©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
The Vacuolar ATPase: Sulfite Stabilization and the Mechanism of Nitrate Inactivation (*)

(Received for publication, August 8, 1994; and in revised form, October 18, 1994)

William J. A. Dschida (§) Barry J. Bowman (¶)

From the Department of Biology, Sinsheimer Laboratories, University of California, Santa Cruz, California 95064

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Using vacuolar membranes from Neurospora crassa, we observed that sulfite prevented the loss of vacuolar ATPase activity that otherwise occurred during 36 h at room temperature. Sulfite neither activated nor changed the kinetic behavior of the enzyme. Further, in the presence of sulfite, the vacuolar ATPase was not inhibited by nitrate.

We tested the hypothesis that sulfite acts as a reducing agent to stabilize the enzyme, while nitrate acts as an oxidizing agent, inhibiting the enzyme by promoting the formation of disulfide bonds. All reducing agents tested, dithionite, selenite, thiophosphate, dithiothreitol and glutathione, prevented the loss of ATPase activity. On the other hand, all oxidizing agents tested, bromate, iodate, arsenite, perchlorate, and hydrogen peroxide, were potent inhibitors of ATPase activity. The inhibitory effect of the oxidizing agents was specific for the vacuolar ATPase. The mitochondrial ATPase, assayed under identical conditions, was not inhibited by any of the oxidizing agents. Analysis of proteins with two-dimensional gel electrophoresis indicated that nitrate can promote the formation of disufide bonds between proteins in the vacuolar membrane. These data suggest a mechanism to explain why nitrate specifically inhibits vacuolar ATPases, and they support the proposal by Feng and Forgac (Feng, Y., and Forgac, M.(1994) J. Biol. Chem. 269, 13244-13230) that oxidation and reduction of critical cysteine residues may regulate the activity of vacuolar ATPases in vivo.


INTRODUCTION

The vacuolar ATPase is a complex proton pump found in many types of membranes in eukaryotic cells. Named after the enzyme found in vacuolar membranes from plants and fungi (Kakinuma et al., 1981; Bowman and Bowman, 1982; Mandala and Taiz, 1986; Randall and Sze, 1986), the enzyme has also been found in many organellar membranes in animal cells such as lysosomes (Galloway et al., 1988; Moriyama and Nelson, 1989a), coated vesicles (Forgac, 1989, 1992; Stone et al., 1989), and chromaffin granules (Nelson, 1992a, 1992b). Plasma membranes of specialized proton-secreting cells also have vacuolar ATPases. Examples are the goblet cells of insect midgut (Wieczorek, 1992), the intercalated cells of kidney tubules (Gluck, 1992) and the osteoclasts surrounding bone (Chatterjee et al., 1993).

The function of the vacuolar ATPase is to generate an electrochemical gradient for protons across the membrane and in many cases to acidify an internal compartment. A major unsolved problem is how a single type of enzyme is regulated to establish different proton gradients across different membranes. For example, the interior of coated vesicles is essentially the same pH as the cytosol while the interior of the lysosome may be 2 pH units more acidic (Forgac, 1989; Mellman, 1992). One possible explanation is that different isoforms encode organelle-specific subunits (Manolson et al., 1994). In both plants and animals evidence has been reported for isoforms of genes encoding subunits of the vacuolar ATPase (Bernasconi et al., 1990; Hasebe et al., 1992; Lai et al., 1988; Peng et al., 1994; Puopolo et al., 1992). As appears to be the case for the osteoclast, however, these isoforms may be specific for different types of cells rather than specifying different organelles within a cell (van Hille et al., 1993). Indeed in Saccharomyces cerevisiae and Neurospora crassa only a single set of genes appears to encode almost all subunits of the vacuolar ATPase (Anraku et al., 1992; Bowman et al., 1992b; Kane and Stevens, 1992; Nelson, 1992a).

Feng and Forgac (1992a, 1992b) have recently suggested that the activity of the vacuolar ATPase may be regulated in vivo by oxidation/reduction of sulfhydryl groups. While exploring the effects of nitrate and sulfite on the vacuolar ATPase from N. crassa we have obtained data that support this idea. As described below, these data show how the activity of the ATPase can be stabilized in vitro and they offer an explanation for the mechanism of nitrate inhibition.

