Mycol. Res. 104 (1) : 81–86 (2000)
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Phosphatase activity of external hyphae of two arbuscular mycorrhizal fungi
E R I K J. J O N E R1* A N D A N D E R S J O H A N S EN2† " Department of Biotechnological Sciences, Microbiology Section, Agricultural University of Norway, P.O. Box 5040, N-1432 Ab s, Norway, # Plant Nutrition, Environmental Science and Technology Department, Risø National Laboratory, DK-4000 Roskilde, Denmark
Extraradical hyphae of Glomus intraradices or G. claroideum were extracted from root free sand of two-compartment pot cultures and used to determine fungal phosphatase activity [p-nitrophenyl phosphate (p-NPP) hydrolysis]. Enzyme activity was assayed with respect to pH, temperature and different fractions of the hyphae (external soluble, wall-bound and internal phosphatases). The results showed an overall maximum enzyme activity at pH 5n2–5n6 for both fungi, with a possible secondary maximum at pH 8n8 for G. claroideum. Of the two fungi tested, G. intraradices had the highest external phosphatase activity in two experiments and the same activity in one experiment, reaching 184 µmol p-NPP hydrolysed mg−" .. h−". Phosphatase activity at pH 5n2 decreased sharply with temperature, with 4n5 and 10n5 % of the enzyme activity remaining at 5 mC relative to that at 37m for G. intraradices and G. claroideum, respectively. Separation of the phosphatase activity into external soluble, wall-bound and internal fractions revealed that up to 70 % of the measured activity was associated with the hyphal wall, and the rest with internal structures. Exuded phosphatases were not found in measurable amounts. The implications of these results on possible hyphal utilization of organic P in soil are discussed.
Arbuscular mycorrhizal (AM) fungi are important to plant phosphorus (P) nutrition in many soils through hyphal transport to the plant (Jakobsen, Joner & Larsen, 1994), but a large pool of P in soil, the organic P, is equally unavailable to both AM and non-mycorrhizal plants (Joner & Jakobsen, 1995 a). Plants growing in soils that do not receive ample amounts of inorganic P fertilizers thus depend to a large extent on mineralization of P from organic material to acquire adequate P (Dalal, 1977). In this context one step in the mineralization process, the action of enzymes that hydrolyse organically bound phosphate in soil (acid and alkaline phosphatase ; i.e. the orthophosphoric monoester phosphohydrolases ; EC 3;1;3;1 and 3;1;3;2, respectively), has received much attention (Skujins, 1976 ; Tarafdar & Claasen, 1988 ; Asmar, Gahoonia & Nielsen, 1995 ; Tibbett, Sanders & Cairney, 1998). Phosphatase activity has sometimes been found to coincide with disappearance of organic P (Tarafdar & Jungk, 1987 ; Helal & Sauerbeck, 1991), whereas others have been unable to establish this relationship (Hedley, Nye & White, 1982 ; Helal & Sauerbeck, 1984 ; Joner & Jakobsen, 1995 b). One likely explanation for the inconsistency in organic P disappearance may be differences in the availability of organic P for enzymatic attack due to adsorption and * Present address, and address for correspondence : Centre de Pedologie Biologique – CNRS, 17, rue N.D. des Pauvres, B.P. 5, F-54501 Vandoeuvreles-Nancy Ce! dex, France. † Present address : Department of Genetics and Microbiology, Royal Veterinary and Agricultural University, Thorvaldsensvej 40, DK-1871 Fredriksberg C, Denmark.
