Ecology Letters, (2005) 8: 1264–1270
doi: 10.1111/j.1461-0248.2005.00831.x
LETTER
Tracking carbon from the atmosphere to the
rhizosphere
Pål A. Olsson* and Nancy
C. Johnson Department of Microbial
Ecology, Ecology Building, Lund
University, SE-223 62 Lund,
Sweden
*Correspondence: E-mail:
[email protected]
Present address: Environmental
and Biological Sciences,
Northern Arizona University,
Abstract
Turnover rates of arbuscular mycorrhizal (AM) fungi may influence storage of soil
organic carbon (SOC). We examined the longevity of AM hyphae in monoxenic cultures;
and we also used 13C incorporation into signature fatty acids to study C dynamics in a
mycorrhizal symbiosis involving Glomus intraradices and Plantago lanceolata. 13C enrichment
of signature fatty acids showed rapid transfer of plant assimilates to AM fungi and a
gradual release of C from roots to rhizosphere bacteria, but at a much slower rate.
Furthermore, most C assimilated by AM fungi remained 32 days after labelling. These
findings indicate that 13C labelled fatty acids can be used to track C flux from the
atmosphere to the rhizosphere and that retention of C in AM fungal mycelium may
contribute significantly to SOC.
Flagstaff, AZ 86011-5694, USA.
Keywords
Arbuscular mycorrhiza, bacteria, 13C, carbon cycling, Glomus intraradices, Plantago
lanceolata, signature fatty acids, stable isotope.
Ecology Letters (2005) 8: 1264–1270
INTRODUCTION
The size and activity of microbial biomass in the soil
regulates the accumulation of soil organic carbon (SOC).
Arbuscular mycorrhizal (AM) fungi are obligate biotrophs
that are well known for their importance in plant nutrition
(Smith & Read 1997) and the formation of soil structure
(Miller & Jastrow 2000). These fungi are also important
regulators of C flux from plants to the soil (Zhu & Miller
2003) because they can consume up to 20% of plant C
(Jakobsen & Rosendahl 1990) and they are often the largest
contributor to soil microbial biomass (Miller et al. 1995;
Olsson et al. 1999).
Mycorrhizal fungi form a variety of structures with
varying function, morphology and longevity. AM fungi
colonize host plant roots from spores, extraradical hyphae
or previously colonized roots. Hyphae grow from colonized
roots into the soil and form an extraradical mycelium
composed of recalcitrant runner hyphae, short-lived absorbing hyphae and persistent reproductive spores (Friese &
Allen 1991; Bago et al. 1998). Intraradical structures form in
the root cortex. These structures include hyphae, arbuscules,
which are critical sites for nutrient transfer between the
symbionts and vesicles, which are lipid-filled storage
structures (Smith & Read 1997). Arbuscules are short-lived
structures believed to have a turnover rate of 1–2 weeks
2005 Blackwell Publishing Ltd/CNRS
(Alexander et al. 1989), while vesicles may persist even in
senescent roots. AM fungi convert plant photosynthate into
lipids, which are transported throughout the intra- and
extraradical mycelium and stored in high concentrations in vesicles and spores (Smith & Read 1997; Bago
et al. 2002).
Recent studies suggest that assimilated C can move very
rapidly from plants into AM fungi and back to the
atmosphere (Johnson et al. 2002). Staddon et al. (2003)
estimated that extraradical AM fungal hyphae turn over
within 5–6 days. However, this study focused solely on C
flux in the extraradical hyphae near the surfaces of plant
roots; therefore, it is of limited value for estimating the
residence time of AM fungal biomass within soils because it
did not account for the dynamics of intraradical mycelium
or of the network of extraradical mycelium that may extend
many cm into the soil volume. The residence time of all AM
fungal tissues must be considered before they can be
accounted for in models of SOC dynamics. Zhu & Miller
(2003) noted that it is unlikely that the estimated turnover
rate of 5–6 days can be generalized to all AM fungal tissues
because AM fungal hyphae are composed of chitin and they
produce glomalin, two compounds that are known to have
extremely long residence times in the soil (Rillig 2004).
