(Received for publication, June 16, 1995; and in revised form, July 26, 1995)
From the
In plants, the proton pump-ATPase (H-ATPase) of
the plasma membrane is encoded by a multigene family. The presence
within an organ of several isoforms prevents a detailed enzymatic
characterization of individual H
-ATPases. We therefore
used the yeast Saccharomyces cerevisiae as a heterologous host
for the expression of PMA2, an H
-ATPase isoform of Nicotiana plumbaginifolia. Yeast transformed by the plant pma2 was still able to grow under conditions where the yeast
ATPase gene (PMA1) was either repressed or deleted. The
transformed yeast strain was resistant to hygromycin, and its growth
was prevented when the medium pH was lowered to 5.0. The N.
plumbaginifolia PMA2 expressed in S. cerevisiae has
unusual low K
for ATP (23
µM) and high pH optimum (6.8). Electron microscopic
examination revealed PMA2 in internal structures of the karmellae type which proliferated when cell growth was arrested, either at a
nonpermissive pH or at the stationary phase in a minimal medium. Under
the latter conditions, subcellular fractionation on sucrose gradients
revealed, in addition to the expected plant PMA2 peak linked to the
plasma membrane fraction, a low density peak containing PMA2 and KAR2,
an endoplasmic reticulum marker. These observations suggest that the
partial internal accumulation of PMA2 occurs in membranes derived from
the endoplasmic reticulum and largely depends on growth conditions.
Solute transport across the plasma membrane of fungi and plants
is powered by a proton motive force built up across the membrane by a
proton pump-ATPase (H-ATPase). In yeast,
H
-ATPase is encoded by two PMA (plasma
membrane H
-ATPase) genes (Serrano et al.,
1986; Schlesser et al., 1988). Only the PMA1 gene,
however, is constitutively expressed and essential to growth (Serrano et al., 1986). This simple situation has made it possible to
characterize the PMA1 enzyme in detail, using biochemical and genetic
approaches (for reviews, see Goffeau and Slayman(1981), Serrano(1989),
and Rao et al.(1992)).
The situation in plants is much more
complex, since there, H-ATPase is encoded by a large
family of genes (for reviews, see Sussman(1994) and Michelet and
Boutry(1995)). In Nicotiana plumbaginifolia, for instance,
nine pma genes have been isolated so far. Four of these have
been characterized more extensively and classified in two gene
subfamilies according to their homology. All four genes are expressed
in most organs of the plant (Perez et al., 1992; Moriau et
al., 1993), although expression of each of them is restricted to
particular cell types (Michelet et al., 1994). (
)The occurrence of gene subfamilies might be linked to
specialization of the H
-ATPases, since plant
H
-ATPases trigger a variety of physiological functions
(Michelet and Boutry, 1995). It can therefore be hypothesized that,
although the basic function of an H
-ATPase is to
couple H
translocation to ATP hydrolysis, its kinetic
parameters and regulatory features might vary according to the isoform.
The simultaneous presence of several isoforms within the same organ,
however, prevents their individual biochemical characterization. This
restriction can be lifted by expressing individual plant pma genes in a heterologous host. Three Arabidopsis thaliana genes encoding the H
-ATPase isoforms AHA1, AHA2, AHA3 (they are called AHA for Arabidopsis H
-ATPase) have been
expressed in the yeast Saccharomyces cerevisiae (Villalba et al., 1992; Palmgren and Christensen, 1994). However none of
these plant H
-ATPases supported yeast growth when
expression of the yeast H
-ATPase was silenced
(Palmgren and Christensen, 1994). This feature prevented the study of
the native Arabidopsis ATPase expressed in a still growing S. cerevisiae. Yeast growth occurred, however, when 92
residues of the AHA2 carboxyl terminus, thought to contain a regulatory
element, were removed (Palmgren and Christensen, 1993).
