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Keywords:

  • Cucumis sativus L. (cucumber) ;
  • Glomus mosseae;
  • 14C-partitioning;
  • leaf age;
  • phosphorus

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

An assessment of the effects of arbuscular mycorrhizal (AM) infection on photosynthesis, carbon (C) allocation, translocation and biomass production of cucumber, grown in sand culture, was made using a previously determined phosphorus (P) supply (0·13 mol m−3 P) which had a significant impact on AM infection. Separation of a direct effect of AM infection from an indirect one due to an enhanced leaf P status was achieved using a comparable non-mycorrhizal treatment (NAM + P) supplemented with extra P (0·19 mol m−3 P). Total leaf P concentration, specific leaf mass, photosynthetic capacity, and incorporation of 14C into non-structural carbohydrate pools were dependent on leaf age. Both maximum and ambient photosynthetic rates were significantly higher in the youngest fully expanded leaves from AM and NAM + P plants which also had the higher leaf P concentrations. There were no differences in the total concentrations of starch, sucrose, raffinose or stachyose in young or old leaves among AM, non-mycorrhizal (NAM) and NAM + P treatments. However, younger leaves of NAM plants showed a shift in 14C-partitioning from stachyose and raffinose synthesis to starch accumulation. Determination of ADP-glucose pyrophosphorylase (AGPase), sucrose synthase and sucrose phosphate synthase enzyme activities revealed that only AGPase activity was correlated with the increased incorporation rate of 14C into starch in young leaves of NAM plants. Although there were significant AM-specific effects on C translocation to the root system, AM plants had similar rate of photosynthesis to NAM + P plants. These results suggest that the increase in photosynthetic rate in leaves of AM-infected cucumber was due to an increased P status, rather than a consequence of a mycorrhizal ‘sink’ for assimilates.


INTRODUCTION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

Arbuscular mycorrhizal (AM) colonization results in an enhanced ‘sink’ demand for carbon (C) in roots ( Snellgrove et al. 1982 ; Douds, Johnson & Koch 1988; Wright, Scholes & Read 1998a, b). This ‘sink’ effect could account for an extra 10–23% drain of C from the host by the AM fungus ( Snellgrove et al. 1982 ; Kock & Johnson 1984; Kucey & Paul 1982; Jakobsen & Rosendahl 1990). It has been hypothesized that this enhanced AM ‘sink’ demand increases the rate at which C is assimilated by the host plant ( Fitter 1991; Wright et al. 1998a , b). Alternatively, other workers have indicated that an increased photosynthetic rate may be due to a mycorrhizal-dependent increase in the P status of AM plants grown at low P ( Allen et al. 1981 ; Fredeen & Terry 1988; Azcon, Gomez & Tobar 1992). Where the P status and biomass of AM and non-mycorrhizal (NAM) plants were similar, differences in C assimilation in leaves were attributed to a P-independent mycorrhizal ‘sink enhancement’ of photosynthesis ( Brown & Bethlenfalvay 1988; Wright et al. 1998a , b). However, the presence of two symbiotic associations in these studies may confound the assessment of a specific mycorrhizal-induced ‘sink’ effect ( Smith & Read 1997). It is also possible, in view of these studies, that there may be an effect due to both an enhanced P status and an AM-dependent ‘sink’ demand.

The assessment of AM- and P-dependent effects on photosynthesis and C allocation in leaves could be confounded by specific interactions with leaf age. Leaf P concentration, C fixation and partitioning of C in individual leaves have, for instance, been reported to vary with leaf age in non-mycorrhizal plants ( Dietz & Heilos 1990; Turgeon 1973). Therefore, it may be necessary to consider differences in leaf physiology within a single plant, as well as between similar leaves from different treatments, to assess the impact of AM colonization.

Variations in P concentration in leaf tissue have been shown to influence the partitioning and flux of C into starch or soluble carbohydrates in non-mycorrhizal plants via their effect on the export of triose phosphate out of the chloroplast ( Herold 1980; Foyer & Spencer 1986; Sharkey 1990; Stitt 1996; Rao 1997). Previous reports on the effect of AM colonization on non-structural carbohydrate concentrations in relation to leaf P status are equivocal ( Brown & Bethlenfalvay 1988; Graham, Duncan & Eissenstat 1997; Wright et al. 1998b ), and appear to be inconsistent with current models dealing with P status and partitioning of C in leaves ( Stitt 1996; Rao 1997). Due to compartmentation and the dynamic nature of C allocation in leaves, it may be necessary to measure the 14C flux into carbohydrate pools, and not the total concentration of non-structural carbohydrates. Previous work has also been largely confined to an assessment of the effects of AM infection on below-ground C metabolism ( Snellgrove et al. 1982 ; Douds, Johnson & Koch 1988; Jakobsen & Rosendahl 1990). To obtain a mechanistic understanding of the way in which AM infection specifically influences shoot metabolism, more 14C labelling studies, coupled with enzymological investigations, are required.

In this study, we have used cucumber, in which AM infection has been associated with a 17–23% increase in C translocation to the root ( Jakobsen & Rosendahl 1990). Initially, the effect of a range of P concentrations on biomass production and photosynthesis of AM and NAM cucumber plants were assessed to identify the P supply at which the greatest effect of AM infection on plant performance was observed. In the second experiment, we used AM and NAM plants grown at this selected P supply, together with a comparable P supplemented non-mycorrhizal treatment (NAM + P), to separate a direct AM effect from an indirect one due to an improved P status. Gas exchange measurements, in combination with 14C-labelling techniques, were used to determine the influence of P status and AM colonization on photosynthesis and incorporation of 14C into non-structural carbohydrate pools in leaves of different ages. Concomitantly, the activity of ADP-glucose pyrophosphorylase, sucrose synthase and sucrose phosphate synthase, which are known to be involved in the regulation of carbohydrate synthesis in leaves ( Stitt 1996), were investigated. In a third experiment, 14C translocation studies were carried out on AM, NAM and NAM + P plants to determine whether AM infection was associated with an increased ‘sink’ demand for assimilates.

