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Carbon Metabolism in Spores of the Arbuscular Mycorrhizal Fungus Glomus intraradices as Revealed by Nuclear Magnetic Resonance Spectroscopy1 *Corresponding author; e-mail ppfeffer/at/arserrc.gov; fax 215–233–6581. Received March 24, 1999; Accepted June 9, 1999.  This article has been cited by other articles in PMC. | |||||||
Abstract Arbuscular mycorrhizal (AM) fungi are obligate symbionts that colonize the roots of over 80% of plants in all terrestrial environments. Understanding why AM fungi do not complete their life cycle under free-living conditions has significant implications for the management of one of the world's most important symbioses. We used 13C-labeled substrates and nuclear magnetic resonance spectroscopy to study carbon fluxes during spore germination and the metabolic pathways by which these fluxes occur in the AM fungus Glomus intraradices. Our results indicate that during asymbiotic growth: (a) sugars are made from stored lipids; (b) trehalose (but not lipid) is synthesized as well as degraded; (c) glucose and fructose, but not mannitol, can be taken up and utilized; (d) dark fixation of CO2 is substantial; and (e) arginine and other amino acids are synthesized. The labeling patterns are consistent with significant carbon fluxes through gluconeogenesis, the glyoxylate cycle, the tricarboxylic acid cycle, glycolysis, non-photosynthetic one-carbon metabolism, the pentose phosphate pathway, and most or all of the urea cycle. We also report the presence of an unidentified betaine-like compound. Carbon metabolism during asymbiotic growth has features in between those presented by intraradical and extraradical hyphae in the symbiotic state. | |||||||
Fossil evidence (Taylor et al., 1995) and molecular phylogenetic studies (Simon et al., 1993) indicate that 400 million years ago the roots of land plants were colonized by ancestors of modern arbuscular mycorrhizal (AM) fungi. Today, more than 80% of land plants still acquire nutrients from soil using arbuscular mycorrhizas, reflecting the evolutionary success of this mutualistic symbiosis (Jakobsen, 1995; Smith and Read, 1997). During arbuscular mycorrhiza formation, plant and fungus integrate structurally and functionally to become a single supraorganism (Azcón-Aguilar and Bago, 1994) whose physiological capacities are superior to those of either organism alone (Jakobsen, 1995). This integration improves nutrient uptake by the plant and allows the heterotrophic, obligately symbiotic fungus to complete its life cycle. Understanding AM physiology has practical and fundamental significance, but requires first a knowledge of the metabolism of each partner. Carbon metabolism of AM fungi has been reviewed by Jakobsen (1995). Lipids are the major form of carbon in AM fungal spores, hyphae, and vesicles (Cox et al., 1975), comprising 45% to 95% of the spore C pool, depending on the species (Beilby, 1983; Jabaji-Hare, 1988; Bécard et al., 1991). Triacylglycerides account for most of the lipids in spores (Beilby and Kidby, 1980; Jabaji-Hare, 1988; Gaspar et al., 1994). During spore germination (AM fungal asymbiotic growth), the triacylglyceride content remains constant for approximately 5 d (Bécard et al., 1991; Gaspar et al., 1994), after which time it decreases (Beilby and Kidby, 1980; Gaspar et al., 1994), probably through the activity of a membrane-bound lipase (Gaspar et al., 1997). At the same time, an increase in phospholipids is observed during germ tube growth (Beilby and Kidby, 1980). This latter result, together with the incorporation of label from 14C-acetate into triacylglycerides during spore imbibition, led Beilby (1983) to suggest that lipid synthesis and breakdown occurred simultaneously during AM fungal asymbiotic growth. The dominant storage carbohydrates detected in AM fungal hyphae and spores are glycogen and trehalose (Amijee and Stribley, 1987; Bécard et al., 1991; Bonfante et al., 1994; Shachar-Hill et al., 1995; Bago et al., 1998; Pfeffer et al., 1999). In vivo 13C-NMR studies revealed that such carbohydrates were formed by the AM fungus Glomus etunicatum shortly after providing colonized leek roots with 13C-labeled Glc (13C-Glc) (Shachar-Hill et al., 1995). Half of the trehalose stored in G. etunicatum spores is broken down during the 5 d after germ tube emergence, indicating that this carbohydrate sustains fungal growth at the very early stages of germination (and