In this
work, we have analyzed the expression in S. cerevisiae of the pma2 H-ATPase from N.
plumbaginifolia. This gene was chosen because it belongs to
another subfamily than the one comprising the A. thaliana genes aha1-3 and N. plumbaginifolia pma4 gene. The two subfamilies diverged before the separation of
current plant families and their divergence might be linked to a
specialization of their products (Moriau et al., 1993). In
addition, we used a recipient S. cerevisiae strain deleted of
its own two H
-ATPase genes (PMA1 and PMA2) to prevent their recombination with the plant pma2, S. cerevisiae being prone to homologous
recombinations that may lead to artefactual data (discussed in Harris et al. (1991, 1993, and 1994)). We show that the plant pma2 can sustain yeast growth. This has enabled us to
characterize biochemically a functional plant H
-ATPase
in a yeast strain growing in the absence of any residual yeast enzyme.
Moreover, analysis of internal structures by subcellular fractionations
and electron microscopy revealed that expression of plant PMA2
H
-ATPase induces proliferation of ER-derived karmellae membranes whose structure and abundance depend on
growth conditions.
The HindIII (blunted by Klenow DNA
polymerase)-BamHI fragment (934 bp) of plasmid
pPS15-P (Supply et al., 1993a), containing the
promoter region of the yeast PMA1 gene extending to 3 bp
upstream from the ATG start codon, was inserted into the EagI
site (blunted by Klenow DNA polymerase) and the BamHI site of
pRS-315, yielding cp(PMA1)pma2.
The 2-µm multicopy plasmid bearing the plant pma2 was obtained by subcloning the SacI-EcoRV fragment of cp(PMA1)pma2 between the SacI-SmaI sites of the Yeplac181 plasmid (Gietz and Sugino, 1988), yielding 2µp(PMA1)pma2.
Plasmid cp(PMA1)PMA1 was obtained by inserting the 4.65-kilobase pair HindIII (blunted by Klenow DNA polymerase)-SmaI fragment of PMA1 contained in plasmid pECPTZ-PMA1 (Supply et al., 1993a) into the unique EcoRI site (blunted by Klenow DNA polymerase) of the Ycp50 plasmid (Johnston and Davis, 1984).
The cp(GAL1)PMA1 plasmid was derived from pRS315GPMA1 ()a pRS-315
plasmid in which the entire coding sequence of PMA1 (the
3.7-kilobase pair ClaI-XbaI fragment) (Supply et
al., 1993a) is under the control of the GAL1-10 promoter. The pRS315GPMA1 BglI fragment containing LEU2, ARSH4, CEN6, and part of amp
was replaced with the corresponding BglI fragment of pRS-316 (Sikorski and Hieter, 1989),
containing URA3, ARSH4, CEN6, and the same
part of amp
.
The
YAK1 strain was obtained as follows: strain YPS14-4
(Ura) was transformed with the
cp(PMA1)PMA1 plasmid. The LEU2 plasmid, also
containing the PMA1 gene, was then lost by successive growth
cycles on a nonselective leucine-containing medium.
The YAK2 strain
was derived from the GPMA1 strain.
The latter has
the same genotype as YPS14-4 except that the PMA1 gene borne
by the pRS-315 plasmid is under the control of the GAL1-10 promoter (pRS315GPMA1). This strain therefore grows only on a
galactose-containing medium. Strain
GPMA1 was transformed
with plasmid cp(GAL1)PMA1 (URA3) and cured
of plasmid pRS315GPMA1 (LEU2) by successive growth cycles on a
nonselective leucine-containing medium.
The YAK1 strain was transformed with either cp(PMA1)pma2 or 2µp(PMA1)pma2. The YAK2 strain was transformed with 2µp(PMA1)pma2. The control strains were obtained by transformation with the pRS-315 plasmid or the Yeplac181 plasmid with no insert.
Strain YAKpma2 was obtained as follows. Strain YAK2, transformed with 2µp(PMA1)pma2, was placed on MGlu-His,Leu,Trp medium at pH 6.5, containing 5-FOA so as to cure the strain of the URA3 plasmid cp(GAL1)PMA1, containing the yeast PMA1 gene. Loss of this plasmid was checked by Southern analysis and Western immunodetection with an antibody raised against the PMA1 protein (Capieaux et al., 1993).
The protein concentration was determined by the method of Lowry et al.(1951) with bovine serum albumin used as a standard.
