Plant, Cell and Environment (1998) 21, 881–891
ORIGINAL ARTICLE
OA
220
EN
Mycorrhizal sink strength influences whole plant carbon
balance of Trifolium repens L.
D. P. WRIGHT, D. J. READ & J. D. SCHOLES
Department of Animal and Plant Sciences, University of Sheffield, Sheffield S10 2TN, UK
ABSTRACT
A comparative analysis of daily carbon (C) budgets and
aspects of the C physiology of clover (Trifolium repens L.)
colonized by vesicular-arbuscular (VA) mycorrhizal fungi
was carried out over a 70 d growth period under conditions designed to ensure that shoots of mycorrhizal (M)
and non-mycorrhizal (NM) plants were of similar nutrient
status. C budgets did not differ on day 24 but by day 42 M
plants had a significantly higher rate of photosynthesis
than their NM counterparts when expressed on a whole
shoot basis or unit dry weight basis. As both sets of plants
were of the same size it was concluded that this greater C
gain was the result of increased sink strength provided by
the mycorrhizal fungus. By day 53 M plants had become
larger than their uncolonized counterparts and a sinkinduced stimulation in the rate of photosynthesis was no
longer apparent. M plants had higher root sucrose, glucose
and fructose pools from day 24. Analyses suggested that
these sugars were utilized for trehalose and lipid synthesis,
for the production of the large extramatrical mycelium
and for the support of the respiratory demands of the M
root system. Increased C allocation to roots of M plants
was associated with a stimulation of the activities of cell
wall and cytoplasmic invertases and of sucrose synthase in
roots colonized by VA fungi. Such increases in enzyme
activity may provide the mechanism enabling increased
partitioning of carbohydrate both to the M root system
and the fungal symbiont.
Key-words: Trifolium repens L.; carbohydrates; carbon partitioning; clover; invertase; photosynthesis; sucrose synthase;
vesicular-arbuscular mycorrhiza.
INTRODUCTION
As obligate symbionts, vesicular-arbuscular (VA) mycorrhizal fungi can exert a substantial effect on the carbon (C)
economy of the autotroph (Smith & Read 1997).
Comparison of the C economies of mycorrhizal (M) and
non-mycorrhizal (NM) roots has shown that support of the
symbiosis requires the transfer of an additional 4–20% of
the total net C fixed by the plant (Pang & Paul 1980; Paul
& Kucey 1981; Kucey & Paul 1982; Snellgrove et al.
Correspondence: D. P. Wright. Fax: 0114 2760159;
e-mail: [email protected]
© 1998 Blackwell Science Ltd
1982; Koch & Johnson 1984; Harris, Pacovsky & Paul
1985; Douds, Johnson & Koch 1988; Wang et al. 1989).
While this demand can be seen as the ‘cost’ of the symbiosis, it has been hypothesized that mycorrhizal colonization
of the root, by increasing its sink strength, may stimulate
the process of C assimilation so that the ‘cost’ imposed on
the plant’s C economy is reduced or eliminated and thus
contribute to the overall benefit derived from association
with mycorrhizal fungi (Fitter 1991; Tinker, Durall &
Jones 1994). Some evidence in support of such a hypothesis was provided in an earlier study (Wright, Scholes &
Read 1998), but the mechanisms involved in enhancement
of sink strength have not been investigated in depth. In our
previous study we demonstrated, using M and NM clover
(Trifolium repens L.) plants with similar foliar nitrogen (N)
and phosphorus (P) contents, size and growth rate, that VA
mycorrhizal colonization stimulated the rate of photosynthesis sufficiently both to compensate for the C requirement of the fungus and to eliminate growth reductions of
the autotroph. We hypothesized that the stimulation in the
rate of CO2 fixation may have been caused by an increased
sink strength arising from the additional C requirement of
the mycorrhizal fungus colonizing the roots of the plant.
To further investigate this hypothesis it is necessary to
carry out comparative analyses of diurnal C budgets of M
and NM plants and to examine the partitioning of C into
different non-structural carbohydrate pools in the leaves
and roots of the autotroph and into fungal-specific compounds within the mycobiont in M roots. To determine
the mechanism by which M colonization stimulates the
movement of C to the root we have investigated the possible involvement of the sucrolytic enzymes, invertase
and sucrose synthase. These enzymes have been shown to
be involved in the regulation of the sink strength of plant
tissues (Ho 1988), but have not been studied in VA
mycorrhizal associations. However, they have been
strongly implicated in the diversion of carbohydrates to
the mycobiont in other plant/fungus associations (Long
et al. 1975; Greenland & Lewis 1983; Scholes et al.
1994; Wright et al. 1995).
In this study we have again manipulated M and NM
clover plants with the aim of producing similar foliar N and
P status, size and growth rate, in order to examine the
hypothesis that the presence of the fungal symbiont fundamentally influences the pattern of assimilation and allocation of C in the plant. Gas exchange techniques have been
881
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used to produce diurnal C budgets for M and NM plants.
The amounts of soluble carbohydrates and starch in the
leaves and roots and of lipid and the fungal sugar trehalose
in the roots were measured. Finally, the roles of invertase
and sucrose synthase in regulating the sink strength of M
compared with NM roots was investigated.
MATERIALS AND METHODS
Growth and maintenance of clover (Trifolium repens L.)
plants was carried out as described in Wright et al. (1998).
Clover seeds were germinated on moist filter paper for 4 d.
