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 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

()

Preparation of Vacuolar and Mitochondrial Membranes

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

Assay of Vacuolar and Mitochondrial Membranes

Two-dimensional Polyacrylamide Gel Electrophoresis

Polyclonal Antibody Production and Western Blot Analysis

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 NaSO. We observed only a modest 5-15% stimulation of ATPase activity (data not shown). The K 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 NaSO 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 and varying amounts of ATP as indicated. 100 mM NaSO 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 NaSO 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 NaATP. 100 mM NaSO 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 (NaSeO), thiophosphate (NaSPO), and dithionite (NaSO) 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 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 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 for 45 min at 25 °C with the indicated amounts of NaSO. Samples were centrifuged for 15 min in a microfuge at 16,000 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 NaSO

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 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 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 -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 and 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()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. ()(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.

()

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

()

E. J. Bowman and A. Steinhardt, unpublished results.

()

V. Melnik and B. J. Bowman, unpublished results.

ACKNOWLEDGEMENTS

REFERENCES

1. Allison, J. P., McIntyre, B. W., and Bloch., D. (1982) J. Immunol. 129, 2293-2300 [Abstract]
2. Anraku, Y., Hirata, R., Wada, Y., and Ohya, Y. (1992) J. Exp. Biol. 172, 67-81 [Abstract/Free Full Text]
3. Arai, H., Pink, S., and Forgac, M. (1989) Biochemistry 28, 3075-3082 [CrossRef][Medline] [Order article via Infotrieve]
4. Bauerle, C., Ho, M. N., Lindorfer, M. A., and Stevens, T. H. (1993) J. Biol. Chem. 268, 12749-12757 [Abstract/Free Full Text]
5. Beltrán, C., Kopecky, J., Pan, Y.-C. E., Nelson, H., and Nelson, N. (1992) J. Biol. Chem. 267, 774-779 [Abstract/Free Full Text]
6. Bennett, A. B., Leigh, R. A., and Spanswick, R. M. (1988) Methods Enzymol. 157, 579-590 [Medline] [Order article via Infotrieve]
7. Bernasconi, P., Rausch, T., Struve, I., Morgan, L., and Taiz, L. (1990) J. Biol. Chem. 265, 17428-17431 [Abstract/Free Full Text]
8. Bowman, B. J., Blasco, F., and Slayman, C. W. (1981) J. Biol. Chem. 256, 12343-12349 [Abstract/Free Full Text]
9. Bowman, B. J., Dschida, W. J., Harris, T., and Bowman, E., J. (1989) J. Biol. Chem. 264, 15606-15612 [Abstract/Free Full Text]
10. Bowman, B. J., Dschida, W. J., and Bowman, E. J. (1992a) J. Exp. Biol. 172, 57-66 [Abstract/Free Full Text]
11. Bowman, B. J., Vázquez-Laslop, N., and Bowman, E. J. (1992b) J. Bioenerg. Biomembr. 24, 361-370 [CrossRef][Medline] [Order article via Infotrieve]
