Impact of peat mining, and restoration on methane turnover

potentials and methane-cycling microorganisms in a northern bog.

Max Reumer1, Monika Harnisz2, Hyo Jung Lee1,3, Andreas Reim4, Oliver Grunert5, Anuliina

Putkinen6, Hannu Fritze6, Paul L.E. Bodelier1, Adrian Ho1#.

1Department of Microbial Ecology, Netherlands Institute of Ecology (NIOO-KNAW),

Droevendaalsesteeg 10, 6708 PB, Wageningen, The Netherlands.

2Department of Environmental Microbiology, University of Warmia and Mazury in Olsztyn,

Prawocheńskiego 1 Street, 10-719 Olsztyn, Poland.

3Department of Biology, Kunsan National University, Gunsan 54150, Republic of Korea.

4Department of Biogeochemistry, Max-Planck-Institute for Terrestrial Microbiology, Karl-

von-Frisch-Str. 10, D-35043 Marburg, Germany.

5Center for Microbial Ecology and Technology, Ghent University, Coupure Links 653, 9000

Gent, Belgium.

6Natural Resources Institute Finland, Latokartanonkaari 9, 00790 Helsinki.

#For correspondence: Adrian Ho (adrian.ho@ifmb.uni-hannover.de).

Current affiliation: Institute for Microbiology, Leibniz University Hannover, Herrenhäuser Str.

2, 30140 Hannover, Germany.

Running title: Methane turnover in restored peatlands.

Keywords: Sphagnum / Methanogenesis / Methane oxidation / Nitrogen fixation/ Land-use

change / nifH.

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Abstract

Ombrotrophic peatlands are a recognized global carbon reservoir. Without restoration and

peat regrowth, harvested peatlands are dramatically altered, impairing its carbon sink

function, with consequences for methane turnover. Previous studies determined the impact

of commercial mining on the peat physico-chemical properties, and the effects on methane

turnover. However, the response of the underlying microbial communities catalyzing

methane production and oxidation have so far received little attention. We hypothesize that

with the return of Sphagnum post-harvest, methane turnover potentials and the

corresponding microbial communities will converge in a natural and restored peatland. To

address our hypothesis, we determined the potential methane production and oxidation

rates in a natural (as a reference), actively mined, abandoned, and restored peatland over

two consecutive years. In all sites, the methanogenic and methanotrophic population size

were enumerated using qPCR assays targeting the mcrA and pmoA genes, respectively. Shifts

in the community composition was determined using Illumina MiSeq sequencing of the

mcrA gene, and a pmoA-based t-RFLP analysis, complemented by cloning and sequence

analysis of the mmoX gene. Peat mining adversely affected methane turnover potentials, but

rates recovered in the restored site. The recovery in potential activity was reflected in the

methanogenic and methanotrophic abundances. However, the microbial community

composition was altered, more pronounced for the methanotrophs. Overall, we observed a

lag between the recovery of the methanogenic/methanotrophic activity and the return of

the corresponding microbial communities, suggesting a longer duration (>15 years) is

needed to reverse mining-induced effects on the methane-cycling microbial communities.

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Importance

Ombrotrophic peatlands are a crucial carbon sink, but this environment is also a source of

methane, an important greenhouse gas. Methane emission in peatlands is regulated by

methane production and oxidation, catalyzed by the methanogens and methanotrophs,

respectively. The methane-cycling microbial communities have been documented in natural

peatlands. However, less is known of their response to peat mining, and the recovery of the

community after restoration. Mining exerts an adverse impact on the potential methane

production and oxidation rates, as well as the methanogenic and methanotrophic population

abundances. Peat mining also induced a shift in the methane-cycling microbial community

composition. Nevertheless, with the return of Sphagnum in the restored site after 15 years,

methanogenic and methanotrophic activity, as well as population abundance recovered well.

The recovery, however, was not fully reflected in the community composition, suggesting >

15 years is needed to reverse mining-induced effects.

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Introduction

Peatlands play a crucial role in the global carbon budget. Although peatlands are a net

carbon sink, northern ombrotrophic peatlands are also a source of methane, a potent

greenhouse gas with a 34-fold stronger radiative forcing effect than carbon dioxide in a 20-

year time scale (1). Peatlands, along with other wetlands contribute approximately 23% of

the total methane budget of 500-600 Tg per annum (2). Methane released from peatlands

would be higher if not for the methanotrophs acting as a natural filter at oxic-anoxic

interfaces to mitigate methane emission (3,4,5); it has been estimated that up to 90% of

biologically produced methane are consumed before being released into the atmosphere in

this environment (6). Net methane released is thus regulated by methane production and

oxidation. Methane-cycling processes, along with the carbon sink function of peatlands can

be significantly affected by peat drainage and mining.

Mining alters the peat physico-chemical properties, increasing the pH and humified

materials by exposing highly decomposed peat, and causes nutrient deficiency for the

microorganisms by removing inorganic compounds (e.g. P, K, and Na), which in turn, exerts a

response in the methane-cycling microorganisms with consequences for methane

production and oxidation (3,4). Depending on the mining method (block-cut and vacuum-

harvested peat) and duration after restoration, these effects can be persistent over

prolonged periods (up to three decades; 4). Among the criteria suggested for monitoring

peat restoration include surveys of biogeochemical cycles, hydrology, nutritional and

chemical status, and microbiological functioning (3,7). While the response of methane-

cycling processes, as well as peat respiration driven by changes in the physico-chemical

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properties and substrate chemistry post-harvest and during restoration have been the focus

of previous studies (e.g. 3,4,8), less is known of the response of the microorganisms

mediating these processes. In particular, the microbial biogeochemistry regulating methane

emission during Sphagnum-dominated peat restoration have so far received little attention

(9,10,11).