Integrative research is needed to reconcile these different
views.
Tracking ecosystem carbon 1265
Our study uses a new method to track C flow into AM
fungi and rhizosphere bacteria that will help to monitor the
fate of organic C in the soil. Neutral lipids are storage
compounds that may comprise up to 20% of the biomass of
hyphae, vesicles and spores of AM fungi (Olsson &
Johansen 2000). The neutral lipid fatty acid (NLFA)
16 : 1x5 and, to a lesser extent, the phospholipid fatty acid
(PLFA) 16 : 1x5 are effective signatures for AM fungi
(Olsson 1999; Van Aarle & Olsson 2003). Other PLFAs are
signatures for rhizosphere bacteria (Frostegård & Bååth
1996). Phospholipids are easily decomposed through
enzymatic actions in soil (White et al. 1979) and thus
PLFAs can be assumed to reflect the occurrence of living
organisms. On the contrary, NLFAs are stored in spores,
which have a long residence time. Thus, it is likely that C is
maintained in the NLFA pool in the mycelium. Degradation
of neutral lipids in soil has not been studied, but it is likely
that they are more persistent than the phospholipids (White
et al. 1979). A recent study indicates that total 13C
incorporation in AM fungal hyphae is correlated to that in
NLFA 16 : 1x5 (Olsson et al. 2005). Consequently, 13C
labelled signature fatty acids can be used to effectively track
C flux from plants to intra- and extraradical AM fungal
tissues and to other rhizosphere microorganisms (Olsson
et al. 2002; Gavito & Olsson 2003).
The objective of this study was to measure the flux of
newly assimilated C into AM fungi and rhizosphere bacteria.
This investigation examined whether residence time of C in
intraradical mycelium differs from that in extraradical
mycelium. Also, we tested the hypothesis that the flow of
newly assimilated C to AM fungi is greater and faster than to
rhizosphere bacteria.
METHODS
Glasshouse study
We grew 20 pots of Plantago lanceolata L. plants colonized
with the AM fungus Glomus intraradices in a greenhouse.
Rectangular pots (13.5 · 11 · 9.5 cm, l · w · h) divided
into two compartments by means of a 25-lm mesh nylon
screen were used as experimental units. The screen allowed
the penetration of mycorrhizal hyphae but prevented the
passage of roots. The rooting compartment (5.5 cm w), was
used to grow host plants. Extraradical mycorrhizal hyphae
were expected to develop from the root compartment
through the mesh and into the side compartment (8 cm w).
The pots were filled with a 1 : 1 mixture of gamma irradiated
(10 kGy) sandy loam soil and quartz sand, with 450 g inside
the nylon mesh and 600 g outside the mesh. In 17 pots, 50 g
of root fragments and rhizosphere soil from cultures of
G. intraradices Schenk & Smith (BEG 87) grown with
Trifolium subterraneum was added to the 450 g of soil mixture
inside the nylon mesh. Instead the three non-mycorrhizal
controls received 50 g of gamma-irradiated soil. All pots
were given 5 ml of a microbial wash (Van Aarle & Olsson
2003). Seeds of P. lanceolata L. were surface sterilized,
germinated on moist filter paper, and five 5-day-old
seedlings were transplanted inside the nylon mesh in each
pot. Pots were watered by weight to 70% of the soil : sand
water holding capacity. Every second week each pot
received 10 mg of N as NH4NO3. The pots were randomly
arranged and regularly redistributed in a greenhouse with an
average 22 C day and 18 C night temperature, a minimum
of 270 lmol m2 s)1 photosynthetic photon flux density
supplemented with 400 W Osram light (Osram AB,
Haninge, Sweden) bulbs as required and 18 h photoperiod.
When P. lanceolata plants were 8 weeks old they were
exposed to a 13CO2 enriched atmosphere (Olsson et al.
