Editor: Christoph Tebbe
Absence of carbon transfer between Medicago truncatula plants linked by a mycorrhizal network, demonstrated in an experimental microcosm
Article first published online: 28 JUN 2008
© 2008 Federation of European Microbiological Societies. Published by Blackwell Publishing Ltd. All rights reserved
FEMS Microbiology Ecology
Volume 65, Issue 2, pages 350–360, August 2008
How to Cite
Voets, L., Goubau, I., Olsson, P. A., Merckx, R. and Declerck, S. (2008), Absence of carbon transfer between Medicago truncatula plants linked by a mycorrhizal network, demonstrated in an experimental microcosm. FEMS Microbiology Ecology, 65: 350–360. doi: 10.1111/j.1574-6941.2008.00503.x
- Issue published online: 11 JUL 2008
- Article first published online: 28 JUN 2008
- Received 3 December 2007; revised 28 March 2008; accepted 1 April 2008.
- arbuscular mycorrhizal symbiosis;
- autotrophic plants;
- in vitro;
- carbon transfer;
- Top of page
- Materials and methods
Carbon transfer between plants via a common extraradical network of arbuscular mycorrhizal (AM) fungal hyphae has been investigated abundantly, but the results remain equivocal. We studied the transfer of carbon through this fungal network, from a Medicago truncatula donor plant to a receiver (1) M. truncatula plant growing under decreased light conditions and (2) M. truncatula seedling. Autotrophic plants were grown in bicompartmented Petri plates, with their root systems physically separated, but linked by the extraradical network of Glomus intraradices. A control Myc-/Nod- M. truncatula plant was inserted in the same compartment as the receiver plant. Following labeling of the donor plant with 13CO2, 13C was recovered in the donor plant shoots and roots, in the extraradical mycelium and in the receiver plant roots. Fatty acid analysis of the receiver's roots further demonstrated 13C enrichment in the fungal-specific lipids, while almost no label was detected in the plant-specific compounds. We conclude that carbon was transferred from the donor to the receiver plant via the AM fungal network, but that the transferred carbon remained within the intraradical AM fungal structures of the receiver's root and was not transferred to the receiver's plant tissues.
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- Materials and methods
Arbuscular mycorrhizal (AM) fungi are obligate root symbionts, able to interconnect different plants through a common mycorrhizal network (CMN) (Read, 1998; van der Heijden et al., 1998; Giovannetti et al., 2004; Leake et al., 2004). These mycelia favor the movement of soil-derived nutrients and plant-derived carbon within the network and possibly between plants (Smith & Read, 1997; Leake et al., 2004; Simard & Durall, 2004). Such a redistribution of elements can have major ecological implications, impacting intra- and interspecies competition between plants (Read, 1998; van der Heijden et al., 1998; Leake et al., 2004; Simard & Durall, 2004).
The transfer of carbon via a CMN has been clearly demonstrated from autotrophic to achlorophyllous plants, linked by an ectomycorrhizal network (Björkman, 1960; McKendrick et al., 2000; Bidartondo et al., 2003), but the carbon transfer between autotrophic plants linked by a CMN remains more questionable. Several studies reported such a transfer from a donor plant to the shoot of a receiver plant with ectomycorrhiza (Finley & Read, 1986; Simard et al., 1997a, b) or AM fungi (Grime et al., 1987; Lerat et al., 2002; Carey et al., 2004). However, in other experiments such a transfer into the receiver plant's shoots could not be detected, either with ectomycorrhiza (Wu et al., 2001) or with AM fungi (Graves et al., 1997; Fitter et al., 1998; Zabinski et al., 2002), not even when a sink was created by shading or clipping the receiver's shoot (Fitter et al., 1998). This suggested that the translocated carbon remained in the intraradical fungal structures of the receiver plant.
Transfer of carbon from an established plant to a seedling via the AM fungal network has also been investigated. It has been observed that seedlings established more easily within an existing mycorrhizal network, most likely because they have a direct access to a large pool of soil nutrients and water through the CMN or even directly from other plants (Smith & Read, 1997; Jakobsen, 2004; van der Heijden, 2004). Moreover, because the mycorrhizal network is already maintained by the surrounding vegetation, the fungus could receive carbon from the already established plants, supporting the hypothesis that the carbon cost of the seedling to the mycorrhizal network is strongly reduced or even zero (Jakobsen, 2004). A direct transfer of carbon from the surrounding established vegetation to seedlings was studied by Grime et al. (1987). They found, by isotopic labeling, that carbon was transferred into the seedling's tissues through a CMN. However, a very recent study showed a flow of carbon from the established plant to the mycelium and to the roots of the seedling, but not in a greater quantity than to the soil mycelium, indicating that the seedling was not a stronger sink for carbon than the soil (Nakano-Hylander & Olsson, 2007).