Ombrotrophic Sphagnum-dominated peatlands are characterized by relatively inhospitable

conditions (e.g. low pH, nutrient poor, antimicrobial properties of Sphagnum; 12,13). These

conditions are in part engineered by the Sphagnum, and likely favored the proliferation of

some methanogens and methanotrophs (13). For instance, methane is primarily produced

by hydrogenotrophic methanogenic archaea, rather than aceticlastic ones in ombrotrophic

peatlands (14,15,16,17,18). These include members of the genera Methanoregulaceae and

‘Candidatus Methanoflorentaceae’ which are predominant, and to a lesser extent,

Methanosarcinaceae (17,19). The methanotrophs in natural peatlands have been well

documented. Using stable isotope labeling approaches and molecular analyses targeting

gene transcripts, the alphaproteobacterial methanotrophs (Type II subgroup) have generally

been shown to form the vast majority of metabolically active methane-oxidizers in many

peatlands; gammaproteobacterial methanotrophs (Type Ia and Ib subgroups) seem to form a

minor fraction of the total active community (12,20,21). Ecologically relevant active

alphaproteobacterial methanotrophs in peatlands include the Methylocystis-Methylosinus

group, as well as members belonging to the family Beijerinckiaceae (Methylocella,

Methyloferula, and Methylocapsa) which are phylogenetically distinct from Methylocystis

and Methylosinus (12). In particular, Methylocystis and Methylosinus have been suggested to

possess ecological traits to survive under unfavorable conditions (22). Moreover,

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diazotrophic methanotrophs play an important role in peatlands, fixing atmospheric N2 to

assimilable nitrogen forms and can contribute > 33% of new N input in developing peatlands

(13,23,24). Hence, diazotrophs are also a relevant microbial guild in peat environments.

Because of the oligotrophic conditions in ombrotrophic peat, many of these microorganisms

are slow-growing, and resist isolation in the laboratory. To circumvent cultivation,

quantitative and qualitative culture-independent approaches targeting the structural genes

of methanogens (i.e., mcrA encoding for the α-subunit of the methyl-coenzyme M

reductase) and methanotrophs (i.e., pmoA encoding for the A subunit of the particulate

methane monooxygenase and mmoX encoding for the α-subunit of the soluble methane

monooxygenase) have been employed to characterize peat-inhabiting methane-cycling

microbial communities (20,25,26,27).

Although mining alters the peat physico-chemical properties, with the return of Sphagnum

and conditions approximating the natural site, we hypothesized that the methanogenic and

methanotrophic community compositions will converge, and the corresponding process

rates will become more similar in the natural and restored sites than in an actively mined or

abandoned site. To address our hypothesis, we determined the potential methane

production and oxidation rates in an active peat mine, drained and abandoned mine, and

restored mine to elaborate the impact of peat mining on methane turnover. A natural (un-

disturbed) peatland, dominated by Sphagnum sp. served as a reference for comparison to

the other sites. Comparing the natural and abandoned/restored sites thus presents an

opportunity to determine the effect of peat mining on methane turnover, and the response

of the associated microbial communities. We enumerated the structural genes, that is, mcrA

and pmoA genes to relate the abundances of the methanogens and methanotrophs to total

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methane produced and oxidized, respectively in the same season over two consecutive

years. In addition, we followed the response of the methanogenic and methanotrophic

community composition, respectively using high throughput amplicon sequencing of the

mcrA gene and a pmoA-based t-RFLP complemented by cloning and sequencing of the

mmoX gene. Our efforts were aimed to determine the impact of peat mining and restoration

on methane turnover, and the response of the methane-cycling microorganisms.

Materials and Methods

Site description and sampling

The sampling sites are regarded as ombrotrophic peatlands as water and nutrients are rain-

fed, and the sites are isolated from the groundwater (Table 1). The abandoned sites were

dammed post-harvest since 2004 and 2009, but water naturally drained from these sites,

and vegetation shifted towards shrub land/forest (Table 1). The restored site was dammed

and remained water-logged since 2000. Like in the natural site, Sphagnum sp. predominate

in the restored site. Besides damming to prevent water outflow, no other restoration

intervention was introduced. The natural peatland, considered as the reference site, was an

undisturbed site and was declared a nature reserve since 1962. In addition, peat was also

sampled from a drained actively mined site managed by Greenyard Horticulture, Poland.

Peat was extracted close to the water table in the actively mined, abandoned, and restored

sites using the “block peat” or surface milling method (Table 1).

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Peat material was collected in mid-August 2015 and mid-June 2016 at the same sampling

sites. The atmospheric temperature recorded at the time of sampling was in the range of

30OC– 35OC and 27OC-32OC in 2015 and 2016, respectively. Peat material was sampled from

the surface peat layer (0-10 cm) from four or five randomly selected plots in each site. The

plots were spaced > 2 m apart, and were considered to be independent from one another.

Three cores (diameter x height; 3.5 cm x 10 cm) were collected from each plot, and

composited after separating the upper 0-5 cm from the subsequent 5-10 cm in 2015. In

2016, the cores were composited without distinguishing the 0-5 and 5-10 cm intervals.

During sampling, vegetation was collected from ~1 m x 1 m grids from which the samples

were collected, and delivered to the laboratory to be identified. In the natural and restored

sites where Sphagnum predominates, Sphagnum as well as the overlaying water were

sampled. Water from an adjacent ditch to the actively mined site was also collected. All

samples were transported to the laboratory in Styrofoam boxes containing cooling blocks.

Samples were immediately processed upon arrival for incubation (in 2015), or kept in the

4OC fridge for two days before incubation (in 2016).

Microcosm setup to determine methane production and uptake rates.

In the laboratory, roots, stones, and other debris were removed by hand from the samples

before incubation. Each microcosm consisted of 7.5 g fresh weight peat in 120 ml bottle

capped with a butyl rubber stopper. To determine methane production rate, microcosm was

flushed with N2 for 30 mins before incubation. To determine methane uptake rate, methane

was added into the microcosm to achieve a headspace concentration of approximately 1-2 v/v

%. The microcosm was shaken (120 rpm) during incubation at 25oC for 7 days. In parallel,

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approximately 5 g of the same material was oven-dried at 65oC until weight remained

constant (approximately 2 weeks) to determine the gravimetric water content, later used to

normalize methane production and uptake rates to per gram dry weight material.