prior to lipid breakdown) (Bécard et al., 1991). CO2 also seems to play an important role in AM fungal axenic growth: Bécard and Piché (1989) found a 10-fold increase in Gigaspora margarita germ tube development when a 0.5% CO2 atmosphere was supplied. Assays of enzymatic activities have indicated the presence of several metabolic pathways during asymbiotic growth of Glomus mosseae (Macdonald and Lewis, 1978) and Gi. margarita (Saito, 1995). Of the glycolytic enzymes, phosphofructokinase (EC 2.7.1.11) and glyceraldehyde-P dehydrogenase (EC 1.2.1.12) have been detected (Macdonald and Lewis, 1978; Saito, 1995). The presence of the tricarboxylic acid cycle (TCA) enzymes malate dehydrogenase (EC 1.1.1.37) and succinate dehydrogenase (EC 1.3.5.1) has also been demonstrated (Macdonald and Lewis, 1978; Saito, 1995). The activity of Glc-6-P dehydrogenase (EC 1.1.1.49) (Macdonald and Lewis, 1978; Saito, 1995) was found to be higher in germinated spores of Gi. margarita than in intraradical hyphae (Saito, 1995), suggesting a role for the pentose phosphate pathway (PPP) in the developing spore. More recently, the presence of the enzyme 3-phosphoglycerate kinase (EC 2.7.2.3) has been revealed by differential RNA display in G. mosseae germinating spores and mycorrhizal roots (Harrier et al., 1998). This enzyme is implicated in both glycolysis and gluconeogenesis. Recently, Pfeffer et al. (1999) used AM monoxenic cultures of Glomus intraradices and Ri T-DNA-transformed carrot (Daucus carota) roots to study C metabolism in the symbiotic state. That study indicated that uptake, transport, and metabolism are very different in the intraradical and extraradical parts of AM fungi in the symbiotic state. Whereas intraradical fungal structures take up carbon (as hexose) and synthesize lipids, the extraradical mycelium does neither. Since earlier results indicated differences in the metabolism of the AM fungus when growing in symbiosis or asymbiotically (Harrison and van Buuren, 1995; Shachar-Hill et al., 1995), the aim of the present work was to extend our knowledge of the sources and fates of C compounds during AM fungal spore germination and the metabolic pathways connecting both. | |||||||
Production and Labeling of Spores Ri T-DNA-transformed carrot (Daucus carota L.) roots colonized by Glomus intraradices Schenck & Smith (DAOM 197198, Biosystematic Research Center, Ottawa) were grown in Petri plates with two compartments as described by St-Arnaud et al. (1996). The roots were confined to one compartment but the fungus was allowed to grow over the divider and into the other compartment. In the following 9 to 10 weeks the fungus grew and sporulated extensively in this fungal compartment. To produce prelabeled spores, 13C1-Glc was added to the medium through a sterile filter (25 mm final concentration) in the compartment containing the mycorrhizal roots 1 to 2 weeks after the fungus had crossed the plastic barrier (Pfeffer et al., 1999).Spores and extraradical hyphae from the fungal compartment were recovered by blending the solidified medium in sodium citrate solution (10 mm) at high speed for 45 s in a commercial blender (Waring). Fungal tissue was collected on a 38-μm sieve and rinsed with water. Spores from the fungal compartments of three plates were combined for each sample. For ungerminated spore samples, fungal tissue was frozen at −80°C immediately after collection. Spores were germinated while incubating in liquid M medium (Bécard and Fortin, 1988) without Suc for 14 d (32°C in 2% CO2, in the dark). When prelabeled spores were germinated, no carbon source was added to the liquid medium. 13C-labeled substrates (99% 13C-enriched) were added as filter-sterilized solutions to the M medium of unlabeled spores for a final concentration of 25 mm for 13C1- or 13C1,2-Glc, 13C1-Fru, or 13C1-mannitol experiments and to 4 mm for 13C1- or 13C2-acetate experiments. For 13CO2 labeling, unlabeled spores were incubated at room temperature in the presence of 2% 13CO2 (generated by addition of a 50% solution of lactic acid to solid K213CO3 in a sealed 6-L desiccator). In all cases more than 80% of the spores germinated within 3 d and formed a macroscopically visible diffuse mycelium during the incubation period. After incubation, the fungal material was recovered and frozen at −80°C until carbohydrate and lipid extraction. Extraction and NMR Sample Preparation Extracts were prepared as previously described (Pfeffer et al., 