Introduction
Nitrogen is a key element controlling the species composition, diversity,
dynamics, and functioning of many ecosystems (Vitousek et al., 1997).
Despite of recent processes in our understanding of nitrogen cycling
activities in soils, fresh and marine waters, and sediments (Francis et al.,
2005; He et al., 2007; Beman et al., 2008; Jia and Conrad, 2009; Konneke et
al., 2005; Nicol and Schleper, 2006), gaps in knowledge associated with
high-temperature ecosystems have prevailed (C. L. Zhang et al., 2008). Recently,
some studies have elucidated nitrogen metabolism and cycling in
high-temperature hot spring ecosystems (Dodsworth et al., 2011b; Nishizawa
et al., 2013; Gerbl et al., 2014). In such systems, there has been evidence
of microbial communities oxidizing ammonia, the first and rate-limiting step
of nitrification (Reigstad et al., 2008; Hatzenpichler et al., 2008). Since
the occurrence of a putative archaeal amoA gene in hot spring environments was
first reported by Weidler et al. (2007) and Spear et al. (2007),
Thaumarchaeota possessing ammonia monooxygenase (AMO) have been obtained
from some terrestrial hot springs in the USA, China, and Russia (Pearson et
al., 2008; C. L. Zhang et al., 2008).
Previous studies targeting ammonia oxidation in hot springs mainly focused
on archaeal amoA gene (AOA) via a variety of culture-independent approaches
(e.g., 16S rRNA clone library, biomarkers; Weidler et al., 2007; Francis et
al., 2007; C. L. Zhang et al., 2008; Jiang et al., 2010; Xie et al., 2014). The
results from these studies suggested that ammonia-oxidizing Archaea (AOA)
may be ubiquitous in high-temperature environments and even more abundant
than their bacterial counterparts, which has led to a hypothesis that
Archaea rather than bacteria drive ammonia oxidation in high-temperature hot
spring environments. This hypothesis, however, still needs to be verified.
Currently, our knowledge about the activity of AOA in such high-temperature
environments is largely constrained, especially due to the data deficiency
of ammonia oxidation rates (Reigstad et al., 2008; Dodsworth et al., 2011b;
Li et al., 2015). In situ incubation experiments are urgently required to verify
the potential activity of AOA and their contribution to ammonia oxidation in
such high-temperature environments.
In this study, we selected the Gongxiaoshe hot spring at Tengchong
geothermal field as a representative site to test the hypothesis that
Archaea rather than bacteria drive ammonia oxidation in high-temperature hot
spring environments. The reasons for choosing the Gongxiaoshe hot spring as
the research site are: (1) Ammonia concentration in the Gongxiaoshe hot
spring water is 102.61 µg L-1, thermodynamically favorable to
ammonia oxidation (Shock et al., 2005); (2) Ammonia-oxidizing Archaea
Candidatus Nitrosocaldus yellowstonii were dominant in the hot spring water and no
AOB amoA genes were detected in the hot spring (Hou et al., 2013), indicating
that the ammonia oxidation driven by Archaea might be active. Here, in
combination of culture-independent (fluorescence in situ hybridization,
quantitative polymerase chain reaction (PCR), and clone library) and culture-dependent (15N pool
dilution technique) approaches, we provide direct evidence that AOA are
indeed responsible for the major portion of ammonia oxidation in
high-temperature hot spring environments.
The Gongxiaoshe hot spring, located in the Ruidian
geothermal area. (a) A full view of the spring; (b) bottom sediments of the hot spring, designated BS; (c) an
enlarged view of the white box from Fig. 1a, surface sediments of the hot
spring; (d) surface sediments of the hot spring designated as SS;
and (e, f) in situ nitrification activity and potential nitrification
activity experiments in the field.
Materials and methods
Site description and chemical measurements
The Gongxiaoshe hot spring is a small pool with a diameter of ∼ 300 cm and a depth of ∼ 130 cm (Fig. 1). Hot spring water in
the pool is well mixed and water chemistry shows no difference in different
areas of the pool (G. Zhang et al., 2008). Sediments of the Gongxiaoshe hot spring
are found to be only present at the margin of the pool and at the bottom of
the pool, representing two typically sedimentary environments in this pool.
