Biol Fertil Soils DOI 10.1007/s00374-014-0902-2
ORIGINAL PAPER
Changes in bacterial and archaeal community assemblages along an ombrotrophic peat bog profile Edoardo Puglisi & Claudio Zaccone & Fabrizio Cappa & Pier Sandro Cocconcelli & William Shotyk & Marco Trevisan & Teodoro M. Miano
Received: 12 November 2013 / Revised: 31 December 2013 / Accepted: 14 January 2014 # Springer-Verlag Berlin Heidelberg 2014
Abstract Peatlands are archives of extreme importance for the assessment of past ecological, environmental and climatic changes. The importance as natural archives is even greater in the case of ombrotrophic peat bogs, where the only inputs are atmospheric in origin. Here we integrated previously published physical and chemical results regarding the solid and liquid phase of peat with a biomolecular microbiological approach to assess the relationships between chemistry and microbial biodiversity along a Swiss bog profile corresponding to approximately 2,000 years of peat formation. The structure of bacterial and archaeal communities was assessed through a polymerase chain reaction-denaturing gradient gel electrophoresis (PCR-DGGE) approach followed by sequencing of PCR-DGGE bands of interest. Both chemical and microbiological data showed a differentiation of properties along the peat profile, with three major zones identified. Both bacterial and archaeal profiles clustered according to the depth (i.e., age) of samples. Among bacteria, E. Puglisi : F. Cappa : P. S. Cocconcelli Istituto di Microbiologia, Università Cattolica del Sacro Cuore, Via Emilia Parmense 84, 29122 Piacenza, Italy C. Zaccone Department of the Sciences of Agriculture, Food and Environment, University of Foggia, Via Napoli 25, 71122 Foggia, Italy C. Zaccone : W. Shotyk Department of Renewable Resources, University of Alberta, 348E South Academic Building, Edmonton, Alberta, Canada T6G 2H1 M. Trevisan (*) Istituto di Chimica Agraria ed Ambientale, Università Cattolica del Sacro Cuore, Via Emilia Parmense 84, 29122 Piacenza, Italy e-mail:
[email protected] T. M. Miano Department of Soil, Plant and Food Sciences, University of Bari “Aldo Moro”, Via Amendola 165/A, 70126 Bari, Italy
Acidobacteria were recovered primarily in the first layers of the profile, whereas methanogenic archaea were more commonly recovered in the deepest part of the core, corresponding to the occurring anoxic conditions. Finally, a number of sequences had low homologies with known species, especially in bacteria: this points to an almost unknown microbial community adapted to the extreme conditions of peat bogs, which are acidic, rich in dissolved organic C, and predominantly anoxic. Keywords Peatlands . Ombrotrophic peat bog . Bacterial community . Archaeal community . PCR-DGGE
Introduction Peatlands are water saturated organic soils (histosols) that represent natural archives of extreme importance for the assessment of past ecological, environmental and climatic changes. Their formation is due to the accumulation of organic material, mainly of plant, but also of animal and microbial origin. These materials are preserved from complete degradation thanks to the low pH and generally anoxic conditions which reduce decomposing activity (Shotyk et al. 1998; Suyama et al. 2008). Among the different kind of peatlands, the most reliable as natural records are ombrotrophic bogs, where the only inputs are represented by wet and dry atmospheric deposition, and whose hydrology is controlled primarily by precipitation and evaporation processes (Damman 1978). Sampling of cores from ombrotrophic bogs allows a reconstruction of thousands of years of environmental changes, including deposition of trace elements, radionuclides and organic contaminants (e.g., Madsen 1981; Martinez-Cortizas et al. 1999; Shotyk 1996; Shotyk et al. 1998; Zaccone et al. 2007a, 2009a) or past climatic or vegetation conditions (e.g.,
