• The flux of pulse-derived 13C from upland pasture plants to the external mycelium of their arbuscular mycorrhizal (AM) symbionts was traced and quantified over a 7-d post-labelling period.
• Mesh cores, which allowed in-growth of native AM mycelium but were impenetrable to roots, were inserted into unlimed and limed plots and the surrounding vegetation was exposed to 13CO2 at ambient CO2 concentrations.
• Release of 13CO2 from cores colonized by AM mycelium peaked 9–14 h after labelling and declined within 24 h after severance of mycelial connections to roots. Between 5 and 8% of carbon lost by plants was respired by AM mycelium over the first 21 h after labelling. Liming increased the amount of carbon fixed by plants and subsequently allocated to fine roots and AM mycelium.
• The results demonstrate for the first time under field conditions that AM mycelia provide a rapid and important pathway of carbon flux from plants to the soil and atmosphere.
If you can't find a tool you're looking for, please click the link at the top of the page to "Go to old article view". Alternatively, view our Knowledge Base articles for additional help. Your feedback is important to us, so please let us know if you have comments or ideas for improvement.
The roots of the dominant plants of semi-natural grasslands are normally heavily colonised by arbuscular-mycorrhizal (AM) fungi (Read et al., 1976; Sparling & Tinker, 1978). These fungi, which are obligate symbionts, are believed to obtain almost all their carbon (C) from their autotrophic plant partners. In laboratory studies AM have been reported to utilise 5–20% of the net C assimilation by the plants (Pearson & Jakobsen, 1993), but the quantities of C required to support their external mycelial systems in nature have not been determined. The mycelial systems of AM fungi extend for several centimetres away from the surface of plant roots into bulk soil from which they obtain nutrients. These mycelial networks provide vital conduits for the translocation of nutrients from soil to plants and for reciprocal movement of C from plant roots through AM hyphae (Smith & Read, 1997).
Despite the evidence that the majority of grassland plants form mycorrhizas and that these associations are a significant drain on plant photosynthate (Pearson & Jakobsen, 1993), C movement from plant roots to AM mycelial systems has been quantified only under laboratory conditions. A number of studies have specifically investigated C fluxes from grassland plants to soil microbiota (Martin & Merckx, 1992; Saggar et al., 1997; Kuzyakov et al., 1999; Stewart & Metherell, 1999), but in circumstances where AM associations were either absent or ignored. As a result, most recent models of C fluxes between plants and the soil have been based upon the assumption that root exudation, sloughed cells and dead roots provide the only significant pathways for the supply of plant-fixed C to the free-living microbial populations in soils (Toal et al., 2000; Kuzyakov et al., 2001). Increasing awareness of the extent of the AM mycelial network in grassland ecosystems (Francis & Read, 1984; Jakobsen & Rosendahl, 1990) exposes the limitations of these models and highlights the need for quantification of its involvement in the plant-soil C flux pathway.
To date it has proved extremely difficult to distinguish between fluxes of C originating in roots and those that occur through their associated mycorrhizal mycelial systems. In this paper we employ a novel mesh-bound soil core system (Johnson et al., 2001) enabling separate determination of these two compartments to provide a time-course analysis of C fluxes to the AM mycelial network. A recently developed mobile laboratory, which provides in situ pulse labelling of 13CO2 at atmospheric CO2 concentrations (Ostle et al., 2000), was used to yield an identifiable pulse of C, which was subsequently traced and quantified through the mycelial network of a naturally occurring population of AM fungal symbionts in a grassland ecosystem.