12. Bowman, E. J. (1983) J. Biol. Chem. 258, 15238-15244 [Abstract/Free Full Text]
13. Bowman, E. J., and Bowman, B. J. (1982) J. Bacteriol. 151, 1326-1337 [Abstract/Free Full Text]
14. Bowman, E. J., and Bowman, B. J. (1988) Methods Enzymol. 157, 562-573 [Medline] [Order article via Infotrieve]
15. Bowman, E. J., Tenney, K., and Bowman, B. J. (1988) J. Biol. Chem. 263, 13994-14001 [Abstract/Free Full Text]
16. Chatterjee, D., Neff, L., Chakraborty, M., Fabricant, C., and Baron, R. (1993) Biochemistry 32, 2808-2812 [CrossRef][Medline] [Order article via Infotrieve]
17. Du, Z. Y., and Boyer, P. D. (1990) Biochemistry 29, 402-407 [CrossRef][Medline] [Order article via Infotrieve]
18. Feng, Y., and Forgac, M. (1992a) J. Biol. Chem. 267, 5817-5822 [Abstract/Free Full Text]
19. Feng, Y., and Forgac, M. (1992b) J. Biol. Chem. 267, 19769-19772 [Abstract/Free Full Text]
20. Feng, Y., and Forgac, M. (1994) J. Biol. Chem. 269, 13224-13230 [Abstract/Free Full Text]
21. Forgac, M. (1989) Physiol. Rev. 69, 765-796 [Free Full Text]
22. Forgac, M. (1992) J. Bioenerg. Biomembr. 24, 339-340 [CrossRef][Medline] [Order article via Infotrieve]
23. Foury, F. (1990) J. Biol. Chem. 265, 18554-18560 [Abstract/Free Full Text]
24. Galloway, C. J., Dean, G. E., Fuchs, R., and Mellman, I. (1988) Methods Enzymol. 157, 601-611 [Medline] [Order article via Infotrieve]
25. Gardlik, S., and Rajagopalan, K. V. (1991) J. Biol. Chem. 266, 16627-16632 [Abstract/Free Full Text]
26. Gluck, S. (1992) J. Exp. Biol. 172, 29-37 [Abstract/Free Full Text]
27. Graf, R., Harvey, W. R., and Wieczorek, H. (1994) Biochim. Biophys. Acta 1190, 193-196 [Medline] [Order article via Infotrieve]
28. Guerrieri, F., and Papa, S. (1982) Eur. J. Biochem. 128, 9-13 [Medline] [Order article via Infotrieve]
29. Hasebe, M., Hanada, H., Moriyama, Y., Maeda, M., and Futai, M. (1992) Biochem. Biophys. Res. Commun. 183, 856-63 [CrossRef][Medline] [Order article via Infotrieve]
30. Hatefi, Y. H., and Hanstein, W. G. (1974) Methods Enzymol. 31, 770-779 [Medline] [Order article via Infotrieve]
31. Hirsch, S., Strauss, A., Massod, K., Lee, S., Sukhatme, V., and Gluck, S. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 3004-3008 [Abstract/Free Full Text]
32. Ho, M. N., Hirata, R., Umemoto, N., Ohya., Y. Takatsuki, A., Stevens, T. H., and Anraku, Y. (1993) J. Biol. Chem. 268, 18286-18292 [Abstract/Free Full Text]
33. Inatomi, K.-I. (1986) J. Bacteriol. 167, 837-841 [Abstract/Free Full Text]
34. Kakinuma, Y., Ohsumi, Y., and Anraku, Y., (1981) J. Biol. Chem. 256, 10859-10863 [Abstract/Free Full Text]
35. Kane, P. M., and Stevens, T. H. (1992) J. Bioenerg. Biomembr. 24, 383-393 [CrossRef][Medline] [Order article via Infotrieve]
36. Kane, P. M., Yamashiro, C. T., and Stevens, T. H. (1989) J. Biol. Chem. 264, 19236-19244 [Abstract/Free Full Text]
37. Kibak, H., Van Eeckhout, D., Cutler, T., Taiz, S. L., and Taiz, L. (1993) J. Biol. Chem. 268, 23325-23333 [Abstract/Free Full Text]
38. Konishi, J., Wakagi, T., Oshima, T., and Yoshida, M. (1987) J. Biochem. 102, 1379-1387 [Abstract/Free Full Text]