complexation in soil, and the various forms of organic P present in different experimental soils (Magid, Tiessen & Condron, 1996). The activity of phosphatases thus may or may not be the rate limiting factor in mineralization of organic P in soil (see Adams & Pate, 1992 ; Joner 1994 ; Joner et al., 1995), but in either case these enzymes still seem to be essential to one step in the process : that where phosphate is finally cleaved from its organic moiety. Phosphatase activity has also been used as a general biochemical indicator in measurements of biological activity, as it is present in many soil organisms, and responds to adverse conditions like pollution and soil degradation (e.g. Kuperman & Carreiro, 1997 ; Trasar-Cepeda et al., 1998). In organisms phosphatase may be used as a general indicator of metabolic activity, and as such it has been used extensively in connection with vital staining of intraradical (e.g. Tisserant et al., 1993 ; Larsen et al., 1996) and extraradical (Zhao et al., 1997 ; Vosatka & Dodd, 1998) AM hyphae. The presence and effect of phosphatases of extraradical hyphae of AM on organic P substrates in soil have recently been reported, though results are contradictory (Tarafdar & Marschner, 1994 b ; Joner et al., 1995). A major problem in detection of phosphatase from external AM hyphae is the obligate symbiotic nature of these fungi which makes it difficult to distinguish hyphal phosphatases from those of plant roots and soil. Compartmented growth systems have been employed to avoid the interfering effect of roots, but hyphal enzyme activity has so far only been measured in soil or soil-sand mixtures after growth under non-sterile conditions
Phosphatases of arbuscular mycorrhizal hyphae where enzymes are adsorbed onto and possibly inactivated by clay, organic materials and microbial proteases (Burns, 1982 ; Dick, Juma & Tabatabai, 1983). Besides inactivation, the intrinsic phosphatase activity of soil organic matter and phosphatases exuded by other microorganisms give rise to a high background activity that renders a small difference between mycorrhizal and non-mycorrhizal treatments (Joner & Jakobsen, 1995 b). Thus, measuring AM hyphal phosphatase activity as the difference between activities in soil from root free compartments with and without AM hyphae is inaccurate and unprecise. Our aim was to verify the existence of extracellular phosphatases of external hyphae of two AM fungi and quantify it on a biomass basis with a low level of interference from soil (intrinsic soil enzyme background, other microbial phosphatases, soil adsorption or microbial degradation of AM phosphatases etc.). This was attempted through measurements of the enzymatic activity of AM hyphae grown in sand-filled hyphal compartments after washing excised mycelium from the sand, a cultivation technique previously used to study hyphal uptake and assimilation of inorganic nitrogen in vitro (Johansen, Finlay & Olsson, 1996). Secondly, we wanted to measure phosphatase activity as influenced by pH and temperature to determine optimum pH for activity of external phosphatases of AM hyphae, and to be able to assess the importance of phosphatases at below-optimum temperatures, as normally encountered in the field, respectively. Thirdly, we wanted to fractionate the enzyme activity between external soluble, wall-bound and internal enzymes to evaluate their relative importance, and determine if they had different pH optima, i.e. if they were indeed different enzymes. Finally, we wanted to compare phosphatase activity of pure mycelium to previous results where phosphatase activity was measured in root-free soil containing AM hyphae. This would indicate to what extent the involved hyphal enzymes can participate in organic P mineralization, and predict if this contribution to the total phosphatase activity of soil is ecologically significant.
MATERIALS AND METHODS Plants and fungi Subterranean clover (Trifolium subterraneum L., cv. Mount Barker) was grown in bags of nylon mesh (20 µm) inoculated with approx. 500 surface sterilized (5 % Chloramin T) spores of Glomus intraradices N. C. Schenck & G. S. Sm. (Danish isolate 28A, BEG 87) or G. claroideum N. C. Schenck & G. S. Sm. (Danish isolate SC09, BEG 14). The growth medium in the mesh bags consisted of equal volumes of Leca2 (beads of expanded clay), sand and Grodania2 (pieces of mineral wool). The mesh bags were placed centrally in 1n5 l pots lined with plastic bags and the space outside the mesh bag was filled with washed quartz sand (1n4 kg, particle size : 0n2–0n6 mm ; 90 %, 0n06–0n2 ; 10 %) to serve as hyphal compartment. All growth media were irradiated (10 kGy, electron beam) to keep the presence of foreign microorganisms at a low level. The hyphal compartment was covered with non-transparent PVC and plastic to reduce microbial contamination and prevent algal growth. Plants were kept in a growth chamber with a 16–8 h
82 light–dark cycle, at 24–15 mC (day–night), and an average photon flux density of 350 µmol m−# s−". A nutrient solution (1 m Ca(NO ) :4H O ; 1 m NH NO ; 1n0 m K SO ; $# # % $ # % 0n8 m MgSO :7H O ; 70 µ (Na HPO :2H O ; 25 µ % # # % # Fe() NaEDTA ; 25 µ H BO ; 5 µ MnSO :H O ; 2 µ $ $ % # ZnSO :7H O ; 0n5 µ CuSO :5H O ; 0n1 µ Na MoO : % # % # # % 2H O ; 4 n CoCl :6H O, adjusted with HCl to pH 6) was # # # supplied daily to maintain the equivalent to 70 % of the water holding capacity of the sand.
