Vanadate Influence on Metabolism of Sugar Phosphates in Fungus Phycomyces blakesleeanus

Milan Žižić; Miroslav Živić; Vuk Maksimović; Marina Stanić; Strahinja Križak; Tijana Cvetić Antić; Joanna Zakrzewska

doi:10.1371/journal.pone.0102849

Abstract

The biological and chemical basis of vanadium action in fungi is relatively poorly understood. In the present study, we investigate the influence of vanadate (V⁵⁺) on phosphate metabolism of Phycomyces blakesleeanus. Addition of V⁵⁺ caused increase of sugar phosphates signal intensities in ³¹P NMR spectra in vivo. HPLC analysis of mycelial phosphate extracts demonstrated increased concentrations of glucose 6 phosphate, fructose 6 phosphate, fructose 1, 6 phosphate and glucose 1 phosphate after V⁵⁺ treatment. Influence of V⁵⁺ on the levels of fructose 2, 6 phosphate, glucosamine 6 phosphate and glucose 1, 6 phosphate (HPLC), and polyphosphates, UDPG and ATP (³¹P NMR) was also established. Increase of sugar phosphates content was not observed after addition of vanadyl (V⁴⁺), indicating that only vanadate influences its metabolism. Obtained results from in vivo experiments indicate catalytic/inhibitory vanadate action on enzymes involved in reactions of glycolysis and glycogenesis i.e., phosphoglucomutase, phosphofructokinase and glycogen phosphorylase in filamentous fungi.

Citation: Žižić M, Živić M, Maksimović V, Stanić M, Križak S, Antić TC, et al. (2014) Vanadate Influence on Metabolism of Sugar Phosphates in Fungus Phycomyces blakesleeanus. PLoS ONE 9(7): e102849. https://doi.org/10.1371/journal.pone.0102849

Editor: Richard A. Wilson, University of Nebraska-Lincoln, United States of America

Received: April 28, 2014; Accepted: June 24, 2014; Published: July 18, 2014

Copyright: © 2014 Žižić et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All relevant data are within the paper.

Funding: This work was supported by Grant of Ministry of Education, Science and Technologic Development of Republic of Serbia, 173040, www.mpn.gov.rs. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Vanadium is widely spread, naturally occurring, multivalent transition metal, existing in nature mostly in 5+ and 4+ oxidation states. It is essential in micromolar concentrations for many eukaryotic organisms, but at higher concentrations it is toxic and can represent an environmental threat [1]. Even though both oxidation states of vanadium can alter enzyme activities, particularly those involved in splitting a phosphate ester linkage [2], such effect is attributed mainly to vanadate (V⁵⁺) and arises from its structural and electronic similarity to phosphate [3], [4]. This analogy is most evident in its tetrahedral, but is also valid for other V⁵⁺ forms [5]. Vanadate is known as a potent inhibitor of many enzymes involved in sugar phosphate metabolism such as: glucose 6 phosphatase [6], phosphoglucomutase, phosphoglyceratemutase [7], [8], fructose 2,6 bisphosphatase [7]–[9]. On the other hand, it has stimulatory effect on glucose 6 phosphate dehydrogenase, ribulose 5 phosphate epimerase, phosphoglucose isomerase and adenylyl cyclase [10].

It has been demonstrated that fungi have the ability to take up and accumulate vanadium [11]. In fact, there are a few edible vanadium-rich mushrooms useful in treatment of some types of cancer, hypertension, hypercholesterolemia and hepatitis [12]. As peculiarity, high content of vanadium was confirmed in three species of Amanita mushrooms [13]. In yeasts, vanadate dimmer acts as inhibitor of glucose 6 phosphate dehydrogenase the first enzyme in phosphate-pentose pathway [14], while the tetramer has inhibitory effect on 6-phosphogluconate dehydrogenase [15]. Concerning other aspects of V⁵⁺ influence on fungal phosphate metabolism, the role of polyphosphates (PolyP) should not be neglected. It was demonstrated that PolyP play a central role in vanadate detoxification in Hansenula polymorpha [16]. Also, V⁵⁺ addition considerably increases PolyP level in vanadate resistant yeasts by inhibiting the enzyme exopolyphosphatase [17]. Nevertheless, the data about vanadate influence on (phosphate) metabolism of fungi are very scarce and were obtained almost solely on two ascomycetous yeasts species: Sacharomyces cerevisiae and H. polymorpha [18], and especially little is known about effects of vanadate on phosphate metabolism of filamentous fungi. It should be noted that due to diversity of fungal species, the results obtained on one fungus (especially yeast), cannot be simply extrapolated to other fungal species.

The action of vanadium on metabolism of zygomycetous fungus P. blakesleeanus, has not been examined so far. It is known, however, that the metabolism of sugar phosphates (SP) in P. blakesleeanus occurs mainly through glycolysis [19], [20] and is controlled at the level of phosphofructokinase (PFK) [21]. It was demonstrated that the cessation of spores dormancy induces strong transient burst of fructose 2,6 bisphosphate (F2,6P) and hexose 6 phosphates [22]. In addition, almost all components of carbohydrate metabolism in the mycelium of P. blakesleeanus appear to be influenced by light growth conditions [23]. Regarding other phosphate compounds, the importance of PolyP in energy storage of P. blakesleeanus should be noted [24]. It was shown that its content in this fungus is dependent on stage and cultivation conditions [25].

The mechanisms of vanadium reduction, entrance into P. blakesleeanus in both (4+ and 5+) oxidation states, and toxicity were proposed in our previous paper [26]. We suggested that V⁵⁺ is physiologically active form, provoking the changes in content of phosphate compounds in mycelium. In this paper, ³¹P NMR spectroscopy and HPLC analysis were used for the investigation of vanadium influence on phosphate metabolism of P. blakesleeanus.

Materials and Methods

Mycelium cultivation and materials

The wild-type strain of the fungus P. blakesleeanus (Burgeff) (NRRL 1555(−)) was used. The mycelium was grown in standard minimal medium, with spore concentration of 10⁶/ml [27], in Erlenmeyer flasks which were shaken and aerated in the growth cabinet with continuous overhead white fluorescent light of 10 W/m², at temperature of 20°C, and ca. 95% relative humidity. Stock solution of 200 mM sodium orthovanadate (Na₃VO₄) was prepared by the method of Gordon [28]. All chemicals were of analytical grade and obtained from Sigma-Aldrich (Taufkirchen, Germany).

³¹P NMR.

For the purpose of NMR measurements, 24 h old mycelium was collected by vacuum filtration, and washed with modified minimal medium without phosphates and microelements (experimental medium). The amount of 0.6 g of fresh weight (FW) mycelia was suspended in 2 ml of aerated experimental medium, and packed in a 10 mm diameter NMR tubes. Sodium orthovanadate (V⁵⁺) was added at the final concentration of 80 µmol/g_FW. For the concentration-dependent investigation final amounts of added V⁺⁵ were 20, 40, 70 and 80 µmol/g_FW; glucose 1 phosphate (G1P), glucose 6 phosphate (G6P), and fructose 6 phosphate (F6P) were added at final concentrations of 24 µmol/g, and 8-Br-cAMP at 80 µmol/g_FW. The measurements were performed using Apollo upgrade, Bruker MSL 400 spectrometer operating at 161.978 MHz for ³¹P. Spectra were accumulated with 14 µs pulse duration (about 45°) and 300 ms recycle time. The assignment of NMR spectra and spectral line intensities evaluation were performed as described previously [25]. For ³¹P NMR analysis of mycelial extracts, the extracts prepared for HPLC experiments were used (without EDTA). SP content was estimated by NTNMR (Tecmag) software, using methylene diphosphonic acid (MDP) signal as an external standard.