MATERIALS AND METHODS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

Growth of plants and mycorrhizal inoculation

All plants were grown in a Vindon growth cabinet (Vindon Scientific Ltd, Diggle, UK) at 20 °C, 70% relative humidity, with a 16 h photoperiod at an irradiance of 350 μmol m−2 s−1 (λ = 400 − 700 nm), giving an integrated value of 20·2 mol m−2 d−1. Cucumber (Cucumis sativus L. Telegraph Improved) seeds were germinated on moist tissue paper in Petri dishes for 2 weeks. Plant roots were inoculated by mixing 3 g of a granular inoculum containing chlamydospores and hyphal fragments of Glomus mosseae (Nicolson & Gerdemann), Gerdemann & Trappe, strain YV (MicroBio, Hemel Hempstead, Hertfordshire, UK), with 1300 cm3 (2·1 kg) of lime-free, washed silica sand (Pioneer Supamix Ltd, Nuneaton, UK) and grown in 12·5 × 15 cm (diameter × height) sections of PVC piping sealed at the bottom, apart from several small drainage holes (3 mm diameter). The same amount of autoclaved (121 °C under steam pressure for 1 h) granular inoculum was added to the non-mycorrhizal treatments. Plants were grown for 50 d and supplied with 350 cm3 Long Ashton nutrient ( Hewitt 1966) solution per week. Pots received additional water as required (100 cm3 per week) in order to keep the surface of the sand moist. Auxiliary shoots and flower buds were removed from plants after bud initiation.

For the first experiment, plants were supplied with 350 cm3 per week of Long Ashton nutrient solution containing a range of P concentrations (0·13–1·17 mol m−3). This was equivalent to 0·67–6·1 mg P kg−1 sand per week, or a total of 10–91 mg P per pot over the 50 d duration of the experiment. The results of this experiment enabled us to determine the P level at which there was the largest mycorrhizal effect on plant growth and photosynthesis. In the second and third experiments, mycorrhizal (AM) and non-mycorrhizal (NAM) plants were supplied with a Long Ashton nutrient solution containing this predetermined P level (0·13 mol m−3 P). An additional non-mycorrhizal treatment (NAM + P) was supplied with 0·19 mol m−3 P, or a total supply of 14·8 mg P per pot over the duration of the experiment, to obtain non-mycorrhizal plants with the same biomass and leaf P concentration as AM plants grown at 0·13 mol m−3 P. The extra amount of P supplied to NAM + P plants was extrapolated from the relationship between total biomass or leaf P concentration versus P supply, using data obtained from the first experiment (see RESULTS, Fig. 1).

image

Figure 1. Mycorrhizal infection (a), total biomass (b), total leaf area (c), root biomass (d), leaf area ratio (e), shoot:root ratio (f), leaf P concentration (g) and maximum photosynthetic rate (h) of mycorrhizal (black bars) and non-mycorrhizal (white bars) cucumber plants 50 d after inoculation. Plants were supplied with a Long Ashton nutrient solution containing various concentrations of P. Histograms (mean ± SD, n = 3) with different letters are significant at P≤ 0·05.

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Measurement of photosynthetic performance

Photosynthetic measurements were made on the youngest, fully expanded leaf (leaf 3, counting from the top of the plant), leaf 5 and leaf 7, 1 week prior to harvest using an infra-red gas analyser with a Parkinson leaf chamber (CIRAS-1, PP Systems, Hitchin, Hertfordshire, UK). Gas exchange measurements were performed between 10:00 and 15:00 h (6–11 h into the light phase) over 4 d in the Vindon plant growth cabinet. No diurnal variations were observed during this sampling period. Leaves were enclosed within the Parkinson leaf chamber and allowed to equilibrate for 5–10 min (after alteration of light level and/or external CO2 concentrations) before light or CO2 response curve measurements were taken. Light response curves (0–1500 μmol photons m−2 s−1) were determined using an external CO2 concentration of 350 μmol mol−1 CO2, a temperature of 20 °C (± 1·5) with a vapour pressure deficit (VPD) of approximately 0·7 kPa, followed by measurement of the response to variations in CO2 concentration at saturating light. Estimates of in vivo Rubisco activity and electron transport capacity were calculated from data on the response of photosynthesis to intracellular CO2, using the method of Watanabe, Evans & Chow (1994). For the second and third experiments, photosynthesis measurements were also performed at the ambient temperature and irradiance (350 μmol photons m−2 s−1) used in the growth cabinet, with an external CO2 concentration of 350 μmol mol−1 and a VPD of approximately 0·7 kPa.

Radioactive 14C-labelling of plants

In experiments 2 and 3, shoots of one plant from each treatment (AM, NAM, NAM + P) were placed in a 75 × 40 × 30 cm Perspex chamber similar to the one described by Snellgrove et al. (1982) . Perspex discs were clamped around the base of the stem to separate the root compartment from the aerial part of the plant. The interfaces between root and shoot compartments, as well as the lid of the chamber, were sealed with fabric-backed duct tape. The chamber was placed in a Vindon growth cabinet maintained under the same environmental conditions as described earlier. Plants were humidified and aerated overnight by circulating air through a flask containing 125 cm3 deionized water, on the inlet side of the chamber. Plant replicates (AM, NAM and NAM + P; n = 4) were labelled at a similar time during the photoperiod (after 8 h in the light phase) as those used for the gas exchange measurements. The water in the flask was replaced with 1 cm3 2·2 MBq NaH14CO3 (558 MBq mmol−1; Sigma Chemical Co, Poole, UK) and 14CO2 was liberated by the injection of 10 cm3 70% (m/v) lactic acid into the flask. The ambient CO2 concentration in the chamber was only increased by approximately 3 μmol mol−1 after the addition of labelled 14CO2. Whole plants were fed for 5 (experiment 2) and 15 min (experiment 3) and the remaining 14CO2 in the chamber was trapped by the addition of 100 cm3 10% (m/v) KOH solution to the labelling flask. Plants underwent a ‘chase period’ in humidified ambient air for 5 min in experiment 2 and 105 min in experiment 3. Leaves 3 (youngest, fully expanded leaf), 5 and 7 (oldest leaf) were removed, split into three similar sub-samples (0·5–2 g FW), immediately frozen in liquid N and either stored at –20 °C for further carbohydrate or enzyme analyses, or dried at 80 °C for 24 h for biomass and nutrient analysis. Roots were sub-sampled, washed in deionized water and either stored in 50% ethanol, for estimation of AM infection, or dried at 80 °C for 24 h for biomass and nutrient determinations.

