. The soil, which is a sandy
loam, is subject to an 8-year crop rotation that minimizes
the potential effects of vegetation. The soil had supported a
crop of potatoes in the previous year. Three individual
soil samples were obtained from the surface 10 cm of each
plot, sieved through a 3.35-mm diameter metal mesh and
stored at 20 1C. Dry weight was determined following
drying at 105 1C for 24 h and total N and total C were
determined on oven-dried soil using an automated Fison
NA 1500 NCS Analyser mass spectrophotometer (Elemental
Microanalysis Ltd, Okehampton, UK). Soil pH was
determined after shaking a soil suspension for 15 min in
deionized water [1 : 2 soil : water (w/v)] and settling for
30 min, and ammonia concentration was determined
colorimetrically by flow injection analysis (FIA star 5010
Analyzer, Tecator) (Allen, 1989).
With the exception of pH, measured soil physicochemical
characteristics showed little variability between plots and no
significant trends were observed, as determined by ANOVA. The
average water, carbon and nitrogen contents across all plots on
an oven dry weight basis were 23.6( 0.4)%, 0.4( 0.02)%
and 7.0( 0.4)%, respectively, and the average ammonia
1
concentration was 1.5( 0.1) mg NH1
4 -N g soil.
To compare with data obtained from the pH-manipulated plots, soil samples from a further 13 sites were
obtained and represented a diverse range of soil, habitat
and pH ranges (Table 1). For all sites, soil was collected from
the top 0–20 cm depth. DNA was extracted from 0.5 g
triplicate samples for all sites using the method of Griffiths
et al. (2000) with some modifications (Nicol et al., 2005).
Primer design
Primers were designed to target Group 1.1c Crenarchaeota
specifically in quantitative PCR (qPCR) and DGGE assays
and to discriminate against other major crenarchaeal or
bacterial groups. To design primers, sequences representing
major crenarchaeal lineages, including 1.1a, 1.1b, 1.1c and
1.1c-associated groups, were retrieved from GenBank and
aligned using CLUSTALW (Thompson et al., 1994) implemented in BIOEDIT sequence alignment editor (Hall, 1999). Using
comparison of consensus sequences within each individual
group and also Primrose (Ashelford et al., 2002), suitable
discriminatory primer sets were identified and tested with
FEMS Microbiol Ecol 70 (2009) 367–376
369
Archaea in acid soil
Table 1. Soil characteristics and locations
Site
Soil type
Habitat type (predominant plant species)
pH
Location
1
2
3
4
5
6
7
8
9
10
11
12
13
Peaty podzol
Peaty podzol
Dystric cambisol
Brown podzol
Dystric cambisol
Dystric cambisol
Clay loam
Eutric cambisol
Peaty podzol
Clay loam
Sandy loam
Sandy soil
Loamy sand
Moorland (Calluna vulgaris)
Forest (Picea abies)
Forest (Pinus sylvestris)
Pasture (Pteridium aquilinum and Festuca ovina)
Forest (Pinus sylvestris)
Forest (Larix ssp.)
Pasture (mixed grasses)
Pasture (Lolium perenne and Trifolium repens)
Pasture (mixed grasses)
Meadow (mixed grasses)
Pasture (mixed grasses)
Pasture (mixed grasses)
Pasture (mixed grasses)
3.9
3.9
4.2
4.3
4.3
4.4
5.0
5.1
5.7
5.8
5.8
6.5
6.6
Cabrach, Scotland
Cabrach, Scotland
Cabrach, Scotland
Abergwyngregyn, North Wales
Cabrach, Scotland
Cabrach, Scotland
Cruden Bay, Scotland
Abergwyngregyn, North Wales
Cabrach, Scotland
Palace Leas, UK
Insch, Scotland
St Fergus, Scotland
Boyndie, Scotland
Table 2. Details of primers used in this study
Primers
Target groups
Sequences (5 0 –3 0 )
Uses
References
27F
A109F
200F
385R
771F
957R
1369F
1492R
Bacteria
Archaea
Group 1.1c Crenarchaeota
Group 1.1c Crenarchaeota
Group 1 Crenarchaeota
Group 1 Crenarchaeota
Bacteria
Universal
AGAGTTTGATCCTGGCTCAG
ACKGCTCAGTAACACGT
AGGAGAGATGGCTTAAAGGGG
GGATTAACCTCRTCACGCTTTCG
ACGGTGAGGGATGAAAGCT
CGGCGTTGACTCCAATTG
CGGTGAATACGTTCYCGG
GYYACCTTGTTACGACTT
PCR
Cloning
qPCR, cloning
qPCR
qPCR, DGGE
qPCR, DGGE, cloning
qPCR
PCR, qPCR
Lane (1991)
Großkopf et al. (1998)
Jurgens & Saano (1999)
This study
Ochsenreiter et al. (2003)
Ochsenreiter et al. (2003)
Suzuki et al. (2000)
Nicol et al. (2008)
Groups 1.1b, 1.1c and 1.1c-associated crenarchaeal templates to verify their specificity for Group 1.1c Crenarchaeota and their ability to exclude 1.1b and 1.1c-associated
Crenarchaeota frequently found in soil (in silico and PCR
of cloned templates from different lineages).