Mycorrhizal colonization of germinated seedlings was
induced by transplantation into seed trays containing autoclaved dune sand mixed with chopped, infected root material. Infected root material was obtained from pot cultures
of clover colonized by VA mycorrhizal fungi isolated from
grassland turfs collected from Aber, North Wales. The aim
was to obtain M plants which were colonized by a population of VA mycorrhizal fungi representative of those seen
in a natural community. Seedlings were maintained in
these trays for 5 weeks to allow colonization of the clover
roots, during which time the plants were watered with distilled water only. NM plants were grown in trays of dune
sand from which root material was omitted and supplemented with 50% Long Ashton nutrient solution to produce seedlings of similar biomass to M seedlings at the end
of the colonization stage. A rhizobial inoculant (Nodulaid,
Elsoms Seeds Ltd, Spalding, Lincolnshire, UK) was added
to the dune sand prior to use to ensure uniform nodulation
of M and NM plants. The seedlings were maintained in a
Fisons Fitotron growth cabinet at 20 °C with a 16 h photoperiod (360 µmol m–2 s–1 irradiance). After the 5 week
colonization stage, M and NM plants were transferred into
individual pots (25 cm long and 7·5 cm in diameter) filled
with autoclaved dune sand. Each pot received 20 cm3 of
50% Long Ashton nutrient solution once weekly and was
watered with distilled water on all other days. Nutrient
analysis of the dune sand showed it to contain only trace
quantities of macronutrients.
Diurnal CO2 exchange of M and NM clover plants:
chamber and system design and operation
To measure diurnal CO2 exchange from the shoot and from
the root environment, the plants were enclosed in sealed
plant chambers similar in design to those of Eissenstat
et al. (1993). The plants were grown in cylindrical plastic
pots 25 cm long (of which the top 1·5 cm was removable)
and with a diameter of 7·5 cm. To measure CO2 exchange
from the root and sand the top 1·5 cm portion of the plastic
pot was removed and the pot lowered into the root compartment (25 cm long and 11 cm diameter) of the plant
chamber. The shoot was enclosed in a clear perspex cylindrical compartment (25 cm long and 15 cm diameter). The
atmospheres in the shoot and root compartments were separated by two perspex sheets which when brought together
formed a central hole (7 mm diameter) through which the
plant stem could pass. These sheets and the shoot and root
compartments were held together by four wingnuts.
Silicon grease was used to create an airtight seal between
each compartment and the two sheets, and around the plant
stem. Turbulent air flow within the shoot compartment was
provided by a fan. The chambers were not temperature
controlled, but the temperature within the chamber was
23 ± 2 °C. Air was supplied to the base of each compartment and exited via a port at the top of each compartment.
Air supplied to the root compartment mixed with the root
atmosphere around the top 1·5 cm of the pot only to minimize disturbance of the root atmosphere.
Eight plant chambers were used simultaneously in an
open gas exchange system. Compressed air was passed
through a pressure regulator to reduce the pressure in the
airline to atmospheric pressure. After passing through a
20 dm3 ballast to mix the air thoroughly the airline was
split into two and passed to one of two manifolds each
housing nine needle valves. One manifold supplied air via
eight needle valves to the eight root compartments, the
other manifold supplied air to the shoot compartments. The
ninth needle valve on each manifold supplied prechamber
reference gas to one of two infrared gas analysers (IRGA)
(LCA4, Analytical Development Company, Hoddesdon,
UK), one to analyse gas flowing from the root compartments and one to analyse shoot compartment gases. The
rate of flow of air through each shoot and root compartment was set using a flow meter (Fisher Controls Ltd,
Croydon, UK), by adjustment of the needle valve supplying each compartment. Flow rates were varied between
200 and 500 cm3 min–1 depending on the size of the plants.
From each root or shoot compartment the analysis gas
passed to a dedicated three-way solenoid valve. The air
passing from each root compartment was dried with silica
gel before the solenoid valve. When the solenoid valve was
switched off the analysis gas from the compartment was
vented to waste. When the solenoid valve was powered the
gas from the compartment was passed to the appropriate
IRGA and analysed against the prechamber reference gas
supplied directly to the IRGA from the manifold. The pair
of solenoid valves controlling the gas flow from the root
and shoot compartments of each plant chamber was
switched on and off simultaneously using a programmable
logic controller (model Melsec FX-48MR, Mitsubishi
Electric). Each pair of valves was powered for 10 min
every 80 min in a continuous loop. During each 10 min
period five separate recordings of the CO2 and H2O concentrations in the reference and sample gas streams of the
root or shoot compartments were stored by each IRGA.
Record collection by each IRGA was initiated by the
Melsec FX-48MR controller. All gas tubing within the system was connected using Bosch Quickfix couplings
(Bosch, Stuttgart, Germany).
Diurnal CO2 exchange from the shoots and roots
of M and NM clover plants
Diurnal CO2 exchange from the shoot and from the root
© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
Carbon allocation in mycorrhizal clover
and sand environments of four M and four NM plants was
measured simultaneously on days 24, 42 and 53.
Conditions in the growth chamber were as described
above. The CO2 concentration of the air supplied to the
plant chambers was 380 cm3 m–3. The plants were sealed
into the chambers and allowed to equilibrate for at least 6 h
prior to initiation of data recording and measurements were
taken over the subsequent 36 h period. After the measurement period the plants were harvested and the leaf, stem
and root material was dried at 70 °C until constant weight
was achieved. The rate of whole shoot and the rate of combined root and sand CO2 exchange was calculated as
µmol CO2 g–1 DW s–1. Using these data the daily C gain
(µmol CO2 plant–1 d–1 or mmol CO2 g–1 DW d–1) of M and
NM plants on days 24, 42 and 53 was determined.