The conflicting results on the carbon transfer between established plants, but also from established plants to seedlings, could be related to the plant–fungus combination and the environmental conditions. It is obvious that the experimental system may also influence the results. Indeed, these studies were performed in pot cultures and some even in the field. Under such conditions, the uptake by the roots of the receiver plant of the labeled CO2 respired by the mycorrhizal donor plant and/or released by the hyphae could not be excluded.
Recently, Pfeffer et al. (2004) investigated carbon transfer using the root organ culture (ROC) system (Declerck et al., 2005) with two transformed carrot roots growing in physically separated compartments (St Arnaud et al., 1996), but linked by a CMN of Glomus intraradices. They demonstrated that carbon was transferred from one metabolically active root to another, but found no label in the receiver's root cells, suggesting that the carbon was stored in the intraradical mycelium. However, the absence of photosynthetic tissues, i.e. a true source–sink relationship, and the hormonal disturbance created in ROC lacking shoots (Fortin et al., 2002) could have represented a serious drawback. Therefore, Pfeffer et al. (2004) suggested to study the transfer of carbon from the fungal cells to the root cells of an autotrophic receiver plant, within a system linking, in vitro, autotrophic plants by a CMN. Recently, Voets et al. (2005) developed such an in vitro cultivation system in which a photosynthetically active plant was associated with an AM fungus. In this system, the transport of elements (phosphorus and radiocesium) from the fungus to the plant was clearly observed (Dupré de Boulois et al., 2006). Therefore, such a system could be adapted for interplant transfer of elements such as carbohydrates by combining two plants interconnected by a CMN.
In this study, the system of Voets et al. (2005) was extended to two photosynthetically active plants, physically separated by a plastic barrier, but connected by a CMN. Using this system, we investigated the capacity of an AM fungus to transfer carbon from a donor plant to a receiver plant, between already established plants, but also from an established plant to a seedling, by means of a 13C tracer. We determined whether this carbon was transferred to the plant vegetative cells by means of analysis of the fatty acids in the receiver's roots.
Materials and methods
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- Materials and methods
Medicago truncatula Gaertn., cv. Jemalong line J5 (Myc+/Nod+) was used as the carbon-donor and carbon-receiver plant in the experiments. Its symbiosis-defective mutant TRV 25 (Myc-/Nod-) was used as control. Both lines were provided as seeds by the Institut National de la Recherche Agronomique (INRA, Dijon, France). Glomus intraradices Schenk & Smith (MUCL 43204), grown in association with Ri T-DNA transformed carrot roots, was purchased from GINCO (BCCM/MUCL, Microbiology unit, Université catholique de Louvain, Belgium, http://www.mbla.ucl.ac.be/ginco-bel) and supplied in Petri plates (90-mm diameter) on the Modified Strullu Romand (MSR) medium (Declerck et al., 1998, modified from Strullu & Romand, 1986).
Disinfection, germination and maintenance of the biological material
Seeds of M. truncatula line J5 (Myc+/Nod+) and TRV 25 (Myc-/Nod-) were surface-sterilized by immersion in sodium hypochlorite 5% for 10 min and rinsed three times with sterilized (121 °C for 15 min) deionized water. Seeds were then plated in Petri plates (90-mm diameter, 15 seeds per plate) on the MSR medium, lacking sucrose and vitamins, and solidified with 3 g L−1 Gel Gro™ (ICN, Biochemicals Inc., Irvine, CA). The Petri plates were incubated in the dark at 27 °C. Seeds germinated within 1–2 days and plantlets were ready for use after 4 days.
The G. intraradices cultures were incubated for 4 months in an inverted position in the dark at 27 °C. Several thousand spores were produced in each Petri plate within this period.
General description of the experimental system for the study of carbon transfer between plants
The experimental system (Fig. 1a) consisted of a bicompartmented Petri plate (St Arnaud et al., 1996), each compartment containing 25 mL of the same medium as described in ‘Disinfection, germination and maintenance of the biological material’. The carbon-donor plant, M. truncatula Myc+/Nod+(J5-D), was inserted into the donor compartment (DC) and associated with AM fungal spores. In the receiver compartment (RC), the carbon-receiver M. truncatula Myc+/Nod+(J5-R) was inserted together with the nonmycotrophic control plant M. truncatula Myc-/Nod- (TRV 25). The TRV25 line was used to examine whether uptake of the 13C, released by the AM fungus in the RC, was important. After the establishment of a fungal network connecting the J5-D to the J5-R plants, the carbon-donor plant was labeled with 13CO2 and the transfer of carbon from the donor to the receiver plant was investigated.