Fractionation of 14C-assimilates

Leaf sub-samples were ground in liquid N and partitioned in 20 cm3 methanol:chloroform:water (12:5:3) according to the protocol of Dickson (1979) and centrifuged at 5000 g for 5 min. The methanol:chloroform:water fraction was retained and the insoluble pellet was washed three times in water to remove any residual soluble carbohydrates. The methanol: chloroform:water-soluble extracts (20 cm3) were partitioned into a methanol-soluble fraction and a lipid fraction by the addition of 5 cm3 chloroform and 4 cm3 water, followed by centrifugation at 5000 g for 5 min. The insoluble fraction was then resuspended in 10 cm3 250 m M acetate buffer (pH 4·5), boiled for 1 h and incubated with 10 units of amyloglucosidase at 55 °C for 4 h to hydrolyse the starch component. The remaining insoluble starch-free fraction was removed by centrifugation at 5000 g for 5 min. Complete hydrolysis of starch was confirmed by the absence of a starch–iodine positive reaction after the addition of a 10% (m/v) iodine solution to the hydrolysed insoluble pellet. Hydrolysed starch in the supernatant was determined as glucose subunits, using the glucose oxidase method (Boehringer Mannheim, Germany). The radioactivity in the starch, insoluble compounds, lipid and methanol-soluble fractions was determined using 1 cm3 of each fraction, in 10 cm3 scintillation cocktail (Ecoscint-A, National Diagnostics, Atlanta, Georgia, USA), using scintillation counting (LKB Rackbeta model 1211, Wallac OY, Turku, Finland). The amount of 14C (kBq) in each fraction was corrected using a 14C quench curve, in which percentage counting efficiency was determined from a known amount of [U-14C]glucose (10 kBq) in a range of chlorophyll solutions in ethanol (1–20 mg cm−3). The total amount of 14C added to the labelling chamber and the amount of label recovered from plants and CO2 traps was used to calculate the percentage 14C recovery. The percentage recovery of 14C (total kBq) from plant fractions and the CO2 trap after labelling was 92–101%.

Identification and quantification of 14C-soluble carbohydrates

Methanol-soluble fractions were reduced to dryness and resuspended in 2 cm3 deionized water. Soluble sugars in the extracts (40 μL) were then separated by paper chromatography using butanol:pyridine:acetic acid:water (45: 30:2·5:22·5), according to Giegenberger & Stitt (1991). Fructose components in the carbohydrates were visualized by spraying with a urea–phosphoric acid stain ( Wise et al. 1955 ). The detection limit of this reagent was approximately 0·4 μg fructose. Carbohydrates were identified by comparison with fructose, sucrose, raffinose and stachyose standards. Another set of chromatograms were exposed to X-ray film (BioMax MR, Eastman Kodak Company, St Louis, Minnesota, USA) for the detection of radioactivity in each sugar pool. Radioactive sugar spots were cut out from the paper chromatograms and 14C incorporation was measured using scintillation counting, as described above.

Collection of phloem exudates and determination of 14C translocation rates

For experiment 3, plants were labelled for 15 min with 14C as described earlier, followed by a 105 min ‘chase period’. Phloem sap was collected from the internodes below leaves 3 and 7 and at the base of the stem after the 14C ‘chase’, 8 h into the photoperiod. As cucumber phloem sap gels oxidatively within 2–4 min on contact with air ( Richardson, Baker & Ho 1982), samples were collected from cut stem surfaces by exuding droplets into graduated microcapillary tubes, while the sap was still fluid. Known volumes (10–40 μL) were then transferred to 100 μL 80% ethanol which precipitated p-protein and prevented gelling of the sap. The sap was frozen in liquid N and stored at −20 °C prior to scintillation counting. Carbon translocation rates (kBq cm−2 h−1) were calculated from the amount of label in phloem sap aliquots using a modified equation for specific mass transfer rate based on that of Richardson, Baker & Ho (1984).

For the determination of phloem area, transverse sections of stem tissue were taken from regions adjacent to those where phloem sap was collected, stained with toludene blue (2% m/v in 50% glycerol, pH 4·5) and viewed under a dissecting microscope at × 20–50 magnification. Internal and external phloem areas were calculated from tracings obtained from photographs of stained sections, using stereological techniques ( Steer 1981).

Calculation of 14C-specific incorporation and incorporation rate

Data obtained from the amount of 14C incorporated into different carbohydrate pools, together with the measured concentrations of carbohydrates in leaves, were used to calculate specific incorporation (kBq in specific carbohydrate pool mg−1 carbohydrate g−1 leaf FW). The rate of incorporation of 14C was calculated as the amount of 14C incorporated into the specific carbohydrate compound per unit time (kBq g−1 leaf FW h−1).

Determination of methanol-soluble carbohydrates

Methanol-soluble carbohydrates were detected and quantified by means of gas–liquid chromatography, using a flame ionization detector (Model 8410, Perkin Elmer, Cambridge, UK). A single glass-lined stainless steel column (2 m long × 3 mm diameter) packed with chromosorb Diatomite ‘C’ (80–100 BSS mesh) as the solid support, which was coated with a non-polar (2% methyl phenyl silicone gum material; SE 52), was used to separate the trimethylsilyl (TMS) derivatives of carbohydrates ( Holligan & Drew 1971). The TMS ester derivatives of carbohydrates were obtained by re-dissolving the dried sample in 425 μL anhydrous pyridine, 50 μL hexamethyldisilazine and 25 μL trimethylchlorosiline, and incubated at room temperature for 3 h. For the separation of TMS derivatives of monosaccharides and disaccharides, a carrier gas (N) with a flow rate of 40 cm3 min−1, was used with a temperature programme of 140 °C for 5 min, 140–220 °C at a ramp rate of 6 °C min−1, 220–290 °C at a ramp rate of 8 °C min−1 and 290 °C for 5 min. For the separation of TMS derivatives of raffinose and stachyose, the programme was modified to a carrier gas (N) flow rate of 80 cm3 min−1 and a temperature profile of 200 °C for 10 min, 200–300 °C at a ramp rate of 20 °C min−1, and 300 °C for 5 min. The concentrations and identification of carbohydrates were determined by comparison with TMS derivatives of standard carbohydrates, which were run after the analysis of three samples.