Determination of soluble sugars and starch in the
leaves and roots of M and NM clover plants
Leaf and root material from five randomly selected M and
NM clover plants were rapidly harvested 9 h into the photoperiod on days 18, 38 and 52, their fresh weight determined, then frozen in liquid nitrogen. The tissue was
extracted in 4 volumes ethanol: 1 volume water,
50 mol m–3 Hepes-KOH (pH 7·4) and 5 mol m–3 MgCl2
(three changes of 2 cm3 each for 45 min) at 80 °C. The
aliquots were combined and dried under vacuum using a
Speedvac centrifuge and refrigerated vapour trap (models
SC110 and RVT100, Savant Industries Inc., Farmingdale,
NY, USA). The extract was resuspended in 1 cm3 of
100 mol m–3 Hepes-KOH (pH 7·4), 5 mol m–3 MgCl2 and
assayed for glucose, fructose and sucrose by enzymelinked assay as described by Scholes et al. (1994). Starch
was determined from the tissue remaining after extraction
of soluble sugars. The tissue was washed in three 1 cm3
aliquots of distilled water, blotted, then ground in 1 cm3 of
500 mol m–3 2-(N-Morpholino) ethanesulphonic acid
(Mes) buffer (pH 4·5) using a glass homogenizer. A
200 mm3 aliquot of the extract was transferred to a screw
cap microcentrifuge tube and autoclaved for 45 min.
Starch was hydrolysed by the addition of 20 units α-amylase and 14 units of amyloglucosidase (Boehringer
Mannheim, UK) for 4 h at 37 °C. The samples were centrifuged (13 000 g for 20 min) and an aliquot assayed for
glucose as described by Scholes et al. (1994). Rehydrolysis
of the pellet with α-amylase and amyloglucosidase confirmed that all the starch had been digested.
Determination of invertase activity in the roots of
M and NM clover plants
Clover roots from M and NM plants were rapidly harvested, their fresh weight determined, then frozen in liquid
nitrogen. The frozen tissue was transferred to a mortar containing liquid nitrogen and a 1 cm3 pellet of extraction
buffer (all as mol m–3: Hepes-KOH (pH 8·0), 50; MgCl2, 5;
ethylenediaminotetraacetic acid (EDTA), 2; MnCl2, 1;
CaCl2, 1; benzamidine, 1; dithiotreitol, 1; phenyl-methyl© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
883
sulphonyl fluoride, 0·1) and ground to a fine powder. Upon
thawing the extract was transferred to a microcentrifuge
tube and the mortar washed with 1 cm3 of extraction buffer.
The total homogenate was mixed thoroughly and divided
into two 1 cm3 aliquots. One aliquot was reserved for total
activity determinations. The second aliquot was centrifuged (13 000 g for 15 min) and the supernatant containing the vacuolar and cytoplasmic invertase activities was
transferred to a new microcentrifuge tube. The pellet containing the cell wall invertase activity was washed three
times, each using 1 cm3 of extraction buffer, before final
resuspension in 1 cm3 of extraction buffer. The reaction
mixture contained 80 mol m–3 citrate/phosphate buffer,
200 mol m–3 sucrose and 50 mm3 of extract in a final volume of 500 mm3. After incubation at 37 °C for 30 min,
100 mm3 of 1000 mol m–3 Tris–HCl (pH 10·0) was added
to neutralize acidic mixtures and inactivate cytoplasmic
invertase prior to heating to 95 °C for 4 min. Two types of
blanks were set up. One was designed to determine the
endogenous sugar level and consisted of boiled extract in
buffer without sucrose. The other, designed to detect any
acid hydrolysis of sucrose during the assay procedure, consisted of buffer and sucrose but no extract. Glucose and
fructose produced in the assay were determined by
enzyme-linked assay, as described by Scholes et al. (1994).
Roots of M and NM plants were harvested on day 0 and the
pH optimum of the cell wall, cytosolic and vacuolar invertases was determined using citrate/phosphate buffers, over
the pH range 3·5–9·0. Roots from five M and five NM
plants were harvested on days 0, 28, 48 and 70 and the
activities of the cell wall, cytosolic and vacuolar invertases
were measured at their pH optima. The activities of the cell
wall, vacuolar and cytoplasmic invertases were also measured in root segments of M and NM plants which were
free from nodules on day 35, to determine how nodulation
might influence the activity of invertase in both root types.
Determination of sucrose synthase activity in the
roots of M and NM clover plants
Sucrose synthase activity was determined in the sucrose
hydrolysis direction. The pH optimum of sucrose synthase
activity from roots of M and NM plants, harvested on day
0, was determined using the supernatant fraction produced
during the extraction procedure described above for vacuolar and cytoplasmic invertase activity measurements. Over
the pH range 5–7 the reaction mixture contained
100 mol m–3 Mes, 1 mol m–3 MgCl2, 400 mol m–3 sucrose,
5 mol m–3 uridine diphosphate (UDP) and 20 mm3 of
supernatant extract in a final volume of 100 mm3. For the
pH range 7–9 the reaction mixture was identical, except
that 50 mol m–3 Hepes-KOH buffer was used. Incubation
was carried out at 37 °C for 30 min before the reaction was
stopped by heating to 95 °C for 4 min. Blanks contained
the same reaction mixture except the extract was inactivated by heating to 95 °C for 4 min. The amount of UDPglucose produced during the incubation was determined in
200 mol m–3 glycine-KOH (pH 8·9), 10 mol m–3 NAD and
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D. P. Wright et al.
0·01 U UDP-glucose dehydrogenase. The activity of
sucrose synthase was determined on days 0, 28, 48 and 70,
using the supernatant fraction extracted from M and NM
roots to measure the activity of the vacuolar and cytoplasmic invertases described above. In addition, the activity of
sucrose synthase from root segments of M and NM plants
which were free of nodules was measured, at its pH optimum, to determined how nodulation influenced the activity of sucrose synthase.