1999) with minor modifications. Samples were lyophilized and ground with a mortar and pestle with acid-washed sand at −20°C in 40 mL of methanol:water (MeOH:H2O) (70:30, v/v). After filtration using Whatman no. 2 filter paper, the methanol was removed by evaporation under reduced pressure and the aqueous solution was freeze-dried. For NMR analysis, the extract was dissolved in 750 μL of 2H water and insoluble matter was removed by centrifugation.To extract neutral lipids and fatty acids, the solid residue remaining after MeOH/H2O extraction was freeze-dried and re-extracted in 30 to 40 mL of boiling isopropyl alcohol for 20 min. After filtering, the solvent was removed by evaporation under a stream of nitrogen. These extracts were dissolved in 2H chloroform for NMR analysis. NMR Spectroscopy Spectra were obtained using a 400 MHz spectrometer (UnityPlus, Varian, Palo Alto, CA) with a superconducting magnet (9.4T, Oxford Instruments, Concord, MA), although several 1H extract spectra were acquired at 600 and 750 MHz on spectrometers (Brüker, Billerica, MA). A 5-mm broad-band probe was used for 13C spectra, while a 5-mm, 1H inverse-detection probe with a Z gradient and broad-band decoupling coils was used for 1H and 1H-13C correlation spectra.13C spectra were accumulated with 80° pulse angles, WALTZ-1H decoupling, and a recycle time of 4.2 s (MeOH extracts) or 13.2 s (isopropyl alcohol extracts). For 1H spectra, 80° pulses and recycle times of 4.5 s were used; when necessary, 12-s recycle times were employed to prevent distortion of the relative intensities of the 1H-12C and 1H-13C signals (London, 1988). Total acquisition times for both 13C and 1H extracts were between 12 and 36 h depending on the concentration of each sample. The identification of peaks in 13C and 1H spectra was made from literature values (Gunstone et al., 1977, for lipids; Fan, 1995, for all other metabolite assignments), via comparison of spectra of purified compounds (e.g. trehalose in Fig. 1A) or natural abundance (n.a.) signals of unlabeled tissue extracts (e.g. Fig. 1B), by spiking extracts with purified compounds, and/or by analyses of 1H and 1H-13C correlation spectra (Pfeffer et al., 1999). 13C chemical shifts were referenced to the signals of C1-trehalose (T1, 94.1 ppm) or chloroform (77.0 ppm). 1H signals were referenced to water (4.67 ppm at 35°C) or to the chloroform peak (7.24 ppm). Carbon and proton shifts were expressed in parts per million with respect to tetramethylsilane at 0 ppm.
Quantification of 13C-Labeling 13C-Isotopic abundance (atom percent 13C) of the labeled positions of a given compound was calculated by comparison with n.a. 13C-isotopic signals of unlabeled positions of the same compound and/or by measurement of the 13C-1H satellites of 1H signals in proton spectra. Specifically, the satellites of the 1H signals coupled to carbons: (a) 1,1′ of trehalose at 5.18 ppm (Fig. 2C, inset); (b) of the unknown betaine at 3.25 ppm (Fig. 2A, inset) for MeOH/H2O extracts; (c) of the methyl group (C16) of fatty acids at 0.88 ppm (not shown); and (d) at the 1 and 3 positions of glyceryl (Gly1, Gly3) (4.2 ppm) for lipid extracts (Fig. 3A, inset) were routinely used. From these measures of absolute 13C content, the 13C content at other positions was then deduced by comparison of the intensities of the respective resonances in the 13C spectra.
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Labeling in Fungal Carbohydrate Representative 13C-NMR spectra obtained for the MeOH/H2O extracts from each treatment are shown in Figure 2. Peaks at 94.1, 73.4, 72.9, 71.9, 70.5, and 61.4 ppm correspond to the chemical shifts of carbons (1,1′), (3,3′), (5,5′), (2,2′), (4,4′), and (6,6′) of trehalose (Fig. 2, T1–T6, compare with Fig. 1A). The predominance of trehalose signals in such spectra is in accordance with previous studies (Bécard et al., 1991, Pfeffer et al., 1999) in which trehalose was the most abundant extracted carbohydrate in AM fungal spores.Levels of 13C-labeling (percent in excess of n.a.) for each C position of fungal trehalose (T1–T6) following various labeling treatments are shown in Table I. In prelabeled, non-germinated spores (Fig. 2A), labeling was found in all C positions, with T1, T6, T5, and T3 having the most incorporation of label. When 13C-prelabeled spores were germinated without exogenous label, a similar pattern was observed in trehalose with slight decreases in T1 and T3 labeling (Fig. 2B; Table I).