Nitrate has long been known as a relatively specific inhibitor of the vacuolar ATPase in many organisms (Bowman and Bowman, 1982; Bowman, 1983; Mandala and Taiz, 1986; Rea et al., 1987; Bennett et al., 1988; Moriyama and Nelson, 1989b; Arai et al., 1989). Its mechanism of inhibition has been unclear. Relatively high concentrations (approximately 50 mM) are typically required for half-maximal inhibition, and the degree of inhibition is strongly dependent on the time of exposure. One possible clue to the mechanism was the observation that incubation of membranes in nitrate, thiocyanate, or iodide caused the dissociation of peripheral subunits of the vacuolar ATPase (Rea et al., 1987; Arai et al., 1989; Bowman et al., 1989; Moriyama and Nelson, 1989b; Kane et al., 1989; Ward et al., 1991). The effectiveness of these anions as inhibitors and in dissociation of subunits followed the Hofmeister series, i.e. SCN > I > NO(3) Cl (Hatefi and Hanstein, 1974). Thus, the suggestion was made by our laboratory and others that nitrate was acting as a chaotropic salt, inhibiting the vacuolar ATPase by disrupting subunit structure (Bowman et al., 1989; Rea et al., 1987).

This explanation is not entirely satisfactory. The concentration of nitrate used to inhibit the ATPase is high but not nearly so high as is typically used for chaotropic dissociation (Hatefi and Hanstein, 1974). The concentrations of nitrate which inhibit activity are often much lower than the concentrations required for dissociation of subunits. Several laboratories have reported that inhibition by nitrate appears to occur by two different mechanisms (Arai et al., 1989; Kibak et al., 1993; Rea et al., 1987). Furthermore, for the vacuolar ATPase in osteoclasts (Chatterjee et al., 1993) and in kidney cells (Wang and Gluck, 1990) nitrate is a potent inhibitor but does not appear to cause the dissociation of subunits from the enzyme.

While nitrate is a chaotrope, it is also an oxidizing agent. As reported below, we have found that the activity of the N. crassa vacuolar ATPase can be stabilized with antioxidants such as sulfite. We have examined the ability of sulfite to prevent inhibition by nitrate and have explored the idea that the mechanism of nitrate inhibition is to cause the formation of disulfide bonds within the vacuolar ATPase.


EXPERIMENTAL PROCEDURES

Materials

Sodium salts of ATP, glutathione (oxidized and reduced forms), cystine, sulfate, sulfite, selenite, thiosulfate, nitrate, nitrite, arsenate, arsenite, bromate, iodate, and tetrathionate were purchased from Sigma. Sodium dithionite was purchased from Fluka Chemie, Buchs, Switzerland, hydrogen peroxide from Mallinckrodt, Paris KN, and dithiothreitol (DTT) (^1)from American Bioanalytical, Natick, MA.

Preparation of Vacuolar and Mitochondrial Membranes

For our initial experiments, vacuolar membranes were prepared as described previously (Bowman and Bowman, 1988). However, we have obtained higher yields of vacuolar membranes by a modification of this method. As in the previous procedure cells were harvested, disrupted with glass beads and centrifuged in a Sorvall GSA rotor for 10 min at 1000 times g, 4 °C to pellet nuclei, cell wall, and other cellular debris. The supernatant was centrifuged in the same rotor for 20 min at 25,000 times g, 4 °C to pellet the vacuoles and mitochondria. In the modified procedure 1 ml of 50% sucrose (in 10 mM Hepes, 1 mM EDTA, 2 mM Na(3)ATP, pH adjusted to 7.4 with Tris base) was layered below 2 ml of the resuspended organellar pellet in thick-walled polycarbonate centrifuge tubes. The organellar suspension was centrifuged for 20 min at 200,000 times g, 4 °C, in a Beckman tabletop ultracentrifuge using the TLA 100.3 rotor. Mitochondria and cell membranes remained above the layer of sucrose while the vacuoles formed a pellet beneath the sucrose. The vacuoles were lysed in 1 mM EGTA (pH adjusted to 7.4 with Tris base) plus 2 mM ATP (EGTA + ATP) and then centrifuged for 5 min at 16,000 times g, 4 °C to pellet contaminating heavy debris, mostly cell wall fragments. The supernatant was removed and centrifuged for 10 min at 150,000 times g, 4 °C to pellet the vacuolar membranes. Vacuolar membranes were washed once by resuspension in EGTA + ATP and centrifugation for 10 min at 100,000 times g, 4 °C. The pellet, containing purified vacuolar membranes was resuspended in EGTA + ATP to a final protein concentration of 5-10 mg/ml, frozen in liquid nitrogen, and stored at -70 °C. With these alterations to the procedure membranes have been prepared in significantly less time; 30 liters of mycelial culture (harvested at 1.0 mg/ml, dry weight) were processed in 3 h. Vacuolar membranes were routinely isolated with specific activities of 2-5 µmol P(i)/min/mg of protein, and protein yields of 3-10 mg of vacuolar membrane protein.