HPLC experiments

For the purpose of High Pressure Liquid Chromatography (HPLC) experiments, 24 h old mycelium was treated with V⁵⁺ or vanadyl sulphate (V⁴⁺) at final concentration of 80 µmol/g_FW for 10 minutes and then washed with deionised water. Control and treated mycelia were suspended in 0.5 M perchloric acid (1∶5 w/v), and homogenized in mortar on ice for 15 min. Obtained homogenate was stirred for 15 min on ice and centrifuged at 10000×g for 12 min. The precipitate was discarded and supernatant was treated with 2M KOH until pH was 7. The aliquots were kept on −20°C and unfrozen just before experiments.

HPLC analyses were performed on a Waters Breeze chromatographic system (Waters, Milford, MA) connected to Waters 2465 electrochemical detector with 3 mm gold working electrode and hydrogen referent electrode. Separation of sugars was performed on CarboPac PA1 (Dionex, Sunnyvale, CA) 250×4 mm column equipped with corresponding CarboPac PA1 guard column at constant temperature of 30°C and at a flow rate of 1.0 ml•min⁻¹. Sugars were eluted using 100 mM sodium hydroxide (phase A) and 500 mM sodium acetate in 100 mM sodium hydroxide (phase B) using following gradient method: linear rise in the first 20 minutes from initial 20 to 40% B; during next 20 minutes fast rise up to 95% of phase B, followed by 15 minute reverse to 20% B with additional 5 minutes for column equilibration. Signals were detected in pulsed amperometric mode (PAD) within 150 ms of integration time using following waveform: E1 = +0.1 V for 400 ms; E2 = +0.75 V for 200 ms; E3 = −0.8 V for 180 ms. Filter timescale was 0.2 s and range was set to 500 nA for the full mV scale.

For the preparation of 100 mM NaOH solution, 5.25 ml of sodium hydroxide solution (50% w/w, low carbonate, J.T. Baker, Deventer, Holland) was diluted to final volume of 1 l with vacuum degassed deionized water. All solutions were prepared in previously vacuum-degassed deionized water. Each component was analyzed quantitatively by the external standard method using pure standards (Sigma Co. St. Louis, MO) as references for concentration and retention time. Data acquisition and quantification were carried out by Empower 2 software (Waters, Milford, MA, USA).

Statistics

The samples were compared statistically using t-test at the 5% level of significance (P<0.05), which were performed with the Sigma Plot program, version 12.3. Data are presented as means ± s.e.m. with n indicating the number of independent experiments.

Results

The ³¹P NMR spectroscopy was used to monitor the action of vanadium on the level of phosphate compounds in fungus P. blakesleeanus. All experiments were performed on 24 h old mycelium i.e., on mycelium in exponential phase of growth, as the effect of vanadate is most pronounced in this phase of development [29]. ³¹P NMR spectra of 24 h old mycelium before, and after addition of 80 µmol/g_FW V⁵⁺ are shown in Figure 1A. Addition of V⁵⁺ induced considerable changes in the intensity of almost all signals in the spectrum (Figure 1A, B), with most pronounced changes in part of the spectrum assigned to the sugar phosphates (2–5 ppm). The signals from G1P, G6P, F6P, fructose 1, 6 phosphate (F1,6P), and fructose 2, 6 phosphate (F2,6P) can be expected in this region [30]–[32]. The intensity of the broad signal centered at about 3.2 ppm increased 60%, indicating changes in SP metabolism after V⁵⁺ addition. Furthermore, rise in the intensity of signal at about 0.3 ppm, which can correspond to second phosphate of F2,6P [30] was noticed. The intensity of signal assigned to the core polyphosphate residues (PPc) increased by 40%, while the intensity of Pi signal decreased by 21.5%, i.e., V⁵⁺ caused a decrease of PPc/Pi signal intensity ratio by 67.8%. Intensity decrease (41%) of the signal originating from the phosphorus atoms of UDPG and NADP(H), together with drastic drop (71%) of signal intensity of second UDPG phosphorus indicates prominent decrease of UDPG level. Vanadate addition also affected level of ATP in the mycelium. Namely, the area of γ-ATP signal, originating from both, ATP and ADP, has increased by 43±4% (n = 7) while that of β-ATP, originating from ATP only, was reduced by 21±7% (n = 7), indicating increase of ATP consumption.

Download:

Figure 1. Influence of vanadate on ³¹P NMR spectrum of P. blakeskeeanus mycelium.

A. Spectra of 24(control), and after addition of 80 µmol/g_FW V⁵⁺. The signals were assigned to the following compounds [25]: 1, sugar phosphate (SP); 2, inorganic phosphates (Pi); 3, γ-ATP; 4, terminal residues of PolyP and pyrophosphate; 5, α-ATP; 6, NADP(H) and UDPG; 7, UDPG (second resonance); 8, β-ATP; 9, penultimate phosphates in PolyP; 10, core PolyP residues (PPc). B. Signal intensities of control and 80 µmol/g_FW V⁵⁺ treated mycelium. Statistically significant differences are marked with asterix (n = 7, P<0.05). N/U designates NADP(H) and UDPG signal. C. Effect of different V⁺⁵ concentrations on ³¹P NMR spectrum. Insert: concentration dependency of SP signal intensity.

https://doi.org/10.1371/journal.pone.0102849.g001

Having in mind that V⁵⁺ uptake occurs only when the amount of V⁵⁺ exceeds the reduction capacity of an enzyme with ferricyanide reductase activity [26], we examined the concentration of externally added V⁵⁺ required to cause the intensity increase in SP part of NMR spectrum of mycelium. Over 20 µmol/g_FW of externally added vanadate was needed to cause initial signal intensity increase, while the increasing concentrations of V⁵⁺ amplified this effect (Figure 1C).

To determine the phosphorylated sugar primarily responsible for observed intensity increase in SP part of ³¹P NMR spectrum, we performed HPLC and ³¹P NMR analysis of control and vanadate treated mycelial extracts. In ³¹P NMR spectra of mycelial extracts after V⁵⁺ addition, new signals, that could be assigned to G6P, F6P, F1,6P, G1P, and F2,6P (Table 1), appeared. SP contents of both, control and V⁵⁺ treated mycelial extracts were estimated by integration of the region from −2 to +6 ppm in ³¹P NMR spectra (SP and Pi signals area), and it was observed that this area almost doubled after V⁵⁺ addition.

Download:

Table 1. Sugar phopsphates content in perchloric acid extracts of P. blakesleeanus mycelium.

https://doi.org/10.1371/journal.pone.0102849.t001

According to HPLC data (Figure 2, Table 1), V⁵⁺ caused the increase of G6P content by 2.3 times. Also, considerable increase was observed in contents of F6P (65%), and slightly less in F1,6P and G1P level (32%). Chromatograms of mycelial extracts after V⁵⁺ treatment differed from controls by noteworthy increase in the intensity of peak with retention time of 31 min, which could represent F2,6P. Furthermore, increasing intensity of peaks with retention times of 17 and 26 min that could originate from glucosamine 6 phosphate (Gluam6P) and glucose 1,6 bisphosphate (G1,6P), respectively, was observed. Lack of standards for above mentioned compounds made usual HPLC data interpretation impossible, but selectivity of HPLC-PAD method narrows down the list of interfering compounds, and makes this assignation plausible [33].