Experiment 1: Carbon transfer from a donor to a receiver M. truncatula plant under decreased light conditions
In the DC, a Myc+/Nod+M. truncatula plant (J5-D) was inserted (Voets et al., 2005) and inoculated with c. 100 spores of G. intraradices following the procedure described by Cranenbrouck et al. (2005). In the RC, two M. truncatula plants were planted, a Myc+/Nod+(J5-R) and a Myc-/Nod- (TRV25), following the design in Fig. 1a. The Petri plates were covered with a dark plastic bag, allowing the roots and the AM fungus to grow in the dark, while the plant shoot developed in the light. The Petri plates were placed in a growth chamber set at 22/18 °C (day/night) with 85% relative humidity, a 16-h photoperiod and a photosynthetic photon flux density (PPFD) of 200 μmol m−2 s−1. Three weeks after association, c. 10 mL of sterilized (121 °C for 15 min) culture medium (MSR without sugar and without vitamins and solidified with 3 g L−1 Gel Gro™) was added in both compartments, to provide the plants with nutrients and to maintain the medium at the level of the top of the partition wall, allowing hyphae to cross from the DC to the RC. Addition of MSR medium was repeated weekly. Roots that passed the partition wall were trimmed. Labeling of J5-D with 13CO2 was performed at week 12, when a profuse network of extraradical hyphae was established in the Petri plates, linking the J5-D to the J5-R plants. In order to create a sink for carbon, 3 days before labeling and until the end of the 6-day chase period, a wooden grid with two layers of plastic gauze was placed over the J5-R and TRV25 plants, reducing the light intensity to half of the initial light intensity (i.e. 100 μmol m−2 s−1) without affecting the light intensity of the J5-D plants. Five replicates were used for labeling.
Experiment 2: Carbon transfer from a donor M. truncatula plant to a M. truncatula seedling
The second experiment was performed under the same growth conditions as experiment 1, with the exception that no plants were inserted into the RC at the beginning of the experiment. In this compartment, only 25 mL of culture medium (MSR without sugar and without vitamins) was added. The weekly addition of culture medium starting from week 3 was therefore restricted to the DC. Between week 5 and 6, mycelium started to cross the partition wall separating the DC from the RC. At week 8, an extensive network of hyphae and spores was produced in the RC of all the Petri plates. A 4-day-old Myc+/Nod+M. truncatula receiver plant (J5-R) and a 4-day-old Myc-/Nod- M. truncatula control plant (TRV 25) were subsequently inserted into the RC following the design in Fig. 1a. Labeling was performed 9 days after planting the seedlings in the RC. This period was based on a preliminary experiment (with four replicates) strictly conducted under the same growth conditions and with the same plant/AM fungus combination. Root colonization was estimated after 7 and 15 days following the method described below. The frequency and intensity of root colonization in the J5-R plants during this preliminary experiment was 31.7±4.1% and 3.0±0.9%, respectively, at day 7 and 65.9±10.2% and 33.0±5.3% at day 15. Because root colonization at day 15 seemed appropriate and a chase period of 6 days was fixed, plants in the experiment were labeled 9 days after planting in the culture system. Five replicates were used for labeling in experiment 2.
For labeling, the J5-D was enclosed in a plastic container (1.5 L, L'Oiselle, France) (Fig. 1b) and the hole in the cap through which the plant was inserted into this container was closed with plasticine. The five replicates of both experiments were placed randomly in the growth chamber and were separated by nonmycorrhizal M. truncatula plants, developing in monocompartmental Petri plates, for the same duration as the plants in the RC (three replicates for each experiment). They served as a control for leakage of 13CO2 out of the plastic container by the hole in the cap. One and a half milliliters of a 0.35 M H3PO4 solution was injected with a syringe through a septum into a small tube in the plastic container. Additionally, 1 mL of a 0.27 M NaH13CO3 solution (99 at%, CK Gas, Products Ltd, Hampshire) was injected in the same tube to create an augmentation of the [CO2] in the container from 380 to 1500 μg L−1. This CO2 concentration was measured using GC (Trace 2000; Thermo Finnigan, Interscience). After 3 h, the plastic containers were removed from the J5-D and the systems were kept in the growth chamber for a chase period of 6 days. The whole labeling procedure was the same for both experiments.