39. Lai, S., Randall, S. K., and Sze, H. (1988) J. Biol. Chem. 263, 16731-16737 [Abstract/Free Full Text]
40. Lübben, M., and Schafer, G. (1987) Eur. J. Biochem. 164, 533-540 [Medline] [Order article via Infotrieve]
41. Mandala, S., and Taiz, L. (1986) J. Biol. Chem. 261, 12850-12855 [Abstract/Free Full Text]
42. Manolson, M. F., Wu, B., Proteau, D., Taillon, B. E., Roberts, B. T., Hoyt, M. A., and Jones, E. W. (1994) J. Biol. Chem. 269, 14064-14074 [Abstract/Free Full Text]
43. Means, G. E., and Feeney, R. E. (1970) Chemical Modification of Proteins , pp. 149-174, Holden-Day, Inc., San Francisco, CA
44. Mellman, I. (1992) J. Exp. Biol. 172, 39-45 [Free Full Text]
45. Moriyama, Y., and Nelson, N. (1989a) Biochim. Biophys. Acta 980, 241-247 [Medline] [Order article via Infotrieve]
46. Moriyama, Y., and Nelson, N. (1989b) J. Biol. Chem. 264, 3577-3582 [Abstract/Free Full Text]
47. Nanba, T., and Mukohata, Y. (1987) J. Biochem. 102, 591-598 [Abstract/Free Full Text]
48. Nelson, N. (1992a) J. Bioenerg. Biomembr. 24, 407-414 [CrossRef][Medline] [Order article via Infotrieve]
49. Nelson, N. (1992b) J. Exp. Biol. 172, 149-153 [Abstract/Free Full Text]
50. Nelson, H., Mandiyan, S., Noumi, T., Moriyama, Y., Miedel, M. C., and Nelson, N. (1990) J. Biol. Chem. 265, 20390-20393 [Abstract/Free Full Text]
51. Peng, S.-B., Crider, B. P., Xie, X.-S., and Stone, D. K. (1994) J. Biol. Chem. 269, 17262-17266 [Abstract/Free Full Text]
52. Puopolo, K., and Forgac, M. (1990) J. Biol. Chem. 265, 14836-14841 [Abstract/Free Full Text]
53. Puopolo, K., Kumamoto, C., Adachi, I., Magner, R., and Forgac, M. (1992) J. Biol. Chem. 267, 3696-3706 [Abstract/Free Full Text]
54. Randall, S. K., and Sze, H. (1986) J. Biol. Chem. 261, 1364-1371 [Abstract/Free Full Text]
55. Rea, P. A., Griffin, C. J., Manolson, M. F., and Sanders, D. (1987) Biochim. Biophys. Acta 904, 1-12 [CrossRef]
56. Schobert, B., and Lanyi, J. K. (1989) J. Biol. Chem. 264, 12805-12812 [Abstract/Free Full Text]
57. Stone, D. K., Crider, B. P., Sudhof, T. C., and Xie, X. S. (1989) J. Bioenerg. Biomembr. 21, 605-620 [CrossRef][Medline] [Order article via Infotrieve]
58. Taiz, L., Nelson, H., Maggert, K., Morgan, L., Yatabe, B., Taiz, S. L., Rubenstein, B., and Nelson, N. (1994) Biochim. Biophys. Acta 1194, 329-334 [Medline] [Order article via Infotrieve]
59. Traut, R. R., Casiano, C., and Zercherle, N. (1988) in Protein Function: a Practical Approach (Creighton, T. E., ed) pp. 101-133, IRL Press, Oxford, UK
60. van Hille, B., Richener, H., Evans, D. B., Green, J. R., and Bilbe, G. (1993) J. Biol. Chem. 268, 7075-7080 [Abstract/Free Full Text]
61. Vasilyeva, E. A., Minkov, I. B., Fitin, A. F., and Vinogradov, A. D. (1982) Biochem. J. 202, 15-23 [Medline] [Order article via Infotrieve]
62. Wang S. Y., Moriyama, Y., Mandel, M., Hulmes, J. D., Pan, Y. C., Danho, W., Nelson, H., and Nelson, N. (1988) J. Biol. Chem. 263, 17638-17642 [Abstract/Free Full Text]
63. Wang, Z. Q., and Gluck, S. (1990) J. Biol. Chem. 265, 21957-21965 [Abstract/Free Full Text]
64. Ward, J. M., Reinders, A., Hsu, H.-T., and Sze, H. (1991) Plant Physiol. 99, 161-169
65. Wieczorek, H. (1992) J. Exp. Biol. 172, 335-343 [Abstract/Free Full Text]