Spores germinated for 14 d showed different 13C-labeling patterns depending on the substrate provided. Spores exposed to 13C1-Glc (Fig. 2C) or 13C1-Fru (not shown) yielded trehalose mostly labeled in T1 (Table I). We observed no evidence of scrambling of the label provided as T1 into the T6 position. These results were confirmed when using 13C1,2-Glc as a substrate (Table I), with most of the label being found in the T1 and T2 positions. Incubation with 13C1,6-mannitol produced no labeling in trehalose or elsewhere (not shown). Spore germination in the presence of either 13CO2 or 13C1-acetate yielded trehalose with the highest labeling found in the T4 position (Fig. 2, D and E; Table I). In these two treatments the T3 position was also highly labeled, T2 and T1 were less so, and T6 and T5 were either minimally labeled or unlabeled. In contrast, upon exposure to 13C2-acetate, detected fungal trehalose displayed a very different labeling pattern, with T4 containing the least amount of label and T6 and T5 having the highest (Fig. 2F; Table I). This latter labeling pattern closely resembles the one observed for prelabeled, germinated spores (Table I). Splittings (multiplets) observed in trehalose peaks, particularly for experiments with 13C2-acetate (Fig. 2F) but also in extracts of 13C1-acetate and 13CO2 labeled samples (Fig. 2, D and E), were due to homonuclear coupling between 13C nuclei at adjacent carbon positions of the same molecules. These couplings arising from multiply-labeled molecules are expected when a product such as hexose is made from more than one molecule of labeled substrate. The 13C spectra of MeOH/H2O extracts also showed evidence of labeling in two amino acids: Glu and Arg. The small signals from Glu (Fig. 2, A, B, and F) showed that it was labeled in γ and α carbons in prelabeled, germinated spores, whereas labeling was only detected in the γ carbon in prelabeled, nongerminated spores. Spores germinated in 13C1-acetate had labeling in the δCO position (182.0 ppm, not shown), and the ones germinated in 13C2-acetate had labeling in the α, γ, and β positions of Glu. The 13C spectra also showed consistent labeling of Arg. Spectra from prelabeled, germinated spores showed labeling at the γ, β, δ, and α carbons of Arg; unlabeled spores germinated in the presence of 13C1-Glc showed labeling at the δ and α carbons; 13CO2 resulted in labeling in the terminal C=N and carboxyl positions; 13C1-acetate labeled δCH2 and carboxyl groups; and 13C2-acetate labeled the γ, α, and β carbons (Fig. 2; Table I). Notably, no Arg signals were found in spectra of prelabeled, non-germinated spores. A single resonance with a chemical shift of 52.9 ppm in the 13C spectra of MeOH/H2O extracts of prelabeled spores (germinated or not) arises from an unknown compound (Fig. 2, A and B, [CH3]3N+). This 13C signal was shown (by 1H and 1H-13C correlation spectra, see Methods) to arise from a carbon with attached protons whose signal in the 1H spectrum is a singlet at 3.25 ppm (Fig. 2A, inset). The 13C and 1H-chemical shifts and 13C-1H-coupling (145 Hz) are similar to those of methyl groups of betaines ([CH3]3N+). Signals from this putative (CH3) 3N+ group were also detectable in spectra of unlabeled spores germinated in 13C1-Glc (Fig. 2C) and 13C1,2-Glc (not shown). Estimations of the 13C content of this group (Table I) were carried out by 1H-13C-satellite measurements (Fig. 2A, inset). The (CH3)3N+ peak of this betaine-like compound showed approximately 8% 13C enrichment in prelabeled spores (either germinated or not), but less than 2% in 13C1-Glc labeling experiments, whereas in other treatments the (CH3) 3N+ peak corresponded to n.a. signals. Labeling in Fungal Lipids A previous study of this system (Pfeffer et al., 1999) showed that when labeled hexose was provided to the host roots during spore formation (corresponding to the prelabeled, nongerminated spores in this study) fungal lipids became labeled both in glyceryl and fatty acid moieties. Here we observed similar lipid labeling (approximately 8% over n.a.) in prelabeled spores, whether germinated or not (Fig. 3A, Glyc1,3:Glyc2 ratio > 2:1; C16:C15 ratio > 1:1). However, only n.a. 13C signals were observed in lipids when any of the labeled substrates was supplied to germinating spores (Fig. 3B, Glyc1,3:Glyc2 ratio = 2:1; C16:C15 ratio = 1:1). | |||||||
The fact that AM fungi do not grow axenically has puzzled and frustrated researchers for over three decades. Limitations in the metabolism or uptake of carbon in the asymbiotic state have been proposed to explain this failure (for review, see Azcón-Aguilar et al., 1998). To test this possibility it is necessary to define which pathways, if any, may be deficient in the germinating spore. To this end, we have compared the labeling patterns in different products of metabolism with the patterns expected from the operation of known metabolic pathways. Figure 4 summarizes our interpretation of the results by showing the flow of labeled C from each substrate provided to each product detected via key intermediate compounds whose presence is implied but which were not detected.