In both compartments, plants were removed from the Petri plates and roots, stems, leaves, flowers and fruits were separated. In the RC, the root systems of the TRV25 and J5-R were carefully separated to avoid any risk of cross-contamination. Using a spatula, small blocks of gelled MSR medium were gently detached and removed from the roots and the Petri plates under a binocular microscope until no gelled MSR medium remained in the Petri plates. The root systems were then separated with forceps and washed gently under tap water to eliminate all possible unattached root fragments (e.g. the roots of J5-R sticking to the root system of TRV25 and vice versa). The same process was followed for the root system in the DC. The gelled MSR medium removed from the DC and RC was further dissolved with the citrate buffer (Doner & Bécard, 1991), to separate the AM fungus from the gelled MSR medium.
Plant material from experiment 1 was oven-dried at 70 °C for 72 h and subsequently weighed. A sample of the dried root systems was taken and fungal colonization was estimated. The remaining dried material was ball-milled to a fine powder and 3–5 mg of the powder was placed in tin capsules and analyzed for 13C enrichment using an Europa Scientific ANCA 20-20/GSL continuous-flow isotope-ratio mass spectrometer (IRMS) (SerCon Ltd, Cheshire, UK). Fungal material, which mostly had a dry weight <3 mg, was freeze-dried and 100 μg of this material was encapsulated in tin capsules and analyzed using continuous-flow IRMS using an ANCA-NT 20-20 Stable Isotope Analyzer interfaced to a solid/liquid preparation module (PDZ Europa Scientific Instruments, Crewe, UK).
In experiment 2, most stems and leaves of the seedlings had a dry weight <3 mg and therefore, all the plant samples of this experiment (leaves, stem and roots of the three plants, except the roots of J5-R and TRV25) and the AM fungus were analyzed with the ANCA-NT 20-20 Stable Isotope Analyzer interfaced to a solid/liquid preparation module.
The δ13C results were expressed relative to the Pee Dee Belemnite standard in parts per thousand (‰). Roots of J5-R and TRV25 were used for analysis of fatty acids and, because of their limited quantity (dry weight of J5-R and TRV25 roots is 7.3±1.7 and 6.28±0.87 mg, respectively), could not entirely be used for 13C analysis.
Plant and AM fungal growth parameters at labeling
In both experiments, the stem height, number of leaves, flowers and fruits of the J5-D, J5-R and TRV 25 plants were evaluated. The root length of the plants was also estimated following the gridline method (see Voets et al., 2005). The number of AM fungal hyphae crossing the partition wall and the presence of the cytoplasmic flow within these hyphae was evaluated under a bright-field microscope at × 125 magnification. In addition, in experiment 2, the hyphal length and number of spores developed by the AM fungus in both compartments was also evaluated following the methods detailed in Voets et al. (2005).
Quantification of the intraradical root colonization
In experiment 1, the intraradical structures in the roots were quantified after staining. Dried root samples were first cleared in 10% KOH and then stained with a blue ink solution [1% HCl with 1% blue ink (Parker)] (Vierheilig et al., 1998). Thirty randomly selected root pieces (10 mm length) were mounted on microscope slides and examined under a bright-field microscope at × 50 or × 125 magnification. The frequency (%F) and intensity (%I) of AM fungal colonization were estimated (Declerck et al., 1996). Also, the percentage of arbuscules (%A) and vesicles (%V) was calculated following the same methodology.
Fatty acid analysis
In experiment 2, neutral lipid fatty acids (NLFAs) were identified in the roots of the plants in the RC, following the method described in Nakano-Hylander & Olsson (2007). The 13C abundance in the NLFAs was determined using the IRMS interfaced to a GC (6890, Hewlett-Packard, Palo Alto, CA), which was equipped with a 50-m column (HP-5, Agilent, Palo Alto, CA) using Helium as the carrier gas.
Measurement of the photosynthetic capacity
In experiment 1, the photosynthetic capacity of the plants in the RC was measured at the time of labeling by means of an infrared gas analyzer (IRGA) LCA-4 (ADC Bioscientific Ltd). This was performed on the last fully developed leaf.
Data analysis was performed with the statistical package statistica® for Windows (StatSoft, 2001). Parameters for internal root colonization were arcsin√(x/100) transformed before analysis. Data that were normally distributed and had homogeneous variances were subjected to an anova. The Tukey honest significant difference (HSD) test was used to identify the significant differences (P<0.05, P<0.01, P<0.001).
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- Materials and methods
Plant and AM fungal growth parameters
In experiment 1, the J5-D, J5-R and TRV25 plants followed a similar growth pattern. The first flowers were observed at week 6 and the first fruits at week 10. No significant differences were measured at the end of the experiment among the stem height, plant dry weight, number of leaves, flowers and fruits, root length and root dry weight between the different plants (results not shown). At the time of labeling, the number of active hyphae crossing the partition wall was 116±32, with a mean diameter of 11.7±0.7 μm per hypha. The decreased light conditions at the end of the experiment, induced in order to create a strong sink for carbon, decreased the photosynthetic activity of the plants in the RC (i.e. J5-R and TRV25) from 14.3±1.7 to 5.4±1.1 μmol m−2 s−1.