The samples from the pool margins and sediments from the bottom of the
spring, designated SS (surface sediments) and BS (bottom sediments),
respectively, were collected using sterile equipment in April 2013. During
transportation, all of the samples were packed with dry ice. They were then
stored in a freezer at -80 ∘C in a lab for further analysis.
Temperature and pH were measured in situ in the hot water spring. Temperature was
determined with an iButton thermometer (DS1922T, Dallas Semiconductor, USA).
The pH was measured using a pH meter (SevenGo™ pH meter SG2,
Mettler Toledo, USA). Water samples for cation and anion analysis were
filtered through a syringe filter with a 0.22 µm filtration membrane;
these samples were diluted 10 times with deionized water and stored in 100 mL polypropylene bottles in the field because an analysis was carried out
after 2 days. The cation concentrations were determined using an IRIS
Advantage ICP-AES, whereas the anion (F-, SO42-, Cl-)
concentrations were determined using the Ion Chromatography System (DIONEX
ICS-1500, Thermo Scientific, USA). The HCO3- concentration was
measured using the Gran titration method (Appelo and Postma, 1996). The
NH4+-N and NO3--N concentrations were determined using a
nutrient analyzer (Micromac-1000, Partech, UK).
15N stable isotope tracing of nitrification activity
Gross N nitrification rates were determined in situ by the 15N pool dilution
technique. All of the nitrification measurements were conducted in 500 mL
polycarbonate culture flasks (Nalgene) with a silicone plug that contained
400 mL of mud (∼ 1/3 sediment by volume). Two subsamples were
collected from the bottom and surface sediments with 350 µL of
K15NO3 (485 µmol L-1, at 10 % 15N). For each
sample, two experiments were conducted to measure the in situ nitrification
activity: A1 (SS slurry +15NO3-) and A2 (BS slurry +
15NO3-). Meanwhile, potential nitrification activity was
determined in the presence of high ammonium concentration: B1 (SS slurry +
15NO3-+14NH4+) and B2 (BS slurry +
15NO3-+14NH4+). Two pairs of duplicate
reactors were set up in four experiments. The reactors were incubated near
the in situ conditions of the hot spring water at 77 ∘C for 30 and 120 min. At certain time intervals (e.g., 30, 120 min), 80 mL aliquots were
collected from the experimental reactors with sterile serological pipettes
and transferred to acid-cleaned 250 mL polypropylene bottles. Prior to
filtration, 40 mL of KCl (3 M) was added to each sample bottle, and the
samples were shaken at 120 rpm for 1 h and then centrifuged at 1600 × g for 10 min (Reigstad et al., 2008). The supernatant was filtered
through a syringe filter containing a 0.22 µm filtration membrane; the
supernatant was subsequently stored in acid-cleaned 60 mL polypropylene
bottles at 4 ∘C, and analysis was performed after 2 days.
In the laboratory, the concentrations of NH4+ and NO3-
in the filtrate were determined by a nutrient analyzer (Micromac-1000, UK).
The NO3-(15NO3- and 14NO3-) ions
of the filtrates were converted to N2O by denitrifying bacteria
(Pseudomonas aureofaciens) lacking N2O reductase activity, and N2O was quantified by
coupled gas chromatography isotope ratio mass spectrometry (GC-IRMS, Thermo
Scientific, USA; Dodsworth et al., 2011a). The ammonia oxidation rates were
calculated using the equations of Barraclough (1991) as were the
concentrations and N isotope ratios of NO3- in the samples
incubated for 30 and 120 min, respectively.
DNA extraction and purification
DNA was extracted by the SDS-based extraction method described by Zhou et al. (1996), with some modifications. Briefly, approximately 5 g samples were
frozen with liquid nitrogen and milled three times. Then the powdered
samples were mixed with 13.5 mL of DNA extraction buffer and 100 µL of
proteinase K (10 mg mL-1) in tubes; these tubes were horizontally
shaken at 225 rpm for 30 min at 37 ∘C. After shaking, 1.5 mL of
20 % SDS was added, and the samples were incubated in a water bath; the
temperature of the water bath was maintained at 65 ∘C for 2 h.