Materials and Methods
The study was undertaken in an upland grassland (the NERC Soil Biodiversity field site, Sourhope, Kelso, Scotland; NGR: NT 854196; altitude: 500 m asl). The community comprises a mixture of 24 higher-plant species, 21 of which typically form AM-mycorrhizas (Harley & Harley, 1987), and is classified as U4d (Festuca-Agrostis-Galium-Luzula multiflora-Rhytidiadelphus loreus subcommunity) under the British National Vegetation Classification (Rodwell, 1992). The site has been under pasture and has received no fertiliser additions for the past 40 yr. It was fenced and excluded from grazing from April 1999 onwards. The soils (Table 1) are developed on locally derived drift from andesitic lavas of Old Red Sandstone Age and are acid brown earths (pH 4.5–5.0) with a mean clay content in the 2 mm fraction of 40%.
Table 1. Physico-chemical characteristics of soil from the FH and Ah horizons in control and limed plots at Sourhope (± SE)
Total C (%)
NA, not analysed.
5.25 (± 0.09)
6.8 (± 0.33)
4.95 (± 0.06)
21.2 (± 2.1)
12.2 (± 0.66)
7.97 (± 0.07)
6.4 (± 0.30)
5.94 (± 0.18)
22.6 (± 1.5)
12.2 (± 0.60)
Design of experimental cores
Soil cores, designed to permit in-growth of mycorrhizal hyphae but exclusion of roots were based on the design of Johnson et al. (2001) and consisted of 22 mm diameter plastic tubing (150 mm deep) with four ‘windows’ cut into the below ground portion of the tube each being covered with 35 µm nylon mesh (Plastok Associates Ltd, Birkenhead, Wirral, UK). The mesh permits in-growth of AM hyphae as well as movement of bacteria, solutes and water between the core and the surrounding bulk soil but is impenetrable to plant roots. The cores were filled with approx. 40 g f. wt of sieved (2 mm) nonsterile soil collected from plots of the same treatments to which they were re-inserted in the field. The cores were maintained free of plants for the duration of the experiments.
Experiment 1: test of the physical effect of core rotation on flux of CO2 from bulk soil to soil cores
To validate the use of mesh core systems to quantify effects of AM fungal mycelium on C flux from plant roots to soil, it was necessary to establish whether rotation of soil cores to sever AM hyphae physically altered the rates of passive diffusion of CO2 into and through the cores from the bulk soil. Core rotation may damage aggregates at the interface between the bulk soil and mesh wall of the cores, and could result in lowered porosity of the soil in this area. Any change in passive diffusion of CO2 could mask or exaggerate estimates of C flux through AM hyphae based on differences between rotated and unrotated cores.
An experiment was therefore conducted to determine the effects of core rotation on rates of passive diffusion of CO2 from the bulk soil into and through soil cores lacking mycorrhizal hyphae. Two cores containing moist cotton wool were inserted into each of six 1 l pots containing intact blocks of Sourhope turf and maintained in a glasshouse. The cotton wool was used to enable mycelial penetration of the mesh walls and subsequent colonisation of the cores during the in-growth period, thus mimicking as closely as possible any changes in soil structural properties at the mesh/bulk soil interface that may occur when the cores are rotated in the field experiments. In addition, the cotton wool and associated fungal mycelium that had penetrated the mesh could be removed prior to exposure of the turf to 14CO2 without causing disruption to the interface between the core walls and bulk soil. This was done after 6 wk incubation in the turfs whereupon the cotton wool was replaced with moist sieved (2 mm) Sourhope soil. One of the cores in each pot was then rotated and the turfs placed in a 26-l clear acrylic gas-tight chamber into which was released 1.5 MBq 14CO2 by addition of a few drops of 10% lactic acid to 14C sodium bicarbonate solution (specific activity 2 GBq mmole−1). The turfs remained exposed to the 14CO2 for 2 h, during which time the box was placed in a growth room (20°C, 18 h day; 15°C, 6 h night). After labelling, an eppendorf tube containing 1 ml 1 M NaOH was placed above the soil surface inside each core and the headspace of the core sealed with a suba-seal. The eppendorf tubes were replaced five times throughout a 46-h postlabelling period. The radioactivity in the NaOH was determined by liquid scintillation counting (Packard Tricarb 1600 TR, Packard Instruments Ltd, Pangbourne, Reading, UK).