containing 8% polyacrylamide as described previously (Nicol et al., 2005). Electrophoresis was performed at 75 V for
960 min in 1 TAE buffer (at 60 1C) and gels were visualized by silver staining (Nicol et al., 2005) before scanning
with Epson GT9600 scanner (Epson, Hemel Hempstead,
Hertfordshire, UK).
PCR and denaturing gradient gel
electrophoresis (DGGE)
Phylogenetic analysis
A nested PCR strategy was used to obtain Group 1.1c
amplicons for DGGE analysis for all samples. First-round
PCR was performed using Group 1.1c specific primer 200F
and the universal primer 1492R (Table 2). These products
were then diluted and used for nested PCR amplification
with Group 1-specific primers 771F and 957R (with GC
clamp) (Table 2). For all standard end-point PCR, amplification was performed in 50-mL reaction mixtures consisting
of 1 mL 50-fold diluted DNA extract (approximately 10 ng
DNA), 1 PCR buffer (Bioline, London, UK), 20 mM
MgCl2, 200 nM each primer, 1 mM dNTPs and 1 U Taq
polymerase (Bioline). Cycling conditions used were 95 1C
for 5 min, followed by 35 cycles of 94 1C for 30 s, 55 1C for
30 s and 72 1C for 2 or 1 min (for first and second round,
respectively), followed by one cycle of 72 1C for 10 min.
DGGE analysis was performed using a 30–70% gradient
Based on the trends of differences in Group 1.1c crenarchaeal communities identified by DGGE analysis, cloning,
screening and sequencing were performed to obtain clones
representative of band positions of interest. Group 1.1c
sequences were amplified from environmental samples with
200F and 1492R primers as described above, generating
amplicons approximately 1300 bp in length. A total of six
clone libraries of 16S rRNA genes were constructed from
soils of pH 4.5, 6.0 and 6.5 (two, one and three clone
libraries, respectively). However, due to poor amplification
from one sample of pH 6.0 soil, a semi-nested PCR was
performed with primers 200F and 957R, generating amplicons approximately 800 bp in length. PCR products were
purified using a NucleoSpin Extract II Kit (Clontech,
Mountain View, CA), cloned into pGEM-T easy vector
system (Promega, Southampton, UK) and transformed into
FEMS Microbiol Ecol 70 (2009) 367–376
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370
L.E. Lehtovirta et al.
XL1-Blue supercompetent Escherichia coli cells (Stratagene,
Cambridge, UK). Clones were screened for inserts using
M13F/M13R vector primers before performing nested PCR
using 771F and 957R-GC primers. DGGE was performed to
screen for clones representing the major environmental
DGGE bands. Selected clones were sequenced and included
in phylogenetic analysis, with LogDet/Paralinear distances
and tree construction performed using PAUP v4.01 (Swofford, 1998) as described previously (Nicol et al., 2005).
Bootstrap support was calculated using for distance (PAUP),
parsimony (PAUP) and maximum likelihood (PHYML; Guindon & Gascuel, 2003) methods (1000, 1000 and 100 bootstrap replicates, respectively).
Crenarchaeota and nontransformed data for total Crenarchaeota and Bacteria. A Student’s t-test was carried out to
compare the abundances of Bacteria and Crenarchaeota.