During this period, the tubes were subjected to gentle end-over-end
inversions every 15 to 20 min. The supernatant fluids were collected after
subjecting the tubes to centrifugation at 6000 × g for 10 min at
room temperature; the collected supernatant tubes were subsequently
transferred into 50 mL centrifuge tubes. The supernatant fluids were mixed
with an equal volume of chloroform: isoamyl alcohol solution (24:1, v/v). The aqueous phase was recovered by centrifugation and precipitated
with a 0.6 volume of isopropanol at room temperature; this process was
carried out for at least 1 h. Crude nucleic acids were obtained by
centrifugation at 16 000 × g for 20 min at room temperature; these
crude nucleic acids were washed with cold 70 % ethanol and resuspended in
sterile deionized water; the final volume of this solution was 100 µL.
The crude nucleic acids were purified with a Cycle-Pure kit (Omega, USA).
These crude nucleic acids were then resuspended in the elution buffer, and
the final volume of the solution mixture was 50 µL; this solution was
stored at -80 ∘C.
PCR and clone library construction
The 16S rRNA gene was amplified with purified genomic DNA as templates using
universal primers. The primer pairs A21F (5′-TTC CGG TTG ATC
CYG CCG GA-3′) and A958R (5′-YCC GGC GTT GAM
TCC AAT T-3′) were chosen for Archaea (Delong, 1992) and
Eubac27F (5′-AGA GTT TGA TCC TGG CTC AG-3′)
and Eubac1492R (5′-GGT TAC CTT GTT ACG ACT T-3′) were chosen for bacteria (Lane, 1991). In a total volume of 50 µL,
the reactions were performed using 1.25 U of Taq DNA polymerase (Takara,
Japan). The amplification conditions were as follows: an initial
denaturation was carried out at 94 ∘C for 4 min, and then, the
same denaturation was continued at 94 ∘C for 1 min. Thereafter,
annealing was carried out at 55 ∘C for 45 s, while extension was
conducted at 72 ∘C for 60 s; the process was repeated for 30
cycles, followed by a final extension step at 72 ∘C for 10 min.
The PCR products were excised after being separated by gel electrophoresis;
a gel-extraction kit (Omega, USA) was used to purify the products in
accordance with the manufacturer's instructions. The purified PCR products
were cloned into pMD20-T vectors (Takara, Japan) and transformed into
competent Escherichia coli DH5α cells. To select the positive clones, colony PCR was
used to determine the presence of correctly sized inserts containing
vector-specific primers M13f (5′-GTA AAA CGA CGG CCAG-3′) and M13r (5′-CAG GAA ACA GCT ATGAC-3′).
Sequencing and phylogenetic analysis
All of the clones were sequenced by the dideoxynucleotide chain-termination
method. In this procedure, an ABI 3730 capillary electrophoresis sequencer
(Applied Biosystem, Inc., USA) was coupled with the T vector universal
primers M13f and M13r. The whole sequence of each clone was spliced using
DNAMAN software (version 6.0), and the vector sequences were deleted; the
presence of chimeras was checked using the Greengenes chimera check tool
(Bellerophon server; Huber et al., 2004). The program DOTUR was used to
determine the operation taxonomic units (OTU) for each sequence; 97 %
similarity was considered as the cutoff for the chimeric sequences. To find
closely related sequences in the GenBank and EMBL databases for phylogenetic
analysis, none of the chimeric sequences were submitted to the Advanced
BLAST search program. Phylogenetic trees were constructed using the
neighbor-joining method and the software MEGA (version 5.05). A bootstrap
analysis was used to provide confidence estimates of the tree topologies.
Amplification of amoA (ammonia monooxygenase subunit A)-related
sequences
Archaeal amoA gene fragments were amplified using the primer pair Arch-amoAF
(5′-STA ATG GTC TGG CTT AGA CG-3′) and
Arch-amoAR (5′-GCG GCC ATC CAT CTG TAT GT-3′; Francis et al., 2005). Bacterial amoA genes were also tested using the
bacterial primer sets amoA 1F (5′-GGG GTT TCT ACT GGT
GGT-3′) and amoA 2R (5′-CCC CTC KGS AAA GCC TTC
TTC-3′; Rotthauwe et al., 1997). PCR cycling was performed by
the method of Francis et al. (2005). In this method, PCR products from SS
and BS were recovered from the gel slices using a gel-extraction kit (Omega,
USA) in accordance with the manufacturer's instructions. The purified PCR
products from each type of sample were cloned into the pMD20-T vectors
(Takara, Japan) and transformed into competent Escherichia coli DH5α cells. Cloning
and sequencing were performed according to the above-mentioned process.