Experiment 2: time-course of 13C released from external AM mycelium following pulse-labelling of plant shoots in the field
In May 1999, 12 subsidiary plots of 40 cm diameter were established in a completely randomised block design on an area of unimproved pasture outside the main experimental plots at Sourhope. The subsidiary plots were not mown for the duration of the experiment, although the vegetation was clipped to a height of c. 15 cm before labelling. Into each plot were inserted four experimental cores (constructed as described above) containing plant-free Sourhope soil. Half of the cores in each plot were rotated c. 45° about their vertical axis to break AM mycelial connections with the host plants immediately before commencing exposure of vegetation to 13CO2.
Labelling of field plots
Nine plots were exposed to 50 atom %13CO2 for 5 h and three plots were supplied with unlabelled gas in order to provide natural abundance data. The gas was supplied at ambient concentrations (c. 350 ppm) using a mobile Stable Isotope Delivery (SID) system (Ostle et al., 2000). The SID system comprised an isotope-mixing unit with flow control to a series of 12 independent clear acrylic labelling chambers, which were pushed approximately 10 mm into the surface soil F-horizon. A flow rate of 6 l min−1 chamber−1 provided a gas residence time of c. 5 mins. In-line computer controlled infrared gas analysers (calibrated for 13CO2) allowed automated measurement of chamber CO2 concentrations and gas flow management.
Time course of 13CO2 released from mesh cores following pulse-labelling
A set of cores (comprising six static and six rotated cores from the labelled plots and three static and three rotated cores from the unlabelled plots) were removed 14, 20 and 38 h after the start of pulse labelling. Each core was placed into sealed (with suba-seals) acrylic vessels and incubated at soil temperature (13°C) in a waterbath 40 mins after removal from the plots. After a further 40 mins, a 12-ml headspace gas sample was removed and the incubation vessel was uncapped. The procedure was repeated so that in total four gas samples were taken from each vessel, 80 mins, 140 mins, 24 h and 7 d after removal of the cores from the plots. The time-course enabled us to compare the near-instantaneous 13C enrichment of respiration and the effects of hyphal severance and their subsequent senescence on the dynamics of plant-derived 13C release from cores with and without active AM mycelium.
The gas samples were stored at room temperature in evacuated glass exetainers (PDZ-Europa Ltd, Crewe, UK) from which the 13C : 12C ratio of the accumulated CO2 was determined by isotope ratio mass spectrometry (IRMS; Micromass TraceGas Pre-concentrator coupled to an Isoprime isotope ratio mass spectrometer, Micromass, Wythenshawe, UK) at the Stable Isotope Facility (SIF), CEH-Merlewood, UK. After the final gas sampling, the soil from the cores was oven-dried (80°C) and the 13C : 12C ratio determined by continuous-flow/combustion/IRMS (Carlo Erba NA1500 elemental analyser coupled to a Dennis Leigh Technology isotope ratio mass spectrometer) at SIF, CEH-Merlewood.
Experiment 3: effect of liming on C allocation from plants to AM mycelium in the field
The second experiment was undertaken between May and July 2000 using control and lime treatments of three blocks of the Soil Biodiversity site at Sourhope, located on a north-east facing slope. The main experimental plots (20 m × 12 m) were arranged in a 5 block × 6 treatment completely randomised design, including control and lime treated plots. The lime (CaCO3; 39% Ca; 0.5% ash soluble in HCl) was manually spread onto the plots as a solid (0.6 kg m−2) during April/May of each year. The whole site was mown on six occasions during the summer and all clippings were removed. Solar irradiance and soil temperature data were collected hourly by an automatic weather station located within the site.