Levene statistics was used to test for the homogeneity of
variances. All statistical analyses were performed using the
SPSS statistics package (SPSS Inc., Chicago, IL).
qPCR
Effects of pH on Group 1.1c crenarchaeal
community abundance
The abundance of 16S rRNA genes from Group 1.1c, total
Group 1 Crenarchaeota and total Bacteria was determined
by qPCR amplification of 16S rRNA genes of extracted
DNA. qPCR was performed using the BioRad MyIQ SingleColor Real-Time PCR Detection System (Biorad, Hertfordshire, UK) with Qiagen QuantiFast SYBR Green PCR Master
Mix (Qiagen, West Sussex, UK). Reaction mixtures of 25 mL
contained 12.5 mL Qiagen Master Mix, 400 nM of each
primer for bacterial (1369F and 1492R) and general crenarchaeal (771F and 957R) assays and 1200 nM of each
primer for the Group 1.1c assay (200F and 385R) (Table 2).
qPCR standards were generated by PCR amplification of
crenarchaeal 16S rRNA genes from extracted DNA. Clones
were screened for Group 1.1c sequences (using specific
primers and confirmation by sequencing) before amplification of the insert and quantification. A bacterial qPCR
standard was generated by amplifying and quantifying the
16S rRNA gene of E. coli, using primers 27F and 1492R (Table
2). For all runs, amplification of a serial dilution of standard
template (in triplicate) from 107 to 101 copies was used to
generate standard curves. Amplification efficiencies for all
reactions ranged from 99.4% to 111.5% with an r2 value of
4 0.99 for standard curve regression. For all qPCR assays, the
following cycling conditions were used: 95 1C for 15 min,
followed by denaturation at 94 1C for 10 s, combined annealing and extension for 30 s and fluorescence reading at 72 1C
for 6 s, followed by 72 1C for 10 min. To test the specificity of
the qPCR, after each run a melting curve analysis was
performed between 72 and 95 1C to determine the presence
of specific products. The PCR products were also subject to
standard agarose gel electrophoresis to ensure amplicons of
the expected size only were produced. Ten nanograms of
extracted environmental DNA was used as template.
Statistical analysis
The effect of pH on gene abundances was analysed by oneway ANOVA of the ln-transformed data for Group 1.1c
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Accession numbers
All sequences have been deposited in the GenBank database
with accession numbers GQ141959–GQ141977.
Results
Abundances of Group 1.1c, total Crenarchaeota and total
bacteria were determined by real-time PCR amplification of
16S rRNA genes using group-specific primers. Group 1.1c
was detected only is soils at pH 4.5–6.0 (detection limit
2.5 104 copies g1 dry soil). The abundance of Group 1.1c
Crenarchaeota was greatest (3.8 105 16S rRNA gene
copies g1 dry soil) at the lowest pH and gene abundance
declined steadily as pH increased (Fig. 1). There was at least
one order of magnitude difference in 16S rRNA gene
abundance between soils at pH 4.5 and 6.0 and, given the
lack of detection at pH values 4 6.5, the difference in
abundance between the two pH extremes studied, 4.5 and
7.5, was more than two orders of magnitude. Statistical
analysis confirmed a significant effect of soil pH on Group
1.1c crenarchaeal abundance (ANOVA, P o 0.001). In contrast
to Group 1.1c Crenarchaeota, the 16S rRNA gene abundances of total Crenarchaeota and total Bacteria did not vary
significantly with soil pH (Fig. 1) (P = 0.536 and P = 0.215,
respectively). Bacterial 16S rRNA gene abundance was
significantly greater than that of Crenarchaeota (Student’s
t-test, P o 0.001) with proportions of total Crenarchaeota
: total Bacteria ranging from 1.1% to 2.2%. Group 1.1c
Crenarchaeota constituted 0.1–1.8% of total crenarchaeal
abundance in plots where quantification was possible.