There were 40–50 randomly selected colonies per sample analyzed for the
presence of insert archaeal amoA gene sequences.
FISH probe and PCR primer pairs used in this study.
Application
Probe/
Specificity
Sequence (5′–3′)
FA(%)/
Reference
primer set
AT(∘C)*
FISH
Cren679
Crenarchaeota
TTTTACCCCTTCCTTCCG
35
Labrenz et al. (2010)
qPCR
518F
Bacteria
CCAGCAGCCGCGGTAAT
57
Muyzer et al. (1993)
786R
GATTAGATACCCTGGTAG
344F
Archaea
ACGGGGCGCAGCAGGCGCGA
60
Øvreas et al. (1998)
518R
ATTACCGCGGCTGCTGG
amo196F
Archaeal
GGWGTKCCRGGRACWGCMAC
60
Treusch et al. (2005)
amo277R
amoA
CRATGAAGTCRTAHGGRTADCC
Clone library
A21F
Archaea
TTCCGGTTGATCCYGCCGGA
55
Delong (1992)
A958R
YCCGGCGTTGAMTCCAATT
Eubac27F
Bacteria
AGAGTTTGATCCTGGCTCAG
55
Lane (1991)
Eubac1492R
GGTTACCTTGTTACGACTT
Arch-amoAF
Archaeal
STAATGGTCTGGCTTAGACG
53
Francis et al. (2005)
Arch-amoAR
amoA
GCGGCCATCCATCTGTATGT
amoA 1F
Bacterial
GGGGTTTCTACTGGTGGT
60
Rotthauwe et al. (1997)
amoA 2R
amoA
CCCCTCKGSAAAGCCTTCTTC
* FA: formamide; AT: annealing temperature.
Quantification of 16S rRNA genes and amoA genes
Archaeal and bacterial populations were determined by quantifying their 16S
rRNA genes with 344F-518R (Øvreas et al., 1998) and 518F-786R primer
pairs (Muyzer et al., 1993), respectively. In addition, the abundance of AOA
and AOB were quantified using amo196F-amo277R (Treusch et al., 2005) and
amoA-1F and amoA-2R (Rotthauwe et al., 1997) primers, respectively. All
sample and standard reactions were performed in triplicate. The SYBR Green I
method was used for this analysis. The 20 µL reaction mixture contained
1 µL of template DNA (10 ng), a 0.15 µM concentration of each
primer, and 10 µL of Power SYBR Green PCR master mix (Applied
Biosystems Inc., USA); this reaction mixture was analyzed with ROX and SYBR
Green I. The PCR conditions were as follows: 10 min at 50 ∘C, 2 min at 95 ∘C; 40 cycles consisting of 15 s at 95 ∘C
and 1 min at 60 ∘C; 15 s at 95 ∘C, 1 min at 60 ∘C, and 15 s at 95 ∘C to make the melting curve (Wang
et al., 2009). Melting curve analysis was performed after amplification, and
the cycle threshold was set automatically using system 7500 software v2.0
Patch 6. The efficiencies of the qPCR runs were 87.8–105.6 %
(R2= 0.992–0.999) for 16S rRNA genes and 102 % (R2= 0.998) for
AOA. Primers targeting different genes are listed in Table 1.
