sion and microdiversity of soil CrenarchaeaG. W. Nicol
et al.
Environmental Microbiology (2006) 8(8), 1382–1393
doi:10.1111/j.1462-2920.2006.01031.x
Crenarchaeal community assembly and microdiversity
in developing soils at two sites associated with
deglaciation
Graeme W. Nicol,1* Dagmar Tscherko,2 Lisa Chang,1
Ute Hammesfahr2 and James I. Prosser1
1
School of Biological Sciences, University of Aberdeen,
Cruickshank Building, St. Machar Drive, Aberdeen,
AB24 3UU, UK.
2
Institute of Soil Science and Land Evaluation, University
of Hohenheim, Emil-Wolff-Straße 27, 70599 Stuttgart,
Germany.
Summary
Non-thermophilic Crenarchaeota are recognized as
ubiquitous and abundant components of soil microbial communities. Previous studies of the foreland of
the receding Rotmoosferner glacier in the Austrian
Central Alps have demonstrated that crenarchaeal
communities in soil at different stages of development are distinct from each other, with Group 1.1b
crenarchaeal populations dominating throughout the
successional gradient, while Group 1.1c crenarchaea
are present in mature soils only. To determine whether
this highly structured succession was unique to the
Rotmoosferner glacier foreland, 1.1b and 1.1c communities were compared with those present along a
successional gradient at Ödenwinkelkees glacier,
125 km away, by denaturing gradient gel electrophoresis (DGGE) of 16S rRNA reverse transcription
polymerase chain reaction products. Similarities in
community structure were observed; 1.1b communities were present throughout both successional gradients (though lacking the defined structure at
Ödenwinkelkees) and 1.1c communities were present
only in mature soil. Comigration of bands on DGGE
gels indicated that a number of similar crenarchaeal
populations were present at both sites. To compare
populations, and examine microscale diversity, 16S
rRNA genes and complete 16S-23S internal transcribed spacer (ITS) regions representing six major
band positions in DGGE analysis were amplified,
cloned and sequenced and represented four 1.1b and
Received 15 November, 2005; accepted 9 March, 2006. *For
correspondence. E-mail [email protected]; Tel. (+44) 1224
272700; Fax (+44) 1224 272703.
two 1.1c lineages. The data provide no evidence of
endemism, but large differences in the rate of
sequence divergence in the ITS region (relative to that
in 16S rRNA genes) were observed. Two of the 1.1b
lineages (each possessing > 98% 16S rRNA gene similarity) had relatively long and highly divergent ITS
sequences. In contrast, two other 1.1b and two 1.1c
lineages (each possessing > 99% 16S rRNA gene similarity) exhibited markedly less variation in their
respective 16S-23S ITS regions. The results reveal
common patterns in the ecology and assembly of
crenarchaeal communities in spatially separated soil
systems and may indicate different evolutionary rates
between soil crenarchaea lineages.
Introduction
Alpine glaciers have been receding since the end of the
‘little ice-age’ in the mid-19th century and present successional gradients of soil and plant community development
over relatively short distances (Tscherko et al., 2003).
They therefore represent suitable environments for studying interactions between ecosystem development and
selection and assembly of complex soil microbial communities, with soil development resulting in successional
assemblages increasing in cell numbers, biomass and
functional diversity (Ohtonen et al., 1999; Sigler and
Zeyer, 2002; Tscherko et al., 2003). Initial microbial colonizers of exposed substrate may contain an inactive component of allochthonous organisms (Jumpponen, 2003),
as well as a greater proportion of fast-growing, opportunistic organisms that are more tolerant to environmental
stress (Sigler and Zeyer, 2004). Recently, Nicol and colleagues (2005) demonstrated a clear succession of
archaeal communities across the forefield of the receding
Rotmoosferner glacier in Austria. Exposed soil substrates
contained communities of soil archaea that were distinct
and presumably adapted to the harsh conditions. These
organisms were then replaced by closely related, intermediate communities before the community structure developed into that similar to mature alpine grassland. These
results indicated that archaeal community succession
(and perhaps microbial community assembly) may proceed in a predictable manner, with communities adapted
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd
Succession and microdiversity of soil Crenarchaea 1383
to different stages of soil development. However, contrasting patterns of replacement and succession of bacterial
communities have been reported across forelands of different glaciers, which may be attributed to differences in
local site characteristics (Sigler and Zeyer, 2002).
Archaea constitute a significant proportion of prokaryotic numbers in mesophilic environments. In particular, a
lineage of crenarchaea termed ‘Group 1’, which forms a
specific but deeply branching association with cultivated
thermophilic crenarchaea, appears to be the most abundant and ecologically diverse (DeLong, 1998). For example, it has been estimated that these organisms represent
approximately 20% of planktonic marine prokaryotes
(Karner et al., 2001) and ≥1% in soil systems (Sandaa
et al., 1999; Ochsenreiter et al., 2003). Archaeal 16S
rRNA gene surveys reveal domination of archaeal communities in many soil systems by mesophilic Crenarchaeota (e.g. Bintrim et al., 1997; Jurgens et al., 1997; Nicol
et al., 2003a; Ochsenreiter et al., 2003), which are more
abundant than euryarchaeal populations in grassland
soils (Nicol et al., 2003b; 2005). Representative
sequences of most major Group 1 lineages have been
recovered from soil samples, but those belonging to the
1.1b lineage appear to be most abundant and ubiquitous,
dominating most soil systems (Ochsenreiter et al., 2003).