2002; Gavito & Olsson 2003) by sealing the pots in a clear,
airtight box to which 125 ml of 13CO2 (99% 13CO2;
Larodan Fine Chemicals, Malmö, Sweden) was injected
using a gas tight syringe through a septum. The CO2
concentration inside the box increased from 190–290 p.p.m.
to around 1100 p.p.m. after injection. Air inside the box was
circulated using a fan. The labelling period lasted 120 min at
which time the CO2 concentration inside the box decreased
to its initial level.
Plants and soil were harvested five times over the
following 32 days after labelling with three replicates at each
sampling time. The duration of the experiment was chosen
because most C was expected to turnover within 1 week
(Staddon et al. 2003). At harvest shoots, roots and soil were
separated. Shoots were oven dried and weighed. A
subsample of roots was stained with trypan blue (0.1% in
lactic acid, glycerol and water 1 : 2 : 2, v/v/v) and per cent
root length colonization was measured using the gridline
intersect method (Giovannetti & Mosse 1980). Soil and root
samples were frozen immediately following harvest and they
were freeze-dried prior to lipid analysis.
Lipid extraction and analysis
Lipids within soil and roots (ground with a pestle in a
microcentrifuge tube with a few grains of sand) were
extracted by vortex mixing (1 min) in a one-phase mixture
of citrate buffer, methanol and chloroform (0.8 : 2 : 1, v/v/v,
pH 4.0). The lipids were fractionated into neutral lipids,
glycolipids and phospholipids on pre-packed silica columns
(100 mg sorbent mass; Varian Medical Systems, Palo Alto,
CA, USA) by eluting with 1.5 mL chloroform, 6 mL
acetone and 1.5 mL methanol respectively. The fatty acid
residues in neutral lipids and phospholipids were transformed into free fatty acid methyl esters and analysed by gas
chromatography using a 50 m HP5 capillary fused silica
column (Hewlett Packard, Palo Alto, CA, USA) with H2 as
2005 Blackwell Publishing Ltd/CNRS
1266 P. A. Olsson and N. C. Johnson
the carrier gas (Gavito & Olsson 2003). The fatty acid
methyl esters were identified and quantified in relation to
the added internal standard (fatty acid methyl ester, 19 : 0).
These were compared with those identified earlier by gas
chromatography–mass spectrometry. No hyphae were
detected in the root-free soil compartment (outside the
nylon mesh) and thus soil from outside the nylon was not
considered in further analyses.
membranes, with rather low specificity as a signature
because of relatively low content in AM fungi and a high
background in soil originating from bacteria (Olsson et al.
1995). Ten bacteria-specific PLFAs (i15 : 0, a15 : 0, i16 : 0,
10 Me16 : 0, i17 : 0, a17 : 0, cy17 : 0, 10 Me17 : 0, 10 Me18 : 0
and cy19 : 0) can be used as indicators of bacterial biomass
(Frostegård & Bååth 1996).
Calculation of excess
Determination of
and fatty acids
13
C
13
C enrichment in crude tissue samples
Freeze-dried and ball-milled root and shoot material (c.
100 lg) was enclosed in tin capsules (crude tissue samples)
and analysed by continuous-flow isotope ratio mass
spectrometry (IRMS) using an ANCA-NT 20–20 Stable
Isotope Analyser interfaced to a solid/liquid preparation
module (PDZ Europa Scientific Instruments, Crewe, UK).