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Aaby 1976; Langdon et al. 2011; van Geel 1978; Tareq et al. 2004; Woillard 1978). The vegetation of peat bogs is generally dominated by Sphagnum species that create an acidic, nutrient-poor and water-saturated environment (Clymo and Hayward 1982) in which few other plants can flourish (van Breemen 1995). Sphagnum species have also been reported to produce phenolic metabolites that can exert anti-microbial activities (Mellegård et al. 2009; Rudolph and Samland 1985): this factor, together with low temperatures and high moisture levels, shapes an extreme environment with low decomposition rates, especially in the anaerobic zone where pores are always filled with water (Clymo 1984; Frolking et al. 2001; Pankratov et al. 2011). One of the most important features of peat bogs, and of peatlands in general, is their major role in biogeochemical cycles related to greenhouse gases (GHG) emissions: albeit covering less than 3 % of the Earth’s terrestrial surface (Kivinen and Pakarinen 1981), peatlands host an estimated 30 % of the global reserves of soil C (Gorham 1991). To better understand their role in the C cycle, several microbiological analyses of bog environments focused on the assessment of methanogenic microbial communities involved in GHG emissions (Dedysh et al. 2006) and on the isolation of related strains (Bräuer et al. 2006) or consortia (Sizova et al. 2003). Variation in microbial communities structure along a depth profile has been widely shown for mineral soils, reflecting changes in water levels, redox conditions and availability of nutrients (Boz et al. 2013; Chaparro et al. 2012; Fierer et al. 2003; Fuka et al. 2008; Rahman et al. 2013). About peat bogs, after the pioneering works of Waksman and Stevens (1929) and Waksman (1930), a number of studies focused on the variation with depth in the composition of microbial communities along the profile, as recently reviewed by Anderson et al. (2013). In particular, most of studies regarded the first 60–70 cm using primers specific for methanogenic populations (Cadillo-Quiroz et al. 2006; Dedysh et al. 2006; Juottonen et al. 2005; Kotsyurbenko et al. 2004), while to the extent of our knowledge the only work analyzing a deep bog core (200 cm) was carried out by Dettling et al. (2007) in order to evaluate methanogenesis and the diversity of methanogens along the profile. Regarding the total bacterial community structure, Dedysh et al. (2006) analysed an 80 cm deep profile, and they found a predominance of Acidobateria, followed by Alphaproteobacteria and Planctomycetes. They also outlined how a high proportion of 16S rRNA sequences could not be assigned to any known phylum, thus showing that peat bogs host a partially unknown microbiota. According to Morales et al. (2006), total bacterial counts were higher but the community was less diverse for samples found at 1 m depth than at the surface, while other studies suggested a general decrease in biomass and diversity with depth (Golovencho et al. 2007; Jaatinen et al. 2007). A
differentiation according to the depth has also been highlighted, with the interface between oxic and anoxic layer richer in methanotrophic bacteria, and the deeper anaerobic horizons characterized by a limited dispersal of strains and a domination of methanogenic strains (Andersen et al. 2013). The present study was carried out with the aim of assessing the total community structure of archaea and bacteria along a 105-cm undisturbed peat core from the Etang de la Gruère (EGr) ombrotrophic bog that has been previously characterized in detail for a number of physical and chemical properties (Zaccone et al. 2007a; 2009a,b). Thus, the main hypotheses tested here were that (1) the structure of total microbial communities would change along the profile reflecting the changing chemical and physical conditions published previously; (2) peat bogs not only host a diversity of microbial species, but that some of these are adapted, almost exclusively, to the unique conditions characteristic of these environments.