Determination of enzyme activities

Leaves (0·5–2 g FW) were ground in liquid N and enzymes extracted in 2 vol 100 mmol m−3 HEPES–NaOH at pH 7·5, containing 4 mmol m−3 MgCl2, 1 mmol m−3 EDTA, 1 mmol m−3 EGTA, 5 mmol m−3 mercaptoethanol and BSA (1 μg cm−3). Extracts were dialysed against the extraction buffer at 4 °C for 4 h, frozen in liquid N and stored at –80 °C. No change in the maximal activity of ADP-glucose pyrophosphorylase (AGPase) was found before and after dialysis. The activity of sucrose synthase (SS) or sucrose phosphate synthase (SPS) could not be checked before dialysis because of the presence of fructosyl monomers and dimers in the extracts, which interfered with the anthrone reagent. Based on the data for AGPase, we assumed that no significant changes in enzyme activities occurred. The protein content was determined by the method of Bradford (1976), using BSA as the standard. For the estimation of SS activity, the reaction rate was measured in the sucrose synthesis direction using a substrate-dependent fixed-time assay ( Huber et al. 1996 ). Reaction mixtures contained 50 mmol m−3 HEPES–NaOH (pH 7·5), 10 mmol m−3 UDP-glucose, 10 mmol m−3 fructose, 1 mmol m−3 MgCl2, 1 mmol m−3 EDTA and 20 μL of enzyme extract in a final volume of 70 μL. For the measurement of SPS, the fructose-6-phosphate-dependent fixed-time reaction was used ( Dancer, Hatzfeld & Stitt 1990). Reaction mixtures contained 50 mmol m−3 HEPES–NaOH (pH 7·5), 2 mmol m−3 UDP-glucose, 2 mmol m−3 fructose-6-phosphate, 10 mmol m−3 glucose-6-phosphate, 1 mmol m−3 EDTA, 1 mmol m−3 MgCl2 and 15 μL of extract in a final volume 70 μL. After 15 min incubation at 25 °C, both reactions were terminated by the addition of 70 μL 30% KOH (m/v). Reaction mixtures from SPS and SS assays were then boiled for 20 min and sucrose determined using the anthrone reagent ( Huber et al. 1996 ). The activity of AGPase was measured in the cleavage direction of ADP-glucose, using 100 μL of the extract, and glucose-6-phosphate was measured in an NAD-dependent dehydrogenase coupled assay, as described by Sowokinos (1976). Assays were initiated by the addition of 0·6 mmol m−3 pyrophosphate (PPi).

Leaf area, nutrient and pigment analyses

Leaf area was determined on individual leaves using a leaf area meter (AM100, Analytical Development Company, Hoddesdon, UK). Total nitrogen was determined on oven-dried leaves using the method of Hendershot (1985). Total P in the same leaf digests was assayed using the ammonium molybdate reaction ( Murphy & Riley 1962). Chlorophyll was extracted by grinding 200 mm2 fresh leaf discs in 80% acetone over ice. Absorbances on the centrifuged extracts were measured at wavelengths of 647 and 663 nm using a Novaspec spectrophotometer (Model 4409, Pharmacia LKB, Biochem Ltd, Cambridge, UK) and chlorophyll concentration calculated according to the equations described by Arnon (1949).

Estimation of mycorrhizal infection

Sub-samples of roots, previously stored in 50% ethanol, were cleared in 2·5% KOH overnight, acidified with 1% HCl for 4 h, and stained using aniline blue (0·05%) in 70% acidified glycerol. The percentage of root length with AM infection was estimated by evaluating 100 random intersections for each root system, using a hairline graticule in the eyepiece of a compound microscope at × 400 magnification ( Brundrett, Melville & Peterson 1994).

Statistical methods

There were three replicate plants for each treatment in experiment 1 and four for each treatment in experiments 2 and 3. Percentage values for mycorrhizal infection and 14C distribution were arcsine-transformed prior to statistical analysis. Data were analysed by multiple analysis of variance (ANOVA), using the SAS statistical package (SAS Institute Inc., Cary, North Carolina, USA). Where ANOVA revealed significant differences between treatments, their means were compared using Fisher’s least significant difference (LSD) test. Regression analysis for the total biomass and leaf P concentration data versus P supply, in experiment 1, was performed using the curve fit regression function y = a ln x + b (SigmaPlot version 4·0, SPSS Inc., San Rafael, California, USA). Correlations between incorporation rate of 14C into non-structural carbohydrate components and P concentration in each leaf were carried out using linear regression analysis.

RESULTS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

Experiment 1

None of the plants exposed to autoclaved inoculum showed AM infection. There were no significant differences in AM infection between inoculated plants supplied with 0·13 and 0·38 mol m−3 P, although AM plants supplied with 0·38 mol m−3 P showed a greater percentage root length infection when compared to AM plants supplied with 0·78 and 1·17 mol m−3 P ( Fig. 1a). A microscopic examination of the root systems of AM plants revealed the presence of arbuscules and vesicles at 0·13 and 0·38 mol m−3 P but not at 0·78 and 1·17 mol m−3 P. A significant effect of mycorrhizal colonization on total plant biomass, root mass, total leaf area, leaf P concentration and maximum photosynthetic rate (Pm) was only found for the 0·13 mol m−3 P supply ( Fig. 1b–d,g,h). Overall, Pm declined as the P concentration increased ( Fig. 1h). These results were paralleled by changes in the ambient photosynthetic rate (Pa), again with only a significantly higher value in AM plants at 0·13 mol m−3 P (data not shown). There was no effect of P supply or mycorrhizal infection on total biomass at 0·78 and 1·17 mol m−3 P ( Fig. 1b). The leaf area ratio (LAR) was not influenced by P supply nor AM colonization ( Fig. 1e). Although the shoot:root ratio was higher in plants supplied with 0·13 mol m−3 P, the allocation of dry mass between the shoot and the root was not influenced by mycorrhizal infection ( Fig. 1f).

On the basis of these results, an external P supply of 0·13 mol m−3 P was used in the second and third experiments, as it was the only level at which an AM-dependent stimulation of photosynthesis and biomass was observed. The amount of extra P (an increase of approximately 46%) required to obtain NAM plants with a similar biomass or leaf P concentration to AM plants at 0·13 mol m−3 P was determined using a natural log curve fit (P≤ 0·01) of the relationships between biomass or leaf P concentration and P supply. Using these relationships, we estimated that a P supply of 0·17–0·22 mol m−3 P (equivalent to a total supply of 13·2 and 17·1 mg P per pot) was required to produce NAM plants with a similar biomass and leaf P concentration as AM plants supplied with 0·13 mol m−3 P. Thus, in the second and third experiments, an average value of 0·19 mol m−3 P was supplied to uninoculated plants (NAM + P) to examine P-independent effects of mycorrhizal infection.