Figure 7: ATP-dependent fluorescence quenching of ACMA in the presence of sealed plasma membrane vesicles from yeast cells expressing only the plant pma2. Sealed plasma membrane vesicles were prepared from strain YAKpma2 (expressing only the plant pma2) and fluorescence quenching measurements were carried out as described under ``Materials and Methods,'' in the absence (A) or presence (B) of 930 µM vanadate. In A, the release of quenching after addition of carbonyl cyanide p-trifluoromethoxyphenylhydrazone has been corrected for the nonspecific effect of the cation carrier on ACMA fluorescence.
Figure 1:
The plant pma2 sustains yeast
growth. A, strains
YAK1((PMA1)PMA1)+Yeplac181 (yeast PMA1 with its own promoter + control vector without an insert,
positive control), YAK2((GAL1)PMA1)+Yeplac181
(yeast PMA1 under the GAL1 promoter + control
vector without an insert, negative control) and
YAK2+2µp(PMA1)pma2 (yeast PMA1 under the GAL1 promoter + plant pma2 under
the yeast PMA1 promoter on the Yeplac181 vector) were grown in
MGal-His,Leu,Ura,Trp liquid medium and spotted at serial 10-fold
dilutions onto solid media MGal-His,Leu,Ura,Trp (Gal) or
MGlu-His,Leu,Ura,Trp (Glu), 20 mM KHPO
buffered at pH 5.0 or 6.5. B, strains YAKpma2
(containing only the plant pma2) and YPS14-4 (control strain
expressing the yeast PMA1 under its own promoter on a
centromeric plasmid) were spread on a solid rich YGlu medium at pH 6.5
containing, in addition, either 100, 200, or 300 µg/ml hygromycin
B.
The plant pma2-expressing yeast cells were examined at
permissive and nonpermissive pH by electron microscopy. To obtain an
adequate number of cells expressing the plant pma2 at a
nonpermissive pH, we transferred 100 ml of a preculture on galactose
medium into 1 liter of glucose medium at pH 4.0. Over 20 h, only
1.5-2 divisions occurred, most likely owing to the progressively
diluted yeast H-ATPase. At this stage, 27% of the
sections displayed karmellae structures (Table 2; Fig. 2A). karmellae are proliferations of
stacked membranes around the nucleus, probably derived from the
endoplasmic reticulum (Wright et al., 1988; Schunck et
al., 1991). To study the same strain grown on glucose at a
permissive pH, we used two sets of conditions. Minimal glucose medium
buffered at pH 6.5 was inoculated at 10
cells/ml from a
galactose preculture. After 24 h, an aliquot was transferred to the
same medium (10
cells/ml) and both cultures were continued
for 16 h. In the first culture (24 h + 16 h), the cells (20
10
cells/ml) were at late stationary phase, whereas
in the second culture (16 h), the cells (17
10
cells/ml) were at late exponential phase. In both cases karmellae were observed, but differing in amount and structure
according to the age of the culture. At the earlier stage (16 h), 7% of
the cell sections displayed thin, stacked karmellae (Table 2; Fig. 2B), whereas at late
stationary phase (40 h), 20% of the cell sections showed crumpled,
slack karmellae (Table 2; Fig. 2C). In
all cases, the karmellae were gold-labeled by anti-plant PMA2
(shown in Fig. 2C for cells at late stationary phase).
Figure 2:
Electron
microscopy of yeast cells expressing the plant pma2 under
permissive and nonpermissive growth conditions. Strain
YAK2+2µp(PMA1)pma2 (expressing the yeast PMA1 under the GAL1 promoter + the plant pma2 under the yeast PMA1 promoter) was grown in 100
ml of MGal-His,Leu,Ura,Trp to a density of 40 10
cells/ml and transferred to a flask containing 1 liter of
MGlu-His,Leu,Ura,Trp at nonpermissive pH 4.0 (inoculation at 4
10
cells/ml) for a 20-h growth period (A) or to a
flask containing 1 liter of MGlu-His,Leu,Ura,Trp + 20 mM KH
PO
buffered at permissive pH 6.5
(inoculation at 10
cells/ml) for a 40-h growth (C). After 24 h of the latter culture, an aliquot was
transferred to fresh MGlu-His,Leu,Ura,Trp, 20 mM
KH
PO
(pH 6.5) (inoculation at 10
cells/ml) and grown for 16 h (B). In D, strain
YAKpma2 (containing only the plant pma2) was grown in
MGlu-His,Leu,Trp + 20 mM KH
PO
buffered at permissive pH 6.5. Cells were harvested and electron
microscopy (A-D) and immunodetection (C and D) were performed as described under ``Materials and
Methods.''