The colonization of G. intraradices in the root systems of the three plants in experiment 1 was estimated at harvest (Table 1). Colonization in the donor and the receiver plant was characterized by intercellular hyphae, numerous arbuscules and intercellular vesicles. In the donor roots, the colonization was uniformly spread while in receiver's roots, colonization was mostly restricted to small heavily colonized fragments, separated by large sections of root without mycorrhizal structures. This was expressed by an intensity of root colonization higher than the frequency (Table 1). The mean frequency and intensity of the internal root colonization were significantly lower in the J5-R as compared with J5-D. Identically, the % arbuscules and vesicles were significantly lower in the roots of the J5-R plants as compared with the J5-D plants. In the root systems of the TRV25 control plant, no mycorrhizal colonization was observed (Fig. 1). At some points, a ramification was detected at the level of the root epidermis, but hyphae failed to form an appressorium and to enter the root.
|%F||96 ± 2||40 ± 14||0|
|%I||78 ± 2||48 ± 9||0|
|%A||26 ± 5||3 ± 2||0|
|%V||15 ± 1||3 ± 1||0|
In experiment 2, the growth parameters of the seedlings (stem height, plant dry weight, number of leaves, root length and root dry weight) at the end of the experiment did not differ between J5-R and TRV25 (results not shown). The AM fungus developed profusely in both compartments. The mean number of spores was 6970±2334 and 2791±787 and the mean hyphal length was 2721±1006 and 1599±208 cm in the DC and RC, respectively. The number of active hyphae crossing the partition wall at the time of labeling was 120±34 with a mean diameter of 10.5±0.8 μm per hypha.
Experiment 1: Carbon transfer from a donor to a receiver M. truncatula plant under decreased light conditions
Six days after labeling of the J5-D plants with 13CO2, a large quantity of 13C was incorporated into the J5-D plant tissues (Table 2). The label was transferred to the roots of the plant and to the extraradical mycelium (ERM), which extended from the DC into the RC. The 13C enrichment of the ERM (δ13C) was 140.7±69.5‰ and 166.9±127.1‰ in the DC and RC, respectively, and did not differ significantly from each other (P=0.85). Because the ERM in the DC and RC formed a continuum, the data presented in Table 2 were summed. The roots of J5-R showed only a slight increase in 13C, not significantly higher (P=0.15) than the 13C enrichment in the roots of the nonmycotrophic control plant (Table 2). In the stem and leaves of both plants developing in the RC (i.e. J5-R and TRV25), no detectable increase of δ13C was observed (Table 2) (P=0.13 and P=0.86, respectively). Also, no 13C enrichment was observed in the leaves of the plants to control the leakage out of the labeling containers (not in the figure), assuring that the labeling containers were hermetically closed.
|J5-D plant||J5-D roots||ERM*||J5-R roots||J5-R stem||J5-R leaves||TRV25 roots||TRV25 stem||TRV25 leaves|
|Sink: established plants||506.0 ± 78.2||159.8 ± 36.8||169.0 ± 63.9||−29.5 ± 0.9||−33.4 ± 0.3||−34.6 ± 1.5||−28.8 ± 0.9||−30.0 ± 0.4||−34.0 ± 0.5|
|Background δ13C value†||−34.6 ± 0.3||−31.3 ± 0.4||−35.0 ± 0.2||−31.3 ± 0.4||−33.2 ± 0.3||−35.0 ± 0.4||−29.6 ± 0.2||−30.6 ± 0.6||−34.1 ± 0.8|
Experiment 2: Carbon transfer from a donor M. truncatula plant to an M. truncatula seedling.