The ratio 13C : 12C of CO2 of the combusted samples (total
C) were determined with a 0.1& precision. Data were
expressed as atomic % of 13C with reference to a sucrose
standard (BDH Laboratory Supplies, Poole, UK), calibrated
against the Pee Dee Belemnite (PDB) standard (the
limestone fossil Belemnitella americana from the Cretaceous
Pee Dee formation in South Carolina, USA; Dawson &
Brooks 2001). The 13C enrichment in fatty acid methyl
esters was determined in the 20–20 IRMS interfaced with a
Hewlett Packard 6890 gas chromatograph. The chromatographic conditions were as described for the lipid analysis
except that He was used as the carrier gas. The effluent from
the capillary column passed through an Al tube with CuO
wires at 860 C, where the fatty acids were converted to
CO2 released into the 20–20 IRMS. The atomic % of 13C
values were calculated based on atomic 13C of the reference
CO2 gas, injected three times at the beginning and end of a
chromatographic run. The reference CO2 was standardized
with the PDB standard using solid/liquid preparation
module. The precision of the reference gas 13C was 0.2&.
Integration for each peak was checked and corrected
manually. The 13C enrichment of fatty acids was calculated
after correction for the C added in the methanolysis step of
the fatty acid analysis procedure. Background 13C levels
were determined from two non-labelled controls (1.153% in
plant shoots, 1.152% in plant roots, 1.135% in NLFA
16 : 1x5 in soil). The 13C enrichment (excess atomic % of
13
C) was calculated by subtracting the natural abundance
(%) of 13C from each fraction.
Signature fatty acids
Neutral lipid fatty acid 16 : 1x5 is a sensitive signature of
AM fungi in both roots and soil (Graham et al. 1995; Olsson
et al. 1995). PLFA 16 : 1x5 is a constituent of AM fungal
2005 Blackwell Publishing Ltd/CNRS
In addition to measuring 13C enrichment in different
fractions, we also determined the total excess 13C. This is
the most appropriate measure to indicate C flow because it
accounts for variation in mass of different fractions and this
is important in systems where biomass varies over time.
Plant excess 13C (lg) was calculated by multiplying the 13C
enrichment with the total biomass C (assuming 50% C of
dry weight). Excess 13C in AM fungi (including both
intraradical and extraradical mycelium) was calculated by
multiplying the 13C enrichment in NLFA 16 : 1x5 with total
amount of AM fungal NLFA 16 : 1x5 (minus background
levels of 16 : 1x5 fatty acids determined from three nonmycorrhizal pots harvested 8 days after labelling) and then
by multiplying with the conversion factor 2.7, which has
been obtained in studies of pure G. intraradices mycelium
grown in liquid culture (Olsson et al. 2005). This conversion
factor had been calculated by comparing the flow of C from
the host plant to the NLFA 16 : 1x5 from measurements of
NLFA 16 : 1x5 in the mycelia and its 13C enrichment
[NLFA 16 : 1x5-C (lg) · 13C enrichment in NLFA
16 : 1x5 (%/100) ¼ C flow to NLFA 16 : 1x5] and then
relating this to the flow of total C to pure mycelia of
G. intraradices from monoxenic cultures [mycelium biomass
C (lg) · total 13C enrichment (%/100) ¼ C flow to
mycelium].
Bacterial excess 13C was calculated by multiplying 13C
enrichment in the bacteria-specific PLFAs with the bacterial
biomass C. The bacterial biomass C was calculated using a
conversion factor of 350 lmol bacterial PLFAs per g
biomass C (Bååth 1994; Frostegård & Bååth 1996),
assuming similar enrichment in the total bacterial biomass
as in PLFAs.
Study of monoxenic AM cultures
In a separate experiment, we visually studied the dynamics
of AM fungi in monoxenic cultures of transformed carrot
roots and G. intraradices in a split-dish system prepared as
described by St-Arnaud et al. (1995). Hyphal growth rate on
solid medium was microscopically studied in a root-free
compartment adjacent to a compartment containing monoxenic AM roots. After the passage of the first hyphae over
the barrier separating the two compartments, the dynamics
Tracking ecosystem carbon 1267
of runner hyphae, branched absorbing hyphae and spores
(Bago et al. 1998) were documented by day-to-day observations over a 15- to 20-day period.
RESULTS
The roots of all of the inoculated plants were colonized with
AM fungi and there was no AM colonization in the nonmycorrhizal controls. Levels of AM colonization was not
significantly different across the five harvests (Table 1).