CiteULike

Complore

Connotea

Del.icio.us

Digg

Reddit

Technorati

This article has been cited by other articles:

				A. Brux, T.-Y. Liu, M. Krebs, Y.-D. Stierhof, J. U. Lohmann, O. Miersch, C. Wasternack, and K. Schumacher Reduced V-ATPase Activity in the trans-Golgi Network Causes Oxylipin-Dependent Hypocotyl Growth Inhibition in Arabidopsis PLANT CELL, April 1, 2008; 20(4): 1088 - 1100. [Abstract] [Full Text] [PDF]

				M. A. Owegi, D. L. Pappas, M. W. Finch Jr.,, S. A. Bilbo, C. A. Resendiz, L. J. Jacquemin, A. Warrier, J. D. Trombley, K. M. McCulloch, K. L. M. Margalef, et al. Identification of a Domain in the Vo Subunit d That Is Critical for Coupling of the Yeast Vacuolar Proton-translocating ATPase J. Biol. Chem., October 6, 2006; 281(40): 30001 - 30014. [Abstract] [Full Text] [PDF]

				X.-H. Weng, M. Huss, H. Wieczorek, and K. W. Beyenbach The V-type H+-ATPase in Malpighian tubules of Aedes aegypti: localization and activity J. Exp. Biol., July 1, 2003; 206(13): 2211 - 2219. [Abstract] [Full Text] [PDF]

				V. F. Rizzo, U. Coskun, M. Radermacher, T. Ruiz, A. Armbruster, and G. Gruber Resolution of the V1 ATPase from Manduca sexta into Subcomplexes and Visualization of an ATPase-active A3B3EG Complex by Electron Microscopy J. Biol. Chem., January 3, 2003; 278(1): 270 - 275. [Abstract] [Full Text] [PDF]

				G. Gruber, H. Wieczorek, W. R. Harvey, and V. Muller Structure-function relationships of A-, F- and V-ATPases J. Exp. Biol., January 8, 2001; 204(15): 2597 - 2605. [Abstract] [Full Text] [PDF]

				J. Wilson, P Laurent, B. Tufts, D. Benos, M Donowitz, A. Vogl, and D. Randall NaCl uptake by the branchial epithelium in freshwater teleost fish: an immunological approach to ion-transport protein localization J. Exp. Biol., January 8, 2000; 203(15): 2279 - 2296. [Abstract] [PDF]

				E. Bowman and B. Bowman Cellular role of the V-ATPase in Neurospora crassa: analysis of mutants resistant to concanamycin or lacking the catalytic subunit A J. Exp. Biol., January 1, 2000; 203(1): 97 - 106. [Abstract]

				H Wieczorek, G Grber, W. Harvey, M Huss, H Merzendorfer, and W Zeiske Structure and regulation of insect plasma membrane H(+)V-ATPase J. Exp. Biol., January 1, 2000; 203(1): 127 - 135. [Abstract]

				C. Landolt-Marticorena, W. H. Kahr, P. Zawarinski, J. Correa, and M. F. Manolson Substrate- and Inhibitor-induced Conformational Changes in the Yeast V-ATPase Provide Evidence for Communication between the Catalytic and Proton-translocating Sectors J. Biol. Chem., September 10, 1999; 274(37): 26057 - 26064. [Abstract] [Full Text] [PDF]

				M. Forgac Structure and Properties of the Vacuolar (H+)-ATPases J. Biol. Chem., May 7, 1999; 274(19): 12951 - 12954. [Full Text] [PDF]

				M. L. Muller, M. Jensen, and L. Taiz The Vacuolar H+-ATPase of Lemon Fruits Is Regulated by Variable H+/ATP Coupling and Slip J. Biol. Chem., April 16, 1999; 274(16): 10706 - 10716. [Abstract] [Full Text] [PDF]

				M. Forgac The Vacuolar H+-ATPase of Clathrin-coated Vesicles Is Reversibly Inhibited by S-Nitrosoglutathione J. Biol. Chem., January 15, 1999; 274(3): 1301 - 1305. [Abstract] [Full Text] [PDF]

				Y. E. Oluwatosin and P. M. Kane Mutations in the CYS4 Gene Provide Evidence for Regulation of the Yeast Vacuolar H+-ATPase by Oxidation and Reduction in Vivo J. Biol. Chem., October 31, 1997; 272(44): 28149 - 28157. [Abstract] [Full Text] [PDF]

				J. J. Tomashek, B. S. Garrison, and D. J. Klionsky Reconstitution in Vitro of the V1 Complex from the Yeast Vacuolar Proton-translocating ATPase. ASSEMBLY RECAPITULATES MECHANISM J. Biol. Chem., June 27, 1997; 272(26): 16618 - 16623. [Abstract] [Full Text] [PDF]

				B. Drukarch, C. A. M. Jongenelen, E. Schepens, C. H. Langeveld, and J. C. Stoof Glutathione Is Involved in the Granular Storage of Dopamine in Rat PC12 Pheochromocytoma Cells: Implications for the Pathogenesis of Parkinson's Disease J. Neurosci., October 1, 1996; 16(19): 6038 - 6045. [Abstract] [Full Text] [PDF]