After 6 days of chase, a marked increase in 13C was found in the tissues of the J5-D plant (Table 3). The 13C was further translocated to the ERM. The 13C enrichment of the ERM in both compartments (DC: δ13C=71.1±35.9‰ and RC: δ13C=63.5±30.1‰) did not differ significantly (P=0.88) and was summed over both compartments (Table 3). In this experiment, the roots of the plants in the RC were not entirely subjected to 13C analysis but NLFAs were extracted and subsequently analyzed for their 13C content. Analysis of the NLFA 16:1ω5, an indicator of AM fungal storage lipids, showed clear 13C enrichment in the J5-R roots (Table 3), most likely induced by the colonization of G. intraradices in the roots. The presence of the NLFA 16:1ω5 was also observed in the roots of TRV25, the nonmycotrophic control plant (Table 3), probably due to small fungal fractions sticking to the root surface. These values did not differ significantly (P=0.82). A slight augmentation of 13C was observed in the 18:2ω6,9 fatty acid extracted from both roots in the RC and these values did not differ significantly between the J5-R and the TRV25 roots (P=0.87) (Table 3). The NLFA 18:2ω6,9 is mostly attributed to the host root cells, but NLFA 18:2ω6.9 extracted from a fungal control sample showed a small fraction of this 18:2ω6.9 (16:1ω5: 695.42 nmol mg−1; 18:2ω6,9: 0.30 nmol mg−1). A very small part of the 18:2ω6,9 fatty acid in the roots is thus assignable to the fungus. A slight 13C enrichment was detected in the stem and the leaves of the J5-R plant, but data were highly variable and not significantly different (P=0.77 and P=0.42, respectively) (Table 3). The 13C enrichment in the stem of the TRV25 plants was less expressed and was even absent in the leaves (Table 3). The δ13C value in the leaves of the plants to control the leakage out of the labeling containers (not in the figure) was close to the background δ13C value.
|J5-D plant||J5-D roots||ERM†||J5-R roots||J5-R stem||J5-R leaves||TRV25 roots||TRV25 stem||TRV25 leaves|
|NLFA 16:1ω5||NLFA 18:2ω6,9||NLFA 16:1ω5||NLFA 18:2ω6,9|
|Sink: seedlings||848.5 ± 178.0||42.7 ± 37.0||63.1 ± 18.6||37.0 ± 27.1||−19.5 ± 8.0||−31.6 ± 0.9||−34.4 ± 1.2||25.6 ± 36.6||−21.1 ± 6.9||−29.7 ± 2.2||−34.9 ± 1.2|
|Background δ13C value||−34.6 ± 0.3||−31.3 ± 0.4||−35.0 ± 0.2||−42.7||−29.8 ± 0.3*||−33.2 ± 0.3||−35.0 ± 0.4||−42.7||−29.6 ± 0.2*||−30.6 ± 0.6||−34.1 ± 0.8|
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- Materials and methods
Suitability of the culture system
In this study, an in vitro culture system in which two autotrophic plants were linked by a common extraradical mycelium of an AM fungus was developed. Numerous active hyphae were crossing the partition wall, interconnecting the carbon-donor and the carbon-receiver plant. This number was important and close to (Dupré de Boulois et al., 2006) or even much higher (Nielsen et al., 2002; Rufyikiri et al., 2002) than previous observations, suggesting the capacity of the AM fungus to transfer elements from the DC to the RC. It was already shown earlier that such active mycelium was responsible for the transfer of nutrients (N, P) (Jin et al., 2005; Dupré de Boulois et al., 2006) and radionuclides (U, Cs) (Rufyikiri et al., 2002; Dupré de Boulois et al., 2006) from a hyphal compartment to a root compartment, or carbohydrates (Pfeffer et al., 2004) from a donor compartment to a receiver compartment in vitro. Thanks to the partition wall separating the two compartments (St Arnaud et al., 1996), the diffusive pathway of carbon from one compartment to the other could be excluded. In addition, the presence of the Myc-/Nod- control plant in the RC allowed a precise control of the 13C, released into the culture medium of the RC by exudation of the hyphae and by decaying hyphae that release their 13C into the culture medium. Therefore, this system allowed a highly controlled investigation of the carbon flow in mycorrhizal networks.
In vitro split plates have been used previously to track the carbon flow between carbon donors and carbon receivers with transformed carrot roots (Pfeffer et al., 2004). It was shown that the mechanisms of carbon uptake and metabolism by the AM fungus are similar in mycorrhizal roots of autotrophic plants and mycorrhizal transformed roots (Pfeffer et al., 2004). However, culture systems with autotrophic plants as a host, in which a continuous carbon supply from the plant shoots via the roots to the fungus is assured, might be better adapted to certain types of studies, certainly when photosynthesis is involved.
The mycorrhizal colonization in the different root systems was identical to previous data (Dupré de Boulois et al., 2006), although the %V and %A were lower in the receiver plants compared with the donor plants (Table 1). This is most likely due to the shorter contact time between the mycelium and the roots in the RC as compared with the DC. In our symbiotic defective plant (TRV25), the fungus did not form appressoria on the root surface, and no hyphae were recorded within the roots, confirming the nonmycotrophy of the mutant (Morandi et al., 2005).