Experiment 2

Plant growth

Plants from the AM and NAM + P treatments had more leaves (a total of 10–11 leaves) than NAM plants (8–9 leaves). Older leaves from NAM plants senesced 1–2 weeks earlier than the same leaves from the AM and NAM + P treatments. The total dry biomass and leaf areas of AM and NAM + P plants were similar, but 30% higher than those of the NAM plants supplied with 0·13 mol m−3 P ( Fig. 2). The shoot:root ratios from the AM and NAM + P treatments were 20% higher than that of the NAM plants. For the NAM + P plants, the shoot:root ratios were 7% lower than for the AM plants (due to a lower root mass in AM plants), but this was not significant. The specific leaf mass (SLM) of younger leaves was significantly lower than that of old leaves in all treatments, although SLM did not vary between treatments with leaves of the same age. The percentage root length colonized by G. mosseae in the inoculated AM treatment (34 ± 5%) was similar to the level (36 ± 4%) of AM infection observed in experiment 1 at 0·13 mol m−3 P ( Fig. 1a). No AM infection was observed in the control treatments (NAM and NAM + P) where autoclaved inoculum was used.

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Figure 2. Biomass (a) of roots (black areas), stems (white areas) and leaves (hatched areas) and total leaf area (b) of mycorrhizal (AM), non-mycorrhizal (NAM) and non-mycorrhizal P-supplemented (NAM + P) cucumber plants 50 d after inoculation. Histograms (mean ± SD, n = 4) for each treatment with different letters are significant at P≤ 0·05.

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Photosynthesis and related parameters

Values for Pm, Pa, total 14C incorporation, electron transport capacity, Rubisco activity and total leaf P concentration all declined with leaf age ( Fig. 3). For leaf 3, AM and NAM + P plants showed an increase in all of these photosynthetic parameters when compared to NAM plants, although these increases were not always significant for the NAM + P treatment. Estimates of Pa and 14C incorporation rates for all the leaves from AM and NAM + P plants were similar, but higher than those measured in NAM leaves. However, these differences were not significant for the Pa measurements taken on leaf 7, in contrast to the 14C data ( Fig. 3e,f). Differences in photosynthetic performance between AM, NAM + P and NAM plants correlated with P concentration, but not with N concentration or dark respiration rate. No significant differences in stomatal conductance, transpiration rate or chlorophyll concentration were found (data not shown).

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Figure 3. Maximum net photosynthetic rate (a), Rubisco activity (b), electron transport capacity (c), dark respiration rate (d), photosynthetic rate under ambient light (e), total 14C incorporation rate (f), total P concentration (g) and total N concentration (h) in different leaves from mycorrhizal (black bars), non-mycorrhizal (white bars) and non-mycorrhizal P-supplemented (hatched bars) cucumber plants 50 d after inoculation. Histograms (mean ± SD, n = 4) with different letters are significant at P≤ 0·05.

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Non-structural carbohydrates

The main non-structural carbohydrates in leaves of cucumber were sucrose, stachyose, raffinose and starch ( Fig. 4). Leaves contained trace amounts of myo-inositol and hexitol, with a similar retention time to dulcitol. Glucose and fructose concentrations varied between 30 and 70 μg g−1 FW, and were similar in all leaves examined. Although the concentrations of starch were similar for all treatments, a significantly higher (1·3–1·4 times) concentration was found in leaf 5 from NAM plants, compared to the AM and NAM + P treatments ( Fig. 4). Neither the soluble carbohydrate nor starch concentrations in leaves ( Fig. 4) showed a decline with leaf age, in contrast to the observed changes in photosynthesis and P concentration ( Fig. 3).

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Figure 4. Concentration of stachyose (a), raffinose (b), sucrose (c), and starch (d) in different leaves from mycorrhizal (black bars), non-mycorrhizal (white bars) and non-mycorrhizal P-supplemented (hatched bars) cucumber plants 50 d after inoculation. Histograms (mean ± SD, n = 4) for each non-structural carbohydrate concentration marked with different letters are significantly different at P≤ 0·05.

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Distribution of 14C between carbon pools

There were differences in 14C distribution between different treatments for leaves 3, 5 and 7 ( Table 1). The highest incorporation of 14C into the methanol-soluble pool was found in the AM and NAM + P treatments ( Table 1). Conversely, incorporation of label into the starch fraction was always higher in leaves from NAM plants. In leaves 3 and 5, the partitioning of 14C into starch was 2–4 times higher in NAM plants than for AM and NAM + P treatments. Partitioning of 14C into starch in NAM leaves declined with leaf age ( Table 1). Incorporation of 14C into the lipid fraction was similar in all leaves ( Table 1). Although a higher percentage of 14C was incorporated into the insoluble/structural fraction in leaf 3, there were no differences between treatments ( Table 1).

Table 1.  Percentage distribution of 14C in different fractions from leaves of mycorrhizal (AM), non-mycorrhizal (NAM) and non-mycorrhizal P-supplemented (NAM + P) cucumber plants 50 d after inoculation
 Leaf number
FractionTreatment357
  1. Leaves were labelled with 14CO2 for 5 min followed by a chase period of 5 min after 8 h in the light. Results are shown as the mean % (n = 4) and numbers in parentheses are the mean arcsine transformations ± SD. Values within a particular fraction with different letters are significant at P≤ 0.05.