There was significantly more PLFA 16 : 1x5 in mycorrhizal
roots (24 nmol g)1, n ¼ 17) than in non-mycorrhizal
controls (2.9 nmol g)1, n ¼ 3). The content of NLFA
16 : 1x5 was 8600 nmol g)1 in the mycorrhizal roots, and
only 13 nmol g)1 in the roots of non-mycorrhizal controls
(Table 1). PLFA 16 : 1x5 in the soil of mycorrhizal and
non-mycorrhizal pots were not significantly different, while
NLFA 16 : 1x5 was 12 times greater in the soil of
mycorrhizal pots (19 nmol g)1) compared with the nonmycorrhizal pots (1.5 nmol g)1). This indicates that in our
experiment, AM fungi are the major source of intraradical
PLFA and NLFA 16 : 1x5 and of extraradical NLFA
16 : 1x5. The enrichment of 13C in these compounds can
therefore be assumed to reflect 13C enrichment in the AM
fungal lipids. Mass balance calculations of NLFA 16 : 1x5
indicates that 82% of the AM fungal lipids where present
inside the roots (intraradical) and 18% in extraradical tissues.
The AM signature fatty acids were enriched with 13C
within 2 days of labelling (Fig. 1a), 13C enrichment in
NLFA 16 : 1x5 in roots (intraradical) and soil (extraradical)
remained stable through the last harvest of 32 days after
labelling and was not significantly influenced by time (oneway ANOVA). In contrast, 13C enrichment decreased with
time in root PLFA 16 : 1x5 (P < 0.05, Fig. 1a) and
increased with time in bacterial PLFAs (P < 0.01, Fig. 1b).
Labelling of soil PLFA 16 : 1x5 probably represents both
the bacterial and AM fungal biomass. This is indicated by
the fact that there was no significant increase of this
signature because of growth of the AM fungus (Table 1).
Nevertheless, the labelling pattern over time resembled that
of AM fungal signature (Fig. 1a) and not of the bacterial
PLFAs (Fig. 1b). The 13C enrichment in 10 bacteria-specific
fatty acids revealed that there was significant transfer of
plant C to the soil bacteria. Enrichment in the bacterial pool
of PLFAs increased asymptotically with little difference after
8 days. The labelling of signature bacterial PLFAs was much
lower than in the AM fungal signature 16 : 1x5 fatty acids.
The 13C enrichment in the shoots decreased from 0.56% to
0.19% during the harvesting period and from 0.46% to
0.27% in roots.
We used conversion factors to calculate the amount of
excess 13C retained in shoots, roots, AM fungi and bacteria;
between 2 and 32 days after pulse labelling (Fig. 2). Linear
regression indicated that 13C in shoots decreased from 7.9 to
3.1 mg; the amount of excess 13C in the bacterial fraction
increased significantly; and there was no significant trend for
plant roots or AM fungi. Overall excess 13C retained in plant
biomass was of two orders magnitude higher than that in
AM fungi, which in turn was one order of magnitude higher
than that in bacteria.
In the monoxenic cultures of G. intraradices, we observed
that runner hyphae grew at a rate of 4.9 ± 0.38 mm day)1
(SE, n ¼ 9) and persisted for the entire observation period.
No runner hyphae or spores degenerated during the
observation period. In contrast, average longevity of the
finely branched absorbing hyphae was only 5.3 ± 0.52 days
(n ¼ 7), after which time spores were formed and finely
branched hyphae degenerated. This observation agrees with
a previous study showing that branched absorbing hyphae
degenerate into empty septate structures after 7 days (Bago
et al. 1998).
DISCUSSION
Our results corroborate the view that AM fungi contribute
to both fast and slow pools of SOC (Zhu & Miller 2003).