Download:

Figure 2. Influence of vanadium on SP content in perchloric acid extract of P. blakeskeeanus mycelia.

A. HPLC chromatograms obtained from control and V⁵⁺ treated mycelia. B. Changes in content of SP caused by V⁵⁺, and V⁴⁺, and control. Statistically significant differences are marked with asterix (n = 7, P<0.05).

https://doi.org/10.1371/journal.pone.0102849.g002

We have already proposed that the addition of vanadyl (V⁴⁺) has no effect on SP metabolism of P. blakesleeanus [26]. This statement was tested here once more, by analyzing mycelial extracts after V⁴⁺ treatment by HPLC (Figure 2B). In line with our previous results, almost no significant differences in chromatograms of V⁴⁺ treated and control mycelia were noticed.

HPLC and NMR experiments performed with mycelial extracts indicate that the main effect of V⁵⁺ on SP content is the increase of G6P level. But, results of ³¹P NMR with fresh mycelium do not support such conclusion, since chemical shift of G6P (about 4.5 ppm) [32] is far from position of the signal most affected by V⁵⁺ (Figure 1C). Therefore, we examined the influence of SP addition to control and V⁵⁺ treated mycelium, to spike position of SP signals in vivo. After addition of G6P and F6P to V⁵⁺ treated mycelia, new signals appeared at 5.1 and 4.8 ppm, respectively, while the addition of G1P increased the intensity of the signal at 3.2 ppm (Figure 3A), indicating that the signal whose intensity increases mostly after V⁵⁺ addition belongs to G1P. Addition of the same amount of the aforementioned SP to control mycelium increased only the intensity of Pi signal. All signals originating from externally added SP disappeared after mycelium washing (results not shown) that strongly suggests that SP do not enter the cells treated with V⁵⁺.

Download:

Figure 3. ³¹P NMR spectra of P. blakeskeeanus mycelium after addition of: A. 24

µmol/g_FW of G6P, F6P and G1P into mycelium treated with 80 µmol/g_FW V⁵⁺; B. 80 µmol/g_FW cAMP, 80 µmol/g_FW V⁵⁺, and 80 µmol/g_FW cAMP+80 µmol/g_FW V⁵⁺. Positions of added compounds are labeled with arrows.

https://doi.org/10.1371/journal.pone.0102849.g003

Data presented so far point to the influence of V⁵⁺on the levels of SP involved in the metabolic reactions of glycolytic and glycogenolytic pathways. As cAMP shows considerable influence on activity of glycogen synthase, glycogen phosphorylases, and bifunctional enzyme phosphofructokinase 2/fructose 2,6 bisphosphatase in many eukaryotic organisms [34], we tested mutual action of cAMP and V⁵⁺. Thus, 80 µmol/g_FW cAMP was added before and after supplementation of mycelium with V⁵⁺. In both cases, addition of cAMP led only to the appearance of the signal at −0.8 ppm, that can be attributed to cAMP itself, meaning that cAMP has no effects on V⁵⁺ metabolic action on mycelium of P. blakesleeanus (Figure 3B).

Discussion

Limited literature data on vanadium metabolism in fungi are obtained almost solely on two ascomycetous yeasts species: S. cereviesiae and H. polymorpha [18]. There is almost no data about vanadium metabolism in filamentous fungi, which make up a vast majority of fungal species, except for two studies on Neurospora crassa concerning vanadate uptake [35], [36], and our previous study on P. blakesleeanus, dealing mostly with V⁵⁺ reduction [26]. Influence of vanadium on fungal phosphate metabolism is even more difficult to interpret, as all available data were obtained on yeast [14]–[18]. Therefore, in this study we demonstrated the effects of V⁵⁺ on metabolism of phosphate compounds in filamentous fungus P. blakesleeanus. The action of V⁵⁺ was monitored by ³¹P NMR spectroscopy, method which enables detection of changes in phosphate metabolite levels in vivo, thus providing an insight to V⁵⁺ action. Results presented here show that V⁵⁺ induces changes in SP, PPc, Pi, ATP, and UDPG levels. HPLC analysis confirmed increased concentrations of G6P, F6P, G1P and F1,6P (in this order), in 24 h old mycelium of P. blakesleeanus.

Glucose 6 phosphate is the first product of glucose metabolism and it is further metabolized in different ways. For P. blakesleeanus mycelium it was shown that major part of glucose metabolism occurs via glycolysis [19]. However, considerable part of G6P in 24 h old mycelium (exponential phase) can also be involved in glycogen synthesis [37]. Namely, depending on the cultivation conditions, glycogen content can be from less than 1% to 10% of the dry mass of mycelia, reaching maximum values during the exponential growth phase [37]. The concentration of G1P after V⁵⁺addition increased by 36% while that of G6P by 2.3 times, according to HPLC experiments. This suggests that V⁵⁺ causes inhibition of the enzyme phosphoglucomutase, which participates in the first reaction of glycogen metabolism, i.e., conversion of G6P to G1P and vice versa, thereby preventing incorporation of G6P into glycogen. This is in accordance with the previously determined inhibitory action of vanadate on this enzyme obtained in rabbit skeletal muscle [38], [39]. Also, presumed increase in level of G1,6P is in accordance with vanadate effect on phosphoglucomutase and glucose 1,6 bisphosphate synthase in rabbit muscle [40]. The increase of Gluam6P concentration, indicated by HPLC, could be an additional proof, as the enzyme from the same group as phosphoglucomutase is responsible for the transformation of Gluam6P amino sugar to glucose-amino 1 phosphate in bacterial cells [41], and in S. cerevisiae [42]. All accumulated G1P could arise from glycogen degradation. Reversible process is also possible but, as can be seen from the ³¹P NMR results, the level of UDPG, main intermediate metabolite in glycogen synthesis, decreased after addition of vanadate.

Considering the stimulative effects of vanadate on cAMP accumulation, shown in rat adypocytes [43] and isolated rat liver cells [44], and consequently its influence on glycogen metabolism via glycogen phosphorylase and glycogen synthase [34], we tested the effect of cAMP on vanadate action. The accumulation of cAMP was not V⁵⁺-dependent, and cAMP had no effect on vanadate metabolic action in mycelium of P. blakesleeanus. These results are in accordance with those obtained in aquatic fungus B. emersonii [45]. Also, cAMP did not cause any changes in the SP region of the ³¹P NMR spectra, which were expected due to its stimulative effect on glycogen degradation determined in sporangiophores of P. blakesleeanus [46].