Sample processing for FISH
To visualize Crenarchaea cells in situ, fluorescence in situ hybridization (FISH) was performed according to the
procedure described by Orphan et al. (2002, 2009). Small aliquots of
sediment were fixed overnight at 4 ∘C using 2 % formaldehyde
in 1× PBS [145 mM NaCl, 1.4 mM NaH2PO4, 8 mM
Na2HPO4 (pH = 7.4)]; these aliquots of sediments were washed
twice with 1× PBS and stored at -20 ∘C in ethanol: PBS
(1:1, v/v) medium. The total supernatant was filtered through a
polycarbonate filter (Millipore) under low vacuum (5 psi; 1 psi = 6.89 kPa). Filters were cut into suitably sized pieces and transferred onto
untreated, round 2.54 cm glass slides. The transfer of filters onto glass
slides was performed according to the procedure described by Murray et al. (1998). In this process, 5 µL of a 1× PBS solution was spotted
onto a glass slide that was scored with a diamond pen prior to mapping, and
half of the freshly prepared filter was used to invert the sample onto the
slide; this inverted sample was then airdried. Prior to FISH, the samples
on the glass slides were treated with an EtOH dehydration series (50, 75,
and 100 % EtOH), dried, and stored at -20 ∘C. Hybridization
and wash buffers were prepared according to the procedure described by
Pernthaler et al. (2001). Here, 20 µL of hybridization buffer containing
35 or 20 % formamide was added to the samples on the glass slides.
FITC-labeled oligonucleotide Cren679 probe described by Labrenz et al. (2010), was added to the hybridization buffer so that the final solution had
a concentration of 5 ng µL-1.
The hybridization mixtures on the slides were incubated for 1.5 h at 46 ∘C in a premoistened chamber. After hybridization, the slides
were transferred into a preheated wash buffer and incubated for an
additional 15 min at 48 ∘C. The samples were rinsed in distilled
water and airdried in the dark. The microscopic images of the hybridized
samples were recorded on a Leica Imager (Leica, DMI 4000B, Germany).
Nucleotide sequence accession numbers
The clone libraries for archaeal communities (21F-958R), bacterial
communities (27F-1492R), and archaeal amoA genes (amoAF-amoAR) were
constructed. All of the small-subunit rRNA gene sequences and the amoA sequences
were deposited in the GenBank/EMBL nucleotide sequence database under the
following accession numbers: from KP784719 to KP784760 for partial 16S rRNA gene
sequences and from KP994442 to KP994448 for the amoA sequences.
Results
Water chemistry
The hot spring water (pH = 7.7) contained Ca (20.25), K (41.97), Mg (3.986), Na (313.3), SiO2
(130.3), HCO3- (963 mg L-1), NH4+-N
(102.61 µg L-1), NO3--N (7.68 µg L-1), F- (9.158 mg L-1), Cl- (418.9 mg L-1), and SO42-
(24.96 mg L-1). The bottom water had a temperature of 77 ∘C, higher than the surface water that had a temperature of 55 ∘C.
This hot spring was previously categorized as a Na-HCO3 spring due to
the high concentration of alkaline metal ions (K, Na, and Ca; G. Zhang et al.,
2008).
Ammonia oxidation rates
In the surface and bottom sediments (without NH4+ stimulation),
the near in situ rates of ammonia oxidation were estimated to be 4.80 ± 0.2
and 5.30 ± 0.5 nmol N g-1 h-1 using 15N-NO3-
pool dilution technique, respectively. In the meantime, the nitrate
concentration increased from 2.84 ± 2 to 3.25 ± 2 µM in the surface sediments and from
2.33 ± 3 to 2.62 ± 3 µM in the bottom sediments, further providing evidence for strong
nitrification activity under in situ conditions in the hot springs. Furthermore, the
potential activity of ammonia oxidation was determined with ammonium
amendments. The nitrate concentration increased significantly upon the
addition of NH4+, and the ammonia oxidation rates recorded in the
surface sediments and bottom sediments (with NH4+) were 5.70 ± 0.6 and 7.10 ± 0.8 nmol N g-1 h-1, respectively.
Archaeal community composition and phylogenetic analysis
A total of 152 archaeal clone sequences of 16S rRNA genes were obtained in
this study. Phylogenetic analysis showed the distribution of the clone
sequences into three monophyletic groups: Thaumarchaeota, Crenarchaeota, and
Euryarchaeota (Fig. 4). In this study, the most abundant archaeal phylum was
Thaumarchaeota. Among them, two phylotypes (SS-A19 and BS-A1) were the most
dominant archaeal lineage, representing 89 and 86 % of the cloned
archaeal sequences in surface and bottom sediments, respectively. These
sequences were closely related to the thermophilic, autotrophic,
ammonia-oxidizing archaeal Ca. N. yellowstonii (de la Tarre et al., 2008).