Effects of soil pH on crenarchaeal
community structure
pH-associated changes in Group 1.1c community structure
were analysed by DGGE of amplified 16S rRNA gene
fragments. A nested PCR approach was used for all samples,
and the reproducibility of DGGE profiles between biological
field triplicates was high at low pH but less so in the neutral
pH (7.0 and 7.5) soils (Fig. 2). This may reflect lower
abundances of Group 1.1c 16S rRNA gene numbers at
neutral pH, resulting in increasingly stochastic amplification. The DGGE profiles from the lowest pH soils were
FEMS Microbiol Ecol 70 (2009) 367–376
371
Archaea in acid soil
4.5
16S rRNA gene abundance
(copies g–1 dry soil)
1010
5.0
5.5
6.0
6.5
7.0
7.5
109
108
I
107
II
III
IV
V
106
105
104
103
4.5
5.0
5.5
1.1c Crenarchaeota
6.0
pH
6.5
7.0
7.5
Fig. 2. DGGE analysis of Group 1.1c Crenarchaeota in soil samples
from plots with pH ranging from 4.5 to 7.5. Band positions I–V indicate
major DGGE band positions, some of which decreased or increased
in intensity with increasing pH, and were subjected to phylogenetic
analysis.
Crenarchaeota
Bacteria
Fig. 1. Quantification of 16S rRNA gene abundance determined using
primers targeting Group 1.1c Crenarchaeota, total Group 1 Crenarchaeota and total Bacteria in Craibstone pH plots. qPCR amplification was
performed on three experimental replicates and three biological field
replicates were analysed for each pH. Error bars represent SEs of means
of biological replicates.
dominated by three major bands, at positions I, II and IV
(Fig. 2). The relative intensity of DGGE band position IV
decreased with increasing pH and the band was not detectable in many neutral pH replicates. A similar impact of pH
was seen for band positions I and II. While the relative
intensity of these bands did not diminish greatly, they were
not detectable in many replicates from soils of pH 6.5 or
higher. Band positions III and V showed a reverse trend,
increasing in intensity as pH increased. They were, however,
absent from pH 7.5 soil, possibly due to low abundance of
1.1c Crenarchaeota, as for most other phylotypes detected
by DGGE at lower pH. Several additional minor bands
appeared sporadically in at least one of the triplicates from
samples of pH 5.5 or above, but these showed no detectable
trend with pH and probably account for a relatively small
part of the community.
Phylogenetic analysis
Clone library analysis and sequencing were performed to
determine the identity of major DGGE bands of interest and
confirm affiliation within Group 1.1c. For five libraries,
1300 bp products were cloned. For one library (pH 6.0), a
nested PCR strategy was required to generate sufficient
product for cloning. Individual clones were then amplified
with the DGGE primer set for screening by DGGE and
comparison with the environmental DGGE profiles. Between two and eight clones comigrating with each DGGE
FEMS Microbiol Ecol 70 (2009) 367–376
band of interest were chosen for sequencing. Where possible, clones with the same band position were selected from
libraries obtained from soil samples of different pH. Phylogenetic analysis was performed on 619 unambiguous positions using variable sites only (65%) with reference
crenarchaeal Group 1.1c sequences from various forest soils,
moorland and alpine soils (Fig. 3). A marker composed of
these clones was run alongside the environmental samples in
DGGE analysis (Fig. 2) and most sequences showing identical migration patterns formed monophyletic groups in
phylogenetic analyses (Fig. 3). Clones sequenced in this
study were closely related to Group 1.1c 16S rRNA gene
sequences from a variety of other acidic soils, including
coniferous forest and mature alpine soils. Phylogenetic
clusters corresponding to band positions I and V contained
DNA sequences from both low and neutral pH soils, while
the other clusters originated from several clone libraries of
one pH. Sequences dominating at one pH range do not
appear to be monophyletic.
Abundance of Group 1.1c Crenarchaeota in other
soil systems
In order to verify findings on the influence of pH on Group
1.1c crenarchaeal abundance at Craibstone, 13 other soils
with different characteristics and varying pH were investigated by qPCR of Group 1.1c, crenarchaeal and bacterial 16S
rRNA genes. The highest abundance of Group 1.1c was
1.3 107 copies g1 dry soil in a Welsh podzol of pH 4.3
(Fig. 4; Table 1). High abundances of Group 1.1c were also
observed in acidic moorland, coniferous and deciduous
forest soils in the Cabrach area (north-east Scotland) (Fig.