The main route of G6P transformation (glycolysis) proceeds via formation of F6P and F1,6P in reactions driven by phosphoglucoisomerase (PGI), and PFK1, respectively, the second one being irreversible. Vanadate addition caused the increase of F6P level by 65% and F1,6P by 32%. This may indicate inhibitory effect of vanadate on PFK1 activity as concentration ratio of G6P:F6P:F1,6P in control was 1∶0.5∶1.2 while, after V⁵⁺ treatment 1∶0.4∶0.7. However, this part of glycolysis in P. blakesleeanus can also include formation of F2,6P (catalyzed by PFK2), which is not a direct participant of this process [22]. Increased level of F2,6P after V⁵⁺addition, indicated by HPLC and ³¹P NMR data, is in agreement with formation of F2,6P established in chicken liver [7] and rat adypocytes [9], where vanadate inhibited fructose 2,6 bisphosphatase (F2,6P₂), an enzyme which catalyses F6P synthesis through dephosphorilation of F2,6P [47]. Formation of F2,6P may also be a consequence of increased participation of glucose in glycolytic pathway caused by V⁵⁺ and would be in accord with previous data obtained on P. blakesleeanus spores [21] and S. cerevisiae [48], where increase in glucose concentration was directly connected with the higher content of F2,6P. It should be mentioned that glucose is present in experimental medium, giving permanent glucose input in ³¹P NMR experiments in vivo. Accumulation of F2,6P could be a trigger for the increase of F1,6P concentration. Namely, micromolar concentrations of F2,6P activate PFK1 in isolated rabbit muscle [47], S. cerevisiae [49] and P. blakesleeanus spores [21]. The noteworthy rise of F1,6P concentration, after V⁵⁺ addition, is in agreement with the data obtained on erythrocytes [8].

The third known way of G6P transformation is through pentose phosphate pathway, with glucose-6-P dehydrogenase (G6PD) as the first enzyme involved. Inhibition of GDPH by vanadate dimer and tetramer was demonstrated in Leuconostoc mesenteroides [50]. However, vanadate monomer does not show such effect in isolated baker yeast [51]. It strongly activates oxidation of glucose by G6PD, with no effects on the rates of G6P oxidation, and according to the authors, V⁵⁺ interacts with the 6-hydroxyl group of glucose rather than with the enzyme directly [52], [53]. The possibility of direct V⁵⁺ interaction with glucose, or replacement of phosphate group in G6P resulting in formation of glucose 6 vanadate (G6V), is unlikely here because significant increase of G6P level was observed after V⁵⁺ treatment (Table 1). Replacement of phosphate group with vanadate in NADP (cofactor in G6PD reaction) was determined in isolated baker yeast cells [51], but the level of NADP in ³¹P NMR spectra of P. blakesleeanus remained almost unchanged after V⁵⁺ treatment (Figure 1), indicating that such reaction is not plausible. Nevertheless, above mentioned modes of V⁵⁺ action cannot be ignored.

Some attention should be paid to the results of spiking of SP ³¹P NMR signals, which appear after V⁵⁺ treatment of P. blakesleeanus mycelium. Namely, in vivo results differ to some extent from those obtained on mycelial extracts, and suggest that the level of G1P instead of G6P is mainly influenced by V⁵⁺ addition. That can be explained by the fact that in vivo system is dynamic and addition of substrates/products can influence direction of glycolytic and glycogenolytic reactions, while cell extracts give insight in SP levels for a single metabolic state of the cells [54].

As it was already said, washing of mycelium after subsequent V⁵⁺ and SP addition, resulted in disappearance of ³¹P NMR signals attributed to added SP. Fungi are known to be absorptive heterotrophs, meaning that SP should be previously digested/hydrolyzed, and then ingested as inorganic Pi and sugar. Presence of acid phosphatase, enzyme(s) that hydrolyze phosphate group, was confirmed in yeast and mycorrhizal fungi, when inorganic phosphate was unavailable in its simplest form [55]. Based on database search, there are two protein sequences in P. blakeesleanus protein database which show homology with acid phosphatase protein isolated from Penicillium chrysogenum. Predicted gene models for these proteins are located in P. blakeesleanus genome at the scaffold_8∶1745265-1746626 and scaffold_2∶2647390-2648609 [56]. These sequences also show high identity (40%) with acid phosphatase from Laccaria bicolor S238N-H82 [57], and putative acid phosphatase from Aspergillus fumigatus Af293 (43%) [58]. Absence of additional signals in SP part of ³¹P NMR spectra of control and their appearance in V⁵⁺ treated mycelium after SP addition, strongly suggests that V⁵⁺ influences the activity of acid phosphatases in P. blakesesleeanus.

³¹P NMR results presented here also show that V⁵⁺ causes considerable increase of PPc signal intensity (PolyP level). Similar effects of V⁵⁺ were described in amoeba Dyctyostelium discoideum [59] and H. polymorpha [18]. Inhibition of exopolyphosphatase by V⁵⁺ was confirmed in yeast [17], [60], and could be responsible for increase of PolyP level observed here. Predicted gene model for exopolyphosphatase in P. blakesleeanus genome is located at scaffold_6: 2343311-2344745. Protein sequence of this enzyme shows the highest identity level (42%) with Mucor circinelloides hypothetical protein with activity of inorganic pyrophosphatase/exopolyphosphatase (GenBank accession number: EPB83848) [52]. Using this protein sequence as query and BLAST [61] for alignment with sequence of purified exopolyphosphatase encoded by PPX1 gene from S. cerevisiae [62], [63], possible homology of these two proteins was obtained (95% of query coverage, 30% of identity and E-value = 10⁻⁴⁰). Since only a part of added/intracellular V⁵⁺ is reduced to V⁴⁺ [26] we propose that remaining V⁵⁺ induces increase of PolyP level due to inhibition of exopolyphosphatase(s) in P. blakesleeanus.

To the best of our knowledge, these are the first data about vanadate action on any aspect of metabolism in filamentous fungi. The action of V⁵⁺ on SP content and phosphate metabolism in general was studied mainly in vitro i.e., on isolated tissues of various types of organisms, or by measuring the enzyme activity. Here, based on increased intensities of SP ³¹P NMR signals in vivo and HPLC experiments, we presumed the influence of V⁵⁺ on enzymes involved in glycolysis and glucogenesis. Furthermore, results of ³¹P NMR indicate V⁵⁺ induced changes in actions of enzymes acid phosphatase and exopolyphosphatase whose presence is yet to be confirmed in P. blakesleanus.

Author Contributions

Conceived and designed the experiments: M. Žižić M. Živić TCA JZ. Performed the experiments: M. Žižić JZ VM SK. Analyzed the data: M. Žižić JZ VM MS. Contributed to the writing of the manuscript: M. Žižić JZ MS VM M. Živić SK.