Mitochondrial membranes were isolated as described previously (Bowman and Bowman, 1988) with specific activities of 2-4 µmol P(i)/min/mg of protein.

Assay of Vacuolar and Mitochondrial Membranes

Frozen aliquots of vacuolar membranes were thawed in room temperature water, diluted in 9 times volume of 1 mM EGTA plus 100 mM NaCl (pH adjusted to 7.4 with Tris base), and incubated for 10 min at 4 °C. This allowed for release and subsequent removal of loosely associated, peripheral proteins. The suspension was centrifuged in a microcentrifuge for 20 min at 16,000 times g, 4 °C. The pelleted vacuolar membranes were resuspended to their original volume in 1 mM EGTA, pH 7.4 (at approximately 5 mg of protein/ml). Membranes were typically diluted in 9 times volume of treatment solution, to a final concentration of 0.5 mg of protein/ml. ATPase reactions were performed by adding 10-30 µl of the resuspended membranes into 0.5 ml of vacuolar ATPase assay mix (10 mM NH(4)Cl, 5 mM Na(2)ATP, 5 mM MgSO(4), 10 mM Pipes, pH adjusted to 7.4 with Tris base). ATPase activity, as determined by P(i) release, was assayed in vacuolar membrane ATPase assay mix for 10 min at 30 °C as described previously (Bowman and Bowman, 1988).

Two-dimensional Polyacrylamide Gel Electrophoresis

To identify polypeptides with intermolecular disulfide bonds, vacuolar membrane proteins were first electrophoresed in their unreduced form and subsequently reduced and run through a second dimension (Allison et al., 1982; Traut et al., 1988). Briefly, membranes were suspended in 9 times volume of ATPase assay mix in 50 mM NaNO(3), Na(2)SO(3), or Na(2)S(4)O(6) for 1 h at 25 °C. Vacuolar membrane proteins were then denatured by treatment in 1% SDS at 70 °C for 20 min. Samples were run in the first dimension through a 3% stacking, 13% resolving polyacrylamide gel (Bowman et al., 1981) until the protein had entered 5.5 cm into the resolving gel. The sample lanes were excised and reduced in a 50 mM Tris buffer solution, pH 8.8, containing 1% SDS (w/v) and 3% beta-mercaptoethanol at 65 °C for 20 min. The gel slices were washed twice in 50 mM Tris buffer, pH 6.8, containing 0.1% SDS for 20 min at 25 °C and placed horizontally in the top of the gel apparatus, perpendicular to their original orientation. A 13% polyacrylamide gel was cast 1.0 cm below the slices. Once the gel was polymerized, the glass plates were loosened, the gel slices gently pushed down against the resolving gel, and the plates retightened. A 0.5% agarose solution was added around the gel slices to seal any discontinuities with the new gel. The reduced proteins were then electrophoresed. The polyacrylamide gels were subsequently processed for silver-staining or Western analysis.