Trehalose as an Indicator of Hexose Metabolism Trehalose became labeled when any of several labeled exogenous precursors was provided to the asymbiotic fungus, demonstrating that it is synthesized during spore germination (Fig. 4, pathway 1). Since trehalose is a Glc dimer, its labeling pattern reflects labeling in hexose within the fungus and therefore can be used to follow hexose uptake and/or synthesis. Previous studies have demonstrated that trehalose levels decrease during spore germination in many fungal species (Thevelein et al., 1982, and refs. therein), among them AM fungi (Bécard et al., 1991) (Fig. 4, pathway 2). Thus, the labeling obtained reflects turnover rather than net production of trehalose (Fig. 4, pathways 1 and 2). This is consistent with observations of rapid synthesis and degradation of trehalose in the symbiotic state (Shachar-Hill et al., 1995) and with the hypothesis by Pfeffer et al. (1999) that trehalose may act to buffer intracellular Glc levels.Hexose Uptake and Utilization When germinating spores were exposed to hexose (13C1 or 13C1,2-Glc, or 13C1-Fru) the labeling was observed in trehalose, Glu, Arg, and in an unidentified betaine-like compound (see below). Thus, hexose can enter and be metabolized in the asymbiotic state via glycolysis (pathway 5 of Fig. 4). There was little or no scrambling from the labeled C position(s) of the hexose substrates tested to other positions of trehalose (Table I), suggesting that hexose is directly incorporated into trehalose (Fig. 4, pathway 1). The levels of labeling in trehalose when 13C-hexose was supplied (5%, Table I) were modest compared with other substrates, indicating that in the germinating spore intracellular hexose reaches only a low fractional enrichment. This may be because its uptake is much less than in the intraradical phase (where it reaches 80% within 24 h, Shachar-Hill, 1995) and/or because it is diluted by unlabeled hexose being synthesized internally from other C sources.Dark Fixation of 13CO2 and Gluconeogenesis The observation of substantial labeling in trehalose when 13CO2 is supplied (Table I) demonstrates a significant rate of dark fixation (presumably by pyruvate carboxylase, Fig. 4, pathway 8a). Moreover, the labeling of mainly T4 and T3 of trehalose is the pattern usually observed in hexose when labeled CO2 is supplied to tissues undergoing gluconeogenesis (Fig. 4, pathways 8a, then 8b and 8c, and then 4; see e.g. Brosnan, 1982). Bécard and Piché (1989) showed that CO2 stimulates the asymbiotic growth of another AM fungus, Gi. margarita, and suggested that CO2 may be a net source of carbon for anabolic processes. Dark fixation as part of gluconeogenesis does not lead to a net gain of C, since the same amount of CO2 is released by decarboxylation (by PEP carboxykinase to produce PEP, Fig. 4, pathway 8c) as is fixed by pyruvate carboxylase. However, labeling in the carboxyl of Arg indicates that CO2 fixation also has an anaplerotic role.Glyoxylate Cycle During spore germination, the synthesis of cell walls and DNA, among other anabolic processes, must create large demands for carbohydrate units. Since the majority of carbon storage in AM fungal spores is as lipid and not as carbohydrate, one would expect the conversion of lipids to hexose to be important at this stage of the fungal life cycle. Acetate was used to test this hypothesis, since it is a precursor of acetyl CoA, itself a key intermediate in fatty acid breakdown.The labeling pattern of trehalose extracted from germinating spores incubated with 13C1-acetate was similar to that observed in 13CO2 labeling experiments (Table I). This could arise from the liberation (as 13CO2) of the labeled C1 carbon of acetate via the TCA cycle (pathway 10 of Fig. 4) followed by dark re-fixation of the released 13CO2, as indicated above. However, the same labeling pattern is also predicted by the conversion of acetate to triose via the glyoxylate cycle (Fig. 4, pathways 9, 4, and 1). The results of labeling with 13C2-acetate suggest that both of these mechanisms occur. The glyoxylate cycle transfers label supplied in C2 of acetate to carbons 6, 1, 5, and 2 of trehalose (following pathways 9, 4, and 1 of