Impacts of degraded blanket bog restoration on culturable and non-culturable soil bacterial and fungal biodiversity and spatiality were assessed. Peat sampled from unvegetated peat areas and from 3 restored vegetation classes, remnant vegetation and grass dominated gullies were subjected to culturable microbial enumeration and rDNA based community profiling and pyrosequencing. No significant differences in bacterial and fungal rDNA-based diversity estimates were detected but culturable counts in bare peat were significantly lower. rDNA sequencing highlighted key bacteria and fungi involved in carbon and nitrogen cycling. The investigation provide much needed information on soil microbial community responses to active peatland restoration.
UK peatlands, that cover about 8% of the land area but store 40–50% of the nation’s terrestrial carbon, are increasingly coming under threat as a result of historical land management, long-term anthropogenic N and S deposition and climate change (Holden et al., 2007). Evidence from data gathered on peat physico-chemistry coupled to primary productivity, hydrology and atmospheric chemistry suggest that what are currently regarded as net sinks risk becoming major sources of carbon (Worrall et al., 2009). In this respect, our limited understanding of the identity, distribution and functioning of key soil microbial drivers of inter-linked C and N cycling in degrading UK moorland systems (Artz, 2009; Littlewood et al., 2010) gives particular cause for concern and additionally remains a fundamental barrier to their restoration and sustainable management.
From a microbiological perspective, plant community, soil temperature, pH and hydrological status are of crucial importance in determining microbial community structure and thus C and N cycling activity in northern peatland ecosystems (Artz, 2009). Decomposition and mineralisation of peat by heterotrophic soil bacteria and fungi is mainly limited to the predominantly oxic surface layers of peatlands, the acrotelm, and periodically the underlying mesotelm (Clymo and Bryant, 2008) during intermittent water table drawdown. The permanently saturated and anoxic catotelm, by definition, supports limited microbial decomposer communities but in cases of severe drought will be progressively exposed to increased oxidative decomposition and respiratory flux. Fungi appear to dominate the surface acrotelm either as litter decomposing saprotrophs or root symbiotic mycorrhizal fungi (Thormann and Rice, 2006; Smith and Read, 2008). Within the mesotelm, fungi, including yeasts, facultative anaerobic bacteria and methanogenic archaea have been detected that were able to mobilise recalcitrant carbon sources through oxidative mineralisation or fermentative processes under water saturated conditions. Bacterial communities also show depth- and pH-related stratification but, unlike fungal counterparts, are regarded as being less influenced by plant community structure (Artz, 2009 and references therein). Artz (2009) has highlighted limited but highly promising studies of functional microbial community ecology that have been achieved through rapid developments in molecular DNA/RNA based barcoding techniques.
The Southern Pennines in Northern England hosts one of the most south-westerly extensions of European upland blanket bog but over 70% has been classified as being in a degraded condition and in extreme cases areas are devoid of vegetation cover exposing unconsolidated bare peat that is often gullied to the gritstone bedrock through severe water erosion (Tallis, 1997; Evans and Lindsay, 2010). Severe ecosystem degradation has, in part, been attributed to long-term historical anthropogenic loading of N, S ozone and metals and heightened susceptibility to climate change drivers (Caporn and Emmett, 2009).
Large-scale moorland restoration efforts in the area have involved lime and fertiliser application allowing lowland nurse grass (inc. Festuca and Agrostis spp.) cultivation to stabilise bare peat and subsequent application of seed and heather (Calluna vulgaris) brash or planting to establish dwarf shrub cover and Sphagnum re-introduction (Moors for the Future Partnership, 2008). Key restoration goals are stabilisation and re-vegetation of bare peat to increase biodiversity and recover hydrological function and lost carbon sequestration potential.
The main aim in the first phase of this study at Holme Moss was to identify acrotelm and mesotelm bacterial and fungal communities in a restoration derived vegetation mosaic based on culture-dependent and -independent molecular DNA-based methodology. The two specific objectives were 1) vegetation-dependent enumeration of culturable bacteria and rDNA-based identification of soil bacterial and fungal community and 2) below-ground GIS-based spatial analysis of plant-soil-microbial community interactions in restoration mosaics.
Materials and Methods
The Holme moss study site, originally established in 2006 (Caporn et al., 2007) is located close to the the northern boundary of the Peak District National Park (53º 31’ 59” N; 01º 51’ 29” W; elevation 524 m a.s.l) in the Southern Pennines. Restoration of extensive bare peat areas was initiated in 2008 with sucessful establishment of nurse grass swards and young (2 year-old) heather (Calluna vulgaris L.) seedlings on formerly bare peat. Sampling close to a transmitter mast allowed inclusion of peat supporting a 25 year-old heather dominated stand established following mast erection and site restoration in 1985 (P. Anderson, pers. comm.). In June 2010, three parallel line transects (approx. 300 m) were established in a due northerly direction to core (12mm dia. x 150mm) sample surface (acrotelm/mesotelm) bare peat and peat under vegetation mosaics encompassing 3 different stages of restoration, namely, nurse grass, young heather and 25-year heather. Peat supporting remnant dwarf shrub communities and naturally vegetated gullies were also sampled.