Materials and methods Site description Peat coring was carried out in 2005 at EGr, which is a wellstudied ombrotrophic bog situated at an altitude of ca. 1,005 m above sea level in a protected area of the Jura Mountains, Northwest Switzerland. The EGr bog represents the longest continuous record of ombrotrophic peat in the northern hemisphere. The climate in this area is moist continental, with an annual average temperature of 5.5 °C and average precipitation of 1,300 mm/year (Steinmann and Shotyk 1997). The vegetation is dominated by Sphagnum species, although small shrubs, stunted trees, sedges and other mosses are also present. The formation of the bog at EGr began during the Late glacial era, ca. 14,500 years ago (Roos-Barraclough et al. 2004), and resulted in a thickness of peat accumulation of approximately 6.5 m. From 1705 until ca. 1900, drainage ditches dug in the eastern part of the bog supplied the mill at Derrière la Gruère with more water. The western system continued to drain the bog until 1987 when the ditches were filled in. Until 1952, the use of lake water by Moulin de la Gruère caused large variations in lake level, probably with consequences for the water table throughout the lagg zone of the peatland Roos-Barraclough et al. (2004). However, no drainage or peat cutting was carried out on the peninsula. As the dome of the bog at EGr is elevated well above the level of the pond, it is assumed that the water table where the core was taken has never been impacted in a significant way. A previous study carried out on a parallel core (2 h) collected in 1991 from the same bog showed that the first ca. 80 cm of peat have an elemental composition in the range of
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48–58 % C, 6–7 % H, 0.6–2.5 % N, 0–0.2 % S, and 33–45 % O (Zaccone et al. 2007b). Peat core sampling and sample preparation A peat profile (core 2 T, 15×15×105 cm), covering the last 2,110±30 14C years BP (Zaccone et al. 2007a, 2009a), was collected with a Wardenaar sampler (Wardenaar 1987) from the central domed area of the peat bog (47°14′22.62″ N; 7°02′ 57.42″ E), where the surface peat layers are clearly elevated up to 4 m beyond the edge of the bog. The core was wrapped in plastic film and packed in a wooden crate, and kept at −18 °C until analyses; later, in the lab, it was sliced frozen into 1± 0.15 cm sections with a stainless steel band saw. In order to avoid any kind of contamination, the outside edges (ca. 2 cm) were systematically discarded. Of the 91 slices obtained, 20 of them were selected for the present study, based on the physical, chemical and spectroscopic features reported in previous investigations on the same core (i.e., Zaccone et al. 2007a, 2009a,b). Some features of the entire 2 T core are summarized in Table 1, while details about samples utilized in the present study are reported in Table 2. Four commercial peat samples were also included in microbiological diversity analyses. These samples, kindly provided by a local fertilizer company, consisted of two blond Estonian peats (P1 and P2), a black German peat (P3) and an Irish peat (P4), and they were chosen in order to highlight the differences in microbial communities with respect to peat samples from other sites, and to serve as controls in the analyses. These controls were immediately brought to the lab and stored at 4 °C for a maximum of 3 weeks until DNA extraction. DNA extraction and PCR-DGGE analysis First, 500-mg peat samples were thawed at room temperature, and the DNA was then extracted with the FastDNA® SPIN kit for soil (MoBio Laboratories, Carlsbad, CA, USA) according to the manufacturer’s instructions. DNA was checked spectrophotometrically for purity (260/280 and 260/230 nm ratios) and for possible shearing and degradation by electrophoresis on 0.8 % and 1 % agarose gels, respectively. Nucleic acids