In May, two stainless steel isotope inclusion rings (405 mm diameter × 200 mm deep) were inserted 70 mm into each of the control and limed plots of three blocks (i.e. 12 rings in total). The vegetation inside the ring and immediately surrounding it was not mown for the duration of the experiment but was clipped to 15 cm height prior to labelling. Twelve cores (constructed as described above) containing sieved soil from either the control or limed plots as appropriate were inserted the same day within each ring and were grouped into six pairs (i.e. a total of 144 cores used for the experiment). The cores were rotated 2 d before labelling (July 2000).
A complete set (i.e. vegetation subsamples, one static and one rotated core, one bulk soil sample) of measurements was taken prior to labelling to provide natural-abundance control values. The vegetation subsamples (c. 2 g) were removed from one segment of each ring (comprising approximately 1/6th of the total area cut to the soil surface) and immediately frozen (–20°C). A bulk soil core (100 mm deep) was removed from a segment of the ring with a 10-mm diameter cork borer and fine roots were extracted by hand from the upper 40 mm, cleaned in water and stored at –20°C. One rotated and one static mesh walled core were removed, incubated and headspace gas samples taken and analysed as described for the first experiment. The acrylic labelling chambers were then sealed onto the stainless steel rings with rubberised bands and all 12 plots exposed to 99 atom %13CO2 at c. 350 ppm for 3.5 h using the mobile SID system described above. Five more complete sets of samples were then taken as described above, 4, 10, 19, 25 and 34 h after the start of labelling.
At the end of the experiment, the remaining vegetation in each ring was harvested and frozen. All frozen samples were subsequently freeze-dried and weighed before analysis by IRMS as described for the first experiment.
Stable isotope calculations and statistical analyses
In the second experiment, the enrichment of 13C in samples is reported in δ13C‰ units (13C : 12C ratio) which were calculated relative to internal gas standards and solid reference material calibrated against the 13C : 12C of Pee Dee Belemnite (PDB) standards.
In the third experiment, 13C enrichments are reported quantitatively (mg C) as pulse-derived 13C (i.e. increase in the amount of 13C in enriched samples relative to their natural abundance value) which were calculated to take into account differences in shoot or root biomass using the equation: Pulse-derived 13C (mg) = [(δCL × tCL) – (δCU × tCU)] × d. wt, where δCL and δCU= atom excess (atom percentage13C) of labelled and unlabelled samples, respectively, tCL and tCU=% total C of labelled and unlabelled samples, respectively, and d. wt = total d. wt of sample in the labelled plot. Atom excess was calculated from the δ13C‰ values using the equation: Sample atom excess (atom percentage13C) = [(Rsample × 10−3) + 1.12372], where Rsample= sample δ13C‰ determined by IRMS and 1.12372 is the atom percentage13C of PDB standard.
Quantification of 13C release from the cores was calculated by combining the 13C : 12C ratios with respiration (release of CO2) rates from the soil-filled cores. The respiration rates were determined by analysis of the peak height data in the gas samples provided by IRMS calibrated against CO2 standards in the range 200–1500 ppm. The loss of pulse-derived 13CO2 through external mycelium was estimated for the first photo-period (0–21 h post labelling) by subtracting the area under the pulse-derived 13CO2-C curve of the static cores from that of the rotated cores, and multiplying by the difference in surface area of the cores compared with that of the isotope inclusion rings. We assumed that the equilibration rates of dissolved 13CO2 from the soil solution (which is likely to be very low in this acid soil) into the vessel headspace was equal for both the static and rotated cores (see Experiment 1). Quantification of pulse-derived 13C to fine roots (to 4 cm depth) within the isotope inclusion rings was estimated using the fine root density measurements obtained from the bulk soil subsamples. The allocation of pulse-derived 13C to fine roots and external AM mycelium was expressed both as a percentage of the total 13C fixed by plant shoots during pulse labelling and the amount of 13C lost from plant shoots (as shoot respiration and translocation) during the 21 h post-labelling period.
Treatment effects were analysed by ANOVA, Tukey multiple comparison tests and t-tests using Minitab 10.5. All data were log or arc-sine transformed to satisfy test assumptions where appropriate.