Fig. 4), and the labeling observed in carbons 3 and 4 can be accounted for by release of label as 13CO2 (via second and further turns of the TCA cycle) and its refixation (Fig. 4, pathways 8a, 8b, 8c, 4, and 1). The very high level of labeling reached in trehalose when 13C2-acetate was provided (close to 80% in T6) indicates a high metabolic flux from acetyl-CoA to hexose. Such a high flux would be expected if most of the intracellular hexose during the asymbiotic phase were made from storage lipids (sequentially following pathways 7, part of 10, 9, and 4 of Fig. 4). Results from prelabeled spores are also consistent with conversion of lipid to trehalose during spore germination. The fatty acids in prelabeled spores are labeled predominantly in the even-carbon positions of the fatty acid chains (Pfeffer et al., 1999) and thus yield C2-labeled acetyl CoA, as would be expected of 13C2 acetate. Indeed, the relative enrichment in different positions of the trehalose from prelabeled, germinated spores (Table I) is similar to that after 13C2-acetate labeling. PPP Gluconeogenic flux from labeled triose to hexose during germination is expected to result in equal labeling in carbons 6 and 1, in carbons 5 and 2, and in 4 and 3. However, we observed markedly asymmetrical trehalose labeling patterns in experiments using both 13C1 and 13C2-acetate and 13CO2 (Table I). Deviation from this symmetrical labeling pattern can arise in hexose due to the action of the PPP (Fig. 4, pathway 3). The scrambling expected from the operation of the PPP can explain the patterns of asymmetry observed in trehalose: C4 > C3 from 13C1-acetate and from 13CO2, but C3 > C4 from 13C2-acetate. The operation of the PPP is also consistent with the labeling of C1 and C2, and the absence of labeling in C5 and C6 of trehalose when 13CO2 or 13C1-acetate were provided. The PPP is known to be active in fungi (for review, see Jennings, 1995) and its action is consistent with recent observations in the symbiotic state (Pfeffer et al., 1999). However, the results of labeling with 13C1,2-Glc (Table I) do not indicate any PPP activity, since in this case the PPP should result in greater labeling in C1 than in C2. Moreover, estimations of PPP activity carried out in this study (by comparing the central peak of the labeled positions T1 or T2 with the single peak of the n.a. positions T6, T5, or T4 when 13C1,2-Glc was provided exogenously) resulted in no apparent activity of this metabolic pathway. Further experiments are required before alternative explanations (such as more than one intracellular hexose pool) can be fruitfully proposed.Amino Acid Synthesis 13C-Labeling of Glu and Arg shows that these amino acids are synthesized during asymbiotic fungal growth. Whereas Glu labeling was also detected during the symbiotic phase (seen in prelabeled, non-germinated spores), Arg synthesis was only detected during germination (Table I). The patterns of labeling in both amino acids from the various labeled substrates are consistent with the usual biosynthetic routes (Fig. 4, pathways 5 and 10 for Glu; pathways 5, 10, 8d, and 11 for Arg). The production and labeling of Arg reveals the presence and operation in the asymbiotic AM fungus of most enzymes of the urea cycle (Fig. 4, pathway 11), although there is no evidence that the last step, which produces urea (through arginase activity), is active.There are a number of possible roles for Arg during AM fungal spore germination. One of them is as a charge balance for polyphosphates (Cramer et al., 1980; Cramer and Davis, 1984; Bücking et al., 1998). However, the state of polyphosphates in mycorrhizal fungi has been the subject of considerable debate (Kulaev, 1979; Martin et al., 1985; Orlovich and Ashford, 1993; Bücking et al., 1998), and in our view, there is still no conclusive evidence that Arg (or other basic amino acids in the fungal vacuole) is bound to these macromolecules. Other possible explanations for the observed formation of Arg during germination include roles in nitrogen storage, as a compatible osmolyte, and as a possible form of N transferred to the host. A Betaine-Like Compound We consistently observed in the spectra of prelabeled