Bacterial and fungal colony forming units (cfu g-1 soil) were enumerated from homogenised cores (n = 3 per vegetation class per transect) via dilution plating on Tryptone soy agar medium or Potato dextrose agar medium with antibiotic selection. DNA was extracted from the middle sample of each triplicate set of core samples using a Powersoil DNA extraction kit. Extracted community DNA was used as a template for polymerase chain reaction (PCR) to enable denaturing gradient gel electrophoresis (DGGE) fingerprinting and sequencing of the bacterial and fungal microbial communities from amplified 16S-rRNA and ITS-rRNA genes, respectively. Amplified DNA was also cloned and randomly selected 16S and ITS clones Sanger sequenced (Hirsch et al., 2010). 16S and ITS templates were also subjected to high-throughput pyrosequencing (Roche 454) to enable massively high sequence coverage (Hirsch et al. 2010). DGGE banding data and 16S and ITS sequences were subjected to phylogenetics coupled to multivariate cluster analyses using appropriate molecular software packages. For spatial interpolation of microbial counts and 16S and ITS richness estimates, point data was modelled on a map using the Inverse Distance Weighting method implemented in ArcGIS.
Results and Discussion
The focus of this study, targeting assessment of blanket bog vegetation restoration, has been on the upper horizons of peat, the acrotelm and mesotelm, supporting plant roots and root associated microbial communities that play a crucial role in primary peatland productivity. Estimated mean counts for both bacteria and fungi were typically within the order of 106 cfu g‑1 soil for all vegetated samples, and transects, except bare peat that supported a mean of 5.6 x 104 and 2.7 x 104 cfu g-1 bacteria and fungi, respectively. Significant reduction in culturable bacterial and fungal counts in non-vegetated bare peat confirmed earlier preliminary findings (Caporn, 2007). The well known “rhizosphere” microbial enrichment response was apparent in the raised count data (Cheng and Gershon, 2007). However, rich and complex DGGE 16S and ITS rDNA banding patterns were observed in all samples. Band ‘richness’ estimations suggested highest bacterial diversity in young and 25-year heather whereas the fungal richness appeared conserved across vegetation and bare peat. Geo-spatial interpolation of bacterial and fungal counts and richness estimates provided a first visualisation of site specific variation. Bacterial richness appeared to be inversely related to fungal richness and lower bacterial and fungal counts were located in bare peat areas compared to other vegetation in the mosaic.
Sanger sequencing of bare and original vegetation generated preliminary information on the differential presence of key phyla of bacteria e.g. Acidobacteria, Actinobacteria, Verrucomicrobia and Cyanobacteria. The former genera have been commonly identified in the periodically oxic mesothelm of Siberian peatlands (Artz, 2009). Acidobacteria have been identified in surface cores of heavy metal enriched moorlands in the Southern Pennines (Linton et al., 2007) that also develop in rhizospheres in mine trailings. Planctomycytes and related Verrucomicrobiales have been shown to be methanotrophs in the oxic acrotelm. Actinobacteria detected are likely to be involved in litter decomposition along with fungi in the oxic acrotelm. Fungal diversity encompassed three of the five subdivisions, Zygomycota, Ascomycota and Basidiomycota. Ascomycete species in bare peat showed close affinity to species in Scottish cutover peat and uncultured ectomycorrhizal Pezizomycotina (Artz et al., 2007). ITS sequences with high affinities to cellulose and lignin degrading basidiomycete yeasts and metal tolerant Verticillium species were identified.
Preliminary analysis of 65,435 bacterial (16S) and 38,130 fungal (ITS) sequences generated via Roche 454 pyrosequencing from the 18 peat cores representing bare peat and the 5 vegetation classes has been extremely informative. Success in sequence assignment to phyla, genera and species levels was much higher for bacteria than fungi. Over 70 % of fungal, compared to 33% of bacterial rDNA sequences remain unidentifiable after BlastN analyses against public sequence databases (e.g., GenBank and UNITE). Of a total of 22 bacterial phyla, Proteobacteria (mean 55%) and Acidobacteria (23%) representation dominated in bare peat and all vegetation classes. The large Proteobacteria phylum contains highly functional soil bacteria involved in carbon, nitrogen and sulphur cycling (Kesters et al., 2006) whilst the latter are common in acidic soils including peatlands (Artz, 2009 and references therein). Diversity estimates highlighted a trend to increasing richness from bare eroded peat, through early restoration, to remnant dwarf shrub and gully classes. Identified fungal sequences highlighted the predominance of Ascomycota that accounted for between 57-87% of ITS sequences. This confirms and extends literature in peatlands (Artz, 2009) in, for example, the identification of the basal class Archaeorhizomyces that have been recently identified associated with the coniferous roots in boreal Eurasian and North American forests (Rosling et al., 2011). Basidiomycetes were the second most common fungal sub-division (8.5-27.6%) with a high predominance in peat supporting gully vegetation. Neocallimastigomycota, a novel phylum containing anaerobic cellulose and lignin degrading ruminant fungi (Griffith et al., 2010), were represented in all peat samples but predominated in peat supporting remnant dwarf shrubs. Limited numbers of sequences of symbiotic arbuscular mycorrhizal fungal phylum Glomeromycota were restricted to gully peat that hosted naturally regenerated acid grass species known form symbiotic arbuscular mycorrhizas (Smith and Read, 2008).
Multivariate analyses of subsampled bacterial and fungal sequences (780 and 701 sequences, respectively), assigned to sequence homology based operational taxonomic units (OTU) through genetic distance estimates, clearly highlighted vegetation class specific selection on both bacterial and fungal communities. PCA ordination confirmed shifts in bacterial and fungal communities with a clear separation of bare peat and peat supporting recent re-establishing grass and heather cover from well establised 25 year restored heather, gully and remnant dwarf shrub vegetation. Further analyses of functional fungal and bacterial grouping will be presented at the congress.
We gratefully acknowledge support from the MMU Dalton Research Institute and small grant funding from Moors for the Future/Yorkshire Peat Partnership. Dr. Scot Dowd (Research and Testing Laboratory, Lubbock, Texas, USA) is acknowledged for the Roche 454 pyrosequencing service.
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