Experiment 1: test of the physical effect of core rotation on flux of CO2 from bulk soil to soil
Core rotation had no physical effect on the amount of 14CO2 that diffused from the bulk soil into the core systems. The amount of 14CO2 released (± SE) from the static cores was 1.19 ± 0.37 pmol cm−2 h−1 while the amount released from the rotated cores was 0.91 ± 0.27 pmol cm−2 h−1, which was not significantly different (P = 0.550). These results confirm that differences in carbon fluxes through rotated and unrotated cores will be due primarily to the effects of severance of AMF hyphae growing from bulk-soil into the cores and not from change in soil porosity at the mesh–soil interface in rotated cores.
Experiment 2: time-course of 13C released from external AM mycelium following pulse-labelling of plant shoots
The δ13C‰ values in the CO2 released from the mesh-bound cores ranged from –20 to –22‰ 80 min after their removal from the unlabelled plots (Fig. 1). No differences were observed in δ13C‰ between the rotated and static cores at this time. The δ13C‰ of CO2 released from rotated cores removed from labelled plots ranged from –13 to –10‰ and was significantly greater than the natural-abundance controls at all three harvests, but did not differ significantly between harvests. The δ13C‰ of CO2 released from static cores removed from labelled plots ranged from 3 to 20‰ (Fig. 1) and was significantly greater than δ13C‰ of CO2 released from both the rotated cores from labelled plots and natural-abundance controls at all three harvests. The greatest enrichment occurred at the first (14 h) harvest after which δ13C‰ declined to c. 3‰ in the second and third harvests (Fig. 1).
The natural-abundance δ13C‰ value of released CO2 increased by c. 2‰ during the 7 d post-removal incubations for all three harvests (Fig. 2a–c). There were no significant differences between the δ13C‰ values from static and rotated cores after removal from the unlabelled plots at any harvest. The δ13C‰ values of CO2 released from both static and rotated cores declined to control levels 7-days after removal from the labelled field plots at all three harvests (Fig. 2a–c).
The variation observed for each group of measurements generally declined throughout the 7 d incubation period. After 7 d incubation the variation in δ13C‰ ranged by only 2‰ from –19 to –21‰. The greatest decreases in δ13C‰ of CO2 occurred between 80 mins and 24 h after removal of static cores from the labelled plots where δ13C‰ declined significantly from 14 to –11‰ at the first harvest and from 4 to -13‰ at the second harvest (Fig. 2a,b). A near-uniform decrease in δ13C‰ between gas samples from the labelled plots was only observed at the third harvest.
Experiment 3: effect of liming on C allocation from plants to AM mycelium in soil
The amount of 13CO2 fixed by plants during the 4 h labelling period ranged from 90 to 420 mg C m−2. Plants in the plots that had received applications of lime fixed 50% more 13CO2 than that of the control (Fig. 3a). This pattern tended to become more apparent throughout the 30 h post-labelling period and was reflected by an overall significant (P < 0.001) increase in pulse-derived 13C due to liming. The increased fixation of 13C in the limed plots was in part explained by the higher productivity of the plants in this treatment, in which the total live above ground biomass increased significantly by over 33% from 197 to 264 g m−2 d. wt. The best-fit relationship (P = 0.008) between shoot biomass and excess 13C in shoots accounted for 45% of the variation. The amount of 13C in shoots from both control and limed plots generally declined throughout the post-labelling period. After 30 h the amount of 13C remaining in shoots had decreased by about 30% regardless of treatment.
The amount of 13C translocated to fine roots (removed to 4 cm depth) was approximately an order of magnitude lower than the amount fixed by the plants and ranged from 8 to 25 mg C m−2 (Fig. 3b). No significant effects of the liming treatment were seen although the mean excess 13C values were greater in this treatment at all time points. The maximum quantity of 13C in fine roots was measured 21 h after the end of labelling, these values being almost double those obtained during the initial 6-h period. No significant interaction between time and treatment was observed although an overall effect of time (P = 0.002) on the pooled values from the control and lime treatments was detected.