Polyclonal Antibody Production and Western Blot Analysis

The cDNA derived from the VMA1 gene, encoding the 67-kDa subunit of the N. crassa vacuolar ATPase (Bowman et al., 1988) was digested with BglII and BamHI restriction enzymes. This generated a 1440-base pair fragment which encoded amino acid residues 18-500 of this 589-residue polypeptide. The cDNA fragment was inserted into the pATH1 vector (a generous gift from C. Yanofsky, Stanford University) behind a 1008-base pair fragment of the inducible Escherichia coli trp promoter (encoding a 37-kDa NH(3)-terminal fragment of the trpE protein). After transformation into E. coli strain AB1899 the overexpressed polypeptide from an induced culture was purified by gel electrophoresis, excised, and homogenized in Freund's complete adjuvant. A New Zealand White rabbit was inoculated with 150 µg of the fusion protein. After three subsequent booster injections using the same amount of antigen in Freund's incomplete adjuvant, an antibody of high titer and specificity against the VMA1 protein was isolated.


RESULTS

Sulfite has been reported to change the kinetic behavior of F-type ATPases (Du and Boyer, 1990; Vasilyeva et al. 1982), archaebacterial ATPases (Inatomi, 1986; Konishi et al. 1987; Lübben and Schafer, 1987; Nanba and Mukohata, 1987; Schobert and Lanyi, 1989), and the yeast vacuolar ATPase (Kibak et al., 1993). We measured the activity of the N. crassa vacuolar ATPase using standard assay conditions (see ``Experimental Procedures'') in the presence and absence of 1-200 mM Na(2)SO(3). We observed only a modest 5-15% stimulation of ATPase activity (data not shown). The K(m) for MgATP was also measured and found to be essentially the same in the absence (0.71 mM) and presence (0.56 mM) of 100 mM sulfite (Fig. 1). Because the effect of sulfite is sometimes pH-dependent (Inatomi, 1986; Schobert and Lanyi, 1989), we measured the activity of the ATPase as a function of pH in the absence and presence of 100 mM sulfite. As shown in Fig. 2, sulfite did not shift the pH optimum but appeared to broaden the pH dependence. Only a 10% stimulation of activity was observed at the pH optimum, but the enzyme was more active, perhaps more stable, in Na(2)SO(3) at the extremes of its pH range.


Figure 1: Effect of sulfite on the substrate affinity of vacuolar membrane ATPase. Vacuolar membranes were assayed for ATP hydrolysis (as described under ``Experimental Procedures'') in ATPase assay mix containing 5 mM MgSO(4) and varying amounts of ATP as indicated. 100 mM Na(2)SO(3) was either present (closed squares) or absent (open circles).




Figure 2: Effect of sulfite on the pH profile of vacuolar membrane ATPase activity. Vacuolar membranes were assayed for ATP hydrolysis in ATPase assay mix adjusted to the indicated pH with HCl or KOH. 100 mM Na(2)SO(3) was either absent (closed squares) or present (open circles).



To explore the possibility that the vacuolar ATPase was more stable when suspended in sulfite we examined activity as a function of time at room temperature. The ATPase activity of N. crassa vacuolar membranes, suspended in 1 mM EGTA, pH 7.4, was essentially unchanged after 24 h at 4 °C (data not shown). If left at room temperature for 24 h, all of the activity was lost. Fig. 3shows the effect of adding ATP or MgATP. MgATP slightly stabilized, while ATP by itself significantly stabilized the activity. In other experiments not shown, the presence of Mg alone slightly accelerated the loss of activity, MgADP had the same protective effect as MgATP, and ADP had the same protective effect as ATP. An even better stabilizing agent than the nucleotides, however, was sodium sulfite (Fig. 3). Even in the absence of ATP the enzyme retained 65% of ATPase activity after 36 h if sulfite was present. We measured the protective effects at different sulfite concentrations and found that 100 mM was optimal in our experimental conditions (data not shown).


Figure 3: Sulfite increases the stability of the vacuolar membrane ATPase. Vacuolar membranes, suspended at 0.5 mg of protein/ml, were incubated at room temperature in either 1 mM EGTA, MgATPase assay mix (see ``Experimental Procedures''), or 1 mM EGTA plus 5 mM Na(2)ATP. 100 mM Na(2)SO(3) was either absent or present (open and closed symbols, respectively). Membrane aliquots from the various treatments were assayed for ATPase activity at 6-h intervals. The activity of the controls at time = 0 was set at 100%.