Sequences within the Group 1.1c lineage are also found
mainly in soil systems, but have a more restricted distribution than the 1.1b lineage (Nicol et al., 2005). Most of
these sequences have been recovered from coniferous
forest soils (e.g. Jurgens et al., 1997; Bomberg et al.,
2003; Yrjälä et al., 2004) and grassland soils (together
with dominating 1.1b sequences) (e.g. Nicol et al., 2003a;
Nicol et al., 2005). These two groups are substantially
divergent with the more abundant 1.1b group more closely
related to the 1.1a crenarchaea (which includes mesophilic marine crenarchaea) (Nicol et al., 2005). Little is
known of the physiological characteristics of non-thermophilic crenarchaea or their contribution to ecosystem functions. However, recent studies indicate a role in global
nitrogen cycling (Venter et al., 2004; Francis et al., 2005;
Schleper et al., 2005; Treusch et al., 2005) and a mesophilic crenarchaeal ammonia oxidizer (Group 1.1a lineage) has been isolated (Könneke et al., 2005).
The internal transcribed spacer (ITS) region between
16S and 23S ribosomal genes in the rrn operon is often
used to differentiate closely related prokaryotes. These
regions vary much more in size and sequence than associated rRNA genes (Ranjard et al., 2000) and are therefore useful in examining microdiversity (Brown et al.,
2005). García-Martínez and Rodríguez-Valera (2000)
used 16S rRNA gene and ITS sequences to study
microdiversity of marine Group 1.1a, but this approach
has not been used to characterize microdiversity in soil
crenarchaea, and its value in determining phylogeny and
microdiversity of Group 1.1b and 1.1c sequences is not
known. The aims of this study therefore were to: (i) determine whether previously observed patterns of archaeal
community succession were typical of developing soils; (ii)
correlate archaeal community structures with previously
determined soil characteristics; and (iii) assess microdiversity and the potential for endemic populations of crenarchaea in spatially separated soil systems.
Results
Sampling site
Samples were obtained from the forefields of the Rotmoosferner and Ödenwinkelkees glaciers in the Austrian
Central Alps, which have been receding since the mid19th century because of increases in atmospheric temperature. Both Rotmoosferner and Ödenwinkelkees
glacier forelands run in a south-to-north direction and
have receded 2 km and 1.5 km, respectively, from their
maximum extension (Tscherko et al., 2003). Samples
were taken across each forefield (i.e. between the glacial
snout and the terminal moraine) and represented a range
of soil ages from recently deglaciated samples (< 10
years) to those exposed for c. 150 years. In addition,
references samples were taken outside glacier foreland
that had been ice-free for c. 9500 years. At both sites,
trends associated with soil and ecosystem development
were observed with increases in nitrogen, carbon and
organic matter and changes in the dominant plant species
composition (Tscherko et al., 2003; 2005).
DGGE analysis of crenarchaeal communities across two
successional gradients
Nicol and colleagues (2005) showed dominance of
archaeal communities by Group 1.1b and 1.1c sequence
types in developing acidic grassland soils across the forefield of the Rotmoosferner glacier. To compare these communities at both glacial sites, separate 1.1b and 1.1c
denaturing gradient gel electrophoresis (DGGE) assays
were used. Initial reverse transcription polymerase chain
reaction (RT-PCR) amplification was performed using specific 1.1b and 1.1c forward primers (G1.1b280f and
FFS200f respectively), with an archaea-specific reverse
primer (Ar9R). Polymerase chain reaction products were
then nested with an archaeal-specific DGGE primer set
(rSAf/PARCH519r) (Table 1). Due to necessary degeneracies, this primer set generates a ‘doublet’ of two closely
positioned bands for each 16S rRNA gene sequence
amplified. However, 1.1c PCR products always migrate to
lower positions than 1.1b products, because of the higher
GC-content. A marker lane containing nested PCR amplicons from clones representative of dominant DGGE band
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
1384 G. W. Nicol et al.
Table 1. Details of PCR primers used in this study.
Primer (5′-3′)
rRNA gene
Position
Target
Use
Reference
A109f
G1.1b280f
FFS200f (Probe FFS-Uni)
rSAf
PARCH519r
PARCH533fb
Ar9R
Ar9Fb
1492r
1514fb
A51r
16S
16S
16S
16S
16S
16S
16S
16S
16S
16S
23S
109–125
265–280
180–200
341–357
519–533
519–533
906–927
906–927
1492–1513
1492–1513
51–71
Archaea
Group 1.1ba
Group 1.1c
Archaea
Archaea
Archaea
Archaea
Archaea
Universal
Universal
Archaea
PCR/sequencing
PCR
PCR/sequencing
PCR
PCR
Sequencing
PCR/sequencing
Sequencing
Sequencing
Sequencing
PCR/sequencing
Großkopf et al. (1998)
This study
Jurgens and Saano (1999)
Nicol et al. (2005)
Øvreås et al. (1997)
Øvreås et al. (1997)
Jurgens et al. (1997)
Jurgens et al. (1997)
Lane et al. (1991)
Lane et al. (1991)
García-Martínez and Rodríguez-Valera (2000)
a. This primer was designed to discriminate 1.1b sequences from 1.1c only, and may not discriminate in the presence of other crenarchaeal
groups.
b. Primer sequence is reverse complement of a primer from referenced study.
positions (Nicol et al., 2005) in archaeal community profiles at Rotmoosferner was also run alongside all DGGE
profiles. Group 1.1b RT-PCR products were produced
from archaeal cDNA from all soil samples at both sites
(deglaciated for 4–9500 years) and four different band
positions were identified in DGGE profiles (A–D, Fig. 1).
Two (A and B) were common to both sites and one was
unique to either Ödenwinkelkees (C) or Rotmoosferner
(D) soils, but with some evidence of band C in the latter.