Sphagnum was rinsed with deionized water and left on a paper towel to drain excess water

before incubation. Sphagnum (0.25-0.30 g fresh weight) was placed in a 120 ml bottle, and

headspace methane was adjusted to approximately 1 v/v%. Incubation was performed under

oxic condition on a shaker (120 rpm) next to a window receiving natural light in the

laboratory at room temperature (22oC-24oC). As such, light intensity may be sub-optimal

during incubation. A portion of the same Sphagnum was oven-dried at 65OC until weight

remained constant (approximately 2 weeks) to determine the gravimetric water content

which was used to normalize methane uptake rate. The peat water fraction (5 ml) was

incubated oxically under an initial headspace methane of approximately 1 v/v% in a 120 ml

bottle. Incubation was performed at 25OC while shaking (120 rm) in the dark. In the peat

incubation, headspace methane concentration was monitored daily for 7 days, while in the

Sphagnum and peat water incubations, headspace methane was followed over 12-10 days.

After incubation, samples were collected and stored in the -20OC freezer till further analyses.

Headspace methane measurement and soil nutrient content.

Headspace methane was determined using an Ultra GC gas chromatograph (Interscience,

Breda, the Netherlands) equipped with a flame ionization detector (FID). Methane

production and uptake rates were respectively determined linearly from methane increase

or decrease during incubation. In particular, the rate of methane production was determined

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after a lag period (1-3 days), which may not reflect in-situ production rate. Methane was

consumed immediately (< 1 day lag period) during oxic incubation. Peat pH was determined

in 1 M KCl (1:5). The nutrient (NOx, NH4+, and PO4

3-) contents in the peat were determined

using a Seal QuA Atro SFA autoanalyzer (Beun-de Ronde B.V., Abcoude, the Netherlands) in 1

M KCl (1:5), as described before (28). NOx is the total of NO2- and NO3

-. The total C and N

content were determined using the Flash EA1112 CN analyzer (ThermoFischer Scientific,

Breda, the Netherlands) after processing the peat as described before (29).

DNA extraction and qPCR assays.

Peat was randomly selected from three plots per site (starting material), and after oxic and

anoxic incubation of the same starting material for DNA extraction. DNA was extracted using

the PowerSoil® DNA Isolation kit (MOBIO, Uden, the Netherlands) according to the

manufacturer’s instructions.

We performed qPCR assays targeting the mcrA, pmoA, and nifH (encoding for the H subunit

of the nitrogenase) genes as proxies to enumerate the abundance of the methanogens,

methanotrophs, and diazotrophs (nitrogen-fixers), respectively in the starting material. In

addition, we performed a qPCR assay specifically targeting the pmoA gene of type II

methanotrophs. The mcrA, pmoA, and type II pmoA qPCR assays were also applied to the

peat after the incubations. The qPCR assays were performed in duplicate per DNA extract,

yielding a total of six replicate per target gene, site, year, and time (before and after

incubation). The primers, primer concentration, and PCR thermal profiles for each assay is

given in Table 2. The qPCR reaction for all assays consisted of 10 µl 2X SensiFAST SYBR

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(BIOLINE, Alphen aan den Rijn, the Netherlands), 3.5 µl of forward and reverse primers each,

1 µl bovine serum albumin (5 mg ml-1; Invitrogen, Breda, the Netherlands) and 2 µl diluted

template DNA, with an exception; in the qPCR assay targeting the nifH gene, 2.5 µl of

forward and reverse primers each were added into the reaction. DNase- and RNase-free

water (Sigma-aldrich Chemie NV, Zwijndrecht, the Netherlands) was added to reach a final

volume of 20 µl, if needed. In a pilot qPCR run, 10-, 50-, and 100-fold dilutions of template

DNA extracts from each site were performed to determine the DNA concentration yielding

the optimal copy numbers of the target genes. Subsequently, DNA extract was diluted 10-

fold and 100-fold for the qPCR assays targeting the nifH and type II pmoA genes, and pmoA

gene, respectively. DNA extract was diluted 10-fold (natural, actively mined, and abandoned

site since 2009), 50-fold (abandoned site since 2004), and 100-fold (restored site) for the

qPCR assay targeting the mcrA gene. The addition of BSA and dilution of template DNA were

performed to relief PCR inhibition. Plasmid DNA isolated from pure cultures were used as

standards for the calibration curve. The qPCR assays were performed using a Rotor-Gene Q

real-time PCR cycler (Qiagen, Venlo, the Netherlands). The specificity of the amplicon was

verified from the melt curve, and further confirmed by 1% gel electrophoresis showing a

single band of the correct size in the pilot qPCR run.

mcrA gene amplification and sequencing

Amplification of the mcrA gene for sequencing, targeted using the mlas/mcrA-rev primer

pair, was performed after Herbold et al., 2015 (33) using a two-step barcoding approach. In

the first PCR, the target gene was amplified with diagnostic primers synthesized with a 16 bp

head sequence 5’-GCTATGCGCGAGCTGC--3’. In the second PCR, amplicon from the first PCR

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was re-amplified with primers that consist of the 16 bp head sequence and included at the 5’

end, a library-specific 8 bp barcode. Each PCR reaction (total volume: 20 µl in the first PCR

and 50 µl in the second PCR) consisted of I x Taq buffer with 1.8 mM MgCl (Roche

Diagnostics, Almere, Netherlands), 0.2 mM dNTPmix (Roche Diagnostics), 0.025 U Taq DNA

polymerase (Roche Diagnostics), 0.2 mg/ml bovine serum albumin (Invitrogen, Breda, the

Netherlands), 1 µM of forward and reverse primers each, and 1 µl of template. When

needed, DNAase- and RNase-free water (Sigma-aldrich Chemie NV, Zwijndrecht, the

Netherlands) was added to achieve the total volume. The PCR thermal profile included an

initial denaturation step at 95OC for 5 mins, followed by 40 cycles (for the first PCR) and 15

cycles (for the second PCR) of 95°C for 1 min, 60OC (for first PCR) or 53°C (for second PCR) for

1 min, and 72°C for 1 min. The final elongation step was 72°C for 5 min. The first PCR

reaction was performed in duplicate, screened by 1% gel electrophoresis, and pooled for use

as a template in the second PCR, which used only one primer (5’-BARCODE-HEAD-3’).