Because sulfite is frequently used as an ``antioxidant'' we tested the ability of other reducing agents to stabilize the activity of the vacuolar ATPase. Dithiothreitol is often used in the preparation of vacuolar ATPases, but at the concentrations effective for other organisms (typically 5 mM) it did not prevent inactivation of the N. crassa ATPase. In the experiments shown in Fig. 4, membranes were suspended in ATPase assay mix in the absence (0 concentration in each panel) or presence (concentrations shown on x axis of each panel) of reducing agents, and incubated at room temperature for 12 h. With no added reducing agent, the membranes lost half of their ATPase activity. Dithiothreitol at high concentrations, e.g. 300 mM, prevented loss of activity. More effective, however, were a group of reducing agents which are smaller than dithiothreitol. Selenite (Na(2)SeO(3)), thiophosphate (Na(2)SPO(3)), and dithionite (Na(2)S(2)O(4)) demonstrated protective effects, the latter being nearly as effective as sulfite, but at 0.1 the concentration. Even reduced glutathione at high concentrations (10 mM) partially prevented loss of ATPase activity. Oxidized glutathione had no effect on activity (data not shown).


Figure 4: Reducing agents stabilize the vacuolar ATPase. Vacuolar membranes, suspended at 0.5 mg of protein/ml, were incubated in ATPase assay mix with the indicated amounts of reducing agents. After 12 h at 25 °C, the samples with no added reducing agents had lost 50% of their initial ATPase activity. At that time samples were centrifuged for 15 min in a microcentrifuge at 16,000 times g. The membrane pellets were resuspended to their original volume with 1 mM EGTA (pH 7.4) and assayed for ATPase activity. The data represent the activity of the membranes after the 12 h incubation relative to the initial activity of the untreated membranes.



If reducing agents prevented loss of ATPase activity, then oxidizing agents might be effective inhibitors of the vacuolar ATPase. Nitrate and thiocyanate, known inhibitors of vacuolar ATPase, are moderately strong oxidizing agents. As shown in Fig. 5, we tested nitrate and several other oxidizing agents, iodate, bromate, arsenite, perchlorate, and hydrogen peroxide, and found all of them to be potent inhibitors of the vacuolar ATPase when incubated in the presence of 5 mM MgATP. To see if the inhibitory effects of these oxidizing agents was specific for the vacuolar ATPase, we also tested these reagents with the mitochondrial ATPase, an enzyme closely related to the vacuolar ATPase in structure and mechanism. As shown in Table 1, under identical assay conditions oxidizing agents which inhibited the vacuolar ATPase had no inhibitory effect on the activity of the mitochondrial ATPase. In fact, bromate had the surprising effect of increasing the ATPase activity of mitochondrial membranes.


Figure 5: Oxidizing agents inactivate the vacuolar ATPase. Vacuolar membranes, suspended at 0.5 mg of protein/ml, were incubated in ATPase assay mix with the indicated amounts of oxidizing agents for 45 min at 25 °C. Samples were centrifuged for 15 min in a microcentrifuge at 16,000 times g. The membrane pellets were immediately resuspended to original volume with 1 mM EGTA (pH 7.4) and assayed for ATPase activity.





Focusing on nitrate, we assayed the ability of sulfite to block inhibition. In Fig. 6, vacuolar membranes were incubated in 50 mM nitrate together with varying concentrations of sulfite. After 1 h at room temperature ATPase activity was assayed. The results showed that increasing levels of ATPase activity were retained with increasing concentrations of sulfite. Inhibition by nitrate was effectively blocked by 100 mM sulfite. We had previously observed (Bowman et al., 1989) that nitrate also caused the release of the peripheral subunits of the ATPase from the vacuolar membrane. In the experiment shown in Fig. 6we measured the relative amounts of peripheral subunits of the ATPase released into the supernatant. The results indicated that sulfite blocked the release of ATPase subunits with the same concentration dependence seen for protection of ATPase activity.