Methanol/water-solubleAM68 (55 ± 2)a73 (59 ± 4)a78 (62 ± 4)a
 NAM52 (47 ± 2)b57 (49 ± 3)b65 (54 ± 2)a,b
 NAM + P70 (57 ± 4)a73 (59 ± 3)a76 (61 ± 3)a
LipidAM8 (16 ± 2)a5 (13 ± 1)a4 (12 ± 2)a
 NAM6 (14 ± 3)a3 (11 ± 3)a4 (12 ± 3)a
 NAM + P9 (17 ± 3)a5 (13 ± 2)a4 (12 ± 3)a
Insoluble/structuralAM13 (21 ± 2)a8 (16 ± 1)b5 (13 ± 2)b
 NAM9 (17 ± 3)a,b6 (14 ± 2)b6 (14 ± 1)b
 NAM + P11 (19 ± 3)a,b8 (16 ± 2)b6 (14 ± 2)b
StarchAM9 (17 ± 3)b13 (21 ± 1)b13 (21 ± 2)b
 NAM36 (37 ± 12)a34 (36 ± 8)a18 (25 ± 6)a,b
 NAM + P9 (18 ± 4)b13 (21 ± 4)b12 (19 ± 5)b
Incorporation rate and specific incorporation of 14C

There was a general trend towards a decrease in the rate and specific incorporation of 14C into non-structural carbohydrates with leaf age, but this was only significant for the incorporation of label into starch in leaves from NAM plants ( Table 2). The incorporation rate and specific incorporation of 14C into stachyose, raffinose and sucrose in leaves 3 and 5 from AM and NAM + P plants was 1·4–2 times higher than for the NAM treatment. In leaf 3, the incorporation rate of label into raffinose in AM leaves was significantly higher than the same leaves from NAM or NAM + P plants ( Table 2). However, the specific incorporation of label into raffinose in the same leaf from NAM + P plants was similar to that of AM plants, because of the slightly lower concentration of raffinose in NAM + P leaves ( Fig. 4). No differences in incorporation rate and specific incorporation of 14C into glucose and fructose occurred in any of the leaves in any treatment. In contrast to the results for soluble carbohydrates, the rate and specific incorporation of 14C into starch was 2–3 times lower in younger leaves from AM and NAM + P plants, compared to the same leaves from NAM plants ( Table 2). For leaves 3 and 5, there was a significant positive correlation between 14C incorporation into stachyose and leaf P concentration ( Fig. 5a,c). There was also a significant relationship (P≤ 0·05) between the incorporation of label into raffinose and P concentration in leaves 3 and 5. Conversely, a significant negative correlation was found between rate of 14C incorporation into starch and leaf P concentration for leaf 3 ( Fig. 5b). The data also indicated that the AM, NAM + P and NAM plants generally fell into two groups. The NAM plants had leaves with a lower P, which was correlated with a higher flux of label into starch and a lower incorporation of label into stachyose. The opposite results were obtained with AM or NAM + P plants ( Fig. 5).

Table 2.  Incorporation rate (kBq g−1 FW h−1) and specific incorporation (kBq mg−1 carbohydrate g−1 F.W) of 14C into non-structural carbohydrate pools in leaves from mycorrhizal (AM), non-mycorrhizal (NAM) and non-mycorrhizal P-supplemented (NAM + P) cucumber plants 50 d after inoculation
  Leaf number
CarbohydrateTreatment357
  1. For individual carbohydrates, values (mean ± SD, n = 4) for either incorporation rate or specific incorporation which have different letters are significant at P≤ 0·05.

StachyoseAM12·3 ± 2·5a11·7 ± 1·9a8·1 ± 2·7a,b
Incorporation rateNAM7·3 ± 1·5b5·8 ± 2·0b5·4 ± 1·5b
 NAM + P11·1 ± 1·8a10·0 ± 1·2a8·7 ± 2·8a,b
Specific incorporationAM4·5 ± 0·5a5·3 ± 1·1a3·2 ± 1·3a,b
 NAM3·2 ± 0·6b3·6 ± 0·9a,b3·4 ± 2·0a,b
 NAM + P5·3 ± 0·9a4·7 ± 0·7a5·3 ± 2·5a,b
RaffinoseAM10·1 ± 1·2a9·8 ± 2·3a,b6·7 ± 2·4a,b
Incorporation rateNAM5·6 ± 1·7b5·3 ± 1·8b3·7 ± 1·5b
 NAM + P5·8 ± 1·9b9·0 ± 3·4a,b6·7 ± 2·8a,b
Specific incorporationAM3·6 ± 0·4a3·2 ± 0·7a,b2·3 ± 1·3a,b
 NAM1·7 ± 0·5b1·8 ± 0·4b1·2 ± 0·4b
 NAM + P2·4 ± 0·7a,b2·8 ± 0·9a2·2 ± 0·5a,b
SucroseAM4·5 ± 1·2a3·3 ± 0·3a2·8 ± 1·4a,b
Incorporation rateNAM2·4 ± 0·7b2·1 ± 0·6b2·3 ± 0·7b
 NAM + P4·8 ± 1·4a3·3 ± 0·8a2·7 ± 0·5b
Specific incorporationAM4·5 ± 0·6a3·3 ± 0·7a,b2·8 ± 1·1a,b
 NAM2·1 ± 0·9b2·2 ± 0·6b2·8 ± 0·2a,b
 NAM + P4·2 ± 1·2a,b3·1 ± 0·3a,b2·3 ± 0·5a,b
StarchAM6·6 ± 2·0c7·2 ± 2·3c4·4 ± 2·5c
Incorporation rateNAM20·5 ± 9·9a12·8 ± 3·1b3·8 ± 1·4c
 NAM + P6·5 ± 2·6c6·9 ± 2·6c5·2 ± 3·3c
Specific incorporationAM0·5 ± 0·2a0·6 ± 0·2b0·4 ± 0·3b,c
 NAM1·5 ± 0·6a 0·7 ± 0·3a,b0·3 ± 0·1c
 NAM + P0·5 ± 0·3b0·5 ± 0·2b0·3 ± 0·2b,c
image

Figure 5. The relationship between the incorporation rate of 14C into stachyose or starch and P concentration in specific leaves from mycorrhizal (●), non-mycorrhizal (○) and non-mycorrhizal P-supplemented (▾) cucumber plants 50 d after inoculation. Data were taken from leaves 3 (a,b), 5 (c,d) and 7 (e,f). Linear regressions in (a) were significant at P≤ 0·01 and in (b) and (c) were significant at P≤ 0·05. The relationships in (d), (e) and (f) were not significant.

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Maximal specific enzyme activities

There were no differences in SPS or SS activities in any of the leaves examined ( Table 3). However, an increased flux of 14C into starch in leaf 3 of NAM plants ( Table 2) was correlated with a significant 1·2–1·6-fold increase in the specific activity of AGPase in leaf 3, but not in leaf 7 ( Table 3).

Table 3.  Specific activity (nkatal mg−1 protein) of starch and sucrose synthesizing enzymes in leaves 3 and 7 from mycorrhizal (AM), non-mycorrhizal (NAM) or non-mycorrhizal P-supplemented (NAM + P) cucumber plants 50 d after inoculation
  Leaf number
EnzymeTreatment37
  1. Values (mean ± SD, n = 4) for each specific enzyme activity with different letters are significant at P≤ 0·05.