At Rotmoosferner (Fig. 1A), band positions A, B and D
were indicative of pioneering, intermediate and mature
archaeal communities respectively. As expected, they
comigrated with the three 1.1b band positions in the
Rotmoosferner-derived marker lane, confirming archaeal
community development previously identified at Rotmoos-
ferner (Nicol et al., 2005). In contrast, Ödenwinkelkees
1.1b community structures lacked the same clear successional sequence (Fig. 1B). Although some variation was
apparent, community structure was more uniform across
the gradient with no clear succession of populations represented by band positions A and B. In the Ödenwinkelkees profiles, there was a small increase in the relative
(within lane) intensity of band C in soil 50 years or older,
indicating that it may be analogous to organisms represented by band D in Rotmoosferner profiles.
The distribution of 1.1c crenarchaea was similar at both
sites with PCR products only detected in soils considered
to be developed, i.e. deglaciated for 135 or 150 years to
9500 years and with soil characteristics typical of alpine
grassland soil (Tscherko et al., 2003). All amplicons
A - Rotmoosferner
M
4/14
20
48
75
135
9500
M
A
B
A
B
C
C
D
D
B - Ödenwinkelkees
M
10
20
50
100
150
9500 M
A
B
A
B
C
C
D
D
Fig. 1. Denaturing gradient gel electrophoresis
analysis of crenarchaeal Group 1.1b communities in soil substrates across successional chronosequences in front of the receding
Rotmoosferner (A) and Ödenwinkelkees (B)
glaciers. Comparison of both profiles revealed
four different dominant banding positions,
labelled A–D in order of migration. Bands in the
marker lane (lane M) comigrating with positions
A, B and D, are representative of pioneering,
intermediate and mature 1.1b sequence
sequences, respectively, previously retrieved
from the Rotmoosferner glacier forefront
[described as positions GFS2, 3 and 4 by Nicol
and colleagues (2005)]. Band positions highlighted with an arrow indicate that a 16S rRNAITS clone sequence was obtained with an identical DGGE migration pattern (after nested
amplification with primers rSAf/PARCH519r)
from the same nucleic acid extract profiled in
that lane.
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
Succession and microdiversity of soil Crenarchaea 1385
A - Rotmoosferner
M
135
B - Ödenwinkelkees
9500
1.1b
150
9500
A
B
M
1.1b
D
E
1.1c
F
migrated to the lower portion of DGGE gels, within the
range of 1.1c marker sequences (Fig. 2). The lack of
amplicons in the region of (dominant) 1.1b sequences
highlights the specificity of this assay. The 1.1c community
was similar in 135- and 9500-year Rotmoosferner
samples (Fig. 2A) but differed in 150- and 9500-year
Ödenwinkelkees samples (Fig. 2B). The 1.1c community
structures at the two sites differed but profiles contained
several comigrating bands indicating the presence of very
closely related sequences.
PCR amplification of 16S-ITS-23S
Comigrating DGGE bands produced from the primer set
used in this study are consistently identical in sequence
(Nicol et al., 2003a,b; 2005), but amplified fragments are
not of sufficient length to provide substantial phylogenetic
resolution. Therefore, nearly full length 16S rRNA genes,
with complete 16S-23S ITS regions, representative of six
selected band positions in DGGE profiles (Figs 1 and 2)
were amplified, cloned and sequenced. Four of the
selected band positions were present at both sites; two
1.1b (A and B) and two 1.1c (E and F). 16S rRNA geneITS sequences, representing one of six banding positions
(Figs 1 and 2) were obtained by screening individual
clones using DGGE. A minimum of six clones were
selected for each position, with at least three clones from
both glaciers selected for comparison of a common band
position (A, B, E or F). As previous analysis indicated that
communities were dominated by 1.1b sequences, clone
libraries for 1.1b and 1.1c sequences were generated
using forward primers A109f and FFS200f, respectively,
Fig. 2. Denaturing gradient gel electrophoresis
analysis of crenarchaeal Group 1.1c communities in mature soil substrates (deglaciated for
135/150 and 9500 years) in front of Rotmoosferner (A) and Ödenwinkelkees (B) glaciers.
Bands in the marker lane (lane M) represent 10
crenarchaeal clones including representatives
of 1.1b and 1.1c sequences (Nicol et al., 2005).
Band positions labelled E and F were present
in profiles from both locations and were
selected for subsequent phylogenetic analysis
of 16S rRNA gene and ITS sequences.
Sequence positions described as A, B and D in
Fig. 1 are also highlighted. Band positions highlighted with an arrow indicate that a 16S rRNAITS cloned sequence was obtained with an
identical DGGE migration pattern (after nested
amplification with primers rSAf/PARCH519r)
from the same nucleic acid extract profiled in
that lane.
1.1c
with archaeal 23S rRNA reverse primer A51r (Table 1).
After screening and selecting clones by DGGE migration
pattern, all ITS regions were sequenced completely and
16S rRNA regions were sequenced for two clones from
each site (four in total for each migration position).
16S rRNA gene sequence analysis
All 16S rRNA gene sequences (≥ 1299 nucleotides) were
compared against the GenBank database using BLAST
searches and those showing highest similarity were
aligned with other representative 1.1b and 1.1c
sequences. Clones grouped by DGGE migration pattern
(A–F) were identical over the 150-nucleotide region amplified for DGGE analysis. Each group possessed 98.3–
99.8% 16S rRNA gene sequence similarity (Table 2) and
formed well-supported monophyletic groups in phylogenetic analysis (Figs 3A and 4A) (hereby referred to as
‘clusters A–F’). Cluster A and B sequences formed distinct
but closely related lineages, with each group possessing
99.8% and 98.3% identity, respectively, and together sharing 97.7% identity. Interestingly, cluster B sequences were
identical to the amplified 16S rRNA gene fragment on a
crenarchaeal fosmid clone (29i4) isolated from soil
(Quaiser et al., 2002) over the region amplified for DGGE.