concentrations were assessed with the Quant-iT™ HS dsDNA Assay kit (Invitrogen, Paisley, UK) in combination with the QubitTM fluorometer (Invitrogen). DNA samples were analysed by a nested polymerase chain reaction-denaturing gradient gel electrophoresis (PCRDGGE) approach targeting the 16S rRNA gene for bacteria and archaea, respectively. The first amplification step was carried out using the primers 338 F/805R and 787 F/1059R amplifying respectively the V3 region of bacteria and the V5 region of archaea (Sousa et al. 2007). The same primers sets were used in the second amplification, but adding a 40-bp GC clamp to the reverse primers. PCR amplification was carried out in 50-μl reaction mixtures using the Promega MasterMix (Promega Corporation, Madison, WI, USA) containing 50 units/ml of Taq DNA polymerase in pH 8.5 reaction buffer, with 400 μM dNTPs. 2.5 μl of DNA template were added in each mixture containing 12.5 μl of Promega MasterMix, 2.5 μl of each primer set (1 μM final concentration), 2.5 μl of MgCl2 (25 mM final concentration) and dH2O to the final volume of 50 μl. The thermal cycling conditions used for the amplification of the 16S rRNA genes were the same for both bacteria and archaea, and were 94 °C for 2 min, 30 cycles of 94 °C for 30 s denaturation, 55 °C for 30 s annealing, 72 °C for 30 s extension, and a final extension step of 72 °C for 10 min. Conditions of the second amplification step were the same, except for the annealing temperature, that was raised to 56 °C, and the extension step time, raised to 60 s. DGGE was carried out using the INGENY phorU electrophoresis apparatus (Ingeny International BV, Goes, The Netherlands). Polyacrylamide gels (6 %) in 1× TAE buffer (40 mM Tris base, 20 mM acetic acid, and 1 mM disodium EDTA, pH 8.2) were prepared with denaturating gradient ranging from 45 to 65 % for bacteria 16S and from 40 to 60 % for archaea. The electrophoresis was run for 18 h at 60 °C and 90 V. Gels were SYBR Green-stained and visualized by UV radiation. Banding patterns of DGGE profiles were analysed with Fingerprinting II software (Bio-Rad Laboratories, Richmond, CA, USA) using the Dice correlation coefficient and the unweighted-pair group method with averages (UPGMA) for the generation of dendrograms. Preliminary analyses were carried out to assess the variability in DGGE patterns among three sub-replicates per sample.
Table 1 Selected chemical and physical features of peat along the 2 T profile
Maximum value Minimum value Avg. value±SD (n=91)
Dry density (g cm−3)
Depth (cm)
Wet density (g cm−3)
Depth (cm)
Ash content (%)
Depth (cm)
Water content (%)
Depth (cm)
pH
0.16 0.03 0.08±0.03
−35.9 living layer
1.06 0.31 0.88±0.13
−89.4 living layer
6.64 1.05 1.90±0.99
−33.4 −68.3
94.8 84.0 90.9±2.7
−68.3 −35.9
3.92 −98.3 3.43 −8.0 3.70±0.13
Data are from Zaccone et al. (2009b)
Depth (cm)
Biol Fertil Soils Table 2 Details about peat samples used in the present study Peat slice no.
Average depth (cm)
Calendar year (AD)a (210Pb)
2 2 2 2
T 01 T 05 T 13 T 16
1.5 −3.5 −11.0 −14.1
2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2
T 27 T 29 T 30 T 33 T 37 T 41 T 46 T 53 T 58 T 60 T 66 T 71 T 76 T 81 T 88 T 90
−26.2 −28.4 −29.6 −33.4 −38.3 −43.3 −49.4 −58.1 −64.4 −67.0 −74.6 −80.7 −86.4 −91.6 −99.5 −102.5
Radiocarbon agea (14C years BP)
Dry densityb (g cm−3)
Ashb (%)
pHb
DNA concentration (ng mg−1, dry weight)
2005±0 2001±0 1982±2 1971±2
0.029 0.042 0.079 0.085
1.80 1.61 2.35 1.43
3.9 3.6 3.5 3.6
26.9 33.0 78.5 47.4
1941±3 1929±6 1919±6 1871±20
0.071 0.101 0.117 0.152 0.133 0.116 0.122 0.082 0.063 0.059 0.053 0.053 0.067 0.078 0.092 0.059
2.07 3.13 3.18 6.64 2.59 3.01 1.74 1.42 1.67 1.30 1.35 1.20 1.33 1.56 1.67 1.66
3.6 3.7 3.5 3.6 3.6 3.7 3.7 3.7 3.7 3.8 3.7 3.9 3.9 3.9 3.9 3.9
93.0 140.9 82.3 43.2 36.8 51.8 29.1 18.4 20.0 14.3 11.4 14.0 9.7 18.3 4.3 13.2
2,110±30
Chemical data have been previously published elsewhere, while DNA concentrations were determined for this study by QBit measurements a
Data from Zaccone et al. (2007a)
b
Data from Zaccone et al. (2009b)