Hyphal extraction Hyphae were extracted from pots when plants were 6–10 wk old. Ten to fourteen days prior to harvesting the hyphae, the sand in the hyphal compartment had been renewed to obtain hyphae of similar age without excessive amounts of spores in the subsequent harvest. Extraction was done by flotation in a large tray, washing out the bulk of the remaining sand in a beaker and finally cleaning the mycelium under a dissecting microscope. The cleaned mycelium was then divided into approximately equal portions for enzyme measurements. The process of hyphal extraction and cleaning took 30–60 min, during which they were kept in tap water at room temperature. Immediately after extraction\cleaning they were subject to enzyme assays.
Phosphatase measurements Phosphatase activity was measured in 0n2 modified universal buffer (MUB)(Tabatabai, 1982) with a final substrate concentration of 1 mg p-nitrophenyl phosphate ( p-NPP) ml−". In the experiments measuring phosphatase activity at different pH and temperature the hyphae were incubated in 3 ml buffer\substrate in 15 ml polypropylene tubes for 1 h in shaking baths (40 rpm), after which 1n6 ml, 1 NaOH was added to stop the enzymatic reaction. Hydrolysed substrate ( pnitrophenol, p-NP) was then determined spectrophotometrically at 405 nm in the supernatant after centrifugation for 10 min at 1400 g. Blanks prepared from buffer, substrate and hyphae, but with NaOH added before the hyphae, were subtracted. Additional blanks of buffer\substrate, but without hyphae and NaOH were also checked. Hyphal dry weight was determined by resuspending, washing and filtering [1 µm polycarbonate membrane filter (Nuclepore, Pleasanton, CA)] the pellet, removing remaining sand grains under a dissecting microscope, drying (80mC, 24 h) and weighing. Determination of pH optimum for phosphatase activity of excised hyphae (external solublejexternally available wallbound phosphatase) was carried out in MUB (37m) adjusted to pH 4n0, 4n6, 5n2, 5n8, 6n4, 7n0, 7n6, 8n2 and 8n8, using three replicate tubes for each fungus and pH. Phosphatase activity as affected by temperature was measured similarly after incubating triplicate tubes with MUB (pH 5n2), substrate and hyphae in waterbaths at 5, 12, 20 and 37m. Soluble external phosphatase activity was measured by incubating four replicate tubes with hyphae of each fungus in MUB (pH 5n2) for 1 h, and filtering before adding substrate to the filtrate. Wall-bound phosphatase activity was obtained in
E. J. Joner and A. Johansen
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RESULTS Excised mycelium of either fungus displayed significant phosphatase activity with maximum activity at pH 5n2–5n8 (Fig. 1). G. claroideum had a possible secondary maximum at pH 8n8. The total phosphatase activity for G. intraradices reached a maximum of 3n08 enzyme units (EU l µmol p-NPP hydrolysed g−" .. min−"), being up to 78 % higher than that of G. claroideum at acid pH, decreasing to 24 % higher at the highest pH tested. Phosphatase activity of both fungi increased exponentially with temperature, though G. claroideum had higher phosphatase activity at 5 and 12m and lower activity at 20mC than G. intraradices (Fig. 2). As in the experiment with different pH, G. intraradices also had a higher phosphatase activity than G. claroideum at 37m, but the relative difference was only 17 %. At 5m the activity was 5 and 11 % of the corresponding activity at 37m for G. intraradices and G. claroideum, respectively.
EU (lmol p -NPP g–1 d.w. min–1)
3·5
Glomus intraradices Glomus claroideum
3·0 2·5 2·0 1·5 1·0 0·5 0·0
4
5
6
7
8
9
pH
Fig. 1. External phosphatase activity of extraradical hyphae of two AM fungi as influenced by pH in the incubation medium. Bars represent ... (n l 3).
EU ( lmol p -NPP g–1 d.w. min–1)
3·5 Glomus intraradices Glomus claroideum
3·0 2·5 2·0 1·5 1·0 0·5 0·0
5
10
15 20 25 Temperature (° C)
30
35
Fig. 2. External phosphatase activity of extraradical hyphae of two AM fungi as influenced by temperature of the incubation medium. Bars represent ... (n l 4).