Determination of trehalose in the roots of M and
NM clover plants
Soluble sugars were extracted from the roots of five M and
five NM clover plants, 9 h into the photoperiod, on days
10, 28, 48 and 70 as described above, except that 4 volumes ethanol: 1 volume water was used and the extract was
resuspended in 1 cm3 of ultrapure water. The neutral fraction of the soluble extract was collected by passing an
aliquot of the extract through a Dowex-50 (100–200 mesh,
8% cross-linked, Sigma Chemical Company, UK) column
situated above a Dowex-1 (100–200 mesh, 8% crosslinked) column. A 0·5 cm3 aliquot of the extract was loaded
on to the Dowex-50 column and eluted through both
columns using 10 0·3 cm3 aliquots of ultrapure water. The
combined fractions were transferred to a 5 cm3 pearshaped flask and dried under a stream of compressed air.
Trimethysilyl carbohydrate derivatives were prepared by
dissolving the dried extract in 0·8 cm3 pyridine and 0·2 cm3
trimethylsilylimidazole (TSIM) and heating the stoppered
flasks to 60 °C for 1 h. Quantitative analysis of trehalose
was carried out using a gas–liquid chromatograph (Varian
3500) equipped with a flame ionization detector and a DB5 non-polar column (25 m × 0·22 mm) (J & W Scientific,
Folsom, CA, USA). The GC temperature settings were:
injector 280 °C, detector 375 °C and column 170 °C
increasing by 5 °C min–1 to 325 °C. Carbohydrate peaks
were identified by comparing their retention times with
prepared standards and quantified by digital integration of
the peak areas. Recovery of a trehalose standard was
93 ± 6%.
Determination of total lipid in the roots of M and
NM clover plants
Plant material from five M and five NM plants was harvested on days 10, 24, 48 and 70, weighed, then placed into
boiling isopropanol for 15 min to inactivate lipophylic
enzymes (Kates 1957), then stored at – 20 °C until extraction. Samples were ground in 2 cm3 2 volumes chloroform:
1 volume methanol using a glass homogenizer. The extract
was centrifuged (3500 g for 10 min) and the supernatant
decanted into a rotary evaporator flask. The pellet was
washed twice using 2 cm3 of 2 volumes chloroform: 1 volume methanol followed by three washes each of 2 cm3 1
volume chloroform: 2 volumes methanol. The combined
supernatants were dried using a rotary evaporator and a
water bath at 45 °C. The dried extract was resuspended in
2 cm3 Folch lower phase solvent (86 volumes chloroform,
14 volumes methanol, 1 volume water) (Folch, Lees &
Sloane-Stanley 1957) and washed twice using Folch upper
phase solvent (3 volumes chloroform, 48 volumes
methanol, 47 volumes water). The upper phase solvents
were decanted to a separate flask and washed with Folch
lower phase solvent. The combined lower phase aliquots
were again dried down using a rotary evaporator, taken up
in 1 cm3 of chloroform and stored at – 20 °C until assayed.
Total lipid was estimated by the acid charring method of
Marsh & Weinstein (1966) using olive oil as a standard. To
avoid oxidation, 50 g m–3 butylated hydroxtoluene was
added to all solvents.
Determination of the length of fungal hyphae in
the sand surrounding M and NM clover roots
The length of external hyphae in the sand surrounding the
roots of M and NM plants was estimated using a method
similar to that of Abbott, Robson & De Boer (1984). On
days 28 and 48 the sand was removed from the pots of five
M and five NM plants and the roots removed. After thorough mixing, a 5 g subsample of sand was placed into a
Waring blender with 200 cm3 of deionized water and
blended at full speed for 20 s. After allowing the suspension to settle for 10 s, a 10 cm3 aliquot was pipetted on to a
millipore filter (0·45 µm pore size, 25 mm diameter). After
filtration the sample was stained with 500 g m–3 trypan
blue in lactoglycerol (1 volume lactic acid, 1 volume glycerol, 1 volume water) for 5 min, rinsed with deionized
water then transferred to a microscope slide and allowed to
dry. Dried filters were mounted in 1 volume glycerol: 1
volume water. Hyphal lengths were estimated using the
line intersect method (Tennant 1975). The number of intercepts over an eyepiece grid of 10 × 10 squares was counted
for 25 fields of view per slide. A 5 g sand subsample was
dried at 80 °C to determine the moisture content of the sand
and hyphal length expressed as m g–1 dry sand.
Colonization of clover roots
The roots of M and NM plants were cleared in 100 kg m–3
KOH overnight. The roots were rinsed in 1 volume HCl: 9
volumes water for 20 min and then placed in 500 g m–3 trypan blue in lactoglycerol (1 volume lactic acid, 1 volume
glycerol, 1 volume water) for 1 week to stain the fungal
structures. The roots were destained using several changes
of 1 volume glycerol: 1 volume water and mounted in glycerol on a microscope slide. The percentage colonization of
roots by M fungi (number of intercepts at which colonization was observed expressed as a percentage of the total
number of observations) was determined using the grid
line intersect method (Giovannetti & Mosse 1980).
Statistical analysis
The data were subjected to one-way analysis of variance
(ANOVA) using the statistical package Minitab 10·2.
© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
Carbon allocation in mycorrhizal clover
885
Table 1. The total dry weight (mg), the dry weight of the leaves, stems and stolons and the roots of mycorrhizal and non-mycorrhizal clover
plants used for diurnal gas exchange analyses. Values represent the mean ± standard error (n = 4)
Day 24
Day 42
Day 53
Tissue type
Non-mycorrhizal
Mycorrhizal
Non-mycorrhizal
Mycorrhizal
Non-mycorrhizal
Mycorrhizal
Total dry weight
Leaf
Stem and stolon
Root
120·1 ± 3·8
40·5 ± 1·6
35·8 ± 2·8
43·9 ± 1·6
116·3 ± 13·8
36·3 ± 3·7
31·3 ± 0·8
48·7 ± 9·9
434·9 ± 89·6
148·6 ± 32·0
145·4 ± 36·6
140·9 ± 26·4
509·0 ± 56·7
150·6 ± 12·1
198·5 ± 18·5
160·0 ± 26·8
766·4 ± 86·7
224·2 ± 27·2
306·4 ± 54·6
235·8 ± 38·6
1105·2 ± 17·3*
1330·6 ± 26·5*
1450·3 ± 37·7
1324·3 ± 46·7
On each day, the data for each parameter for mycorrhizal and non-mycorrhizal plants were compared by one-way ANOVA. * P < 0·05.
RESULTS
Diurnal C budgets of M and NM clover plants
The dry weight of the leaves, stems and stolons, and roots
and the total dry weight of M plants were not significantly
different from those of NM plants on days 24 and 42
(Table 1). However, by day 53 leaf dry weight and total dry
weight of M plants were significantly increased compared
with those of NM plants (P < 0·05).
On day 24 there were no significant differences in the
net CO2 assimilation, the amount of CO2 respired by the
shoots or roots or in the net C gain of whole M compared
with NM plants expressed on a per plant or per unit dry
weight basis (Tables 2 & 3).
By day 42 significant differences in diurnal CO2
exchange of M and NM plants were apparent (Fig. 1). At
this time, the amount of CO2 assimilated by the shoots of M
plants, expressed per plant or per unit dry weight, was significantly greater than that assimilated by the shoots of NM
plants, such that the diurnal C gain of M plants was significantly greater than that of NM plants (P < 0·05) (Tables 2 &
3). Also at this stage, the amount of C respired by the M root
system, expressed per plant, was significantly increased
compared with that respired by NM root systems, but was
not significant when expressed per unit dry weight
(P < 0·05). The mean net C gain of the M plant and fungus
was greater than that of the NM plants, expressed on either
basis, although these differences were not significant.
By day 53 the C gain of the shoots, root system respiration and the C gain of the M plant and fungus had become
significantly greater than those of NM plants when
expressed on a per plant basis (P < 0·05) (Table 2).
However, when these data were recalculated on a unit dry
weight basis no significant differences between M and NM
plants were observed (Table 3).
The amount of soluble carbohydrates, starch,
total lipid and trehalose in M and NM clover
plants
The amount of sucrose and of glucose and fructose in the
leaves of M plants was not significantly different to that in
the leaves of NM plants at any time during the experiment
(Fig. 2a & b). In contrast, the amount of sucrose was significantly increased in M compared with NM roots from
day 38 onwards (P < 0·01) (Fig. 2c). M roots also contained a significantly higher amount of glucose and fructose compared with NM roots throughout the duration of
the experiment (P < 0·05) (Fig. 2d). With the exception of
day 18, starch accumulated to a significantly higher
amount in the leaves of M clover compared with those of
NM plants (P < 0·05) (Fig. 3). However, accumulation of
starch in the roots of M plants was only significantly higher
on day 52 (P < 0·01) (Fig. 3).
The amount of the fungal sugar trehalose in the roots of M
plants was variable (Fig. 4). However, during the experiment
Table 2. Diurnal carbon budgets (µmol CO2 plant–1 d–1) of mycorrhizal (M) and non-mycorrhizal (NM) clover plants (per plant basis). Data
represent the mean ± standard error (n = 4)
Day 24
Parameter
Non-mycorrhizal
Net CO2 assimilation
184 ± 25
Dark shoot respiration
36 ± 4
Carbon gain of shoot
148 ± 21
Root and sand respiration 127 ± 18
Carbon gain of
21 ± 6
whole plant (NM) and
plant and fungus (M)
Day 42
Day 53
Mycorrhizal
Non-mycorrhizal
Mycorrhizal
Non-mycorrhizal
Mycorrhizal
195 ± 9 ns
34 ± 5 ns
160 ± 7 ns
138 ± 12 ns
22 ± 11 ns
768 ± 99
89 ± 12
679 ± 91
328 ± 39
351 ± 75
1179 ± 165*
125 ± 12 ns
1054 ± 155*
502 ± 88*
552 ± 157 ns
1365 ± 217
247 ± 29
1118 ± 205
685 ± 90
433 ± 175
2215 ± 290*
336 ± 40 ns
1879 ± 274*
990 ± 130*
889 ± 122*
On each day, the data for each parameter for mycorrhizal and non-mycorrhizal plants were compared by one-way ANOVA. ns, not significant; *
P < 0·05.
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D. P. Wright et al.
Table 3. Diurnal carbon budgets (mmol CO2 g–1 DW d–1) of mycorrhizal (M) and non-mycorrhizal (NM) clover plants (unit dry weight
basis). Data represent the mean ± standard error (n = 4)
Day 24
Parameter
Non-mycorrhizal
Net CO2 assimilation
Dark shoot respiration
Carbon gain of shoot
Root and sand respiration
Carbon gain of
whole plant (NM) and
plant and fungus (M)
4·56 ± 0·60
0·92 ± 0·11
3·64 ± 0·49
3·04 ± 0·42
0·60 ± 0·27
Day 42
Mycorrhizal
Day 53
Non-mycorrhizal
5·46 ± 0·47 ns
0·97 ± 0·17 ns
4·49 ± 0·37 ns
3·48 ± 0·53 ns
1·01 ± 0·31 ns
5·41 ± 0·71
0·62 ± 0·07
4·79 ± 0·67
2·53 ± 0·58
2·26 ± 0·74
Mycorrhizal
7·76 ± 0·52*
0·83 ± 0·04*
6·93 ± 0·51*
3·33 ± 0·23 ns
3·60 ± 0·77 ns
Non-mycorrhizal
6·14 ± 0·83
1·10 ± 0·03
5·04 ± 0·83
2·95 ± 0·14
2·78 ± 1·04
Mycorrhizal
6·45 ± 0·45 ns
1·02 ± 0·10 ns
5·43 ± 0·47 ns
3·23 ± 0·95 ns
2·20 ± 0·42 ns
On each day, the data for each parameter for mycorrhizal and non-mycorrhizal plants were compared by one-way ANOVA. ns, not significant;
* P < 0·05.