Sucrose phosphate synthaseAM1·2 ± 0·2a1·0 ± 0·1a
 NAM1·0 ± 0·1a0·8 ± 0·2a
 NAM + P1·2 ± 0·2a1·2 ± 0·2a
Sucrose synthaseAM0·6 ± 0·3a0·5 ± 0·2a
 NAM0·8 ± 0·2a0·8 ± 0·2a
 NAM + P0·6 ± 0·1a0·8 ± 0·2a
ADP-glucose pyrophosphorylaseAM0·5 ± 0·1b0·8 ± 0·2a
 NAM0·9 ± 0·1a0·9 ± 0·1a
 NAM + P0·6 ± 0·05b0·7 ± 0·2a,b

Experiment 3

Plant growth and AM infection

The percentage AM infection (40 ± 6%) in inoculated plants was similar (P≤ 0·01) to the values (34 ± 5%) observed in experiment 2. The results for total and individual biomass components, leaf N and P concentrations and photosynthetic rates for AM, NAM and NAM + P were also similar to those obtained previously (data not shown).

14C in phloem sap

The 14C transfer rates decreased from the top to the base of the stem but phloem area increased from approximately 1 mm2 at internode 3 to approximately 10 mm2 at the base of the stem. Thus, the total amount of 14C translocated was similar at all sampling points. Carbon transfer rates calculated from samples collected at internode 3 from AM plants were 26 and 53% higher than those calculated for NAM + P and NAM plants, respectively ( Table 4). Differences in 14C transfer rates between AM and NAM + P plants at internode 7 were not significant. At the base of the stem, the 14C transfer rates in AM plants were 28 and 52% greater than those estimated for NAM + P and NAM plants, respectively.

Table 4.  Transfer rates of 14C (kBq cm−2 phloem area h−1) from leaves of mycorrhizal (AM), non-mycorrhizal (NAM) and non-mycorrhizal P-supplemented (NAM + P) cucumber plants measured at different sampling points. Leaves were labelled with 14CO2 for 15 min followed by a chase period of 105 min
 Sampling point
TreatmentInternode 3Internode 7Base of stem
  1. Values (mean ± SD, n = 4) with different letters are significant at P≤ 0·05.

AM312 ± 56a158 ± 34c66 ± 10d
NAM145 ± 15c89 ± 24d29 ± 8f
NAM + P230 ± 24b141 ± 23c47 ± 7e

Soluble carbohydrates represented 60–70% of the total 14C activity present in phloem sap. Stachyose was the major carbohydrate translocated in cucumber plants, comprising 48–61% of the total sugars transported, followed by raffinose (approximately 32%).

DISCUSSION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES

The impact of mycorrhizal infection on both photosynthesis and the partitioning of C between the major carbohydrate pools in leaves of cucumber can be attributed to an enhanced P concentration. This was not due to the absence of a significant C drain in AM plants as a 28% increase in translocation was detected in the present study, compared to NAM + P plants. This estimate is comparable with previous work on cucumber (17–26%; Jakobsen & Rosendahl 1990), but somewhat higher than a number of other plants previously examined (10–15%, Snellgrove et al. 1982 ; Kock & Johnson 1984; Kucey & Paul 1982; Wright et al. 1998b ). These observations are consistent with the studies of Allen et al. (1981) and Azcon et al. (1992) , but contrast with those of Wright et al. (1998a , b), which concluded that alterations in C metabolism in AM Trifolium repens was due to an AM fungal ‘sink’ demand. Previous assessments that have proposed a direct role for the fungal ‘sink’ in the stimulation of photosynthesis where nodulated AM and NAM leguminous plants have been compared ( Brown & Bethlenfalvay 1988; Wright et al. 1998a , b) could, however, be complicated by synergistic interactions between N fixation and mycorrhizal-dependent increases in P. A greater ‘sink’ demand for assimilates in AM nodulated plants, compared to NAM nodulated plants, could be a result, in part, of a higher rate of N fixation, through an increase in the availability of P ( Smith & Read 1997). These conclusions also contrast with other studies, in which mycorrhizal-associated increases in photosynthetic rate were attributed to alterations in stomatal conductance ( Green, Stodola & Augé 1998), increased leaf N or specific leaf mass ( Fredeen & Terry 1988). This does not rule out the possibility that in plants such as cucumber, which have a high demand for P ( Robinson & Decker-Walters 1997), a nutrient response could over-ride any specific sink effect due to AM infection. Under natural situations, with less P-demanding species, there could be a more direct effect of AM infection. Estimates of the carbon drain to AM roots in natural systems may be as high as 30–55% ( Clapperton & Read 1992). However, in natural systems, these estimates may be complicated by C transfer between plants via a common mycorrhizal network ( Robinson & Fitter 1999).

The separate influences of a fungal C ‘sink’ demand, or an AM-enhanced P status, on C metabolism are also difficult to assess due to the complex nature of the AM symbiosis. Some of these complications have been overcome by using split-root systems ( Douds et al. 1988 ), or through the establishment of a non-mycorrhizal P response growth curve that permits comparisons to be made with AM plants ( Eissenstat et al. 1993 ). A variation of the latter approach ( Snellgrove et al. 1982 ; Kock & Johnson 1984; Syvertsen & Graham 1990; Wright et al. 1998a ), and the one used in this study, was to compare AM plants with P-supplemented NAM plants that are similar in size and P status as AM-colonized plants. The amount of extra P (0·06 mol m−3, equivalent to a total of 0·15 mol m−3 P over 50 d) added to the nutrient solution supplied to NAM cucumber plants to obtain the same biomass, leaf area and leaf P concentration as AM plants was similar to the amount of extra P (0·12–0·2 mol m−3 P) supplied to NAM Glycine max and Sorgham bicolor plants ( Pacovsky, Bethlenfalvay & Paul 1986). These authors suggest that it might be necessary to establish separate ‘controls’ for each plant parameter of interest because AM and NAM + P plants may differ in some P-dependent physiological and morphological characteristics. In this study, we only used one P supply for the establishment of a P-supplemented non-mycorrhizal control because the biomass, leaf area, shoot: root ratio and leaf P concentration of NAM plants supplemented with extra P (NAM + P) were similar (3–7% variation for all growth parameters) to AM plants. This allowed us to assess the influence of AM-enhanced ‘sink’ demand on C metabolism, independently of P status.