Relative P-ase activity (% of max.)
parallel tubes by incubating hyphae with buffer\substrate, removing the hyphae by filtration after 1 h, and subtracting the values for soluble phosphatase activity. In both cases, pNP was measured at different times from 0 to 180 min after filtration. Activity of internal phosphatase was determined by macerating hyphae of G. intraradices with acid-washed quartz sand in 1 ml buffer (MUB, pH 7n2) in a ball mill (1 min). The suspension of macerated hyphae and sand was washed from the ball mill container into a syringe with a buffered (MUB, pH 7n2) suspension of polyvinyl pyrrolidone (20 g l−"), ascorbic acid (5 m) and NaCl (100 m) to a total volume of 4 ml. This volume was either filtered (Sartorius Minisart2plus, 0n2 µm cellulose acetate filter unit fitted on the syringe) to yield soluble internal phosphatase, or not (wall-boundj soluble internal phosphatase). Subsamples (0n15 ml) of either of these fractions were transferred to tubes with 1n5 ml buffer\substrate (MUB, pH 5n2, 6n4, 7n6 or 8n8 ; n l 3), and incubated in a shaker at 37m for 1 h. Results are presented as means of three or four parallel measurements, and estimated sample variation by the ...
100 Pellet Extract
80 60 40 20 0
5
6
7 pH
8
9
Fig. 3. Relative difference in soluble internal phosphatase (P-ase) activity (extract) and soluble internaljwall-bound phosphatase activity (pellet) of G. intraradices hyphae as influenced by pH in the incubation medium. Bars represent ... (n l 3).
Macerated hyphae incubated with substrate (wall-boundj soluble internal phosphatase) displayed enzymatic activity that decreased with increasing pH (Fig. 3). The relative difference between pH 5n2 and pH 8n8 being higher when hyphae were macerated (89 % decrease) than when hyphae were intact (67 %, see Fig. 1). The phosphatase activity in corresponding filtrates of macerated hyphae (soluble internal phosphatase only) also decreased with increasing pH, and comprised 29 % of the activity in unfiltered samples at pH 5n2 and 76 % at pH 8n8. Soluble external phosphatase activity [measured in buffer (pH 5n2) incubated with hyphae of either fungus for 1 h prior to removing hyphae and adding substrate] was too low to be distinguished from that of the background auto hydrolysis after a chase period of 3 h (Fig. 4). Similar measurements 0–3 h after incubation of hyphae in buffer and substrate showed high activity of wall-bound phosphatase at the time when hyphae were removed, but again no significant increase during the chase period.
Phosphatases of arbuscular mycorrhizal hyphae
250
Glomus intraradices Wall bound Soluble
200 P-ase activity (lmol p-NPP g–1 d.w.)
84
150 100 50 0
250
Glomus claroideum
200 150 100 50 0 0
25
50
75 100 Time (min)
125
150
175
Fig. 4. Extracellular phosphatase (P-ase) activity in incubation medium (pH 5n2) recorded in a 180 min chase period after incubation with hyphae of two AM fungi with (wall-bound P-ase) or without (soluble P-ase) p-NPP for 60 min prior to the chase period. Blanks not subtracted. Bars represent ... (n l 3).