there was on average 5 µmol trehalose g–1 DW root or
10 µmol glucose equivalent g–1 DW root which represents
between 7 and 17% of the total soluble carbohydrate
(sucrose, glucose and fructose) pool in the roots of M plants
during the course of the experiment. Trehalose was not
detected in the roots of NM plants. No other carbohydrate or
polyol peaks were observed which were unique to M roots. In
general, the amount of total lipid in the roots of M plants was
two to three times higher than the amount present in NM
clover roots throughout the experiment (P < 0·05) (Fig. 4).
Invertase activity in the roots of M and NM clover
plants
The pH optima for the cell wall, vacuolar and cytoplasmic
invertases were pH 4·5, 5·5 and 7·5, respectively, in both M
and NM roots (Fig. 5). All assays were subsequently car-
Figure 1. The diurnal rate of whole shoot CO2 exchange and the
diurnal rate of root and sand respiration of mycorrhizal (M; l) and
non-mycorrhizal (NM; ¡) clover plants. Data represent the
mean ± standard error of M and NM plants measured
simultaneously on day 42 (n = 4).
Figure 2. The amount of sucrose and the sum of glucose and
fructose in the leaves and roots of mycorrhizal (M; n) and nonmycorrhizal (NM; o) clover plants on days 18, 38 and 52.
Samples were taken 9 h into the photoperiod. Data represent the
mean ± standard error (n = 5).
ried out at these pH optima. The activity of cell wall invertase was significantly higher in M roots compared with
NM roots throughout the duration of the experiment
(Fig. 6). However, although initially significantly higher
(P < 0·05) in M roots, the activity of vacuolar invertase was
similar in the roots of M and NM plants by the end of the
experiment (Fig. 6). The activity of cytoplasmic invertase
was significantly higher in the roots of M compared with
NM plants throughout the experiment (P < 0·05) (Fig. 6).
To determine whether nodulation influenced the activity of
the individual invertases, the activity of these enzymes was
assayed in non-nodulated root segments from both sets of
plants on day 35. In the absence of nodules the activities of
the cell wall and cytoplasmic invertases were still significantly increased in root segments from M compared with
NM plants, confirming that these enhanced activities were
due to the presence of the mycorrhizal fungus (Table 4).
However, the activity of the vacuolar invertase of non© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
Carbon allocation in mycorrhizal clover
887
quent assays were performed at this pH (Fig. 7). Sucrose
synthase activity of M roots was significantly higher than
that of NM roots on days 0 and 28 (P < 0·001) (Fig. 8).
However, the activity of sucrose synthase of M roots was
not significantly different from that of NM roots on days
48 and 70 (Fig. 8). Even in the absence of nodules the
activity of sucrose synthase in M root segments was significantly higher than that in NM root segments on day 35
(P < 0·001) (Table 4).
The length of external hyphae in the sand from
pots of M and NM clover plants
Figure 3. The amount of starch in the leaves and roots of
mycorrhizal (M; n) and non-mycorrhizal (NM; o) clover plants
on days 18, 38 and 52. Samples were taken 9 h into the
photoperiod. Data represent the mean ± standard error (n = 5).
The length of external hyphae in the sand of pots which
contained M plants was significantly higher than that in
pots containing NM plants throughout the experiment
(P < 0·001). The length of external hyphae was 20·5 ±
2·2 m hyphae g–1 DW sand and 1·2 ± 0·6 m hyphae g–1 DW
sand in M and NM pots, respectively, on day 28 and was
Figure 4. The amount of trehalose and the amount of total lipids
in the roots of mycorrhizal (M; l) and non-mycorrhizal (NM; ¡)
clover plants on days 10, 28, 48 and 70. Data represent the
mean ± standard error (n = 5).
nodulated M root segments was not significantly different
from that of non-nodulated NM root segments (Table 4).
Sucrose synthase activity in the roots of M and
NM clover plants
Sucrose synthase from both sets of plants exhibited a broad
peak of activity with a pH optimum of 7·5 and all subse© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
Figure 5. Influence of pH on the activity of invertase in the roots
of mycorrhizal (M; l) and non-mycorrhizal (NM; ¡) clover
plants: (a) total invertase activity of the whole root homogenate;
(b) invertase activity associated with the cell wall; and (c) cytosolic
invertase activity. The pH optima for the cell wall, vacuolar and
cytoplasmic invertase enzymes were 4·5, 5·5 and 7·5, respectively.
Data represent the mean ± standard error (n = 5).
888
D. P. Wright et al.
Table 4. The activity of sucrose synthase and invertase enzymes
from non-nodulated mycorrhizal and non-mycorrhizal clover root
segments on day 35. Data represent the mean ± standard error
(n = 5)
Activity (µmol glucose
equivalent g–1 DW root min–1)
Enzyme
Non-mycorrhizal Mycorrhizal
Invertase
Cell wall bound pH 4·5
Vacuolar pH 5·5
Cytoplasmic pH 7·5
Sucrose synthase
4·15 ± 0·21
1·89 ± 0·25
3·48 ± 0·27
7·02 ± 0·22
8·98 ± 0·61***
1·18 ± 0·37 ns
4·95 ± 0·69*
24·2 ± 1·98***
One-way ANOVA was performed on the data. ns, not significant; ***
P < 0·001, * P < 0·05.