The adequate time of labeling in the second experiment was chosen in order to conciliate the age of the seedlings and the level of intraradical root colonization. Because the estimation of root colonization is a destructive treatment, the intraradical root colonization at the time of labeling could not be determined. Therefore, a preliminary experiment was performed under the same conditions and with the same plant/AM fungus combination, in order to determine the adequate time of labeling. In this preliminary experiment, 31.7±4.2% of the roots of all the seedlings presented intraradical root colonization, 7 days after planting, with the presence of many arbuscules. This confirmed the observations of Read et al. (1976) that seedlings already become mycorrhized in their cotyledon stage. After 15 days, this level increased upto 65.9±10.2%, with numerous arbuscules and vesicles observed. It was therefore considered that the seedlings presented adequate root colonization at the moment of labeling (9 days after planting in the culture system). The higher root colonization of the J5-R plant in experiment 2 after a shorter contact time between AM fungi and plant, compared with experiment 1, could be explained by the considerably larger root system of the receiver plants in the first experiment, as compared with the young plants in experiment 2. This probably resulted in a dilution effect of the root colonization value estimated on 30 randomly chosen 10-mm-long root pieces.
Carbon is transferred from the donor plant into the roots of the receiver plant but remains in the fungal tissues
Thirty-two percent of the assimilated 13CO2 was translocated from the labeled shoots to the roots of the donor plant in experiment 1, while this was 5% in experiment 2. The 13C was subsequently translocated to the associated AM fungus. An equal enrichment in 13C was measured in the ERM on both sides of the partition wall, compared with the J5-D roots. This indicates that the flow of 13C from the donor plant to the AM fungus was probably not, or only slightly, counterbalanced by a flow of 12C from the receiver plant, suggesting the creation of an adequate sink in both experiments.
The roots of the receiver plants presented a small enrichment in 13C, most probably stored in the AM fungal structures. Even though 40% of the receiver's root system was colonized by the AM fungus, only a small number of vesicles were observed. It is accepted that the vesicles are the major storage organs for lipids (Bago, 2000), and their small number could therefore explain the small 13C enrichment of the roots. This hypothesis is further supported by the observations of Fitter et al. (1998), showing that the quantity of carbon transferred from a donor plant (Cynodon dactylon) to a receiver plant (Plantago lanceolata) was positively correlated to the number of vesicles in the roots of the receiver plant. The roots of the nonmycotrophic control plant (TRV 25) also presented traces of 13C enrichment. This suggested that hyphal secretion and turnover in the RC could have released 13C, taken up by the receiver and control plants. We could not rule out the possibility of small hyphal fragments sticking to the root surface of the controls. This further indicated that fungal carbon released into the substrate should be systematically considered when studying carbon transfer between plants. Therefore, the combination of a mycotrophic and a nonmycotrophic mutant of the same plant species in the RC may help to differentiate indirect carbon uptake (by the medium/substrate) from the direct carbon uptake, transferred by the AM fungus. Grime et al. (1987) considered a similar combination of a mycotrophic and a nonmycotrophic plant and found that the label in the nonmycorrhizal control plant was negligible compared with the 14C values in the surrounding colonized plants. Simard et al. (1997a), however, planted an AM control plant together with plants linked by an ectomycorrhizal fungus. They found that 18% of the label, transferred from the donor to the receiver, was recovered in the AM control, which indicates the contribution of ectomycorrhizal fungal carbon release in their experiment.
The control plants placed between the culture systems during the labeling procedure had δ13C values close to background in both experiments, indicating that no refixation of 13CO2 leaked out of the labeling container seemed to have occurred. The small increase of δ13C in the leaves of the receiver plant is thus most likely due to the natural variation among plant replicates in their δ13C value, as suggested by Fitter et al. (1998).
Analysis of the fatty acids 16:1ω5 and 18:2ω6,9
The 13C analysis of the entire root system did not resolve the question of whether carbon transferred by the fungus was released into the receiver's tissues. Therefore, a more profound analysis of the root system was required. Extraction and analysis of the 13C enrichment in the NLFA 16:1ω5, which dominates the neutral lipid fraction in AM fungi (Olsson & Johansen, 2000) and the NLFA 18:2ω6,9, a fatty acid present in plant storage lipids but only present in very low levels in AM fungi (Olsson et al., 2005), was performed in the roots of J5-R and TRV 25 of the second experiment.