DISCUSSION This report presents evidence that phosphatases produced by external hyphae of AM fungi can hydrolyse extracellular phosphate ester bonds. Due to the difficulty in collecting sufficient extraradical fungal biomass to perform experiments on their physiology, previous measurements of phosphatase activity of AM hyphae in root-free soil have been made in the presence of soil, and have been based on the assumption that differences in phosphatase activity between root-free soil with and without AM hyphae are caused by direct exudation of phosphatases into the soil or externally available wall-bound phosphatase on the AM mycelium (Tarafdar & Marschner, 1994 b ; Joner et al., 1995). Using this approach it can not be precluded that AM hyphae induce a production of phosphatases by other organisms in the soil due to e.g. competition for available P. This may still be the case, but as we have shown here AM hyphae will contribute if present in the soil. The present experiments were not carried out under strictly axenic conditions. The mycelium used for phosphatase measurements was thus undoubtedly colonized by other micro-organisms. The extent to which this influenced enzyme activity was not measured, and thus remains uncertain. We do, however, have indications that the influence of other organisms was small. First, the microbial contamination pressure was low, the time for growth of contaminants between harvests short, and no organic nutrients were supplied. Secondly, hyphal samples from parallel pots were subject to phospholipid fatty acid analysis, showing that the presence of other microorganisms was negligible. Similar results were obtained in a study of external AM hyphal N metabolism using the same experimental system (Johansen, Finlay & Olsson, 1996). Thirdly, in connection with the latter report, we have carried out experiments on uptake of "&N by excised hyphae where
antibiotics (penicillin and streptomycin) were added to avoid bacterial growth. In these experiments, there was no influence of bacterial presence (A. Johansen, unpublished results). Lastly, rhizosphere bacteria commonly display an optimum phosphatase activity in the alkaline range (Tarafdar & Claassen, 1988), as opposed to the AM fungi analysed here. Based on these observations, we attribute the bulk of the phosphatase activity measured in these experiments to enzymes possessed by the mycorrhizal mycelium. Saprotrophic soil fungi commonly have their highest external phosphatase activity at acid pH, some as low as pH 3 (Casida, 1959). Ectomycorrhizal fungi examined by Antibus, Kroehler & Linkins (1986) had a pH optimum between 4n5 and 5n0. Concerning AM fungi, only rhizosphere soil with hyphae has been assayed for phosphatase pH optimum. Growing wheat in symbiosis with G. mosseae, Dodd et al. (1987) found highest rhizosphere phosphatase activity at pH 5n0–5n5. In good accordance with this, our measurements showed an optimum pH for two other Glomus species between pH 5n2 and 5n8. This was below the pH of the medium that the fungi were grown in (pH 6), and the bulk pH of the soil from which one of the fungi was isolated (pH 6 for G. intraradices, pH of site of origin for G. claroideum is unknown). It is thus uncertain whether the pH of the medium determines optimum pH of external phosphatases, or if this is strictly a genotypic trait of the fungus in question. Due to the potential of AM hyphae to acidify their micro-environment known as the hyphosphere (Li, George & Marschner, 1991), the phosphatases of these fungi may still be adapted to optimal functioning in their native environment. The internal phosphatases were also found to have their highest activity at acid rather than alkaline pH, but the activity at high pH was substantial, and internal phosphatases constituted 75 % of the activity at pH 8n8. This is in accordance with the use of alkaline phosphatase activity as an indicator of metabolic activity of intraradical AM hyphae (Tisserant et al., 1993 ; Ezawa, Saito & Yoshida, 1995). The pH optimum of infection-specific soluble internal phosphatases of AMcolonized roots have been found to be in the range of pH 7–8 (Gianinazzi-Pearson & Gianinazzi, 1978 ; Ezawa & Yoshida, 1994). Clearly, more careful cell fractionation or protein purification could demonstrate if internal enzymes in extraand intraradical fungal structures are the same. External soluble phosphatase was not detected, but it may be that detached hyphae perform differently in this respect compared to a functional mycelium connected to roots, and that previously exuded phosphatase was washed away during hyphal extraction. Phosphatase activity is commonly measured at 35mC, being close to the optimum temperature for enzyme functioning. Temperatures in soil are considerably lower, particularly in temperate and subarctic regions where the content of soil organic matter (and thus organic P) is high and mineralization rates are low (Read, 1991). If phosphatases are important in the mineralization of organic P under these conditions, it is crucial that the enzymes remain active at low temperatures. As shown here, an enzyme assay at 37m is not valid to estimate the activity at 5m : G. claroideum had a phosphatase activity at 5m that was twice as high as G. intraradices, though the latter