Amplicon from the second PCR was also screened by 1% gel electrophoresis. PCR product

was purified using the QIAquick Purification kit (Qiagen, Venlo, Netherlands), and the

concentration was determined using the Fragment Analyzer (Advanced Analytical,

Heidelberg, Germany). Equal amounts of the purified PCR products were pooled (10 ng/ul)

prior to sequencing. High-throughput sequencing was performed by LGC Genomics (Berlin,

Germany) using Illumina MiSeq V3 chemistry generating 300 bp paired-end reads.

pmoA-based terminal restriction fragment length polymorphism (t-RFLP)

Detailed methodology for the pmoA-based t-RFLP had been described elsewhere (34).

Briefly, the pmoA gene was amplified from each DNA extract (n=3) from peat sampled in

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2015 with the primer combination A189f/Mmb661r. The forward primer A189f was FAM-

labelled. The amplicon was digested using the restriction endonuclease MspI, and the t-RFs

were separated using the ABIPrism 310 (Applied Biosystems, Darmstadt, Germany). The

GeneScan 3.71 software (Applied Biosystems, Darmstadt, Germany) was used to determine

the length of the t-RFs by comparison with an internal standard (MapMarker 1000,

Bioventures, Murfreesboro, USA). In silico comparative gene sequence analysis with an

extensive pmoA clone library was performed to bin and assign the t-RFs as described in

detail before (34,35,36). Unidentifiable t-RFs (i.e. tRFs 69, 242, and 143) and t-RFs

comprising of < 2% of the total peak area were excluded from the t-RFLP analysis.

Cloning and sequencing of the mmoX gene

The mmoX clone libraries were constructed from the DNA extract of all peat material

sampled in 2015 using the primer pair mmoX-206F/mmoX-886R with primer concentrations

and PCR thermal profile as given in Table 2 according to Liebner and Svenning (2013) (27).

The PCR reaction consisted of 25 µl Masteramp 2X PCR Premix F (Epicentre Illumina, WI,

USA), 5 µl bovine serum albumin, BSA (5 mg ml-1; Invitrogen, Breda, the Netherlands), 0.5 µl

of each primer, 0.5 µl of TAQ Polymerase (ThermoFischer Scientific, Uden, Netherlands), 1 µl

of template DNA, and 17.5 µl DNAase- and RNase-free water (Sigma-aldrich Chemie NV,

Zwijndrecht, the Netherlands), giving a final volume of 50 µl. The PCR amplicon was verified

on 1% agarose gel electrophoresis. Three PCR reactions were performed for each DNA

extract, and subsequently pooled per site during the PCR clean-up step using the

GenEluteTM Gel Extraction Kit (Sigma-aldrich Chemie NV, Zwijndrecht, the Netherlands).

The purified PCR amplicon was cloned into the pGEM-T vector (Promega, Mannheim,

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Germany) and transformed into competent cells (E.coli JM109; Promega, Mannheim,

Germany). The transformants were screened (blue-white colonies) and confirmed by colony

PCR using the primers T7/M13rev_29, before sequencing. Sequencing was performed at

ADIS (MPI for Plant Breeding, Cologne, Germany). Sequences were assembled using the

SeqMan software (DNA-Star software package, Lasergene, USA) whereby the vector

sequence was trimmed. Sequences were compared and identified against the GenBank

database using the NCBI BLASTn function. Sequences were deposited at the European

nucleotide archive (EMBL-ENA) under the project accession number PRJEB22776.

Statistical analysis

Significant differences (p<0.01) in the physico-chemical parameters (n=3) and process rates

between sites per year (n=4 or 5) were determined by analysis of variance, ANOVA using

SigmaPlot version 13.0 (Systat Software Inc, USA).

The mcrA sequencing reads were assembled using the ‘make.contigs’ command in Mothur

version 1.3.3 (37). The assembled contigs were then sorted, and the barcodes and primers

were removed. Putative chimeric reads were removed using USEARCH 6.0, and the

nucleotide sequences were translated into amino-acids and frame shifting reads were

corrected using the FRAMEBOT program in FunGene Pipeline, RDP (38;

http://fungene.cme.msu.edu/FunGenePipeline). The inferred amino acid sequences were

aligned and clustered into Operational Taxonomic Unit (OTU) with an 89% identity cutoff

value, which have been considered to be species-level affiliation for the methanogen (39).

Representative mcrA nucleotide sequences were assigned to their taxonomic affiliations by

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BLASTn comparisons to GenBank nonredundant (nr) database, and the results were then

imported to MEGAN software version 5.11.3 with the Lowest Common Ancestor (LCA) and

default parameters. As with the t-RFs of the pmoA gene (after normalization to the overall

signal intensity), constrained correspondence analysis (CCA) for the mrcA gene sequences

were produced in the R statistics software environment (40) using the package Vegan

version 2.3.0 (41). The measured environmental parameters (NH4+, NOx, PO4

-3, and pH) were

used as constraints in the CCA. The mcrA gene sequences were deposited at the European

nucleotide archive (EMBL-ENA) under the project accession number PRJEB22766.

Results

The abiotic environment, and methane production and oxidation potentials.

The peat material generally showed consistent physico-chemical properties and methane

turnover trends between sites over two years. Given that the peatlands were located within

the same area, we do not anticipate large temperature or climatic variations between sites,

affecting the process rates. Discrepancies in physico-chemical parameters and methane

turnover rates between sites are thus largely attributable to the impact of peat mining and

restoration.

Peat physico-chemical properties were largely comparable in 2015 and 2016, but values

significantly varied between sites. Peat pH was altered during restoration, showing

significantly higher values (p < 0.01; Table 3) in the actively mined (pH ~ 3.5), and abandoned

(pH ~ 4.8) sites when compared to the natural and restored sites (pH 2.7 - 2.8). More

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pronounced for total N, both total N and C were significantly higher in the abandoned sites,

which resulted in significantly lower C:N ratio in these sites (Table 3). While NO x (total of

NO2- and NO3

-) varied between sites, NH4+ concentration was significantly higher particularly

in the actively mined site (Table 3). PO43- concentration was significantly higher in the

restored site in 2015, but because of high variability, the trend was not replicated in 2016

(Table 3). With the exception of NH4+ and PO4

3- concentrations in the restored site in 2015,

the physico-chemical properties in the natural and restored sites were comparable. Likewise,

the physico-chemical properties in both the abandoned sites were not significantly different.