spores (either germinated or not) and of spores germinated in the presence of 13C-Glc, 1H and 13C signals from an unidentified compound. Because of a number of spectroscopic characteristics we believe that these signals arose from the nitrogen-bonded methyl groups of a betaine-like molecule. The reasons for this tentative assignment include: (a) the spectral position of the 1H and 13C signals compared with known betaines, (b) the absence of homonuclear couplings in the 1H signal, (c) the size of the coupling constant between 1H and 13C, and (d) the liberation of trimethylamine upon heating (not shown). The fact that labeling of this compound was not observed in 13C-acetate-labeling experiments indicates that it is not made from lipids. The labeling obtained when 13C1-Glc was used is consistent with the origin of methyl groups through non-photosynthetic one-carbon metabolism from hexose via Ser (Fig. 4, pathways 5 and 6).Absence of Storage Lipid Synthesis None of the substrates provided during spore germination resulted in detectable labeling of lipids. Since there is clear evidence that acetyl-CoA becomes significantly labeled by at least 13C-acetate, we would expect to detect labeling if 1% or more of the lipid had been synthesized during the incubation period. We therefore believe that, in contrast to trehalose, storage lipids are not synthesized to any great extent during asymbiotic fungal growth. Beilby (1983) found modest labeling of triacylglycerides when 14C-acetate was provided to G. caledonium spores during spore imbibition. This difference may reflect the much greater sensitivity of radiotracer methods or the different timing of the experiments. Beilby's data also showed a consistent increase in the labeling of the phospholipid fraction during germination, which is indicative of membrane synthesis. Since our isopropyl alcohol extracts contained predominantly neutral storage lipids (Pfeffer et al., 1999), we believe that during germination, lipid synthesis is largely or entirely confined to membrane production.Carbon Metabolism in the Asymbiotic versus the Symbiotic AM Fungus Previous studies have shown that during symbiosis, intraradical and extraradical parts of AM fungi are very different in their C metabolism: substantial hexose uptake occurs in the intraradical phase (Shachar-Hill et al., 1995; Saito, 1995, Solaiman and Saito, 1997; Pfeffer et al., 1999) and here also is the site of storage lipid synthesis. In contrast, extraradical hyphae are unable to take up hexose and do not synthesize storage lipids (these are transported from the intraradical to the extraradical mycelium, Pfeffer et al., 1999). When growing asymbiotically, AM fungi have characteristics in common with each of the two metabolic states described above. That is, both hexose and acetate can enter the AM germ tubes and be metabolized; however, the synthesis of storage lipids is either blocked or greatly reduced in the asymbiotic fungus. Since gluconeogenesis and the glyoxylate cycle involving lipid breakdown are active pathways in asymbiotic germ tubes, it may be that a transition from catabolism to anabolism of lipids (initiated when intimate contact is established with the host plant) is essential to the AM fungus in order to fully develop and complete its life cycle. | |||||||
ACKNOWLEDGMENT We thank Dr. Kathleen Valentine of the University of Pennsylvania NMR facility for obtaining some of the 13C-1H satellite spectra. | |||||||
Footnotes 1This work was supported in part by grant no. 97–35107–4375 from the National Research Initiative Competitive Grants Program/U.S. Department of Agriculture and via a fellowship under the Organisation for Economic Co-operation and Development Co-operative Research Program: Biological Resource Management for Sustainable Agriculture Systems (to G.B.). The research utilized in part the Resource for Solid-State NMR of Proteins at the University of Pennsylvania: A National Institutes of Health Supported Research Center (grant no. P41RR09731 from the Biomedical Research Technology Program, National Center for Research Resources, National Institutes of Health). | |||||||
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