This fragment also possesses the full-length associated
ITS sequence and was included in comparisons of 16S
rRNA gene and ITS sequences. Cluster C and D
sequences (from Ödenwinkelkees and Rotmoosferner
respectively) formed well-supported but distinct groups
within the 1.1b lineage, both groups sharing only 93.9%
identity. Cloned sequences with position E and F migra-
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
1386 G. W. Nicol et al.
Fig. 3. Distance analyses of cloned 16S rRNA gene and ITS sequences from organisms placed within the 1.1b crenarchaeal lineage. Bootstrap
support was calculated 1000 times and multifurcating branches indicate that the relative branching order could not be resolved in the majority of
bootstrap replicates. Scale bars indicate 0.01 substitutions per site. Clones are described by location (Rot, Rotmoosferner; Oden, Ödenwinkelkees), DGGE migration position (A–D), length of exposure of the soil substrate (75/100/135/9500 years) and replicate number (i, ii and iii). All
outgroup sequences were pruned for presentation.
A. Phylogenetic tree of 16S rRNA gene sequences with DGGE migration patterns A–D. Sequences highlighted with A–D in superscript denote
sequences from other studies that are identical, over the 150 bp fragment amplified for DGGE analysis, to those of Rot/Oden sequences A–D
respectively. The tree was rooted with the 16S rRNA gene sequence of Cenarchaeum symbiosum (Group 1.1a).
B. Phylogenetic tree of ITS sequences of clones with 16S rRNA gene sequences migrating to position ‘C’ in DGGE analysis. Clones with a fully
sequenced 16S rRNA gene portion (Fig. 3A) are underlined. The tree was rooted using cluster D ITS sequences.
C. Phylogenetic tree of ITS sequences of clones with 16S rRNA gene sequences migrating to position ‘D’ in DGGE analysis. Clones with a fully
sequenced 16S rRNA gene portion (Fig. 3A) are underlined. The tree was rooted using cluster C ITS sequences.
Table 2. Variability of 16S rRNA gene and ITS sequences within the six monophyletic crenarchaeal lineages A–F.
16S rRNA gene sequence
ITS sequence
Sequence group
Lengtha,b
% Identityc
Length rangeb
% Identityd
A
B
C
D
E
F
1375
1375
1376
1375
1300
1299
99.8
98.3
99.6
99.2
99.0
99.7
741–923 (6)
644–927 (7)
251–255 (6)
240 (6)
203 or 206 (12)
220–230 (12)
55.0
18.4
92.0
97.0
96.0
85.0
(4)
(5)
(4)
(4)
(4)
(4)
(4.5)
(2.9)
(2.0)
(0)
(0.7)
(1.1)
a. Length from first nucleotide after primer site A109f (A–D) or FFS200f (E–F) to the end of the 16S rRNA gene. All 16S rRNA gene sequences
within each cluster were identical in length.
b. Numbers of sequences compared are given in parentheses.
c. Ratio of conserved nucleotide positions in all Ödenwinkelkees and Rotmoosferner 16S rRNA gene sequences to the total number of nucleotide
positions within each cluster.
d. Mean (and standard deviation) of the number of conserved nucleotide positions in all sequences compared with the length of each sequence.
tion patterns separated into distinct and well-supported
clades within the 1.1c lineage, as expected.
ITS variability and phylogenetic analysis
Internal transcribed spacer sequences of the six
sequence clusters varied substantially in length and
sequence (Table 2). All were screened using tRNAscanSE 1.21 (Lowe and Eddy, 1997) and no tRNA genes were
detected. Cluster A and B ITS sequences (which together
formed a distinct monophyletic lineage in 16S rRNA gene
analysis) were much longer and length and sequence
varied more than C–F. The high variability in length and
sequence of B clone sequences prevented alignment of
homologous positions representing a significant proportion of the ITS sequences (101 aligned positions from ITS
sequences, 644–927 nucleotides in length). Phylogenetic
trees using this relatively short alignment were considered
uninformative, producing topologies clearly inconsistent
with the level of similarity between some sequences
(based on percentage similarity alone) (Table 3B). Cluster
A sequences also showed very high variability. No out-
Table 3. Percentage similarity of ITS sequences of cluster A and B clones.
A
RotA-75iiba
RotA-75iic
OdenA-100iia
OdenA-100iiiaa
OdenA-100iiib
RotA-75iiaa
55.0
66.6
72.6
84.6
74.4
RotA-75iiba
RotA-75iic
OdenA-100iia
OdenA-100iiiaa
62.7
59.1
57.5
62.1
59.8
71.2
62.4
81.0
93.5
83.4
B
RotB-75iib
RotB-135iia
OdenB-100iiaa
OdenB-100iiba
OdenB-100iic
29i4a,b
RotB-75iiaa
84.3
46.1
29.7
48.0
30.0
46.8
RotB-75iib
RotB-135iia
OdenB-100iiaa
OdenB-100iiba
OdenB-100iic
46.1
29.4
48.4
29.7
45.3
23.7
81.4
23.8
36.1
24.6
98.2
23.6
24.7
35.8
23.6
a. Corresponding 16S rRNA gene sequence represented in Fig. 3A.
b. The ITS sequence of crenarchaeal genomic fragment 29i4 (Quaiser et al., 2002) was included in this analysis as the corresponding 16S rRNA
gene sequence is identical over the region amplified by DGGE analysis with Rot/Oden cluster B sequence.
Percentage values between each pair of sequences were calculated from the ratio of identities to the length of the longer of the two sequences.