The potential methane production and oxidation rates were initially determined in the 0-5

and 5-10 cm peat intervals in 2015. With the exception of the methane production rate in

the abandoned site since 2004, both methane production and oxidation rates were

comparable in these layers (Figure S1). Therefore, peat from the upper 10 cm were

homogenized and incubated as a single layer in 2016. Although values differed, methane

production rate was consistently higher in the restored site sampled in 2015 and 2016.

Appreciably higher methane production rate was detected after a lag of one day in the

restored site in 2015; in 2016, methane production was detected only in this site (Figure 1).

The lower methane production in 2016 may be attributable to the sampling time (June), in

conjunction with the beginning of the vegetation growing season (May), as documented

before (15). Consistent in peat material sampled in 2015 and 2016, methane oxidation rate

was significantly higher in the natural and restored sites, while values in the actively mined

and abandoned sites were comparably low (Figure 1). Methane uptake was immediately

detected in the natural and restored sites during incubation, indicating the presence of a

readily active population of methanotrophs in these peatlands. A lag of 1-2 days was

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detected in the other peat samples. Sphagnum and peat water exhibited comparable

methane oxidation rates in 2015 and 2016, but values were lower than in the peat material

(Figure S2).

The mcrA, pmoA, and nifH gene abundances.

The mcrA and pmoA genes were enumerated using qPCR assays to be used as proxies for

methanogenic and methanotrophic abundances, respectively. Because of the predominance

of alphaproteobacterial methanotrophs in peatlands (12), we also target the pmoA gene of

this group using the type II qPCR assay (Table 2; 31). The mcrA gene abundance was up to

four orders of magnitude higher in the natural and restored sites than in the other sites

which showed comparable values (Figure 2a). The trend was similar, but less pronounced for

the pmoA gene, exhibiting significantly higher gene abundances in the natural, actively

mined, and restored sites than in the abandoned sites. However, the pmoA gene abundance

in the restored site sampled in 2016 was highly variable, and values were not significantly

different from the abandoned site since 2004 (Figure 2b). Although the type II pmoA gene

abundances were relatively higher in the peat sampled in 2016, values were generally

comparable across sites with an exception. In 2015, peat sampled from the restored site

harbored a significantly higher abundance of the type II pmoA gene than in the other sites

(Figure 2c). Moreover, the nifH gene was enumerated to monitor the abundance of the

diazotrophs during peat restoration. The nifH gene was detected in significantly higher

abundance in the natural and restored peatlands than in the abandoned sites (Figure 2d).

Generally, the abundance of the targeted genes showed comparable trends in peat sampled

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from 2015 and 2016, in that, values were significantly higher in the natural and/or restored

peatlands, while targeted gene copy numbers were lower in the abandoned sites (Figure 2).

The mcrA and pmoA gene diversity.

The methanogenic and methanotrophic community composition were characterized based

on the mcrA and pmoA gene amplicons, respectively. The mcrA gene sequence analysis

revealed the presence of 15 OTUs affiliated to the methanogens; one OTU could not be

assigned, but this comprised of a minor fraction of the total number of reads in all samples

(<0.5%) (Figure S3). Among the methanogen-affiliated OTUs, Methanoregula-affiliated OTU

was overwhelmingly represented in the natural (~95% relative abundance) and restored

sites (~75% relative abundance), and comprised of ~45% relative abundance of the total

methanogenic community in the actively mined site (Figure S3). While unclassified

Methanomicrobiales- and Methanosarcina- affiliated OTUs were predominant in the

abandoned site since 2009 (~91% relative abundance), these OTUs along with

Methanomassiliicoccus- and Methanolinea-affiliated OTUs form the majority of sequences

detected in the abandoned site since 2004 (~83% relative abundance; Figure S3).

Discrepancy in the methanogenic community composition between sites was visualized as a

correspondence analysis using the environmental variables (NH4+, PO4

3-, NOx, and pH) as

constraints (Figure 3), with pH being the only parameter significantly (p<0.01) affecting the

community composition in all sites. The methanogenic community composition in both the

abandoned sites grouped together, and could be clearly distinguished from the community

composition in the natural and restored peatlands, which separated along CCA axis 1 (Figure

3). The community composition in the natural and restored sites clustered closely together,

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but did not overlap. There appears to be a shift in the methanogenic community

composition after peat mining and abandonment, and the return to a community structure

resembling the natural peatland after restoration.

Although we were able to retrieve pmoA amplicons in the first PCR cycle, re-amplification of

an aliquot of the first PCR reaction with barcoded primer was unsuccessful, even after

optimization steps. Therefore, we characterized the diversity of the pmoA gene using t-RFLP.

The assignment of the t-RFs was based on comparison to a comprehensive pmoA clone

library (> 5000 clones; 36). After standardization, and normalization of the t-RFLP profiles to

overall signal intensity as described in Lüke et al. (2010; 2014) (35,36), major identifiable t-

RFs were t-RF 349 (affiliated to type Ia methanotrophs and pmoA2 ), t-RF 514 (affiliated to

the type Ia methanotroph, Methylobacter), t-RF 240 (affiliated to type Ia methanotrophs), t-

RF 225 (affiliated to type Ib methanotrophs), t-RF 74 (affiliated to the type Ib

methanotrophs, Methylocaldum and Methylococcus), and t-RF 245 (affiliated to type II

methanotrophs). The t-RFLP analysis revealed a predominance of t-RF 245 affiliated to type

II methanotrophs in all sites (40-50% of total community composition), and was most

pronounced in the restored and abandoned site since 2004, comprising of more than 80%

relative abundance (Figure S3). Differences in the methanotrophic community composition

was further determined by a correspondence analysis using measured environmental

variables (NH4+, PO4

3-, NOx, and pH) as constraints, which was not significantly correlated to

the community composition (p > 0.01) together explaining only 20.3% of total variance

(Figure 3). Because of the large variability between and within sites, no clear separation of

the community composition was discernable (Figure 3). However, the community

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composition in the natural and abandoned site since 2009 tended to separate along CCA axis

1 (Figure 3).