Figure 6: Sulfite blocks ATP-dependent nitrate-inactivation. Vacuolar membranes, resuspended at 0.5 mg of protein/ml, were incubated in ATPase assay mix plus 50 mM NaNO(3) for 45 min at 25 °C with the indicated amounts of Na(2)SO(3). Samples were centrifuged for 15 min in a microfuge at 16,000 times g. The membrane pellets were immediately resuspended to original volume with 1 mM EGTA (pH 7.4) and assayed for ATPase activity. The data represent the activity of the membranes after one h of treatment relative to the initial activity of the untreated membranes. The supernatants from the various treatments were examined by polyacrylamide gel electrophoresis. Release of the peripheral subunits of the ATPase (the V1 sector) was analyzed by gel densitometry. Shown is the amount of these subunits relative to the amount observed in the absence of Na(2)SO



A straightforward interpretation of these results is that ATPase activity can be inhibited by the oxidation of specific residues within the enzyme and that reducing agents prevent this oxidation. Feng and Forgac (1992a, 1992b) have shown that the vacuolar ATPase of bovine coated vesicles can be reversibly inhibited by reaction of cystine in the medium with Cys-154 in the A subunit. We incubated N. crassa vacuolar and mitochondrial membranes with cystine and observed that the vacuolar ATPase was inhibited while the mitochondrial ATPase was unaffected (Table 1). Tetrathionate, another reagent which promotes the formation of disulfide bonds (Means and Feeney, 1970), had the same effect as cystine.

In several important aspects inhibition by cystine and nitrate were different. First, as has been shown by many laboratories, inhibition by nitrate is strongly promoted by the presence of ATP (Arai et al., 1989; Bowman et al., 1989; Moriyama and Nelson, 1989b; Rea et al., 1987). By contrast ATP prevented inhibition by cystine (Feng and Forgac, 1992b). As shown in Table 2, after 4 h in 1 mM cystine ATPase activity was completely inhibited. This inhibition could be partially prevented (nearly 50%) with the inclusion of 5 mM ATP. By contrast, enzyme incubated with nitrate in the absence of ATP lost little activity. Inclusion of ATP with nitrate caused a complete loss of activity. Inclusion of sulfite (100 mM) along with cystine blocked the inhibition but did not reverse it if added after the inactivation (data not shown).



Second, inhibition by cystine was reversible by DTT, even after 24 h, while inhibition by nitrate was irreversible. Table 3shows the effect of DTT on vacuolar membranes treated with oxidizing agents. DTT reactivated enzymes that had been treated with tetrathionate or cystine. DTT did not reactivate nitrate or arsenite treated membranes. Similarly, sulfite at high concentrations (100 mM) blocked the inhibition but did not reverse it (data not shown). Third, we also observed that inhibition by cystine, unlike nitrate, did not cause dissociation of the peripheral subunits of the ATPase. In fact incubation in cystine protected the ATPase against nitrate-induced dissociation of the peripheral V(1) subunits (data not shown).



The hypothesis that nitrate oxidizes the enzyme and causes the formation of disulfide bonds was tested by analyzing vacuolar proteins after two-dimensional polyacrylamide gel electrophoresis. The membranes were first incubated in the absence or presence of 50 mM nitrate. The polypeptides were then separated in the first dimension in the absence of reducing agent. Mercaptoethanol was added and the polypeptides were electrophoresed in the second dimension. All polypeptides are predicted to lie on a diagonal, unless they have initially been cross-linked by disulfide bonds, in which case they will migrate faster in the second dimension and appear as off-diagonal spots (Allison et al., 1982; Traut et al., 1988). After incubation in tetrathionate, arsenite, or nitrate the vacuolar membranes showed a prominent off-diagonal polypeptide of approximately 70 kDa that was not seen in the control (data not shown). Since inhibition was correlated with the appearance of the off-diagonal spot, we tested whether the 67-kDa subunit of the vacuolar ATPase was involved in this oxidation. Using a polyclonal antibody, we identified the 67-kDa subunit in the diagonal, but the off-diagonal polypeptide was only faintly labeled. This experiment was repeated several times with the same result. Thus, we were not able to identify definitively the off-diagonal spot. However, the data clearly indicated that incubation in nitrate could promote the formation of disulfide bonds.