The seven archaeal OTUs found here belonged to Crenarchaeota, which contains
sequences recovered from hydrothermal vents and hot spring environments. In
addition, two phylotypes (BS-A47 and BS-A8) that were branched with
uncultured sequences belonged to Desulfurococcales, which was also recovered from sediments of
the hot spring. Euryarchaeota also occurred in both the sediments, but with
relatively low abundances. Phylotype BS-A80 is associated with Geoglobus ahangari, which
belongs to Archaeoglobales and is capable of oxidizing organic acids (Kashefi et al.,
2002). SS-A12, which represents four clones recovered from the surface
sediments, showed 93 % similarity to an uncultured archaeal clone that was
recovered from the Spring River. SS-A47 belonged to the Thermoplasmatales that were 96 %
similar to their nearest neighbor sequence, which were collected from the
Spring River. The other euryarchaeotal sequences BS-14 and BS-A80 were
similar to their uncultured counterparts (from 96 to 99 % identity), which
were mostly recovered from high-temperature geothermal environments.
Community analysis of AOA
A total of 113 archaeal amoA gene fragments were obtained from the two samples.
They were all branched within the four distinct clusters of archaeal amoA
sequences: cluster Nitrosopumilus, Nitrososphaera, Nitrosotalea, and Nitrosocaldus (Fig. 5). Nitrosopumilus cluster contained
phylotypes SS-AOA-4 and
BS-AOA-22, which branched with large numbers of sequences recovered from the
sediments and water samples in the marine or fresh environments. The other
clade, cluster Nitrososphaera, has two phylotypes representing 44 sequences. OTU BS-AOA-62
contained 18 sequences, which was closely related to sequences from soil.
The clone SS-AOA-76 clustered within clade Nitrososphaera and showed up to 99 % sequence
identity to an uncultured archaeon clone GHL2_S_AOA_19 (JX488447) obtained from lake
sediment.
Cluster Nitrosotalea had one phylotype (SS-AOA-65) with 11 sequences (12 % of the total
sequences). The closely related sequences in this cluster included
characteristic crenarchaeotal group sequences that were obtained from alpine
soil (with 98 % identity). Another clone, MX_3_OCT_18 (DQ501052), from estuary sediment was
96 % similar.
Cluster Nitrosocaldus contained two phylotypes (BS-AOA-15 and SS-AOA-50) with 34
sequences (30 % of the total sequences). They were closely related to the
geothermal water sequences, with 95–99 % similarity. Furthermore, cluster
Nitrosocaldus mainly represented previously described ThAOA/HWCG III (Prosser and Nicol,
2008). Notably, the recently reported amoA gene sequence of Ca. N. yellowstonii
(EU239961; De la Torre et al., 2008) showed 85 % sequence identity to
clones BS-AOA-15 and SS-AOA-50.
(a) Gross ammonia oxidation rates calculated
from 15N-NO3- pool dilution experiments on amended (added
14NH4+) or unamended SS and BS sediment slurries. This defines
that the amendment with “15NO3” represents in situ nitrification
activity, while 15NO3 plus 14NH4 is considered as
potential nitrification activity. Bars represent the mean and standard error
of the mean (n= 3) for 30 and 120 min incubation. (b) Abundance
of archaeal 16S rRNA genes and archaeal amoA genes for SS and BS samples
collected from the Gongxiaoshe hot spring. Data are expressed as gene copies per
gram of sediment (dry weight). Error bars represent the standard deviation
of the mean (n= 3).
Quantitative PCR
The qPCR results (Fig. 2b) indicated that the abundance of the archaeal 16S
rRNA gene in the two samples was similar, ranging from 1.28 to
1.96 × 107 gene copies g-1 of dry weight of sediments.
However, the abundance of the bacterial 16S rRNA gene varied greatly,
ranging from 6.86 × 106 to 4.25 × 108 gene copies g-1 of dry weight of sediments (Fig. S2 in the Supplement). The
copy numbers of archaeal amoA genes in the surface and bottom sediments are
2.75 × 105 and 9.80 × 105 gene copies g-1
sediment, respectively. The copy numbers of the archaeal 16S rRNA genes in
the bottom sediments were significantly higher than those of the bacterial
16S rRNA genes, with a ratio of 28.57. However, in surface sediments, the
ratio of bacterial 16S rRNA genes to archaeal 16S rRNA genes is 3.32.