As some methanotrophs possess only the mmoX gene (e.g. Methylocella, Methyloferula; 12),

we determined the composition of the mmoX-harbouring methanotrophs using cloning and

sequence analysis. Besides Methylocella-like and Methyloferula-like methanotrophs, the

mmoX gene sequences detected were affiliated to Methylocystis and Methylosinus, together

forming the majority of alphaproteobacterial methanotrophs in all sites (Figure S4). Hence,

the t-RFLP, and cloning and sequence analysis of the mmoX gene were in agreement with

the qPCR analysis which showed numerical abundance of the type II pmoA gene.

Discussion

Methane dynamics and the abiotic environment post-harvest

The surface peat layer is a dynamic area where rapid changes in physico-chemical conditions

occur (42), significantly affecting the methane production and oxidation rates, as well as

respiration (8,17,43). The methanotrophs localized at the surface peat, as well as in the

Sphagnum, and planktonic methanotrophs in the overlaying water play a crucial role as a

methane bio-filter, consuming methane before release into the atmosphere (4,5). However,

methane can escape via ebullition bypassing the methanotrophs. Edaphic factors controlling

methane production were found to be significantly correlated to methane oxidation,

suggesting that methane oxidation was primarily fueled by substrate availability and was

dependent on methanogenesis (3,4,17). Therefore, sites producing methane is anticipated

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to show proportional methane oxidizing potential. This appeared to be the case in the

actively mined, abandoned, and restored sites; in the natural peatland, low methane

production did not correspond with the significantly higher methane oxidation rate detected

(Figure 1). The physico-chemical properties of the natural peatland and restored site were

largely similar (Table 3), hence disproportional methane production and oxidation activities

in the natural site is likely caused by differences in the biotic components e.g.

methanotrophic community composition and/or abundances (see below). Moreover, we

cannot exclude seasonal variation affecting the methane production and oxidation rates

caused by fluctuations of the water table layer. Nevertheless, we showed that trends in

methane turnover potentials were at least consistent during summer over two years.

In the actively mined and abandoned sites, methane production was low or was not

detected post-harvest. Mining and abandonment significantly reduces substrate quality and

bioavailability by exposing the humified fraction and already decomposed peat material

(3,8). Not surprisingly, methane turnover in these sites were significantly lower than in the

restored sites where Sphagnum naturally revegetated more recently than in the natural site,

which provides a supply of readily metabolized organic substrate (Figure 1). Besides

affecting the availability of organic substrates, mining also removes inorganic nutrients (e.g.

Mg, Na, P), causing nutrient limitation which appears to affect the rates of processes

dependent on the nutrients (e.g. Na limiting methane production; 4,44). Substrate and

micronutrient deficiencies caused by mining, coupled to the drier condition and hence better

aeration (gravimetric water content; 68-81%, as opposed to 89-91% in the natural and

restored sites) in the surface peat after drainage likely attributed to the significantly lower

methane production potential in the actively mined and abandoned sites (Figure 1). The

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abandoned sites were also characterized by significantly higher pH, and total carbon and

nitrogen concentrations (Table 3), with the markedly higher total nitrogen content

decreasing the C:N ratio, indicating readily decomposable material (45). However, the use of

the C:N ratio as a proxy for decomposability is disputable (29), and the total C and N

determined here also include the recalcitrant fractions not bioavailable to the

microorganisms. Therefore, mining alters the peat physico-chemical characteristics, but

methane-cycling processes were restored to even higher values than in the natural peatland

after 15 years.

Response of the methane-cycling microbial community abundance to mining and restoration.

Peat mining constraints the total microbial biomass by reducing the bioavailability of

substrate and nutrients (3,4,46). In these studies, the total microbial biomass significantly

decreased in the actively mined or abandoned sites, when compared to the natural or

restored sites. Although the total microbial biomass could be correlated to the carbon

dioxide emission (3,4), the methods used to quantify microbial biomass (e.g. fumigation-

extraction technique) were unable to discriminate between microbial groups catalyzing

methane production and oxidation from the total community. Applying qPCR assays

specifically targeting the mcrA and pmoA genes, we enumerated the abundance of the

methanogens and methanotrophs, respectively. Assuming that one methanogen harbors a

single mcrA gene copy (47), and that a methanotroph harbors two pmoA gene copies (48),

mining dramatically decreased the methanogenic population size by around an order of

magnitude, and to a lesser extent in the methanotrophic population (1 x 106 – 1 x 107 cells g

dw-1 depending on the sampling year; Figure 2) when compared to the natural peatland.

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Although changes in the pmoA gene copy numbers were statistically significant,

methanotrophs appeared to be less sensitive to mining and abandonment, with the

alphaproteobacterial (type II) methanotrophic population seemingly increased after

abandonment in 2015 (Figure 2). Regardless, both methanogenic and methanotrophic

populations recovered well in the restored site, showing comparable or even higher mcrA or

pmoA gene copy numbers than in the natural peatland. Similarly, the diazotrophic

abundance, indicated by the nifH gene, significantly decreased after mining, but population

recovered in the restored site. The diazotrophs have been recognized as a key microbial

group in ombrotrophic peatlands, being an important source of assimilable carbon and

nitrogen to sustain Sphagnum growth (23,49,50). Hence, recovery of the diazotrophic

abundance is a relevant indicator for Sphagnum re-vegetation. Overall, more pronounced for

the methanogens, mining compromised the methanogenic, methanotrophic, and

diazotrophic abundances, but population size fully recovered after restoration.