The amount of 13C evolved from mesh cores due to C translocation by AM mycelium (i.e. the difference in excess 13C between static and rotated cores) varied considerably throughout the postlabelling period ranging from 0.1 to 5.5 mg CO2-C m−2 (Fig. 3c). The maximum values occurred 20 and 34 h after the start of labelling in the limed treatment, and 20 h after the start of labelling in the controls. In both treatments a substantial decline was seen 25 h after the start of labelling. The pattern followed closely the level of solar radiation (Fig. 3c), the peak excess 13C values tending to occur at the lowest irradiance levels.
Both fine roots and AM mycelium provided a similar demand for the 13C fixed by plant shoots (Table 2). Allocation of C into fine roots was 5.2 and 5.8% of the amount fixed in the control and limed treatments, respectively, while corresponding values for AM mycelium were 3.9 and 6.2%. When expressed as a proportion of the amount of 13C lost by plants (i.e. by shoot respiration and translocation) throughout the post-labelling period, fine roots accounted for 7.3 and 7.6% and AM mycelium accounted for 5.4 and 7.7% in the control and limed treatments, respectively (Table 2).
Table 2. Allocation of 13C to fine roots (to 4 cm depth) or external arbuscular mycorrhizal (AM) mycelium as a percentage of the amount of label either fixed by plants during pulse-labelling or lost (by shoot respiration and translocation) during 24 h post-labelling with 13CO2 (± SE)
External AM mycelium
Amount fixed (%)
Amount lost (%)
Amount fixed (%)
Amount lost (%)
Values between control and limed treatments are not significantly different (P > 0.05).
5.2 (± 1.5)
7.3 (± 2.6)
3.9 (± 1.8)
5.4 (± 2.6)
5.8 (± 2.4)
7.6 (± 3.3)
6.2 (± 1.7)
7.7 (± 1.9)
Our results indicate that within 21 h of pulse-labelling a grassland sward in the field with 13CO2 between 3.9 and 6.2% of the fixed C can be passed through the external mycelium of the AM fungal symbionts to the atmosphere. This equates to 5.4 and 7.7% of the fixed 13C lost by shoot respiration and translocation during this period. These results provide the first confirmation that amounts of C allocation to AM mycelium similar to those reported from pot experiments (Paul & Kucey, 1981) can be achieved under field conditions. They provide striking evidence that external AM mycelium is an important conduit of C flow from plant to atmosphere, even in pastures (such as at Sourhope) that are dominated by graminaceous species.
We were able, in the first experiment, to discount the potential confounding factor of differential changes in soil structural properties at the mesh:bulk soil interface as a result of core rotation. However, these concerns may still be valid in soils prone to smearing, such as those with a high clay content, and preliminary experiments would need to be undertaken before this technique was extended to different soil types. This study does not enable us to identify with certainty the pathways through which C moves from the plant to the atmosphere above the cores. These could involve direct respiratory losses from AM mycelium, microbial breakdown of mycorrhizal exudates and AM mycelium within the cores and direct transfer of root exudation products from the bulk soil into the cores by free-living saprotrophic fungi. However, the rapidity of both the transfer of 13CO2 through the cores and the decline in 13CO2 release associated with destruction of connections to host plants provides strong evidence that direct respiratory losses by AM fungi may be of greatest importance. A similarly rapid decrease in respiration rates has been observed upon excision of ectomycorrhizal mycelium from host Pinus sylvestris roots (Söderström & Read, 1987). More work is clearly needed to identify the pathways of C transfer from AM fungi into other components of the microflora.