References

1. Rehder D (1992) Structure and function of vanadium compounds in living organisms. BioMetals 5: 3–12.
- View Article
- Google Scholar
2. Tsiani E, Fantus IG (1997) Vanadium compounds, biological actions and potential as pharmacological agents. TEM 8: 52–58.
- View Article
- Google Scholar
3. Rehder D (2008) Bioinorganic vanadium chemistry. J. Wiley and Sons, Chichester, New York.
4. Chasteen ND (1990) Vanadium in biological system. Kluwer Academic Publishers, Dordrecht, Netherlands.
5. Crans D, Smee JJ, Jason JS, Gaidamauskas E, Yang L (2004) The chemistry and biochemistry of vanadium and the biological activities exerted by vanadium compounds. Chem Rev 104: 849–902.
- View Article
- Google Scholar
6. Stankiewicz PJ, Tracey AS, Crans DC (1995) Inhibition of phosphate-metabolizingenzymes by oxovanadiumcomplexes. Met Ions Biol Syst 31: 287–324.
- View Article
- Google Scholar
7. Rider MH, Bartrons R, Hue L (1990) Vanadate inhibits liver fructose-2,6-bisphosphatase. Eur J Biochem 190: 53–56.
- View Article
- Google Scholar
8. Benabe J, Echegoyen LA, Pastranall B, Martinez-Maldonado M (1987) Mechanism of inhibition of glycolysis by vanadate. Journal of Biological Chemistry 262: 9555–9560.
- View Article
- Google Scholar
9. Gomez-Foix AM, Rodriguez-Gil JE, Fillat C, Guinovart JJ, Bosch F (1988) Vanadate raises fructose 2,6-bisphosphate concentrations and activates glycolysis in rat hepatocytes. Biochem J 255: 507–512.
- View Article
- Google Scholar
10. Rehder D (2003) Biological and medical aspects of vanadium. Inorg Chem Commun 6: 604–617.
- View Article
- Google Scholar
11. Kalac P, Svoboda L (2000) A review of trace element concentrations in edible mushrooms. Food Chem 69: 273–281.
- View Article
- Google Scholar
12. Han C, Cui B, Qu J (2009) Comparison of vanadium-rich activity of three species fungi of basidiomycetes. Biol Trace Elem Res 127: 278–283.
- View Article
- Google Scholar
13. Berry RE, Armstrong EM, Beddoes RL, Collison D, Ertok SN, et al. (1999) The structural characterization of amavadin. Angew Chem Int Ed 38: 795–797.
- View Article
- Google Scholar
14. Cohen MD, Sen AC, Wei C-I (1987) Vanadium inhibition of yeast glucose 6 dehydrogenase. Inorg Chim Acta 138: 179–186.
- View Article
- Google Scholar
15. Crans D, Willging EM, Butler SR (1990) Vanadate tetramer as the inhibiting species in enzyme reactions in vitro and in vivo. J Am Chem Soc 112: 427–432.
- View Article
- Google Scholar
16. Mannazzu I, Guerra I, Strabbioli R, Masia A, Maestrale GB, et al. (1997) Vanadium affects vacuolization and phosphate metabolism in Hansenula polymorpha. FEMS Microbiol 147: 23–28.
- View Article
- Google Scholar
17. Lorenz B, Muller WE, Kulaev IS, Schroder HC (1994) Purification and characterization of an exopolyphosphatase from Saccharomyces cerevisiae. J Biol Chem 269: 198–204.
- View Article
- Google Scholar
18. Mannazzu I (2001) Vanadium detoxification and resistance in yeast: a minireview. Ann Microbiol 51: 1–9.
- View Article
- Google Scholar
19. Rudolph H, Furch B (1970) Untersuchungen zur Aktivirung von sporenhomogenaten durch Warmebehandlung. Arch Mikrobiol 72: 175–181.
- View Article
- Google Scholar
20. Rua J, De Cima S, Del Valle P, Gutierrez-Larranzar M, Busto F, et al. (2008) Glycogen and trehalose mobilization by acetic acid in Phycomyces blakesleeanus: dependence on the anion form. Res Microbiol 159: 200–206.
- View Article
- Google Scholar
21. Van Laere A (1983) Stimulation of phosphofructokinase from Phycomyces blakesleeunus and some other fungi by micromolar concentrations of fructose 2,6-bisphosphate. J Gen Microbiol 129: 3281–3285.
- View Article
- Google Scholar
22. Van Laere A, Van Schaftingeen E, Hers H-G (1983) Fructose 2,6-bisphosphate and germination of fungal spores. Proc Natl Acad Sci USA 80: 6601–6605.
- View Article
- Google Scholar
23. Diez M, Rua J, Busto F, Soler J (1988) Analysis of the effects of light on carbohydrate metabolism in Phycomyces sporangiophores and mycelia. Int J Biochem 20: 801–806.
- View Article
- Google Scholar
24. Stanić M, Zakrzevska J, Hadžibrahimović M, Žižić M, Marković Z, et al. (2013) Oxygen regulation of alternative respiration in fungus Phycomyces blakesleeanus: connection with phosphate metabolism. Res Microbiol 164: 770–778.
- View Article
- Google Scholar
25. Živić M, Zakrzewska J, Žižić M, Bačić G (2007) 31P NMR study of polyposphate levels during different growth phases of Phycomyces blakesleeanus. Antonie van Leeuwenhoek 91: 169–177.
- View Article
- Google Scholar
26. Žižić M, Živić M, Spasojević I, Bogdanović Pristov J, Stanić M, et al. (2013) The interactions of vanadium with Phycomyces blakesleeanus mycelium: enzymatic reduction, transport and metabolic effects. Res Microbiol 164: 61–69.
- View Article
- Google Scholar
27. Sutter RP (1975) Mutations affecting sexual development in Phycomyces blakesleeanus.. Proc Natl Acad Sci USA 72: 127–130.
- View Article
- Google Scholar
28. Gordon J (2001) Use of vanadate as protein-phosphotyrosine phosphatase inhibitor. Methods Enzymol 201: 477–482.
- View Article
- Google Scholar
29. Žižić M (2013) Metabolism and metabolic effects of vanadium in fungus Phycomyces blakesleeanus. PhD Thesis. University of Belgrade.
30. Okar DA, Kakalis LT, Narula SS, Armitage IM, Pilkis SJ (1995) Identification of transient intermediates in the bisphosphatase reaction of rat liver 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase by ³¹P NMR spectroscopy Biochem J. 308: 189–195.
- View Article
- Google Scholar
31. Evans FE, Kaplan NO (1977) ³¹P nuclear magnetic resonance studies of HeLa cells. Proc Natl Acad Sci USA 74: 4909–4913.
- View Article
- Google Scholar
32. Barany M, Glonek T (1984) Identification of diseased states by Phosphorus-31 NMR. In David D. Gorenstein, editor. Phopsphorus-31 NMR. Principles and applications. Academic Press. 511–545.
33. Dionex website: http://.dionex.com/en-us/webdocs/5047-AN122_LPN1029-03.pdf. Accessed August 2010.
34. Voet D, Voet J (1995) Biochemistry. J. Wiley and Sons, Chichester. 497–507.
35. Bowman BJ (1983) Vanadate uptake in Neurospora crassa occurs via phosphate transport system. J Bacteriol 153: 286–291.
- View Article
- Google Scholar
36. Bowman BJ, Allen KE, Slayman CW (1983) Vanadate-resistant mutants of Neurospora crassa are deficient in a high-affinity phosphate transport system. J Bacteriol 153: 292–296.
- View Article
- Google Scholar
37. Rua J, Busto F, De Arriaga D, Soler J (1993) Glycogen in Phycomyces blakesleeanus: influence of growth conditions and nutrient limit action. J. Gen. Microbiol 139: 217–222.
- View Article
- Google Scholar
38. Climent F, Bartrons R, Pons G, Carreras J (1981) Effect of vanadate on phosphoryl transfer enzymes involved in glucose metabolism. Biochem Bioph Res Co 101: 570–576.
- View Article
- Google Scholar
39. Carreras J, Climent F, Bartrons R, Pons G (1982) Effect of vanadate on the formation and stability of the phosphoenzyme forms of 2,3-biphosphoglycerate-dependent phosphoglycerate mutase and of phosphoglucomutase. Biochim Biophys Acta 705: 238–242.
- View Article
- Google Scholar
40. Carreras M, Bassols AM, Carreras J, Climent F (1988) Effect of vanadate on the glucose-l,6-bisphosphate synthase and glucose-l,6-bisphosphatase activities of phosphoglucomutase. Arch Biochem Biophys 264: 155–159.
- View Article
- Google Scholar
41. Jolly L, Pompeo F, Van Heijenoort J, Fassy F, Mengin-Lecreulx D (2000) Autophosphorylation of phosphoglucosamine mutase from Escherichia coli. J Bacteriol 182: 1280–1285.
- View Article
- Google Scholar
42. Milewski S, Gabriel I, Olchowy J (2006) Enzymes of UDP-GlcNAc biosynthesis in yeast. Yeast 23: 1–14.
- View Article
- Google Scholar
43. Schwabe U, Puchstein C, Hannemann H, Söchtig E (1979) Activation of adenylate cyclase by vanadate. Nature 277: 143–145.
- View Article
- Google Scholar
44. Catalan RE, Martinez AM, Aragones MD (1980) Effects of vanadate on the cyclic AMP-protein kinase system in rat liver. Biochem Bioph Res Co 96: 672–677.
- View Article
- Google Scholar
45. Vandercammen A, Francois MJ, Torres BB, Maia JCC, Hers H-G (1990) Fructose 2,6 bisphosphate and carbohydrate metabolism during the life cycle of the aquatic fungus Blastocladiella emersonii. J Gen Microbiol 136: 137–146.
- View Article
- Google Scholar
46. Tu JC, Malhotra SK (1977) Plasma membrane of Phycomyces: sequential changes in the structure during spore formation. Cytobios 20: 121–132.
- View Article
- Google Scholar
47. Van Schaftingeen E, Hers HG (1981) Inhibition of fructose-1,6-bisphosphatase by fructose 2,6-bisphosphate. Proc Natl Acad Sci USA 78: 2861–2863.
- View Article
- Google Scholar
48. Francois J, Van Schaftingen E, Hers HG (1984) The mechanism by which glucose increases fructose 2,6 bisphosphate concentration in Saccharomyces cereviseae. A cyclic AMP-dependent activation of phosphofructokinase 2. Eur J Biochem 164: 369–373.
- View Article
- Google Scholar
49. Bartrons R, Van Schaftingen E, Vissers S, Hers HG (1982) The stimulation of yeast phosphofructokinase by fructose-2,6-bisphosphate. FEBS Lett 143: 137–140.
- View Article
- Google Scholar
50. Crans CD, Schelble MS (1990) Vanadate dimer and tetramer both inhibit glucose-6-phosphate dehydrogenase from Leuconostoc mesenteroides. Biochemistry 29: 6698–6706.
- View Article
- Google Scholar
51. Crans CD, Simone MC, Blanchards SJ (1992) Chemically induced modification of cofactor specificity of glucose-6-phosphate dehydrogenase. J Am Chem Soc 114: 4926–4928.
- View Article
- Google Scholar
52. Nour-Eldeen FA, Craig MM, Gresser JM (1985) Interaction of inorganic vanadate with glucose-6-phosphate dehydrogenase. J Biol Chem 260: 6836–6842.
- View Article
- Google Scholar
53. Dreuckhammer GD, Durrwachter RJ, Pederson LR, Crans CD, Daniels L, et al. (1999) Reversible and in situ formation of organic arsenates and vanadates as organic phosphate mimics in enzymatic reactions: Mechanistic investigation of aldol reactions and synthetic applications. J Org Chem 54: 70–77.
- View Article
- Google Scholar
54. Shanks JV, Bailey JE (1988) Estimation of intracellular sugar phosphate concentrations in Saccharomyces cerevisiae using ³¹P nuclear magnetic resonance spectroscopy. Biotechnol Bioeng 32: 1138–1152.
- View Article
- Google Scholar
55. Aleksieva P, Spasova D, Radoevska S (2003) Acid phosphatase distribution and localization in the fungus Humicola lutea. Z. Naturforsch. 58c: 239–243.
- View Article
- Google Scholar
56. Grigoriev IV, Nordberg H, Shabalov I, Aerts A, Cantor M, et al. (2011) The Genome Portal of the Department of Energy Joint Genome Institute. Nucleic Acids Res doi:10.1093/nar/gkr947.
57. Martin F (2008) The genome of Laccaria bicolor provides insights into mycorrhizal symbiosis. Nature 452(7183): 88–92
- View Article
- Google Scholar
58. Neirman WC (2005) Genomic sequence of the pathogenic and allergenic filamentous fungus Aspergillus fumigatus. Nature 438: 1151–6.
- View Article
- Google Scholar
59. Klein G, Cotter DA, Martin J-B, Satre M (1989) Vanadate, an inhibitor of growth, development and endocytosis in Dictyostelium discoideum amoebae. J Cell Sci 94: 127–134.
- View Article
- Google Scholar
60. Guranowski A, Starzyńska E, Barnes LD, Robinson AK, Liu S (1998) Adenosine 5′-tetraphosphate phosphohydrolase activity is an inherent property of soluble exopolyphosphatase from yeast Saccharomyces cerevisiae. Biochim Biophys Acta 1380: 232–238.
- View Article
- Google Scholar
61. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215: 403–410.
- View Article
- Google Scholar
62. Wurst H, Kornberg A (1994) A soluble exopolyphosphatase of Saccharomyces cerevisiae. Purification and characterization. J Biol Chem 269: 10996–11001.
- View Article
- Google Scholar
63. Wurst H, Shiba T, Kornberg A (1995) The gene for a major exopolyphosphatase of Saccharomyces cerevisiae. J Bacteriol 177: 898–906.
- View Article
- Google Scholar