Structural genes appear to correlate well to the activity catalyzed by the respective encoded

enzymes, as has been shown for methanogenesis and methane oxidation (30,51,52,53), as

well as other microbial mediated processes e.g. 2-methyl-4-chlorophenoxyacetic acid, a

herbicide, degradation (54) and N-cycling processes (55). Determining the gene abundances

before and after incubation, we relate the magnitude change in the mcrA and pmoA genes

to the total methane produced and oxidized, respectively (Figure 4). Integrating all replicate

from all sites showing activity, a highly positive significant correlation (p<0.01) was found for

the magnitude of change in the mcrA gene abundance and methane production, suggesting

a coupling of methanogenic growth and activity (Figure 4). Although the magnitude change

in the total pmoA gene abundance was not significantly correlated (p > 0.01; Figure 4) to the

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total methane oxidized during incubation, a significantly positive correlation (p < 0.01) was

detected between the magnitude of change in type II pmoA gene abundance and methane

oxidized (Figure 4). The significant correlation between methane oxidation and type II pmoA

gene abundance, rather than the total pmoA gene abundance suggests the active role of

type II methanotrophs under the incubation conditions. Alphaptroteobacterial

methanotrophs belonging the Methylocystis-Methylosinus group, as well as genera

Methylocella, Methylocapsa, and Methyloferula seemingly form the predominantly active

community in many acidic peat environments (25,27,56), although the presence of active

gammaproteobacterial methanotophs cannot be excluded (43,57).

Response of the methane-cycling microbial community composition to mining and

restoration.

Consistent with previous studies, we detected a predominance of Methanoregula, a genus

represented by hydrogenotrophic methanogens which showed a preference for acidic

condition, in the natural and restored peatlands (11,17,18,19). Methanoregula-like

methanogens decreased with mining and abandonment, while the relative abundance of

Methanosaeta-like and Methanosarcina-like methanogens increased. Methanosaeta and

Methanosarcina are closely related belonging to Methanosarcinales, with Methanosaeta

comprising of aceticlastic methanogens while members of Methanosarcina (e.g. M. barkeri)

show metabolic flexibility, capable of hydrogenotrophic and aceticlastic methanogenesis

(58,59). The apparent shift from a predominance of hydrogenotrophic to aceticlastic

methanogens suggests a physiological change presumably driven by nutritional status,

including substrate quality and availability, and vegetation type following peat mining.

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However, the community composition of the methanogens in the natural and restored

peatlands became similar, with Methanoregula-like methanogens dominating the

community (Figure 3), suggesting the recovery of the methanogenic community composition

with Sphagnum revegetation.

The methanotrophic community composition was not statistically distinct between sites but,

a higher mean relative abundance of alphaproteobacterial methanotrophs were detected in

the restored and abandoned site since 2004. Despite comparable methane oxidation rate

and total methanotroph abundance in the natural and restored sites, peat abandonment

and restoration may have structured the methanotrophic community composition over time

(Figures 3 and S3). In contrast, the methanotrophic community composition was found to be

similar in a natural peatland and a site undergoing 10-12 years restoration in a forestry-

drained peat (9). However, it should be noted that the authors targeted the pmoA gene

using a rather specific primer pair (A189f/A621r; 60), limiting the pmoA gene coverage to

alphaproteobacterial methanotrophs exclusively. Regardless, as shown by the t-RFLP

analysis and supported by the qPCR assays, all sites harbored predominantly

alphaproteobacterial methanotrophs, thought to be the active population in widespread

peat environments (13).

Conclusion

Supporting our hypothesis, results showed that methane turnover potentials and the

associated microbial abundance recovered post-harvest with the return of Sphagnum, but

we found less compelling evidence for the converge of the methane-cycling microbial

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community composition in the natural and restored sites. A shift in the microbial community

composition may alter the collective traits of the contemporary community (61,62,63), with

implications for their response to future disturbances. Our findings could be further

substantiated by field-based flux measurements in future studies.

Acknowledgements

We are grateful to Iris Chardon and Marion Meima-Franke for excellent technical assistance.

We thank Marcin Harnisz (Greenyard Horticulture, Poland) for his invaluable help during

sampling. AH was financially supported by the BE-Basic grant F03.001 (SURE/SUPPORT). This

publication is publication nr. XXX of the Netherlands Institute of Ecology.

All authors have seen and approved the final version submitted. All authors declared no

conflict of interest.

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Figure legends

Figure 1: Methane production (a) and uptake rates (b) in peat sampled in 2015 and 2016

(mean ± s.d; n = 4 or 5). Methane production was not detected in the natural peatland,

actively mined, and abandoned sites in 2016. These sites were not included in the analysis of

variance. Upper and lower case letters indicate the level of significance (p<0.01) in methane

production and uptake rates between sites per year. Note the different scale along the y-

axis.

Figure 2: Numerical abundance of the mcrA (a), pmoA (b), type II pmoA (c), and nifH (d)

genes in peat sampled in 2015 and 2016 (mean ± s.d; n = 6). Two qPCR reactions were

performed for each DNA extract (n = 3), yielding a total of six reactions per target gene, site,

and year. The lower detection limit was approximately 103-104 copy number of target

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molecules g dw-1, depending on the qPCR assay. Upper and lower case letters indicate the

level of significance (p<0.01) between sites per year for each target gene.

Figure 3: Correspondence analysis showing the response of the methanogenic (a) and

methanotrophic (b) community composition to peat abandonment and restoration post-

harvest. The methanogenic and methanotrophic community composition were derived from

Illumina MiSeq amplicon sequencing of the mcrA gene and a pmoA-based t-RFLP analysis,

respectively. The analysis was performed for each DNA extract (n=3) of the starting material

sampled in 2015 per site. The distribution and affiliation of the OTUs and t-RFs for the mcrA

and pmoA genes are respectively given in Figure S3. Green triangle: natural peatland; dark

blue diamond: actively mined site; light blue inverted triangle: abandoned site since 2009;

grey square: abandoned site since 2004; red square: restored site.

Figure 4: Correlation between total methane produced (a) and consumed (b,c) and the

magnitude change in the mcrA and pmoA gene abundances, respectively, during incubation

(10 days). Replicate from all sites showing methane production and oxidation were

incorporated into the correlation. The magnitude change in the mcrA and type II pmoA gene

abundances were significantly correlated (p<0.01) to the total methane produced and

oxidized, respectively. Note the different scales in the x- and y-axes

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Table 1: Description of sampling sites. Identification of vegetation coverage determined during 2016 sampling, courtesy of Rutger Wilschut, Wageningen, The Netherlands.