We were unable to detect any 13C-enrichment in the static or rotated cores at the end of either experiment. This may indicate that the C content of the mycelium largely consists of rapidly turning-over current assimilate rather than compounds used for C storage, as has been demonstrated in studies of ectomycorrhizal systems (Söderström & Read, 1987). However it may also, at least in part, reflect the very high background of unlabelled C associated with this organically rich soil (mean total C = 25%) against which detection of the much smaller amounts of 13C-enriched material arising from short-duration 13CO2 pulse-labelling will inevitably be difficult.
The proportions of fixed 13C released to the atmosphere via AM mycelia were similar to those that were allocated to fine roots removed from the surface soil horizons suggesting that, in the short term (0–24 h after labelling), these components are of equal importance as C sinks. However, the proportion of C allocated into AM fungi is likely to have been underestimated in this study as our estimates do not account for C contained in the internal fungal structures. Amongst these, the vesicles in particular are known to be large sinks for C-storage products (Cooper & Lösel, 1978). In addition, the core systems, whilst providing a relatively noninvasive method for selectively controlling AM colonisation, are likely to have lower hyphal densities than are found in the bulk soil as the density of hyphae declines away from the root surface (Jakobsen et al., 1992).
The accumulation of C in fine roots occurred at a slower rate than the loss of C from the colonised cores. The maximum 13C values from AM respiration occurred 9 h and 15 h after labelling in the first and second experiments, respectively, while transfer into fine roots peaked after 21 h. Similarly, in ectomycorrhizal systems, C transfer from Pinus sylvestris L. seedlings to Suillus bovinus (L.):Kuntze mycelium has been shown to occur 8–24 h after photosynthesis (Leake et al., 2001), many hours earlier than the maximum 14C accumulation in roots. The data we now report provide strong evidence that mycorrhizas primarily use current plant assimilate (Wright et al., 1998) and that C transfer to mycorrhizal mycelium in the short term may be of importance. The data also highlight the value of detailed investigations of the short-term dynamics of C transfer from plants to soil. Many pulse-labelling experiments have either used relatively long fixation periods or have taken samples many hours after cessation of labelling (Stewart & Metherell, 1999), and so may miss the maximum outputs of C from roots and AM fungi.
Addition of lime for 1.5 yr to the upland grassland studied here increased by 50% the quantity of 13C assimilated by plant shoots and resulted in a significant increase in the overall amount of 13C in the shoots during the postlabelling period. The increased assimilation was in part (r2 = 45.1%) explained by the higher productivity in these plots. Other factors, such as plant species composition, the relative proportion of fixed C derived from either soil respiration or from the atmosphere and the proportion of new:old biomass may also have affected the 13C signatures. The percentage of fixed C allocated to fine roots and AM mycelium was generally greater in limed than in control plots. Although there are no studies specifically addressing the impact of lime on respiration rates by AM fungi, bulk soil respiration rates are commonly reported to be higher following addition of lime to grasslands (Hopkins, 1997). Further work is required to investigate the effects of longer-term additions of lime both on the functioning and species composition of AM fungi.
The results presented here provide for the first time qualitative and quantitative data on C dynamics of natural communities of AM fungi in acid organic upland grasslands under field conditions. The data confirm observations from pot-based studies indicating that AM fungi provide a sufficiently important pathway for flow of current assimilate through the soil system to necessitate their inclusion in any models describing C cycling in grasslands. The use of 13CO2 labelling methodologies (Ostle et al., 2000) in combination with novel mesh bound cores are here demonstrated to be powerful tools for the study of in situ AM functioning.
We extend our thanks to Dr Andy Stott, Darren Sleep and Helen Grant (SIF, CEH-Merlewood) for efficient analysis of 13C samples, Irene Johnson for assistance during long-periods of fieldwork, Dr Sarah Buckland (Soil Biodiversity Site Manager) and Dr Patricia Bruneau, Stirling University. This work was funded by the Natural Environment Research Council under the Soil Biodiversity Thematic Programme (Grant numbers: GST/02/2117 and GST/02/2115).