Figure 6. The activity of cell wall, vacuolar and cytoplasmic
invertase enzymes from the roots of mycorrhizal (M; l) and nonmycorrhizal (NM; ¡) clover plants on days 0, 28, 48 and 70. Data
represent the mean ± standard error (n = 5).
43·6 ± 4·6 m hyphae g–1 DW sand and 3·6 ± 1·5 m hyphae
g–1 DW sand in M and NM pots, respectively, on day 48.
Figure 7. Influence of pH on the activity of sucrose synthase in
the roots of mycorrhizal (M; l, ▲) and non-mycorrhizal (NM; ¡,
▲) clover plants. The reaction mixture was buffered over the pH
range 5–7 using 100 mol m–3 Mes (¡, l) and over the pH range
▲, ▲). Data represent the
7–9 using 50 mol m–3 Hepes-KOH (▲
mean ± standard error (n = 5).
Mycorrhizal colonization and nodulation of
clover roots
At least 83% of the root length was colonized during this
experiment. Numerous vesicles were also observed within
colonized roots. Non-mycorrhizal controls remained free
of infection. We have previously demonstrated that the dry
weight of nodules present on the roots of M plants was not
significantly different from that on the roots of NM plants
on days 18 and 51 (Wright et al. 1998).
DISCUSSION
We have previously shown, using M and NM clover plants,
with similar foliar P and N concentrations, that mycorrhizal colonization resulted in a stimulation of the rate of
photosynthesis of young leaves but that the additional C
gained was not converted to biomass production (Wright
et al. 1998). We hypothesized that the stimulation in the
Figure 8. The activity of sucrose synthase in the roots of
mycorrhizal (M; l) and non-mycorrhizal (NM; ¡) clover plants
on days 0, 28, 48 and 70. Data represent the mean ± standard error
(n = 5).
© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
Carbon allocation in mycorrhizal clover
rate of photosynthesis was due to the additional fungal sink
arising through mycorrhizal colonization of the roots of the
autotroph. In this present study, we have further examined
CO2 fixation by M and NM clover plants and the partitioning of C within the association to determine the extent and
the mechanism underlying this effect.
Analysis of diurnal C budgets revealed that on day 24 no
significant difference occurred between M and NM plants
but by day 42 the shoots of M plants had a significantly
higher C gain than those of their NM counterparts even
though their dry weights were not significantly different.
This suggests that the additional assimilate gain was used
to support the growth and maintenance of the fungal symbiont. The additional amount of CO2 respired by M compared with NM root systems as a percentage of the net
amount of CO2 assimilated by M plants was 15% on day
42. This is in good agreement with previous studies which
have shown that the additional amount of 14C respired by
roots of 14CO2-labelled M plants ranged from 4 to 20% of
the total net CO2 fixed by the plant (Pang & Paul 1980;
Paul & Kucey 1981; Kucey & Paul 1982; Snellgrove et al.
1982; Koch & Johnson 1984; Harris et al. 1985; Douds
et al. 1988; Wang et al. 1989).
By day 53, that the C budgets were still significantly different between M and NM plants when expressed on a per
plant basis, could be attributed to the fact that M plants
had, for the first time, become significantly larger than
those in the NM condition. The fact that the growth of M
plants did eventually outpace that of plants grown in the
NM condition could indicate that the superior potential for
nutrient capture was eventually expressed in colonized
plants and may have masked any sink effects. In the only
other study to examine diurnal C budgets of M plants, Peng
et al. (1993), using Citrus volkameriana mycorrhizal with
Glomus intraradices, observed no stimulation in the rate of
photosynthesis in M leaves, although a higher rate of respiration in M roots resulted in a lower diurnal C gain in such
plants and a reduction in the growth of M compared with
NM plants.
In this study we have demonstrated that the pools of
sucrose, starch and glucose and fructose in the leaves of M
and NM plants were similar. However, the pools of sucrose
and glucose and fructose were consistently increased in M
compared with NM roots. We suggest that these data indicate that the pattern of C allocation within M plants was
altered so that an increased proportion of C assimilated
was partitioned to the roots of M plants. Several studies
have reported a decline in the root/shoot ratio upon mycorrhizal colonization, particularly in non-matched M and
NM plants (Hayman & Mosse 1971; Sanders 1975; Smith
1982; Hall, Johnstone & Dolby 1984; Thomson, Robson &
Abbott 1986; Bass & Lambers 1988; Fredeen & Terry
1988; Smith & Gianinazzi-Pearson 1990). A smaller
root/shoot ratio may occur as a result of increased partitioning of C to the fungus at the expense of root production
whilst maintaining an overall neutral effect on C allocation
within the plant. However, contrary to the above studies,
both in this and in our previous study (Wright et al. 1998),
© 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 881–891
889
we have shown that there was no significant difference in
the dry weight of M and NM roots. We suggest that the
increased pools of C in M roots were sufficient both to support root growth and the growth, maintenance or storage of
C in the fungal partner. The latter is supported by three
pieces of evidence. First, the fungus-specific sugar trehalose was consistently present in M roots corresponding
to up to 17% of the soluble carbohydrate pool of these
roots. Trehalose has previously been observed in fungal
spores and roots of other VAM associations (Becard et al.
1991; Schubert, Wyss & Wiemken 1992; Shacher-Hill
et al. 1995). As the total soluble carbohydrate pool of M
plants was itself considerably larger than that of NM
plants, the amount of trehalose observed may represent a
considerable allocation of the plant’s C to the fungus, particularly in comparison with the C budget of NM plants.