Since no significant variability among sub-replicates was found (data not shown), analyses were carried out on a single replicate per depth, as often done in previous studies on peat bog cores (Dedysh et al. 2006; Morales et al. 2006). Individual bands were cut out from the gels, placed in 50 μl sterile dH2O and incubated overnight at 4 °C. Two microliters of the eluate was re-amplified with the original primer pairs and amplification products were checked by agarose gel electrophoresis to make sure that a single band was collected. PCR products were purified using the Wizard SV Gel and PCR Clean-Up System (Promega) and subjected to sequencing at the BMR genomics facilities (Padova, Italy). The resulting sequences were aligned against the Ribosomal Database Project (RDP) database (http://rdp.cme.msu.edu) using the Naïve Bayesian classifier (Wang et al. 2007). All 16S rRNA sequences were also screened for possible presence of chimera with the DECIPHER software (Wright et al. 2011). After screening for good quality read levels (Phred score >16), 20 sequences in total for bacteria and 19 for archaea were retained. Among these, one bacterial and one archaeal sequence were identified as chimeras and thus excluded from the concomitant analyses for assessment of the closest reference sequences on the RDP website, and phylogenetic trees building. Sequence match on RDP was evaluated with Sab
(similarity) score, a value between 0 and 1 showing the identities of sequences as compared to the ones deposited in RDP, where 1 is the complete homology. Sequences were deposited in GenBank under the accession numbers KF482411–KF482429 for bacteria and KF482430– KF482447 for archaea. Multiple alignments of the identified sequences and closer hits on RDP database were carried out with the MUSCLE algorithm (Edgar 2004), and phylogenetic trees were constructed on the aligned sequences with the PhyML (Phylogeny Maximum Likelihood) approach (Guindon et al. 2003) by applying the Shimodaira– Hasegawa [SH]-aLRT test. Alignments and tree generation were carried out with the SeaView software (Gouy et al. 2010) after trimming the reference sequences to the region aligned with the short peat sequences.
Results and discussion DNA quantification and PCR-DGGE fingerprint Storage and handling of samples was carried out in order to minimize possible effects on microbial properties. Commercial peat samples were kept only a few days at 4 °C
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and then immediately subjected to DNA extraction. For the peat samples from the EGr bog, a long-term storage at −20 °C was mandatory, but it is not supposed to have affected the microbial community profiles (Lauber et al. 2010; Rubin et al. 2013). Furthermore, losses or migration of porewaters during the core preparation and possible cross contaminations have been avoided by cutting the core when it was still frozen and discarding the outer edges of each slice, respectively. DNA quantification data, as determined by QBit fluorometric method, are reported in Table 2, along with chemical and physical parameters previously determined in the same samples. DNA concentration data ranged from a minimum of 4.3 to a maximum of 140.9 ng DNA mg−1 of peat (dry weight). A trend was evident, with an increase in concentrations in the first 28 cm (where the maximum peak was measured) and a decrease in the following 30 cm, with concentrations finally remaining minimal and stable from 67 cm onwards. These results are in agreement with previous evidences indicating a maximum of microbial biomass at intermediate depths of the so-called mesotelm (Clymo and Bryant 2008), i.e., the interface between oxic and anoxic layers. Here, the simultaneous presence of CH4 and O2 creates ideal conditions for microbial species, especially methanotrophs (Andersen et al. 2013). DNA concentration seems to show a similar trend when compared with the ash distribution, with corresponding peaks shifted only a few cm deeper (28 vs. 33 cm of depth, respectively), but this correlation was not statistically significant (R=0.415, p=0.069). On the opposite, significant correlations (p