To examine the density of membranes expressing the plant PMA2, protoplasts were prepared from the three sets of cells described above, and their lysates were submitted to subcellular fractionation by centrifugation on a sucrose gradient. Equivalent material from a control strain without plant pma2, whose growth is prevented in the three growth conditions was tested for comparison.
With cells transferred to the nonpermissive pH, a large peak of ATPase activity was observed between 24 and 46% sucrose (d = 1.0990-1.2079), with a maximum at 36% (d = 1.1562) (Fig. 3A, continuous line). A similar peak was observed when the plant PMA2 was immunodetected, its maximum shifted toward a slightly lower density than the maximum observed with the control strain lacking plant pma2 (Fig. 3A, dashed line and immunodetection). At a permissive pH, however, whether the cells were grown in glucose for 16 (Fig. 3B) or 40 h (Fig. 3C), two distinct peaks of ATPase activity were seen. One peak occurred at a high density (46-52%, d = 1.2079-1.2406). For the second ATPase peak (24-34% sucrose, d = 1.0990-1.1464), its height depended on the growth stage, with approximately four times more ATPase activity after 40 h of growth. Western blot analysis showed a direct correlation between the ATPase activity and the quantity of PMA2 (Fig. 3, B and C). The ATPase activity increase observed in the light peak at late stationary phase paralleled the increase in proliferating internal membranes. This suggests that the low density fraction might contain these membranes, presumably developing from the secretory pathway. To further test this possibility, we immunodetected KAR2, a resident protein of the endoplasmic reticulum lumen (Rose et al., 1989). The distribution of this marker paralleled that of the plant PMA2 in both ATPase peaks. This suggests the higher density peak (46-52%, d = 1.2079-1.2406) probably corresponds to the endoplasmic reticulum, whose density is close to that of the plasma membrane (Antebi and Fink, 1992).
Figure 3:
Subcellular distribution of
H-ATPase in yeast cells expressing the plant pma2 under nonpermissive and permissive growth conditions. The
supernatant of a 450
g
5 min centrifugation
(S
) prepared from strains YAK2+Yeplac181 (control
without plant pma2, dashed line) and
YAK2+2µp(PMA1)pma2 (strain expressing the
plant pma2, continuous line) was layered on a
18-54% (w/w) sucrose gradient and centrifuged for 16 h at 100,000
g. After fractionation, enzymes were assayed as in the
Methods section. Growth conditions were as in Fig. 2. A,
nonpermissive pH 4.0 (20 h). B, permissive pH 6.5 (16 h). C, permissive pH 6.5 (40 h). Top panel, the
distribution of ATPase activity (
) and the sucrose percentage
(w/w) (
) are indicated. Middle panel, aliquots (100
µl for anti-plant PMA2, 5 µl for anti-yeast PMA1, or 10 µl
for anti-yeast KAR2) were analyzed by Western blotting (aliquots
numbered 15 are missing for B and C, and aliquot 13 is missing for C for the cells expressing only yeast PMA1). Bottom
panel, protein distribution.
For the control strain without the plant pma2, which does not grow under these conditions, very little ATPase activity was observed, whatever the culture time on glucose medium (Fig. 3, B and C, dashed line). Interestingly, KAR2 also appeared in the lower density fractions of these cells, after 40 h, indicating that the appearance of this peak does not depend on expression of the plant PMA2.
The generation time of
this new strain (YAKpma2) was 4 h on both minimal and rich medium. For
the control strain expressing yeast PMA1 under its own
promoter, the doubling time was 2.5 h on minimal medium and 1.5 h on
rich medium. We reproduced with YAKpma2 our previous results showing
the dependence of growth on the pH (results not shown). We also tested
the strain's resistance to hygromycin B, an aminoglycoside
antibiotic used to select yeast mutants with a partially defective
plasma membrane H-ATPase (Mc Cusker et al.,
1987). The rationale is that uptake of the drug should be less
efficient if a mutation (or, as here, the presence of a heterologous
H
-ATPase) results in a lower transmembrane
electrochemical proton gradient. And indeed, the YAKpma2 strain could
still grow in the presence of 300 µg/ml hygromycin, a
growth-inhibiting concentration for the control strain (Fig. 1B).