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
Succession and microdiversity of soil Crenarchaea 1387
RotA-75iiia
OdenA-100iiia
OdenA-100ii
51
RotA-75iiib
A
GFS2-20i (AY601287) A
97
SCA1145 (U62811) A
TRC23-38 ( AF227644) A
GFS2-48iii ( AY601288 )A
100
76
RotB-75iiia
OdenB-100iia
GFS3-75ii (AY601289) B
29i4 (AJ496176) B
OdenC-100iia
RotB-135ii
B
OdenB-100iib
67
OdenC-100iib
GFS3-75iib (AY601290 )B
100
OdenC-9500iia
GFS3-135i (AY601291) B
80
GRU16 (AY278083) B
71
OdenC-9500iib
S248-6 (AY278099 ) B
OdenC-100iia
100
OdenC-9500iia
OdenC-100iic
C
OdenC-9500iib
0.01
ROB1A9 (AY278074 )C
84
62
OdenC-9500iic
OdenC-100iib
B ITS phylogeny of C sequence group
TRC23-10 (AF227640) C
SCA1154 (U62814)
100
100
97
54d9 (AJ627422)
ROB1A9 (AY278074 )
RotD-9500ia
66
68
RotD-9500ia
RotD-9500iiib
GFS4-135i (AY601293)D
D
94
99
RotD-9500ib
D
RotD-9500iiia
RotD-9500iiia
62
RotD-9500ib
Gitt-GR-78 (AJ535123 )D
RotD-9500ic
100
SCA1173 (U62818)
SCA1151 (U62813)
RotD-9500iiic
TRC132-9 (AF227639)
0.01
RotD-9500iiib
0.01
A 16S rRNA gene phylogeny for A-D sequence groups
group sequence with a reasonable level of homology was
available to root a phylogenetic tree and percentage
similarity values are presented in Table 3A. In contrast,
although all cluster C and D 16S rRNA gene sequences
together share less than 94% sequence identity, the cor-
C ITS phylogeny of D sequence group
responding ITS sequences for these two groups together
exhibit greater sequence similarity (57.6% ± 1.6% mean
identity) than within either cluster A or B ITS sequences.
Comparison of phylogenies for cluster C sequences
(Fig. 3A and B) indicated that ITS sequences (92% iden-
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
1388 G. W. Nicol et al.
tity) had slightly greater resolution, with the relative
branching order of sequences determined in the majority
of bootstrap replicates with ITS sequences only. This was
also observed when only ITS sequences with corresponding 16S rRNA gene sequences were analysed without
the additional ITS sequences shown in Fig. 3B. ITS
sequences of cluster D shared > 97% similarity and did
not contain greater discriminatory information than corresponding 16S rRNA gene sequences (Fig. 3A and C).
For the two 1.1c sequence groups, cluster F ITS
sequences showed greater variation than cluster E
sequences (85.0% versus 96.0%) but comparison of 16S
rRNA and ITS phylogenies indicates that phylogenetic
resolution within ITS sequences was not substantially
greater. Within cluster E, most ITS and 16S rRNA gene
sequences showed multifurcation from one node (Fig. 4A
and C). Within cluster F, ITS sequences did not show
greater resolution than corresponding 16S rRNA gene
sequences (with regard to relative branching of
sequences) although greater separation is apparent within
ITS sequences (Fig. 4A and B). This was also observed
when only ITS sequences with corresponding 16S rRNA
gene sequences were analysed without the additional
ITS sequences shown in Fig. 4B. For all clusters, ITS
RotF-135iia
OdenF-150iia
67
OdenF-150iic
OdenF-150iiib
RotF-135iic
RotF-9500iiia
78
RotF-9500iiic
72
100
RotF-9500iiib
OdenF-150iiia
99
100
FFSB1 (X96688)
GFS8-9500i (AY601302)
GFS9-9500iii (AY601304)
FFSB10 (X96695)
99
FFSB11 (X96696)
100
FFSB4 (X96691)
FFSB7 (X96694)
52
GFS7-9500iii (AY601300)
OdenF-150iia
69
RotF-9500iiia
66
72
0.01
0.01
RotF-135iib
B ITS phylogeny of F sequence group
OdenE-150iiia
OdenE-150iia
RotE-135iic
76
RotE-9500iiib
RotE-9500iiic
OdenF-150iiia
OdenE-150iib
RotE-9500iiia
OdenE-150iiia
RotE-135iia
RotE-9500iiia
F
RotE-135iia
100
OdenF-150iiic
RotE-135iib
RotF-135iia
68
93
OdenF-150iib
FHMa5 (AJ428027)
100
94
61
62
E
OdenE-150iia
A 16S rRNA gene phylogeny for E and F sequence groups
100
OdenE-150iiib
OdenE-150iiic
OdenE-150iic
0.01
C ITS phylogeny of E sequence group
Fig. 4. Distance analyses of cloned 16S rRNA gene and ITS sequences from organisms placed in two clusters within the 1.1c crenarchaeal
lineage. Tree construction and clone designation are as described in Fig. 3. All outgroup sequences were pruned for presentation.
A. Phylogenetic tree of 16S rRNA gene sequences with DGGE migration patterns E and F. The tree was rooted using sequences FFSB6 and
FFSA1 that are placed within a 1.1c-associated lineage distinct from 1.1c sequences (Nicol et al., 2005).
B. Phylogenetic tree of ITS sequences of clones with 16S rRNA gene sequences migrating to position ‘F’ in DGGE analysis. Clones with a fully
sequenced 16S rRNA gene portion (Fig. 4A) are underlined. The tree was rooted using cluster E ITS sequences.