[ref1] 1. Rehder D (1992) Structure and function of vanadium compounds in living organisms. BioMetals 5: 3–12.
View Article
Google Scholar

[2] View Article

[3] Google Scholar

[ref2] 2. Tsiani E, Fantus IG (1997) Vanadium compounds, biological actions and potential as pharmacological agents. TEM 8: 52–58.
View Article
Google Scholar

[5] View Article

[6] Google Scholar

[ref3] 3. Rehder D (2008) Bioinorganic vanadium chemistry. J. Wiley and Sons, Chichester, New York.

[ref4] 4. Chasteen ND (1990) Vanadium in biological system. Kluwer Academic Publishers, Dordrecht, Netherlands.

[ref5] 5. Crans D, Smee JJ, Jason JS, Gaidamauskas E, Yang L (2004) The chemistry and biochemistry of vanadium and the biological activities exerted by vanadium compounds. Chem Rev 104: 849–902.
View Article
Google Scholar

[10] View Article

[11] Google Scholar

[ref6] 6. Stankiewicz PJ, Tracey AS, Crans DC (1995) Inhibition of phosphate-metabolizingenzymes by oxovanadiumcomplexes. Met Ions Biol Syst 31: 287–324.
View Article
Google Scholar

[13] View Article

[14] Google Scholar

[ref7] 7. Rider MH, Bartrons R, Hue L (1990) Vanadate inhibits liver fructose-2,6-bisphosphatase. Eur J Biochem 190: 53–56.
View Article
Google Scholar

[16] View Article

[17] Google Scholar

[ref8] 8. Benabe J, Echegoyen LA, Pastranall B, Martinez-Maldonado M (1987) Mechanism of inhibition of glycolysis by vanadate. Journal of Biological Chemistry 262: 9555–9560.
View Article
Google Scholar

[19] View Article

[20] Google Scholar

[ref9] 9. Gomez-Foix AM, Rodriguez-Gil JE, Fillat C, Guinovart JJ, Bosch F (1988) Vanadate raises fructose 2,6-bisphosphate concentrations and activates glycolysis in rat hepatocytes. Biochem J 255: 507–512.
View Article
Google Scholar

[22] View Article

[23] Google Scholar

[ref10] 10. Rehder D (2003) Biological and medical aspects of vanadium. Inorg Chem Commun 6: 604–617.
View Article
Google Scholar