Table 1 Root colonization (%) by the AM fungus Glomus intraradices in experimental plants
Roots
Colonization
NM
2 days
4 days
8 days
16 days
32 days
37
24
35
28
31
0
±
±
±
±
±
4.2
5.0
3.7
2.5
8.4
Soil
PLFA 16 : 1x5
NLFA 16 : 1x5
PLFA 16 : 1x5
NLFA 16 : 1x5
2.9
18
46
19
22
22
13
5900
13000
7500
8500
10000
0.69
0.79
0.92
0.75
0.74
0.67
1.5
14
16
13
21
21
±
±
±
±
±
±
0.70
2.5
15
2.6
2.3
4.1
±
±
±
±
±
±
5.4
480
3500
680
920
2500
±
±
±
±
±
±
0.13
0.05
0.04
0.18
0.06
0.07
±
±
±
±
±
±
0.17
1.7
4.4
4.5
5.7
4.8
Amounts (nmol g)1) of the AM fungal signature PLFA and NLFA 16 : 1x5 in roots (intraradical) and soil (extraradical) in the mycorrhizal
treatments and the NM control. Values given are mean ± SE.
PLFA, phospholipid fatty acid; NLFA, neutral lipid fatty acid; AM, arbuscular mycorrhizal; NM, non-mycorrhizal.
2005 Blackwell Publishing Ltd/CNRS
1268 P. A. Olsson and N. C. Johnson
Figure 2 Excess 13C retained in shoots (open squares), roots
(closed squares), arbuscular mycorrhizal (AM) fungus (circles) and
bacteria (triangles) as calculated from the 13C enrichment in solid
samples (plant material) or signature fatty acids (microbial
fractions). Regression coefficients (r2) were: root 0.10 (n.s.), shoot
0.35 (P < 0.05), AM fungi 0.17 (n.s.) and bacteria 0.58 (P < 0.01).
Figure 1
13
C enrichment in the arbuscular mycorrhizal fungal
signature 16 : 1x5 was: (a) determined in roots (closed symbols)
and soil (open symbols), and in the neutral lipid fatty acid (squares;
NLFA) 16 : 1x5 and phospholipid fatty acid (circles; PLFA)
16 : 1x5 fractions. 13C enrichment in bacterial fatty acids (b) was
determined as a mean of 10 bacteria-specific PLFAs (i15 : 0, a15 : 0,
i16 : 0, 10 Me16 : 0, i17 : 0, a17 : 0, cy17 : 0, 10 Me17 : 0, 10 Me18 : 0
and cy19 : 0). 13C enrichment denotes the excess 13C when the
natural background has been subtracted as determined in nonlabelled systems. Regression lines were fitted to log-transformed
data and the data are presented on a log-scale.
During microbial growth, almost 50% of C substrate is
normally used for energy production and the rest is used for
biomass components such as storage (e.g. Sylvia et al. 1998).
Microbial C used for energy production is rapidly respired,
often within 1–2 days after application of the substrate (Tsai
et al. 1997). This is in accordance with the finding that AM
fungi released 13CO2 within 9–14 h after labelling of field
vegetation (Johnson et al. 2002); and indicates that recently
assimilated C is mainly used for energy production.
However, the C respired represents only part of the C that
has been transferred from plants to AM fungi; a large
fraction is incorporated into fungal structural and storage
compounds. AM fungal neutral lipids are usually stored in
intraradical vesicles or in spores and make up a large
2005 Blackwell Publishing Ltd/CNRS
proportion of the AM fungal biomass (Olsson & Johansen
2000; Bago et al. 2002; Olsson et al. 2002). In G. intraradices,
50–70% of the neutral lipids are the fatty acid 16 : 1x5
(Graham et al. 1995; Olsson & Johansen 2000; Olsson et al.
2002). Dynamics of this compound in our experimental
system indicates that the C assimilated by AM fungi resides
for much longer than 5–6 days, probably in intraradical
vesicles and in intra- and extraradical spores. The finely
branched hyphae, commonly formed along the runner
hyphae, develop rapidly on new hyphae and degenerate
within a few days. Our measurements of C flux together
with the fact that these fine hyphae are emptied of
cytoplasm followed by cross-wall formation (Bago et al.