4; Table 1, sites 1, 2, 3, 5 and 6). Group 1.1c at these sites
accounted for between 36% and 69% of total Crenarchaeota
and the 16S rRNA gene copy numbers ranged from 8.8 105
to 9.3 106 copies g1 dry soil. A grassland soil (site 9),
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372
L.E. Lehtovirta et al.
Fig. 3. Phylogenetic analysis of cloned 16S rRNA
gene sequences (highlighted in bold) of Group
1.1c Crenarchaeota with DGGE migration
positions I–V (see Fig. 2). Sequence names are also
prefixed with the replicate number and pH of the
soil from which they were derived. CS indicates
that the sequence was obtained from Craibstone
soil. Reference sequences are presented as ‘name
(accession number) origin’. Multifurcating
branches indicate where the relative branching
order could not be resolved in the majority of
bootstrap replicates. Values at nodes represent
the lowest percentage value of distance,
parsimony and maximum likelihood bootstrap
analyses (1000, 1000 and 100 bootstrap
replicates, respectively). LogDet/Paralinear
pairwise distances of unambiguously aligned
positions were calculated using variable sites only.
The tree was rooted using Groups 1.1a and 1.1b
16S rRNA gene sequences.
Discussion
Effect of pH on the microbial community size
and structure
Real-time PCR quantification of 16S rRNA genes demonstrated little variation and no apparent trends in total
crenarchaeal abundance with soil pH but greater abundance
of Group 1.1c Crenarchaeota in low pH soils. This is
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1011
16S rRNA gene abundance
(copies g–1 dry soil)
which was located in the vicinity of these sites and possessed
the same parent material, had a higher pH and low
abundance of Group 1.1c, which comprised only 0.9% of
the total crenarchaeal community at this site. Although
Crenarchaeota were detected in all soils investigated, Group
1.1c were below detection at several sites of near-neutral pH
(Fig. 3). Abundances of total Crenarchaeota and Bacteria
varied between soils by up to two orders of magnitude.
1010
109
108
107
106
105
104
3.9 3.9 4.2 4.3 4.3 4.4 5.0 5.1 5.7 5.8 5.8 6.5 6.6
(1) (2) (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13)
1.1c Crenarchaeota
Crenarchaeota
Bacteria
pH (site)
Fig. 4. Quantification of 16S rRNA gene abundance of Group 1.1c
Crenarchaeota, total Group 1 Crenarchaeota and total Bacteria in a
range of soils with contrasting habitat and soil properties. Error bars
represent SEs of means of three biological replicates. Soil characteristics
are described in Table 1.
FEMS Microbiol Ecol 70 (2009) 367–376
373
Archaea in acid soil
consistent with results of earlier ecological soil surveys, in
which Group 1.1c Crenarchaeota have been found in the
most acidic soil sites (Nicol et al., 2005; Oline et al., 2006;
Bomberg & Timonen, 2007). Soil pH has previously been
described as a major factor in determining microbial community structure and abundance (Fierer & Jackson, 2006)
and Group 1.1c appear to support this view. Nicol et al.
(2008) demonstrated a pH-related decrease in crenarchaeal
amoA gene abundance, in the soil pH plots investigated in
this study, which corresponded to an increase in soil pH and
a simultaneous structural change in crenarchaeal community (Nicol et al., 2008). Whereas their observations could be
explained in terms of established mechanisms for inhibition
of ammonia oxidation at low pH, nothing is known of
Group 1.1c function, and associated amoA genes have not
been identified. Comparative abundances of archaeal amoA
genes from the study by Nicol et al. (2008) and 16S rRNA
genes presented here are similar, but differences do exist in
the overall trends, with crenarchaeal amoA abundance
decreasing with increasing pH while 16S rRNA gene copies
remain relatively constant. This may indicate that other
Crenarchaeota are present, particularly in high pH soils that
do not possess ammonia monooxygenase genes or may
indicate issues with primer specificity or bias. Our data
agree with previous estimates of crenarchaeal 16S rRNA
gene abundances (around 107 copies g1 dry soil) and ratios
of Crenarchaeota to Bacteria in soil environments (around
1%) (Buckley et al., 1998; Ochsenreiter et al., 2003). The
lower abundance of Group 1.1c Crenarchaeota at neutral pH
was not associated with a general reduction in prokaryote
abundance, but with changes in relative abundance.