E. J. Joner and A. Johansen had the highest activity at 37m. The question of adaptation to cold environments for the fungal partner of the AM symbiosis is largely unresolved, but a variation seems to exist. Zhao et al. (1997) recently published results where the viability of external hyphae of two Glomus species was evaluated with two vital stains. Their values varied between 35 and 85 % viable hyphae at their first harvest three weeks after inoculation. In contrast, our previous measurements of several Glomus species grown in the same experimental system showed 99 % of the hyphae to be viable, as evaluated using the same two staining procedures (Joner, Johansen & Thingstrup, unpublished results). The divergence of these findings are probably related to hyphal age and that our system employed re-growth into sand from a pre-existing mycelium. A tentative calculation of AM hyphal contribution to soil phosphatase activity can be made assuming that a hyphal phosphatase activity of approx. 1n5–3 µmol p-NPP hydrolysed g−" min−" (pH 5n2), as reported here, is a valid estimation of hyphal phosphatase activity for two different Glomus species (G. caledonium and G. invermaium) in one of our previous experiments (Joner et al., 1995). In that experiment 23 m hyphae g−" soil comprised approx. 0n1 mg AM hyphae [adopting the following approximate values : mean hyphal diam. of 5 µm, 17 % dry matter in hyphae and a specific weight of 1n3 g cm−$ (Hanssen, Thingstad & Goksøyr, 1974 ; Fægri, Torsvik & Goksøyr, 1977)] and the overall soil phosphatase activity was 8 nmol p-NPP hydrolysed g−" min−". The fungi may thus have contributed 0n15–0n3 nmol p-NPP hydrolysed g−" min−", or potentially 2–4 % of the phosphatase activity of the soil. These figures are potential contributions, as they disregard that enzymes are inactivated in soil (Burns, 1982), so that only a fraction may be detectable in a phosphatase assay of a soil slurry. The low potential and further inactivation may explain why no increases in soil phosphatase activity were found due to AM hyphae in our previous experiments (Joner & Jakobsen, 1995 b ; Joner et al., 1995). Contradictory to our previous reports, Tarafdar & Marschner (1994 a, b) have repeatedly found increased soil phosphatase activity in root free soil in the presence of AM hyphae. The increase in acid phosphatase activity was up to 1n6 nmol p-NPP hydrolysed g−" min−" mediated by a hyphal length of 4n5 m g−" soil in the former case and about 1 nmol p-NPP hydrolysed g−" min−" increase mediated by 2 m hyphae g−" soil in the latter case. The reasons for these discrepancies are difficult to explain. Disregarding soil inactivation, the hyphal biomass in Tarafdar & Marschner’s experiments, which seem to be lower than in our previous experiments, has had a 10-fold higher enzyme activity (using a similar, but reversed calculation, as above) than what was measured for excised hyphae in the present experiment. As we have only examined two fungi, and as inter-fungal differences seem common (Dodd et al., 1987 ; present results), we can not exclude that differences between fungi may account for a part of this difference. Another possible explanation is that the enzyme synthesis is regulated by induction and repression, according to the presence of substrate (organic P esters) and product (inorganic P), respectively. In the present experiment, a low concentration of inorganic P was present during hyphal
85 growth, but no organic P. Our previous experiments in soil as well as Tarafdar & Marschner’s experiments featured supply of both organic and inorganic P, but actual concentrations and availability of the various P fractions were not recorded in a manner that allows comparisons. Thus, enzyme regulation mechanisms may also be part of the answer to the diverging results. Previous measurements of soil phosphatase as influenced by AM hyphae have employed soil microtome cutting to obtain a high resolution in rhizosphere gradients regarding the distance from roots (Tarafdar & Marschner, 1994 b ; Joner et al., 1995). Thus, when phosphatases have been measured in soil with extensively disrupted hyphae, internal phosphatases leaking from fungal cytoplasm might give rise to an erroneously high activity, particularly in the alkaline range where our measurements showed that approx. 75 % of the phosphatase activity belonged to the soluble internal fraction. Even at acid pH where the internal phosphatases comprised about 30 % of the total activity, it should be considered as a source of error in such measurements. Browsing through approx. 50 reports on phosphatase activity in soil or living fungal material from the last 25 years (since the p-NPP-method was introduced), it seems that fungal phosphatase activity in general is about three orders of magnitude higher than that of soil when compared on a dry weight basis. Assuming that the fungal biomass in soil is around 0n1–1 mg g−" soil, the fungal contribution to total soil phosphatase activity may constitute somewhere between 1 and 10 %. On the upper end of this scale the fungal contribution would be of significant ecological importance, particularly if extracellular enzymes originating from fungal hyphae are stabilized in soil so that they would later comprise part of the indigenous soil phosphatase activity. The AM fungal biomass is, however, not likely to constitute more than 0n1 mg g−" soil, so unless higher phosphatase activity than reported here is found in other AM fungi, or can be induced in response to available substrate, the AM fungal contribution to overall phosphatase in soil seems to be low. The work reported here was funded by a Norwegian Research Council NKJ-grant.
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