FISH
FISH was used to analyze the relative abundance of Crenarchaea in two
samples. As expected, most metabolically active Crenarchaea cells and
aggregated cells were detected by FISH probes (Cren679; Fig. 3). Based on
the qPCR results, a high abundance of Crenarchaea in the hot spring
sediments harbored amoA genes, providing strong evidence supporting the
important role of Crenarchaea in the oxidation of ammonia.
Epifluorescence photomicrograph of Crenarchaeota cells
and cell aggregates. (White and red arrows show the cells and carbonate
crystals, respectively. Scale bar corresponds to 20 µm.)
Discussion
Environmental factors affecting the occurrence of ammonia-oxidizing
microorganisms
Temperature is likely a very important factor influencing microbial
community structure. This interpretation is supported by the results of qPCR
(Figs. 2b and S2). The sediment samples from the bottom of pool (T= 77 ∘C) are dominated by Archaea, whereas the sediment samples from
the margin of pool (T= 55 ∘C) are dominated by bacteria. In
addition, no AOB were detected in both bottom and margin samples, indicating
that it might be difficult for AOB to inhabit in high-temperature hot spring
environments (Lebedeva et al., 2005; Hatzenpichler et al., 2008).
Additionally, the abundance of AOA amoA gene in bottom sediments is slightly
higher than that in margin sediments, reflecting that although AOA can adapt
to a wide range of temperature, higher temperature could be more favorable
to the growth of AOA (de la Torre, et al., 2008; Hatzenpichler et al., 2008;
Jiang et al., 2010).
Ammonia concentration may be another factor that influences the potential
activity of AOA and AOB in hot springs. Because AMO in AOA has a much higher
affinity for the substrate compared to a similar process in AOB, the ability
of AOA to compete for ammonia in oligotrophic hot spring environments is
also substantially higher than that of AOB (Hatzenpichler et al., 2008). In
the Gongxiaoshe hot spring, the ammonia concentration of 102.61 µg L-1
is lower compared to other hot springs with high ammonia concentrations.
This relatively low ammonia concentration may possibly be responsible for
the absence of AOB in the Gongxiaoshe hot spring.
Archaeal phylogenetic tree based on 16S rRNA gene
sequences, including various 16S rRNA gene clones obtained from the
Gongxiaoshe hot spring sediments (SS and BS) and cited some sequences from
Hou et al. (2013; in red). The tree is constructed using the
neighbor-joining method, and bootstrap confidence values over 50 % (1000
replicates) are shown. The scale bar represents the expected number of
changes per nucleotide position.
Composition and abundance of AOA
The rarefaction curves (Fig. S3) for archaeal 16S rRNA genes and amoA genes in
the surface and bottom sediment samples reached a plateau, and their
coverage values were relatively high (89–99 %). This result indicated that
a large part of the archaeal/amoA diversity at this spring was probably included
in the archaeal/amoA clone libraries. The majority of archaeal sequences were
closely related to Ca. N. yellowstonii, a known AOA, which may be responsible
for the oxidation of ammonia in this spring.
In this study, phylogenetic analyses of archaeal amoA genes showed that
Candidatus Nitrosocaldus yellowstonii dominated in both of the samples. This result
also agreed with previous hot spring observations reported by Dodsworth et al. (2011b) and Hou et al. (2013). According to the sequences retrieved from
NCBI, Nitrosotalea and Nitrososphaera clusters were closely related to the cluster soil. One possibility is
that some of the amoA genes obtained in this study may derive from soil AOA,
particularly those sequences in cluster Nitrosotalea and cluster Nitrososphaera, which have been widely
found in sediments and soils. Those AOA from soil might have evolved
multiple times and have adapted to high-temperature environments. Based on
the analysis of the real-time PCR and FISH methods, our data indicate that
the abundance of AOA is relatively high in both samples. The archaeal amoA gene
copy numbers ranged from 2.75 to 9.80 × 105 per gram dry
weight of sediments in this study. This is comparable to the abundance in
other hot water springs [104–105 copies g-1 (Dodsworth et
al., 2011b)], but is lower than the abundance of the archaeal amoA gene in
nonthermal environments, such as paddy rhizosphere soil [106–107 copies g-1 (Chen et al., 2008)] and marine
sediments [107–108 copies g-1 (Park et al., 2008)]. The bacterial amoA genes were not
detected, indicating that AOB is absent or is a minority in this hot spring
ecosystem. A predominance of archaeal amoA genes versus bacterial amoA genes
indicated that ammonia oxidation may be due to the activity of Archaea in
the Gongxiaoshe hot spring.