[25] View Article

[26] Google Scholar

[ref11] 11. Kalac P, Svoboda L (2000) A review of trace element concentrations in edible mushrooms. Food Chem 69: 273–281.
View Article
Google Scholar

[28] View Article

[29] Google Scholar

[ref12] 12. Han C, Cui B, Qu J (2009) Comparison of vanadium-rich activity of three species fungi of basidiomycetes. Biol Trace Elem Res 127: 278–283.
View Article
Google Scholar

[31] View Article

[32] Google Scholar

[ref13] 13. Berry RE, Armstrong EM, Beddoes RL, Collison D, Ertok SN, et al. (1999) The structural characterization of amavadin. Angew Chem Int Ed 38: 795–797.
View Article
Google Scholar

[34] View Article

[35] Google Scholar

[ref14] 14. Cohen MD, Sen AC, Wei C-I (1987) Vanadium inhibition of yeast glucose 6 dehydrogenase. Inorg Chim Acta 138: 179–186.
View Article
Google Scholar

[37] View Article

[38] Google Scholar

[ref15] 15. Crans D, Willging EM, Butler SR (1990) Vanadate tetramer as the inhibiting species in enzyme reactions in vitro and in vivo. J Am Chem Soc 112: 427–432.
View Article
Google Scholar

[40] View Article

[41] Google Scholar

[ref16] 16. Mannazzu I, Guerra I, Strabbioli R, Masia A, Maestrale GB, et al. (1997) Vanadium affects vacuolization and phosphate metabolism in Hansenula polymorpha. FEMS Microbiol 147: 23–28.
View Article
Google Scholar

[43] View Article

[44] Google Scholar

[ref17] 17. Lorenz B, Muller WE, Kulaev IS, Schroder HC (1994) Purification and characterization of an exopolyphosphatase from Saccharomyces cerevisiae. J Biol Chem 269: 198–204.
View Article
Google Scholar

[46] View Article

[47] Google Scholar

[ref18] 18. Mannazzu I (2001) Vanadium detoxification and resistance in yeast: a minireview. Ann Microbiol 51: 1–9.
View Article
Google Scholar

[49] View Article

[50] Google Scholar

[ref19] 19. Rudolph H, Furch B (1970) Untersuchungen zur Aktivirung von sporenhomogenaten durch Warmebehandlung. Arch Mikrobiol 72: 175–181.
View Article
Google Scholar

[52] View Article

[53] Google Scholar

[ref20] 20. Rua J, De Cima S, Del Valle P, Gutierrez-Larranzar M, Busto F, et al. (2008) Glycogen and trehalose mobilization by acetic acid in Phycomyces blakesleeanus: dependence on the anion form. Res Microbiol 159: 200–206.
View Article
Google Scholar

[55] View Article

[56] Google Scholar

[ref21] 21. Van Laere A (1983) Stimulation of phosphofructokinase from Phycomyces blakesleeunus and some other fungi by micromolar concentrations of fructose 2,6-bisphosphate. J Gen Microbiol 129: 3281–3285.
View Article
Google Scholar

[58] View Article

[59] Google Scholar

[ref22] 22. Van Laere A, Van Schaftingeen E, Hers H-G (1983) Fructose 2,6-bisphosphate and germination of fungal spores. Proc Natl Acad Sci USA 80: 6601–6605.
View Article
Google Scholar

[61] View Article

[62] Google Scholar

[ref23] 23. Diez M, Rua J, Busto F, Soler J (1988) Analysis of the effects of light on carbohydrate metabolism in Phycomyces sporangiophores and mycelia. Int J Biochem 20: 801–806.
View Article
Google Scholar

[64] View Article

[65] Google Scholar

[ref24] 24. Stanić M, Zakrzevska J, Hadžibrahimović M, Žižić M, Marković Z, et al. (2013) Oxygen regulation of alternative respiration in fungus Phycomyces blakesleeanus: connection with phosphate metabolism. Res Microbiol 164: 770–778.
View Article
Google Scholar

[67] View Article

[68] Google Scholar

[ref25] 25. Živić M, Zakrzewska J, Žižić M, Bačić G (2007) 31P NMR study of polyposphate levels during different growth phases of Phycomyces blakesleeanus. Antonie van Leeuwenhoek 91: 169–177.
View Article
Google Scholar

[70] View Article

[71] Google Scholar

[ref26] 26. Žižić M, Živić M, Spasojević I, Bogdanović Pristov J, Stanić M, et al. (2013) The interactions of vanadium with Phycomyces blakesleeanus mycelium: enzymatic reduction, transport and metabolic effects. Res Microbiol 164: 61–69.
View Article
Google Scholar

[73] View Article

[74] Google Scholar

[ref27] 27. Sutter RP (1975) Mutations affecting sexual development in Phycomyces blakesleeanus.. Proc Natl Acad Sci USA 72: 127–130.
View Article
Google Scholar

[76] View Article

[77] Google Scholar

[ref28] 28. Gordon J (2001) Use of vanadate as protein-phosphotyrosine phosphatase inhibitor. Methods Enzymol 201: 477–482.
View Article
Google Scholar

[79] View Article

[80] Google Scholar

[ref29] 29. Žižić M (2013) Metabolism and metabolic effects of vanadium in fungus Phycomyces blakesleeanus. PhD Thesis. University of Belgrade.

[ref30] 30. Okar DA, Kakalis LT, Narula SS, Armitage IM, Pilkis SJ (1995) Identification of transient intermediates in the bisphosphatase reaction of rat liver 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase by ³¹P NMR spectroscopy Biochem J. 308: 189–195.
View Article
Google Scholar

[83] View Article

[84] Google Scholar

[ref31] 31. Evans FE, Kaplan NO (1977) ³¹P nuclear magnetic resonance studies of HeLa cells. Proc Natl Acad Sci USA 74: 4909–4913.
View Article
Google Scholar

[86] View Article

[87] Google Scholar

[ref32] 32. Barany M, Glonek T (1984) Identification of diseased states by Phosphorus-31 NMR. In David D. Gorenstein, editor. Phopsphorus-31 NMR. Principles and applications. Academic Press. 511–545.

[ref33] 33. Dionex website: http://.dionex.com/en-us/webdocs/5047-AN122_LPN1029-03.pdf. Accessed August 2010.

[ref34] 34. Voet D, Voet J (1995) Biochemistry. J. Wiley and Sons, Chichester. 497–507.