Sample(year since

abandonment)

Coordinates(sampling

site)

Depth of peat deposits

(m)

Water table layer

(m below surface)β

Depth of peat

harvested (m)

Excavation method

Post-excavation restoration activity

Characteristic vegetation coverage

Natural peatland(reference)

53°54’24”N,19°41’41”E

(Zielony Mechacz)

4.00 Higher or equal to the litter

layer

n.a. n.a. n.a.# Sphagnum sp. (S. fimbriatum, S. flexuosum, S. fallax), Orthotrichum lyellii.

Actively minedpeatland

54°6’53”N,19°49’35”E(Jozefowo)

4.40 ~1.5 2.06 “Block peat” method, followed by surface milling.

n.a. No vegetation

Abandoned peatland (since 2009)

54°6’41”N,19°49’34”E(Jozefowo)

1.95 0.7 - 1.5 1.65 Surface milling. Site was dammed to prevent water outflow*, and leveled. No vegetation was introduced.

Phragmites australis,Salix sp., Betula pendula, Asteraceae, Agrostis sp.

Abandoned peatland (since 2004)

54°7’30”N,19°49’35”E(Jozefowo)

3.40 0.7 - 1.5 3.10 “Block peat” method, followed by surface milling.

Site was dammed to prevent water outflow*. No vegetation was introduced.

Cerastium fontanum, Phragmites australis, Equisetum arvense, Agrostis sp.

Restored peatland(since 2000)

54°15’34”N,19°44’0.4”E(Rucianka)

3.00 Higher or equal to the litter

layer

2.70 “Block peat” method, followed by surface milling.

Site was dammed to prevent water outflow. No vegetation was introduced.

Sphagnum sp. (S. fimbriatum, S. flexuosum, S. capillifolium), Orthotrichum lyellii.

Abbreviations; n.a.: not applicable.#The pristine site was declared a natural reserve since 1962. Prior to this, there was no peat excavation activity.*Although dammed, water did not remain in these sites, and were naturally drained without any intervention.

41

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948949950

β Due to the excavation method used, the ground water table layer is lowered as a result of peat excavation. The water table layer at the time of sampling is provided by Greenyard Horticulture, Poland.

Table 2: Primers, and PCR thermal profile used in this study. Abbreviation; n.a.: not applicable.

Primer combination(forward/reverse)

Primer concentrations(forward/reverse)

PCR thermal profile* Data acquisition

Target gene References

mlas/mcrA-rev 700 nM/875 nM 95OC, 10s; 60OC, 10s; 72OC, 25 s 72OC, 8 s mcrA (30)

A189f/Mb661r 875 nM/875 nM 94OC, 10s; 62OC, 10s; 72OC, 25 s 87OC, 8 s pmoA (28,31)

II223f/II646r 525 nM/525 nM 94OC, 10s; 60OC, 10s; 72OC, 25 s 87OC, 8 s Type II pmoA (31)

IGK3/DVV 375 nM/375 nM 95OC, 10s; 58OC, 10s; 72OC, 25 s 81.5OC, 8 s nifH (32)

mmoX-206F/mmoX-886R 330 nM/330 nM 94OC, 60s; 60OC, 60s; 72OC, 60 s n.a. mmoX (27)

*The PCR thermal profile indicate the temperature and time for denaturation, annealing, and elongation.

42

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956957958959960961962963964965966967968

Table 3: Selected peat physico-chemical properties (mean ± s.d.; n = 3) determined in 2015 and 2016 (in bold). Upper and lower case letters indicate level of significance at p < 0.01 between sites in 2015 and 2016, respectively.

Sites pH# Total N(µg g dw-1)

Total C(µg g dw-1)

C:N NOx (NO2- + NO3

-)#

(mg g dw-1)NH4

+#

(mg g dw-1)PO4

-3#

(mg g dw-1)

Natural peatland(reference)

2.83 ± 0.03A

2.72 ± 0.06a6.30 ± 0.41A

6.22 ± 0.46a404.94 ± 0.65A

408.02 ±11.60a64.37 ± 4.08B

65.74 ± 2.96b0.230 ± 0.165A,B

0.004 ± 0.005a0.390 ± 0.114A

0.014 ± 0.006a0.002 ±0.001A

0.007 ± 0.007a

Actively minedpeatland

3.86 ± 0.35B

3.50 ± 0.12b8.75 ± 0.62B

9.75 ± 1.11a471.71 ± 16.42B

475.85 ± 13.72c54.05 ± 2.90B

49.21 ± 5.49b0.007 ±0.004A

0.076 ± 0.062a,b2.118 ± 0.812B

2.432 ± 0.565b0.013 ± 0.007A

0.023 ± 0.014a

Abandoned peatland (since 2009)

4.83 ± 0.13C

4.75 ± 0.08c18.98 ± 1.25C,D

18.45 ± 1.78b468.26 ± 4.63B

439.64 ±1.81b24.74 ± 1.75A

23.97 ± 2.28a0.178 ± 0.062B

0.058 ± 0.016b0.305 ± 0.070A

0.003 ± 0.000a0.001 ± 0.000A

0.002 ± 0.001a

Abandoned peatland (since 2004)

4.80 ± 0.17C

4.82 ± 0.20c23.74 ±3.34D

21.60 ± 1.57b438.15 ±14.72B

448.98 ± 5.77c,b18.74 ± 3.06A

20.87 ± 1.85a0.047 ± 0.015B

0.040 ± 0.037a,b0.378 ± 0.063A

0.004 ± 0.003a0.001 ± 0.001A

0.002 ± 0.001a

Restored peatland(since 2000)

3.19 ± 0.05B

2.82 ± 0.14a11.58 ± 3.92A,B,C

6.60 ±1.59a436.78 ± 48.45A,B

426.89 ± 4.63a40.78 ± 15.34A,B

67.06 ± 14.50b0.184 ± 0.155A,B

0.043 ± 0.037a,b1.491 ± 0.342B

0.349 ± 0.277a0.042 ± 0.009B

0.065 ± 0.044a

#determined in 1 M KCl (1:5 ratio).

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