DISCUSSION

In chloroplast F-type ATPase (Du and Boyer, 1990; Vasilyeva et al., 1982), archaebacterial ATPase (Inatomi, 1986; Lübben and Schafer, 1987; Schobert and Lanyi, 1989), and yeast vacuolar ATPase (Kibak et al., 1993) the rate of ATP hydrolysis versus time often exhibits a biphasic pattern. A fast initial rate is sustained for a few seconds or minutes, followed by a significantly slower rate. Addition of sulfite to the assay mixture causes the fast initial rate to be sustained and can also reactivate enzyme in which the rate had slowed. This behavior has been explained by postulating that during ATP hydrolysis an inhibited form of the ATPase with tightly bound ADP accumulates. In the presence of sulfite the tightly bound ADP is released (Du and Boyer, 1990). With the exception of the enzyme from S. cerevisiae this kind of kinetic behavior has not been reported for vacuolar ATPases. In our analysis of the N. crassa vacuolar ATPase sulfite was not observed to significantly stimulate hydrolysis or to change the K(m) for ATP. We suggest that an inhibited form of the vacuolar ATPase with tightly bound ADP does not significantly accumulate in our assay conditions and that the effects of sulfite we have observed have a different mechanistic basis.

Our data indicated that sulfite significantly slows the inactivation of the enzyme but cannot reactivate after ATPase activity is lost. Most importantly, sulfite prevented inhibition of the enzyme by nitrate. We propose a mechanism which may explain why nitrate and related compounds inhibit the vacuolar ATPase. These compounds act not as chaotropes, as we and others originally suggested (Bowman et al., 1989; Rea et al., 1987) but as oxidizing agents, promoting the formation of disulfide bonds (Means and Feeney, 1970; Gardlik and Rajagopalan, 1991; Guerrieri and Papa, 1982). In this report we have shown that other oxidizing agents, e.g. bromate, perchlorate, iodate, arsenite, and tetrathionate are also potent inhibitors of the vacuolar ATPase. Sulfite stabilizes the vacuolar ATPase and blocks inhibition apparently because it is a good reducing agent (Means and Feeney, 1970). Reducing agents that are larger molecules than sulfite, such as dithiothreitol, are not as effective in protecting the N. crassa enzyme, but are effective in stabilizing vacuolar ATPase in mammalian cells (Feng and Forgac, 1992a). Smaller sized reducing reagents, e.g. dithionite and selenite protect the N. crassa enzyme nearly as well as sulfite. These results suggest that the oxidation site is partially buried within the N. crassa enzyme.

A model consistent with these results is shown in Fig. 7. In the absence of ATP or other nucleotides, the active site of the enzyme can be occupied by cystine which can form a disulfide bond with a cysteine residue, via thio-disulfide exchange. Forgac's laboratory has reported that both cystine and N-ethylmaleimide bind to Cys-254 in the 67-kDa subunit of the bovine vacuolar ATPase (Feng and Forgac, 1992a, 1994). Enzyme inhibited by cystine does not dissociate and, in fact, can be readily reactivated by dithiothreitol. When ATP is bound, a conformational change occurs which makes the enzyme susceptible to nitrate and other oxidizing agents. The data indicate that nitrate can cause intermolecular cross-linking of polypeptides by disulfide bonds, but we have not directly demonstrated that this cross-linking causes inhibition or dissociation. We suggest that a disulfide bond is formed, probably within or between subunits of the enzyme, quickly followed by dissociation of the peripheral sector (Bowman et al., 1989). Thus, inhibition by nitrate is effectively irreversible. By keeping the sulfhydryl groups of cysteine residues reduced, sulfite can block inhibition by either nitrate or cystine.


Figure 7: Model for the mechanism of inhibition of vacuolar ATPase by cystine and nitrate. The ATPase is depicted as being composed of two sectors. After exposure to nitrate the peripheral, ATP-binding sector, can dissociate from the integral membrane sector. Sulfhydryl groups within the enzyme are represented by SH. Further details are given in the text.