[ref35] 35. Bowman BJ (1983) Vanadate uptake in Neurospora crassa occurs via phosphate transport system. J Bacteriol 153: 286–291.
View Article
Google Scholar

[92] View Article

[93] Google Scholar

[ref36] 36. Bowman BJ, Allen KE, Slayman CW (1983) Vanadate-resistant mutants of Neurospora crassa are deficient in a high-affinity phosphate transport system. J Bacteriol 153: 292–296.
View Article
Google Scholar

[95] View Article

[96] Google Scholar

[ref37] 37. Rua J, Busto F, De Arriaga D, Soler J (1993) Glycogen in Phycomyces blakesleeanus: influence of growth conditions and nutrient limit action. J. Gen. Microbiol 139: 217–222.
View Article
Google Scholar

[98] View Article

[99] Google Scholar

[ref38] 38. Climent F, Bartrons R, Pons G, Carreras J (1981) Effect of vanadate on phosphoryl transfer enzymes involved in glucose metabolism. Biochem Bioph Res Co 101: 570–576.
View Article
Google Scholar

[101] View Article

[102] Google Scholar

[ref39] 39. Carreras J, Climent F, Bartrons R, Pons G (1982) Effect of vanadate on the formation and stability of the phosphoenzyme forms of 2,3-biphosphoglycerate-dependent phosphoglycerate mutase and of phosphoglucomutase. Biochim Biophys Acta 705: 238–242.
View Article
Google Scholar

[104] View Article

[105] Google Scholar

[ref40] 40. Carreras M, Bassols AM, Carreras J, Climent F (1988) Effect of vanadate on the glucose-l,6-bisphosphate synthase and glucose-l,6-bisphosphatase activities of phosphoglucomutase. Arch Biochem Biophys 264: 155–159.
View Article
Google Scholar

[107] View Article

[108] Google Scholar

[ref41] 41. Jolly L, Pompeo F, Van Heijenoort J, Fassy F, Mengin-Lecreulx D (2000) Autophosphorylation of phosphoglucosamine mutase from Escherichia coli. J Bacteriol 182: 1280–1285.
View Article
Google Scholar

[110] View Article

[111] Google Scholar

[ref42] 42. Milewski S, Gabriel I, Olchowy J (2006) Enzymes of UDP-GlcNAc biosynthesis in yeast. Yeast 23: 1–14.
View Article
Google Scholar

[113] View Article

[114] Google Scholar

[ref43] 43. Schwabe U, Puchstein C, Hannemann H, Söchtig E (1979) Activation of adenylate cyclase by vanadate. Nature 277: 143–145.
View Article
Google Scholar

[116] View Article

[117] Google Scholar

[ref44] 44. Catalan RE, Martinez AM, Aragones MD (1980) Effects of vanadate on the cyclic AMP-protein kinase system in rat liver. Biochem Bioph Res Co 96: 672–677.
View Article
Google Scholar

[119] View Article

[120] Google Scholar

[ref45] 45. Vandercammen A, Francois MJ, Torres BB, Maia JCC, Hers H-G (1990) Fructose 2,6 bisphosphate and carbohydrate metabolism during the life cycle of the aquatic fungus Blastocladiella emersonii. J Gen Microbiol 136: 137–146.
View Article
Google Scholar

[122] View Article

[123] Google Scholar

[ref46] 46. Tu JC, Malhotra SK (1977) Plasma membrane of Phycomyces: sequential changes in the structure during spore formation. Cytobios 20: 121–132.
View Article
Google Scholar

[125] View Article

[126] Google Scholar

[ref47] 47. Van Schaftingeen E, Hers HG (1981) Inhibition of fructose-1,6-bisphosphatase by fructose 2,6-bisphosphate. Proc Natl Acad Sci USA 78: 2861–2863.
View Article
Google Scholar

[128] View Article

[129] Google Scholar

[ref48] 48. Francois J, Van Schaftingen E, Hers HG (1984) The mechanism by which glucose increases fructose 2,6 bisphosphate concentration in Saccharomyces cereviseae. A cyclic AMP-dependent activation of phosphofructokinase 2. Eur J Biochem 164: 369–373.
View Article
Google Scholar

[131] View Article

[132] Google Scholar

[ref49] 49. Bartrons R, Van Schaftingen E, Vissers S, Hers HG (1982) The stimulation of yeast phosphofructokinase by fructose-2,6-bisphosphate. FEBS Lett 143: 137–140.
View Article
Google Scholar

[134] View Article

[135] Google Scholar

[ref50] 50. Crans CD, Schelble MS (1990) Vanadate dimer and tetramer both inhibit glucose-6-phosphate dehydrogenase from Leuconostoc mesenteroides. Biochemistry 29: 6698–6706.
View Article
Google Scholar

[137] View Article

[138] Google Scholar

[ref51] 51. Crans CD, Simone MC, Blanchards SJ (1992) Chemically induced modification of cofactor specificity of glucose-6-phosphate dehydrogenase. J Am Chem Soc 114: 4926–4928.
View Article
Google Scholar

[140] View Article

[141] Google Scholar

[ref52] 52. Nour-Eldeen FA, Craig MM, Gresser JM (1985) Interaction of inorganic vanadate with glucose-6-phosphate dehydrogenase. J Biol Chem 260: 6836–6842.
View Article
Google Scholar

[143] View Article

[144] Google Scholar

[ref53] 53. Dreuckhammer GD, Durrwachter RJ, Pederson LR, Crans CD, Daniels L, et al. (1999) Reversible and in situ formation of organic arsenates and vanadates as organic phosphate mimics in enzymatic reactions: Mechanistic investigation of aldol reactions and synthetic applications. J Org Chem 54: 70–77.
View Article
Google Scholar

[146] View Article

[147] Google Scholar

[ref54] 54. Shanks JV, Bailey JE (1988) Estimation of intracellular sugar phosphate concentrations in Saccharomyces cerevisiae using ³¹P nuclear magnetic resonance spectroscopy. Biotechnol Bioeng 32: 1138–1152.
View Article
Google Scholar

[149] View Article

[150] Google Scholar

[ref55] 55. Aleksieva P, Spasova D, Radoevska S (2003) Acid phosphatase distribution and localization in the fungus Humicola lutea. Z. Naturforsch. 58c: 239–243.
View Article
Google Scholar

[152] View Article

[153] Google Scholar

[ref56] 56. Grigoriev IV, Nordberg H, Shabalov I, Aerts A, Cantor M, et al. (2011) The Genome Portal of the Department of Energy Joint Genome Institute. Nucleic Acids Res doi:10.1093/nar/gkr947.

[ref57] 57. Martin F (2008) The genome of Laccaria bicolor provides insights into mycorrhizal symbiosis. Nature 452(7183): 88–92
View Article
Google Scholar

[156] View Article

[157] Google Scholar

[ref58] 58. Neirman WC (2005) Genomic sequence of the pathogenic and allergenic filamentous fungus Aspergillus fumigatus. Nature 438: 1151–6.
View Article
Google Scholar

[159] View Article

[160] Google Scholar

[ref59] 59. Klein G, Cotter DA, Martin J-B, Satre M (1989) Vanadate, an inhibitor of growth, development and endocytosis in Dictyostelium discoideum amoebae. J Cell Sci 94: 127–134.
View Article
Google Scholar

[162] View Article

[163] Google Scholar

[ref60] 60. Guranowski A, Starzyńska E, Barnes LD, Robinson AK, Liu S (1998) Adenosine 5′-tetraphosphate phosphohydrolase activity is an inherent property of soluble exopolyphosphatase from yeast Saccharomyces cerevisiae. Biochim Biophys Acta 1380: 232–238.
View Article
Google Scholar

[165] View Article

[166] Google Scholar

[ref61] 61. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215: 403–410.
View Article
Google Scholar

[168] View Article

[169] Google Scholar

[ref62] 62. Wurst H, Kornberg A (1994) A soluble exopolyphosphatase of Saccharomyces cerevisiae. Purification and characterization. J Biol Chem 269: 10996–11001.
View Article
Google Scholar

[171] View Article

[172] Google Scholar

[ref63] 63. Wurst H, Shiba T, Kornberg A (1995) The gene for a major exopolyphosphatase of Saccharomyces cerevisiae. J Bacteriol 177: 898–906.
View Article
Google Scholar

[174] View Article

[175] Google Scholar

Figures

Abstract

Introduction

Materials and Methods

Mycelium cultivation and materials

31P NMR.

HPLC experiments

Statistics

Results

Discussion

Author Contributions

References

³¹P NMR.