1998) suggest that C is retracted from degenerating hyphae.
The only fraction where we observed a decrease in C
retention during the experimental period was in the PLFA
16 : 1x5. This signature inside roots probably reflects
arbuscule formation (Van Aarle & Olsson 2003) and may
therefore result from membrane lipids that are degraded
when arbuscules turnover. Arbuscules are known to live
only 1–2 weeks (Alexander et al. 1989).
Although fine absorptive hyphae turnover rapidly, much
of the C contained within AM fungi is translocated into
spores or persistent hyphal networks that may remain in the
soil for long periods. The close correlation we found in 13C
enrichment of the signature lipid NLFA 16 : 1x5 of
extraradical and intraradical hyphae indicate that lipids are
translocated rapidly between fungal structures within the soil
and roots levelling out any imbalance because of new input
or consumption. Indeed, Bago et al. (2002) observed high
rates of lipid transport in hyphae of G. intraradices. Their
Tracking ecosystem carbon 1269
estimate of 4 lm s)1 means that lipids are transported more
than a cm h)1. We suggest that analysis of the dynamics of
signature fatty acids is an integrative approach to accurately
assess rates of C flux through intraradical and extraradical
AM tissues that are dispersed through biologically meaningful volumes of soil. Staddon et al. (2003) concluded that
there was a rapid turnover of AM fungal C, but only hyphae
were collected in their study. This means that any
disappearance of C from the hyphae in their study may be
due to translocation of lipids into spores.
We present data here for only one AM fungal species and
it is clear that the physiology of AM fungal taxa may vary
widely. For example, one family the Gigasporaceae, does
not form intraradical storage vesicles, but instead forms
extraradical auxiliary cells; thus, a higher amount of the C
may be expected to be transported to the extraradical
mycelium in members of this family. This was observed in a
comparison of lipids in Scutellospora calospora, a Gigasporaceae, with G. intraradices, a Glomaceae (Van Aarle & Olsson
2003). This means that the soil C turnover may differ
depending on whether Gigasporaceae or Glomaceae
dominates the ecosystem. Also, the soil environment as a
whole may influence the rate of AM fungal C turnover. In
our system we inoculated the soil with a microbial wash, but
populations of soil microorganisms were probably still
reduced when compared with natural environments. This
means that in field the turnover may be more rapid than in
our system because of a more rapid flux of C to other
trophic levels. In particular soil animals missing in most
experimental systems may have a fundamental influence.
For example, collembola may feed on the hyphae of G.
intraradices (Klironomos & Ursic 1998). Isotopic labelling of
signature fatty acids could also be used to study C turnover
in field-based experiments; however, it is not possible to
estimate total C turnover unless the extent of individual
mycelia can be determined. Carbon turnover may therefore
be difficult to separate from translocation in mycelia in
natural systems.
It is likely that AM turnover rates will differ among
ecosystems depending on longevity of roots and hyphal
networks (Langley & Hungate 2003). For example, AM
fungal tissues might be expected to store more C in perennial
grasslands with persistent hyphal networks compared with
tilled agricultural systems without these networks. Future
research needs to quantify turnover rates of AM hyphae in
conjunction with hyphal architecture from a variety of
environments before reliable estimates of the contribution of
this important group of fungi to SOC can be achieved.
ACKNOWLEDGEMENTS
We are grateful for financial support from Formas and
Crafoord Foundation, the National Science Foundation
(DEB 0316136), and the Fulbright Commission of Sweden.
Helpful suggestions on a previous version of this manuscript by Bruce Hungate, George Koch, Adam Langley and
Mike Miller are gratefully acknowledged.
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Editor, John Klironomos
Manuscript received 14 June 2005
First decision made 17 July 2005
Manuscript accepted 16 August 2005