The 13C enrichment of 16:1ω5 in the roots of J5-R indicated 13C allocation from the ERM to the intraradical mycelium (IRM). However, this 13C enrichment was slightly lower than the total 13C enrichment in the ERM, indicating that carbon allocation through mycelial networks was not directed towards the seedling. These results were in agreement with the observations of Nakano-Hylander & Olsson (2007), who observed that a seedling was not a higher sink for carbon than was the soil. The presence of the NLFA 16:1ω5 in the roots of TRV25 was not expected and could most probably be attributed to some traces of fungal material sticking to the root surface. Indeed, because the AM fungus had an abundant growth in the RC, with the production of c. 3000 spores and almost 16 m of mycelium, the possibility of small fragments of hyphae sticking to the TRV25 roots after washing could not be excluded. This could have caused a detectable amount of the NLFA 16:1ω5 in the roots of TRV25. No clear background δ13C value of the NLFA 18:2ω6,9 of the roots in the RC could be measured. Therefore, the 13C enrichment in these roots was estimated in relation to the δ13C of the entire roots. Even if the different carbon compounds in a root differ slightly in δ13C background value (O'Leary, 1981; Ballentine et al., 1998; Hobbie & Werner, 2004), these differences are mostly limited [e.g. 2.5‰ difference between cellulose and lignin in roots (Hobbie & Werner, 2004)]. Also, because the J5-R and TRV25 are clones (Morandi et al., 2005), the difference in background δ13C value between the NLFA 18:2ω6,9 and the total δ13C of the respective roots should be in the same line. The slight augmentation of δ13C in the NLFA 18:2ω6,9 was probably due to a combination of several factors, because G. intraradices is known to also contain a small fraction of the NLFA 18:2ω6,9 (Pfeffer et al., 2004; Olsson et al., 2005), a slight increase of the 13C enrichment in 18:2ω6,9 in both roots was expected. However, the direct uptake of the 13C from the culture medium in the RC, released by the AM fungal hyphae, could also have caused the slight 13C enrichment. In addition, arbuscules are known to have a mean life span of 1–2 weeks (Alexander et al., 1989). The turnover of fungal biomass should therefore also be mentioned as a possible pathway for carbon transfer. Such carbon transfer has been proposed in Orchidaceae, where there is massive intracellular colonization, followed by hyphal collapse, which releases the carbon into the plant root cells (Bidartondo, 2005). A possible direct transfer from the AM fungus to the plant cells is also part of the hypotheses (Grime et al., 1987; Carey et al., 2004).
Pfeffer et al. (2004) found the presence of two separate pools in the NLFA 18:2ω6,9: one pool accounting for 75% of the fatty acid that was totally unlabeled and another pool accounting for 25% that was enriched with 13C and could be attributed to the fungus. These authors thus did not detect any direct uptake of 13C from the culture medium. However, AM fungal exudates released into the culture medium by Glomus sp. have been found to consist of low-molecular-weight sugars, organic acids as well as some high-molecular-weight compounds (Toljander et al., 2007). It has been observed that some plant roots are able to take up sugar compounds from the soil, as seen in maize (Kuzyakov & Jones, 2006), and from the culture medium, as seen in mixotrophic cultures in general (Kozai et al., 1997). Therefore, we cannot exclude the possibility of a direct uptake of carbon by the roots of the plants in our culture system that have a high carbon-demand and that are grown in a synthetic medium where all the compounds are easily accessible. It could, therefore, be hypothesized that part of the 13C enrichment in the NLFA 18:2ω6,9 was attributed to the direct uptake of 13C from the culture medium.
From the four above-mentioned pathways of 13C transfer that might have caused the 13C enrichment in the NFLA 18:2ω6.9: (1) the small fraction of 18:2ω6,9 in the AM fungus, (2) the direct uptake of 13C from the medium by the plant roots, (3) the arbuscular turnover in the plant cells and (4) the possible direct transfer from fungus to plant, the last two factors impossibly could have occurred in the TRV25 mutant. Because the 13C enrichment in the J5-R roots is identical to the enrichment in the TRV25 roots, we can actually prove that the direct carbon transfer from the AM fungus to the plant cells and the arbuscular collapse in the plant cells might not have played an important role.
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The results presented in this paper highlighted the complexity of carbon labeling studies and the different pathways involved in the transfer of carbon from one autotrophic plant to another, via a common AM fungal network. While a sink was created by shading the receiver plants or planting seedlings in the culture plates, no evidence for the existence of a carbon flow from the donor to the receiver plant by means of the AM fungal mycelium was found. Our results therefore confirm previous results, indicating that carbon transferred from autotrophic donors to autotrophic receivers remains in the receiver's roots and that a transfer to the shoots is unlikely to occur (Robinson & Fitter, 1999; Zabinski et al., 2002; Pfeffer et al., 2004; Fitter, 2006).
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This research has been supported by a grant of the ‘Fonds Spéciaux de Recherche (FSR)’ of the Université catholique de Louvain and by the Belgian Science Policy–Program Science for a Sustainable development under contract number SD/BD/05A. The authors wish to thank G. Duc from INRA (Dijon, France) for providing the seeds of M. truncatula. Thanks are also due to K. Coorevits, C. Detez and M. Ghanem for technical assistance.
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