C. Phylogenetic tree of ITS sequences of clones with 16S rRNA gene sequences migrating to position ‘E’ in DGGE analysis. Clones with a fully
sequenced 16S rRNA gene portion (Fig. 4A) are underlined. The tree was rooted using cluster F ITS sequences.
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
Succession and microdiversity of soil Crenarchaea 1389
sequences from one location or individual sample did not
necessarily group or show greater similarity to each other
than sequences from other locations or samples.
Discussion
Comparison of crenarchaeal succession using DGGE
analysis
Several studies have revealed associations between
some crenarchaeal populations and plant rhizospheres
(e.g. Simon et al., 2000; Sliwinski and Goodman, 2004).
Bulk soil and rhizosphere crenarchaeal assemblages are
distinct, but individual plant species do not appear to
select for specific crenarchaeal assemblages (Sliwinski
and Goodman, 2004) or to be the major drivers for crenarchaeal communities in successional alpine soils (Nicol
et al., 2005). However, successional changes in crenarchaeal community structure and the contrast in the successional characteristics at both sites correlate with
previously measured soil properties (Tscherko et al.,
2003; 2005). Both chronosequences exhibit typical
changes in characteristics associated with soil development, such as a decrease in pH and increases in total
nitrogen and organic carbon (Tscherko et al., 2003;
2005). However, gradients of carbon and nitrogen were
greater over the Rotmoosferner foreland. Also, discriminant analysis of measured microbial processes (nitrogen
mineralization, ammonium oxidation, arginine deaminase)
and enzyme activities (urease, protease, xylanase, phosphatase, arylsulfatase) revealed clear functional differences, with Rotmoosferner soils grouping into four distinct
successional stages whereas Ödenwinkelkees chronosequences showed little change in function (Tscherko et al.,
2003). These data correlate with changes in 1.1b community across each successional stage with distinct pioneering, intermediate and mature community structures
identified across Rotmoosferner, but only small variation
across Ödenwinkelkees and no distinct succession
stages. If soil crenarchaea (particularly, Group 1.1b crenarchaea) are involved in ammonia oxidation, as recent
evidence suggests (Treusch et al., 2005), it is interesting
to note that changes in ammonium concentrations correlated with changes in crenarchaeal community structure.
In the Rotmoosferner successional soils pioneering (cluster A), intermediate (cluster B) and mature (cluster D) 1.1b
crenarchaeal sequences coincided with soil ammonium
concentrations of 0.1–0.9, 7.9–9.9 and 9.9–18.3 µg NH4+N g−1 respectively. In contrast, ammonium concentrations
in Ödenwinkelkees samples varied between 0.9 and
3.8 µg NH4+-N g−1 across the entire successional
sequence (Tscherko et al., 2003; 2005). These data may
provide evidence for distinct evolutionary lineages reflecting ecological differentiation of crenarchaeal populations
in ammonia oxidation processes.
Distribution of 1.1c sequences was more restricted
than that of 1.1b sequences and, at both sites, the
former were only detected with specific primers in mature
soil (≥ 135 years). This correlates with 1.1c distribution
across the Rotmoosferner glacier foreland (Nicol et al.,
2005) and provides further evidence that this lineage has
more limited ecological distribution than the 1.1b lineage,
which is found more often in archaeal 16S rRNA gene
surveys in soil.
Phylogenetic analysis of 16S rRNA genes from selected
1.1b and 1.1c sequence groups
Sequence clusters A, B, E and F were detected at both
sites and two 1.1b sequence clusters, C and D, were
unique to either site, potentially representing ecotypes
with similar functional characteristics, adapted to mature
soil conditions. Although cluster C and D sequences may
be derived from functionally similar organisms, phylogenetic analysis revealed them to be relatively distinct from
each other, within the context of the Group 1.1b lineage.
Analysis of ITS sequences and crenarchaeal
microdiversity
Intergenic sequences of clones derived from a limited
number of soil samples were analysed to evaluate their
usefulness in examining microdiversity. The relative rate
of divergence between 16S rRNA gene and ITS
sequences varied considerably between the six groups
examined in detail and, consequently, phylogenetic resolution of the 16S-23S rRNA ITS region for different soil
crenarchaea groups, relative to the 16S rRNA gene, also
varied considerably. The ITS sequences of clusters A and
B were, although relatively large, highly variable in both
length and sequence. As phylogenetic analysis of 16S
rRNA gene sequences places these two sequence groups
in a monophyletic group, large but highly variable ITS
sequences appear to be a trait of the ‘A/B’ lineage within
the 1.1b crenarchaea. Although these ITS sequences
(particularly of cluster B) were too divergent for useful
phylogenetic analysis, they were highly discriminatory and
may be useful in examining microdiversity. The inability to
construct useful ITS phylogenies, however, resulted from
relatively large divergence between sequences grouped
together and phylogenetic comparison of ITS sequences
may only be possible for groups with identical 16S rRNA
gene sequences. ITS sequences of cluster C, D, E and F
sequences varied less than clusters A and B in size and
sequence. Although phylogenetic analysis might resolve
sequences within these groups with slightly more resolution than corresponding 16S rRNA sequences, it is
arguable whether analysis of ITS sequences provides
substantially greater resolution within individual groups.
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
1390 G. W. Nicol et al.
Similar variation in the relative rate of divergence of 16S
rRNA gene and ITS sequences has been observed within
the predominantly marine Group 1.1a crenarchaea.