The phylogenetic tree of archaeal amoA genes is cloned
from the Gongxiaoshe hot spring sediments (SS and BS). The tree is
constructed using the neighbor-joining method, and bootstrap confidence
values over 50 % (1000 replicates) are shown. The scale bar represents the
expected number of changes per nucleotide position.
The role of AOA in the nitrification of terrestrial geothermal
environments
In the surface and bottom sediments (without NH4+), the ammonia
oxidation rates calculated from the 15N-NO3- pool dilution
data were 4.80 ± 0.2 and 5.30 ± 0.5 nmol N g-1 h-1,
respectively. The ammonia oxidation rates recorded in the surface sediments
and bottom sediments (with NH4+) were 5.70 ± 0.6 and 7.10 ± 0.8 nmol N g-1 h-1, respectively. Moreover, the rates
reported here were comparable with those observed in the two US Great Basin
(GB) hot springs [5.50–8.60 nmol N g-1 h-1 (Dodsworth et al.,
2011b)] and in two acidic (pH = 3, T= 85 ∘C) Iceland hot
springs [2.80–7.00 nmol NO3- g-1 h-1 (Reigstad et al.,
2008)]. However, the rates reported in this study were lower than those
observed in some wetland sediments and agricultural soils [85–180 nmol N g-1 h-1 (White and Reddy, 2003; Booth et al., 2005)].
The ammonia oxidation rates in bottom sediments (without NH4+)
were slightly higher than those observed in surface sediments (without
NH4+). This result agrees with the distribution of archaeal amoA
genes, which were found to be in higher abundance in the bottom sediment
than in the surface sediment. High abundance of ammonia-oxidizing Archaea
corresponds to high ammonia oxidation rates, which were consistent with the
results reported by Isobe et al. (2012). Compared with the incubation
experiments unamended with NH4+, the ammonia oxidation rate
appeared to be stimulated after amendment with NH4+ (1 M). There
are indications that the ammonia concentration is an important factor
affecting the rates of nitrification (Hatzenpichler et al., 2008).
To understand the relationship between the ammonia oxidation rates and
abundances of amoA in the two samples, we specifically estimated the
contribution of archaeal cells to nitrification. By assuming two amoA copies per
cell (Bernander and Poplawski, 1997) and by comparing the ammonia oxidation
rates with the qPCR results of AOA amoA per gram (however, some uncertainties of
this method may still exist, with respect to the stage of cell cycle and the
diversity of Archaea), the cell-specific nitrification rates were estimated
to be 0.410 and 0.790 fmol N cell-1 h-1
in the surface and bottom sediments of the hot spring, respectively. These
results are much higher than those for AOA in US hot springs [0.008–0.01 fmol N cell-1 h-1 (Dodsworth et al., 2011b)]. It is interesting that
although the GB hot spring possesses higher amoA gene copies
(3.50–3.90 × 108 gene copies g-1 of dry weight) and higher
NH4+ concentration (663 µg L-1), it exhibits a lower
cell-specific nitrification rate than the Gongxiaoshe hot spring. This may imply
that both the abundance of AOA and the NH4+ concentration are not
important factors that control the cell-specific nitrification rates in
high-temperature hot spring environments. The difference in cell-specific
nitrification rates between the Gongxiaoshe hot spring and the GB hot
spring may reflect the difference of AOA population structure in those two
hot springs (Gubry-Rangin et al., 2011; Pester et al., 2012). In line with
this AOA heterogeneity, cell-specific nitrification rates do not reflect the
overall AOA abundance or NH4+ concentration in these AOA-dominated
hot springs. Alves et al. (2013) reported a similar case where soil
dominated by AOA (clade A) exhibited the lowest nitrification rates, in
spite of harboring the largest AOA populations. These results also suggest
the importance of cultivation studies for comparative analysis of
environmentally representative AOA in a wide variety of hot springs.