Puopolo and Forgac(1990) reported that ATPase from mammalian coated vesicles, dissociated with high concentrations of iodide, could be reassembled if the iodide was removed in the presence of the reducing agent beta-mercaptoethanol. Although we have not attempted such experiments with the N. crassa ATPase such results are consistent with our model. The model postulates that inhibition by nitrate and dissociation occur in distinct steps to account for differences between the nitrate effect on ATPase activity and nitrate-induced dissociation of peripheral subunits (Arai et al., 1989; Kibak et al., 1993; Rea et al., 1987). ATPases from different organisms may differ in the rate at which oxidation is followed by dissociation.

One appeal of this explanation for nitrate inhibition is that it can explain the specificity of nitrate for vacuolar ATPases as opposed to F-type ATPases. The A and B subunits of the vacuolar ATPases contain several cysteine residues, three of which are conserved in all sequenced A subunits (Taiz et al.(1994) and references therein) and one of which is conserved in the B subunits (Puopolo et al.(1992) and references therein). By contrast, the homologous alpha and beta subunits of F-type ATPases have fewer cysteines, in several cases none. The residues targeted by nitrate might also be in other subunits of the vacuolar ATPase. The 54-kDa subunit of the S. cerevisiae (Ho et al., 1993) has 6 cysteines, but the sequence for the homologous subunit in other organisms has not yet been reported. Subunit C has no cysteines common to S. cerevisiae and bovine cells (Nelson et al., 1990; Beltrán et al., 1992). The sequence of subunit D has not yet been reported for any vacuolar ATPase. Subunit E has no conserved cysteines when S. cerevisiae (Foury, 1990), bovine (Hirsh et al., 1988), Manduca sexta (Graf et al., 1994), and N. crassa(^2)sequences are compared. Among the membrane associated subunits at least two cysteine residues are conserved in the 40-kDa subunits from S. cerevisiae (Bauerle et al., 1993), bovine cells (Wang et al., 1988), and N. crassa. (^3)(Because of a possible error in the published bovine sequence, discussed in Bauerle et al.(1993), there is probably a third conserved cysteine near the N terminus of this subunit.)

If the cysteine residues in the 67-kDa subunit are the targets of nitrate inhibition, then the recent data of Taiz et al. (1994) are of particular interest. In this report the three conserved cysteines in the vacuolar ATPase from S. cerevisiae were each changed to serine residues. Cys-254, which corresponds to the residue that binds N-ethylmaleimide and cystine in the bovine vacuolar ATPase (Feng and Forgac, 1992a, 1994) was changed to serine without loss of ATPase activity. Furthermore, the altered ATPase had the same sensitivity to nitrate as the wild type. Changing either of the other conserved cysteines (Cys-284 > Ser, or Cys-538 > Ser) inactivated the ATPase. It is because of these data that we suggest (Fig. 7) that inhibition by nitrate occurs at a different site than that affected by cystine.

Our results suggest that sulfite will be useful in development of procedures to purify this complex and sometimes unstable enzyme. In our current protocols we often observe a separation of the integral membrane and the peripheral components during purification on sucrose gradients (Bowman et al., 1992b). Preliminary results indicate the enzyme stays intact in the presence of sulfite. Of broader significance, the results support proposals from other laboratories (Feng and Forgac, 1992a, 1992b, 1994; Kibak et al., 1993) that within the cell, the redox state of the immediate environment may play a key role in regulating the activity of vacuolar ATPases.


FOOTNOTES

*
This work was supported by United States Public Health grants GM28703 and GM08132. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Current address: Division of Geological and Planetary Sciences, California Institute of Technology, Pasadena, CA 91125.

To whom correspondence should be addressed. Tel.: 408-459-2245; Fax: 408-459-3139.

(^1)
The abbreviations used are: DTT, dithiothreitol; Pipes, 1,4-piperazinediethanesulfonic acid.

(^2)
E. J. Bowman and A. Steinhardt, unpublished results.

(^3)
V. Melnik and B. J. Bowman, unpublished results.


ACKNOWLEDGEMENTS

We thank Henrik Kibak and Lincoln Taiz for helpful discussions, and Nora Vázquez-Laslop for assistance in makingantibody. We also thank Emma Jean Bowman for advice on both the experiments and the manuscript.


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