Even at low pH, Group 1.1c Crenarchaeota accounted for
only a modest proportion of total Crenarchaeota. This is
perhaps unsurprising as this study was performed on an
arable soil, and Group 1.1c populations have been shown to
be a minor component of total archaeal communities in
arable and pasture soils (Nicol et al., 2003, 2008). The field
site was selected due to the long-term maintenance of a pH
gradient with other factors (e.g. vegetation) remaining
constant and previous detection of Group 1.1c (Nicol et al.,
2008). Comparison of a range of soils from other sites
revealed higher abundance of Group 1.1c at acidic forest
sites and in moorland, where they constituted a significant
component or even the majority of crenarchaeal community. They were also absent or below detection in soils near
neutral pH (pH 4 5.8), which agrees with the qPCR data
from the pH-adjusted plots, where Group 1.1c was below
detection at pH 4 6.5.
At lower soil pH values in the Craibstone plots, the
biological replicates were consistent and trends in relative
band intensities in DGGE profiles suggest adaptation of
different phylotypes within Group 1.1c to specific pH
ranges. The decrease in the reproducibility between repliFEMS Microbiol Ecol 70 (2009) 367–376
cates from the higher pH soils is probably indicative of low
template copy numbers, which leads to stochastic amplification (Ou et al., 2005). Group 1.1c 16S rRNA gene abundance
was below the detection limit of 2.5 104 copies g1 dry soil
by qPCR (which corresponds to 10 copies per qPCR reaction). It is likely that PCR products were only detected in
DGGE analysis because all the samples were subjected to
nested amplification (i.e. another 35 cycles of PCR).
Phylogenetic analysis
DGGE bands with different migration behaviour fell into
distinct monophyletic groups, which were consistent for
comigrating clones from different clone libraries, and relationships were therefore seen between phylogenetic groups
and soil pH. Thus, clones corresponding to DGGE positions
I and V (Figs 2 and 3) were derived from both pH 4.5 and
6.5 soils, while other DGGE band positions contained clones
from one pH only. Clones associated with DGGE positions
III and V may represent phylotypes adapted to soils of
higher pH, although they are polyphyletic. The most closely
related database sequences originate from a moorland–
coniferous forest transect and developed alpine soils, which
have pH lower than any studied here (Nicol et al., 2005,
2007), suggesting that these Crenarchaeota may be able to
inhabit soils over a wide acidic pH range.
Potential biases and methodological limitations
The abundance of different microbial groups was estimated
by the qPCR analysis of 16S rRNA genes. A recognized
limitation of this approach is variable 16S rRNA gene copy
number per genome, which in bacteria ranges from 1 to 13
depending on ecological strategy (Klappenbach et al., 2000).
While Euryarchaeota may possess up to four copies of 16S
rRNA genes per genome, thermophilic Crenarchaeota only
have one copy (Lee et al., 2009). The genomes of
N. maritimus and Cenarchaeum symbiosum, which are
currently the only available nonthermophilic crenarchaeal
genomes, each have only one ribosomal operon. This
suggests that crenarchaeal 16S rRNA gene copy numbers
measured by qPCR may be representative of cell numbers in
the studied soils. In addition, primer biases cannot be
entirely ruled out. In a mixed template population (e.g.
environmental DNA), the choice of primers can introduce a
stronger bias than the template (Suzuki & Giovannoni,
1996). The efficiency of nucleic acid extraction and reproducibility of standard curve reconstruction may affect the
outcome of qPCR. Because the same extraction method was
used for all samples, and biological replicates were similar, it
is unlikely that significant bias resulted from the nucleic acid
extraction procedure. Also amplification efficiencies, linear
regression coefficients, melting curves and reproducibility of
all qPCR assays were optimal.
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Published by Blackwell Publishing Ltd. All rights reserved
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374
L.E. Lehtovirta et al.
Potential function of Group 1.1c Crenarchaeota
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Acknowledgements
L.E.L. is funded on a NERC Postgraduate Studentship and
G.W.N. by an NERC Advanced Fellowship (NE/D010195/1).
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