(Received for publication, August 8, 1994; and in revised form, October 18, 1994)
From the
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.
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.
Mitochondrial membranes were isolated as described previously
(Bowman and Bowman, 1988) with specific activities of 2-4
µmol P/min/mg of protein.
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 Na
SO
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 Na
SO
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 Na
SO
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
(Na
SPO
), and dithionite
(Na
S
O
) 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 Na
SO
. 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
Na
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 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.
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.