When the growth of these two P-equivalent treatments (AM and NAM + P) were compared, a potential ‘dry weight loss’ in AM plants due to a C drain by the fungus, as suggested by Stribley, Tinker & Rayner (1980), was not observed. The reduced root biomass in AM plants (7%), could reflect this ‘dry weight loss’ to the fungus, although these differences were not significant. Whilst the absence of a significant dry weight loss could indicate that the fungus is a relatively small ‘sink’ for assimilates, significantly higher 14C shoot–root translocation was observed in AM cucumber plants, and this suggests that the supply of translocated C is sufficient to support both root and fungal growth and respiration. Despite lower C translocation rates and an expected lower root respiration rate ( Jakobsen & Rosendahl 1990) in NAM + P plants, biomass production was similar to AM plants. As there were also no differences in biomass allocation, this indicates that the greater respiratory demand in infected plant roots may be compensated for by reductions in other metabolic costs, such as those associated with P uptake and utilization ( Douds et al. 1988 ; Smith & Read 1997), under these conditions.

Whilst it could also be argued that the results of the first experiment at high P supply showed no direct AM effect, due to the absence of any significant increase in biomass or photosynthesis, this is complicated by the well-described inhibitory effect of P supply on mycorrhizal colonization ( Fig. 1). The total level of AM infection was reduced at high P supply and arbuscules were not observed, indicating that the effectiveness of the symbiosis may also be reduced. Therefore, experiments on AM symbiosis, even where relatively high levels of infection are maintained at high P supply ( Peng et al. 1993 ), may not have assessed the full metabolic impact of AM infection. Despite the lower root infection (40%), the magnitude of an AM ‘sink’ effect in our study was similar to that reported in other studies on cucumber, where the percentage infection was reported as approximately 90% in 24-d-old plants ( Jakobsen & Rosendahl 1990). These differences may reflect differences in the metabolic activity of the AM fungus, with a higher activity in the present study compensating for a lower root length infection.

Generalizations about the absence of an indirect mycorrhizal effect to include all situations may be complicated by the effects of irradiance ( Pearson, Smith & Smith 1991). Variations in irradiance had no effect on total infection of Allium porrum, but the frequency of arbuscules and vesicles increased with light levels up to 515 μmol m−2 s−1 (~26 mol m−2 d−1). These results could indicate that the manifestation of a ‘sink’ effect on photosynthesis will be greater at irradiances somewhat higher than those used in the present study (350 μmol m−2 s−1; ~20 mol m−2 d−1). Of the previous studies that have attributed the stimulation of photosynthesis in AM plants to a sink effect, most have used irradiances (350–503 μmol m−2 s−1; ~21–25 mol m−2 d−1) comparable to those used in the present study (see Azcon et al. 1992 ; Wright et al. 1998a , b). Further studies are therefore required to determine whether there is evidence of an increased ‘sink’ activity at high irradiances.

Data obtained from 14C-labelling experiments in this study also indicates that changes in C-partitioning and rate of 14C-incorporation into non-structural carbohydrate pools in young leaves were correlated with leaf P concentration. The addition of extra P to NAM + P plants, as well as a mycorrhizal-enhanced P status in AM plants, altered the partitioning of fixed C from starch accumulation in NAM plants to raffinose and stachyose synthesis in AM and NAM + P plants. Clearly, these alterations in the partitioning of C are not simply predictable on the basis of measurements of total non-structural carbohydrate concentrations.

The relationship between C assimilation and leaf P status suggests a relief of feedback inhibition by enhanced P status on C assimilation in AM and NAM + P plants. Many non-mycorrhizal plant studies have highlighted the regulatory role of P on photosynthesis via effects on electron transport capacity and Rubisco activity ( Fredeen et al. 1990 ; Sharkey 1990; Rao 1997). Such a proposal is also consistent with the altered partitioning of C in AM and NAM + P plants. The accumulation of starch in NAM leaves could reflect alterations in C allocation between the cytosol and the chloroplast due to a limited influx of P into the chloroplast to counterbalance the efflux of fixed C via the P translocator ( Herold 1980; Foyer & Spencer 1986; Stitt 1996; Rao 1997). The increased incorporation rate of 14C into starch in NAM plants is also consistent with an increase in the observed maximal specific activity of AGPase. Similar results have been shown in non-mycorrhizal plants grown under P-limiting conditions ( Priess 1982; Nielson et al. 1998 ). The proposed consequence of a mycorrhizal-enhanced P status in leaves could be an increased incorporation rate of fixed C into soluble carbohydrates in the cytosol and a subsequent increase in phloem loading for long-distance transport to the fungal and plant C ‘sink’.

Correlations between C metabolism and leaf P concentration in individual leaves, reported in the present study, highlight the potential modifying influence of leaf age and the importance of assessing these parameters in individual leaves and not on a bulk leaf sample where these effects might be masked. For example, in leaf 3 of cucumber, we observed mycorrhizal P-related changes in several leaf parameters based on a leaf-by-leaf analysis, whilst similar correlations were not evident in older leaves. Alterations in photosynthesis and C partitioning in older leaves that are independent of P concentration ( Fig. 5) could be due to differences in the partitioning of metabolically active P between the vacuolar, chloroplastic and cytosolic compartments ( Foyer & Spencer 1986), even where the bulk leaf P concentrations are the same. Alternatively, P-independent changes in C partitioning may be due to differences in the development of leaves from AM, NAM + P and NAM treatments. Other studies have shown, for instance, that photosynthetic capacity, C partitioning and leaf P concentration are dependent on leaf expansion and leaf age in non-mycorrhizal systems ( Turgeon 1973; Dietz & Heilos 1990).

In conclusion, our results indicate that P status, together with leaf age, influences C assimilation and partitioning in mycorrhizal plants. A causal relationship between an increased photosynthetic rate and an AM-enhanced ‘sink’ demand for assimilates ( Fitter 1991; Wright et al. 1998b ) was not evident in AM cucumber plants in this study despite a significant increase in the translocation rate of C from the shoot to roots. The effect of an enhanced AM ‘sink’ demand and P status on C metabolism may, however, vary depending on the species of mycorrhizal fungus used as well as host-specific factors including variations in leaf development, the size(s) of various ‘sinks’ for assimilates, and the overall responsiveness of the host to mycorrhizal colonization.

REFERENCES

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. REFERENCES
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