García-Martínez and Rodríguez-Valera (2000) analysed
48 cloned 16S-ITS sequences amplified from Mediterranean and Antarctic waters and sequences fell within one
of four 1.1a clusters in phylogenetic analysis. Sequences
in three of the clusters possessed greater variation over
the ITS region, but in one cluster (‘Crena-S1’), 16S rRNA
gene sequences varied more than corresponding ITS
regions (93.3% versus 98%), with 24 clones represented
by eight different 16S rRNA gene sequence types but only
three ITS sequence types (22 of which were identical).
Soil-derived 1.1b fosmid clones 54d9 (Treusch et al.,
2005) and 29i4 (Quaiser et al., 2002) reveal a potentially
interesting correlation between ITS length and gene density. Fosmid 29i4 fell within cluster B (Fig. 3A), with a
relatively long ITS sequence (828 bp), as for the other six
‘B’ sequences (622–920 bp). The ITS sequence of fosmid
54d9, however, was much shorter (217 bp) and grouped
with cluster C sequences in 16S rRNA gene phylogeny
(Fig. 3A), which also possess relatively short ITS
sequences (204–206 bp). The overall gene content of
these two genomic fragments differs substantially (C.
Schleper, pers. comm.), with 29i4 (having a longer ITS
sequence) and 54d9 (shorter ITS sequence) containing
69.5% and 83.2% predicted coding sequence respectively. The larger, non-coding ITS sequences may therefore reflect larger amounts of non-coding sequence at the
genome level within the 1.1b lineage.
Even though relatively few clones were analysed from
a limited number of samples, there was no evidence for
endemic populations of crenarchaeal groups, grouped
arbitrarily on the basis of an identical 150 bp 16S rRNA
gene fragment. For example, there are several examples
of some clones from each site representing one sequence
group (e.g. RotB-75iiia and OdenB-100iia) that are more
closely related than two clones from the same individual
soil sample (OdenB-100iia and OdenB-100iib). Glacial
sites were only 125 km apart and much larger distances
may be required for geographic isolation of divergent
populations.
These results reveal similarities in the successional
dynamics of soil crenarchaea communities. The broad
architecture of spatially separated soil microbial communities may develop in a predictable manner with crenarchaeal succession showing correlation with previously
measured soil characteristics and functional processes.
Experimental procedures
Study areas, chronosequences and soil sampling
Both glaciers are located in the Austrian Central Alps. The
foreland of the Rotmoosferner glacier (46°50′N, 11°03′E) is
located in the Ötz valley at an altitude of 2280–2450 m, with
an annual mean temperature of −1.3°C (1997–1998), an
annual mean precipitation of 820 mm (1970–1996) and snow
coverage from mid-October to late May (Tscherko et al.,
2003). The foreland of the Ödenwinkelkees glacier (47°07′N,
12°28′E) is approximately 125 km east of the Rotmoosferner
glacier and located near the Großglockner range at an altitude of 2068–2150 m, with an annual mean temperature of
−0.3°C, annual mean precipitation of 2397 mm (1980–1999)
and snow coverage from mid-October to late May (Tscherko
et al., 2003). Soil parent material at both sites was mainly
neoglacial moraine rubble (mica-schist and granite with local
grains of carbonate minerals) and fluvio-glacial sands, with
developing soils leptic regosols (Tscherko et al., 2003). Triplicate soil samples were analysed for each successional site.
An individual soil sample consisted of 5–10 small cores,
collected from the top 10 cm of soil within a 2 m × 2 m area,
which were subsequently sieved (< 2 mm) to remove stones
and root material and stored at −20°C. Relative soil ages had
been determined from photographic evidence and reports
from the Austrian Alpine Club (Edmaier and Jung-Hüttl, 1996;
Slupetzky, 2000).
Extraction of nucleic acids
DNA and RNA were coextracted as previously described
(Nicol et al., 2005) using a method based on that of Griffiths
and colleagues (2000). Briefly, 0.5 g of soil was placed in a
2 ml screw-cap Blue Matrix tube (Hybaid, Ashford, Middlesex, UK) with 0.5 ml of extraction buffer [120 mM potassium
phosphate buffer (pH 7.8), 5% (w/v) hexadecyltrimethylammonium bromide (CTAB), 0.35 M NaCl] and 0.5 ml of
phenol : chloroform : isoamyl alcohol (25:24:1). Cells were
lysed in a Hybaid Ribolyser (Hybaid) for 30 s at speed 4.0.
After centrifugation at 16 000 g for 5 min, the aqueous
phase was removed and extracted with 0.25 ml of
chloroform : isoamyl alcohol (24:1) followed by further centrifugation at 16 000 g for 5 min. The aqueous phase was
removed and total nucleic acids were precipitated by adding
two volumes of 30% (w/v) PEG 6000 in 1.6 M NaCl and
leaving on ice for 2 h. Precipitated nucleic acids were pelleted
by centrifugation at 16 000 g for 10 min and washed in 1 ml
of ice-cold 70% (v/v) ethanol before further centrifugation at
16 000 g for 5 min. The ethanol wash was poured off, residual
liquid removed by pipette and pellets dried by warming for
approximately 1 min at 55°C in a hot-block. Pellets were resuspended in 50 µl sterile deionized H2O.
Primer design
A primer capable of discriminating between Group 1.1b and
Group 1.1c soil sequences was designed by aligning in
BioEdit Sequence Alignment Editor (Hall, 1999) >150
sequences placed throughout 1.1b and 1.1c lineages recovered from various environments. These included bulk and
rhizosphere soils (including those recovered previously from
Rotmoosferner forefield soils), subsurface, freshwater and
insect gut samples, before identifying regions of sequence
(and potential primer sites), which were conserved but unique
to the 1.1b lineage. Primer G1.1b280f (5′-GGGCTCTGAG
AGGAGR-3′) was the closest to the start of the 16S rRNA
© 2006 The Authors
Journal compilation © 2006 Society for Applied Microbiology and Blackwell Publishing Ltd, Environmental Microbiology, 8, 1382–1393
Succession and microdiversity of soil Crenarchaea 1391
gene of all positions identified and was chosen to maximize
the amount of sequence information recovered.