Second, the total lipid pool of M roots was also consistently higher than that of NM roots and was probably primarily accounted for by the large quantity of fungal
membranes and lipid-rich vesicles present within the
highly colonized M root systems. An increase in the lipid
content of M roots has previously been observed (Cooper
& Losel 1978; Pacovsky & Fuller 1988). Peng et al. (1993)
showed that M fibrous roots of Volkamer lemon had significantly higher fatty acid concentrations than NM fibrous
roots and estimated that 10% of the additional root respiration, associated with mycorrhizal colonization, was
attributable to construction of lipid-rich M roots. Third, the
production of large amounts of mycorrhizal hyphae
observed in this study suggest the allocation of a considerable amount of C from the autotroph to the fungal partner.
Reports of the effect of mycorrhizal colonization on the
carbohydrate pools of M and NM plants with similar foliar
nutrient content are mixed. In Allium porrum mycorrhizal
with Glomus mosseae, the amount of glucose and fructose
was similar in the roots of M and NM plants with similar
foliar P concentrations, although the amount of sucrose in
M roots tended to be increased (Amijee, Stribley & Tinker
1993). When Glycine max was grown in association with
Glomus fasciculatum, the amounts of sucrose, starch and
anthrone-positive sugars in the leaves and roots were found
to be either similar to or lower than those in NM plants
supplemented with P (Pacovsky 1989). Higher amounts of
starch were also observed in leaves of G. max mycorrhizal
with G. mosseae compared with NM, P-supplemented
leaves (Brown & Bethlenfalvay 1988). In a variety of citrus genotypes, Graham, Duncan & Eissenstat (1997)
showed that the total non-structural carbohydrate pool
increased from the least to the most mycorrhizal dependent
genotype. At high P supply, when the biomass and P status
of M and NM plants were similar, the roots and leaves of
the most mycorrhizal-dependent genotypes had lower
sucrose pools but higher reducing sugar and starch pools
than their NM counterparts.
Several possible mechanisms have been proposed to
explain the increased sink strength arising from mycorrhizal colonization of the roots of plants. These include
rapid utilization of C and its conversion into fungal-specific
890
D. P. Wright et al.
compounds (Bevege, Bowen & Skinner 1975; Losel &
Cooper 1979) and increased respiration by the M root
system (Pang & Paul 1980; Snellgrove et al. 1982; Harris
et al. 1985), both of which occurred in this study. However,
our results indicate that increased activity of sucrolytic
enzymes represents a further mechanism contributing to the
increased sink strength arising from mycorrhizal colonization of the roots. We have demonstrated that both the activities of the cell wall and cytoplasmic invertases and sucrose
synthase were stimulated solely in response to mycorrhizal
colonization. These enzymes have been implicated in the
regulation of the sink strength of plant tissues (Ho 1988;
Sung, Xu & Black 1989; Wang et al. 1993; Sung et al.
1994; Zrenner et al. 1995). It is interesting that the largest
stimulation in the activities of these enzymes in M roots
occurred at, or just prior to, a large increase in the production of trehalose and lipids in these roots and a stimulation
in the rate of CO2 assimilation on day 42.
There are no previous reports of alterations in the activity of invertase or sucrose synthase in response to VA mycorrhizal colonization. Schaeffer et al. (1995) observed no
stimulation in the activity of acid invertase in roots of
Picea abies, ectomycorrhizal with Amanita muscaria or
Cenococcum geophilum. However, stimulation of the
activity of cell wall invertase has been widely reported in
biotrophic infections of plants where it has been strongly
implicated in the diversion of carbohydrates to the fungus
(Long et al. 1975; Greenland & Lewis 1983; Scholes et al.
1994; Wright et al. 1995).
Whilst it is believed that the ultimate source of C for the
fungus is sucrose, current models favour the transfer of
hexoses formed as a result of the action of acid invertases
upon sucrose exported from root cortical cells into the
apoplast (Smith & Smith 1990). In addition, in this type of
VA mycorrhizal association, it has been suggested that
there may be spatial separation of nutrient transfer, with C
uptake occurring via intercellular hyphae and P transfer
occurring across the arbuscular interface (GianinazziPearson et al. 1991). The stimulation of cell wall invertase
activity observed in this study is certainly consistent with
apoplastic hydrolysis of sucrose to hexose prior to uptake
via intercellular hyphae. However, localization of this
activity to cell walls adjacent to intercellular hyphae is
required. The stimulation of the activities of cytoplasmic
invertase and sucrose synthase within root cells may also
provide hexoses for uptake by either the intercellular
hyphae or the arbuscules, although this would first require
the transport of the hexoses to the apoplast. Alternatively,
these enzymes may produce hexoses to provide energy to
drive the uptake of nutrients, such as P, from the arbuscules.
In conclusion, the results of these experiments suggest
that the stimulation in whole shoot CO2 assimilation of M
plants observed by day 42 was due to the additional fungal
sink arising from mycorrhizal colonization of the roots of
the autotroph. The increased sink activity of M roots
appears to be mediated by a stimulation in the activities of
sucrose synthase and cytoplasmic and cell wall invertases,
which occurred upon colonization, resulting in the partitioning of the additional C fixed to the mycorrhizal root
system. To further confirm the notion that sink strength has
a major influence upon the partitioning of assimilates in M
plants it would be desirable to carry out direct measurements of the fluxes of C out of source leaves in nutritionally matched M and NM plants.
ACKNOWLEDGMENT
This research was funded by a NERC grant (GR3/9258).
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Received 6 March 1998; received in revised form 3 June 1998;
accepted for publication 4 June 1998