RT-PCR amplification of crenarchaeal 16S rRNA genes
for DGGE analysis
Reverse transcription of extracted 16S rRNA was performed
as previously described (Nicol et al., 2005). Total nucleic
acids were treated with RQ1 DNase (Promega, Southampton, UK) before producing archaeal cDNA using Superscript
II RNase H– reverse transcriptase (Invitrogen, Paisley, UK)
according to the manufacturer’s instructions, using primer
Ar9r to select for archaeal rRNA. Two negative controls were
performed with all reactions [no template (water only) and
template but no RT enzyme]. Archaeal 16S rRNA cDNA from
Rotmoosferner soil samples used in this study was generated
for analysis in a previous study (Nicol et al., 2005). Cycling
conditions for amplifying 16S rRNA cDNA of 1.1b (primer set
G1.1b280f/Ar9R) or 1.1c organisms (primer set FFS200f/
Ar9R) were 95°C for 5 min; followed by five cycles of 94°C
for 30 s, 55°C for 30 s, 72°C for 1 min; followed by 30 cycles
of 92°C for 30 s, 55°C for 30 s, 72°C for 1 min; followed by
72°C at 10 min. Nested PCR amplifications using primers
rSAf/PARCH519r for DGGE analysis used the same conditions except that the annealing temperature was 63°C.
DGGE analysis
Denaturing gradient gel electrophoresis was performed using
a DCode Universal Mutation Detection System (Bio-Rad,
Hertfordshire, UK) as described previously (Nicol et al.,
2005). Gels contained a linear gradient of 45–70% denaturant and were electrophoresed in 7 l of 1× TAE buffer at a
constant temperature of 60°C for 900 min at 100 V. Gels were
silver-stained as previously described (Nicol et al., 2005)
before scanning using an Epson GT9600 scanner with transparency unit (Epson, Hemel Hempstead, Hertfordshire, UK).
Cloning and sequencing of crenarchaeal 16S rRNA genes
and ITS regions
Partial 16S rRNA genes (≥ 1299 nucleotides) with complete
ITS regions were amplified using an archaea-specific
(reverse) primer near the start of the 23S rRNA gene in
combination with the archaea-specific 16S rRNA gene primer
A109f or FFS200f (to target specifically the 1.1c crenarchaeal
community). Cycling conditions were 95°C for 5 min; followed
by five cycles of 94°C for 30 s, 55°C for 30 s, 72°C for 2 min;
followed by 30 cycles of 92°C for 30 s, 55°C for 30 s, 72°C
for 2 min; followed by 72°C at 10 min. Polymerase chain
reaction products selected for cloning were purified before
ligating into pGEM-T Easy vector (Promega) and transformed
into XL1-Blue supercompetent Escherichia coli cells (Stratagene, Cambridge, UK). Transformants were screened for
inserts by colony PCR using vector primers M13f/M13r. To
obtain 16S rRNA-ITS clones representative of DGGE migration positions A–F, individual M13f/M13r PCR products were
used as template for PCR with DGGE primer set rSAf/
PARCH519r. Amplicons of individual clones were then com-
pared with the corresponding community DGGE profile for
screening purposes.
To sequence both 16S rRNA and ITS portions of each
clone, the M13f/M13r PCR products were sequenced along
both strands in eight sequencing reactions using primers
A109f (or G1.1c180f), PARCH519r, PARCH533f (reverse
complement of PARCH519r), Ar9R, Ar9F (reverse complement of Ar9R), 1492r, 1513f (reverse complement of 1492r)
and A51r (Table 1) and assembled using Sequencher 4.1
(Genes Codes Corporation, MI, USA). The ITS regions of
some position A and B clones could not be sequenced in
their entirety because of their comparatively long length
(> 900 nucleotides). Therefore, to complete the contiguous
sequence, individual forward and reverse internal primers
were designed for each ITS region after initial 1513f and A51r
sequencing reactions. Additional clones were sequenced
only over the ITS region (after initial screening and selection
by DGGE migration pattern).
Sequence analysis
All 16S rRNA gene sequences were aligned manually using
a secondary structure alignment of archaeal sequences
downloaded from the Ribosomal Database project II (Cole
et al., 2003) using BioEdit. ITS sequences were aligned
using ClustalW (Thompson et al., 1997) implemented in BioEdit before making manual adjustments. Using unambiguously aligned positions only, LogDet/Paralinear distances
(Lake, 1994) were calculated using variable positions (Lockhart et al., 1996) estimated from a maximum-likelihood model
implemented in PAUP v4.01 (Swofford, 1998). Bootstrap support was calculated 1000 times and phylogenetic trees were
constructed by the neighbour-joining method (Saitou and
Nei, 1987) with multifurcation indicating where the relative
branching order could not be determined in the majority of
re-samplings. Sequences of chimeric origin were checked by
analysing alignments using Ballerophon (Huber et al., 2004)
and partial treeing analysis. ITS regions were analysed for
the presence of tRNA genes using tRNAscan-SE 1.21 (Lowe
and Eddy, 1997).
Accession numbers
All sequences have been deposited in the GenBank database with accession numbers DQ278116–DQ278163.
Acknowledgements
This study was supported by the UK Natural Environment
Research Council (Award NER/A/S/2000/01128). The
authors would like to thank Andreas Richter (University of
Vienna) for providing Ödenwinkelkees soil samples and Professor Christa Schleper (University of Bergen) for helpful
discussions.
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