The results of subcellular fractionation and electron microscopy were the same for the PMA1-null strain as for the strain not yet cleared of the yeast PMA1 gene but grown on minimal glucose medium, both strains containing the plant pma2 on a multicopy plasmid; a low density ATPase peak was present (results not shown), and proliferation of karmellae was observed in stationary phase (Table 2; Fig. 2D). Gold labeling by anti-plant PMA2 revealed the presence of PMA2 in the proliferating membranes.
To prepare on a large scale membranes
intended for enzymatic characterization, we ground the cells with glass
beads and used a classic method for plasma membrane purification, i.e. shifting a crude membrane fraction (15,000 g 40 min) to a pH (pH 5.2) at which mitochondrial and
possibly other membranes become aggregated and precipitate while plasma
membranes do not (Goffeau and Dufour, 1988). The yeast cells were grown
in rich glucose medium (exponential phase), harvested, and, before
homogenization, incubated for 15 min in 250 mM glucose, known
to activate the yeast H
-ATPase (Serrano, 1983), or 250
mM sorbitol, used as a control. With cells expressing the
yeast PMA1, the specific ATPase activity was increased in the
plasma membrane-enriched fraction and activated by incubation with
glucose (Table 3), but when membranes prepared from the strain
expressing only the plant pma2 were used, glucose failed to
activate the ATPase. This could be expected since glucose activation
has been shown to involve the regulatory carboxyl-terminal region of
PMA1 (Portillo et al., 1989), whose sequence is totally
different from the corresponding region of the plant PMA2. Another
observation was more surprising; the specific ATPase activity of the pma2-expressing strain was lower in the plasma membrane
fraction than in the crude membrane preparation. This was not due to an
accumulation of PMA2 in internal membranes, since karmellae structures barely arise under the conditions used (exponential
growth in a rich medium) (Table 2). Gel electrophoresis of the
various fractions (Fig. 4A) indicated that the plasma
membrane fraction was in fact enriched in plant PMA2 (2.4-fold), albeit
less so than the plasma membrane fraction of the yeast PMA1-expressing strain in yeast PMA1 (5.6-fold). We therefore
characterized in more detail the plant PMA2, and the yeast PMA1 as a
control, in a crude membrane fraction. The electrophoretic analysis (Fig. 4A) also shows that the plant PMA2 migrated
faster than the yeast PMA1, although they have similar predicted
molecular weight. Western blot analysis with plant PMA2-specific
antibodies revealed, in the PMA2-expressing strain (Fig. 4B,
lane 3), a band with the same electrophoretic mobility as the band
revealed in a plasma membrane fraction isolated from N.
plumbaginifolia (lane 2). To rule out the possibility
that a mutation occurred in the N. plumbaginifolia pma2 expressed in yeast, we completely sequenced pma2 from the
2µp(PMA1)pma2 plasmid independently retrieved
from five YAKpma2 strains. No alteration was found. The different
electrophoretic mobility between PMA1 and PMA2 could thus be attributed
to post-translational modifications.
Figure 4:
Polypeptide composition of plasma
membranes of a yeast strain expressing the plant pma2. A, the YAKpma2 strain (expressing only the plant pma2) (lanes 1 and 3) and the YPS14-4 strain
(expressing only the yeast PMA1) (lanes 2 and 4) were grown in the rich YGlu medium at pH 6.5 to a density
of 120 10
cells/ml. A crude membrane fraction
resulting from the 15,000
g
40 min
centrifugation (P15/40, lanes 1 and 2) and a
plasma membrane fraction (PM, lanes 3 and 4)
(purified by acid precipitation of contaminating material) were
electrophoresed (70 µg) on a 7% polyacrylamide standard gel and
stained with Coomassie Blue. The 94- and 64-kDa molecular mass markers (MW) are shown in the left. PMA1 and PMA2 are
indicated. B, purified plasma membranes (20 µg) from the
control strain YAK1+pRS-315 (expressing only the yeast PMA1) (lane 1), from N. plumbaginifolia (2), and from strain
YAK1+2µp(PMA1)pma2 (lane 3) were
electrophoresed, transferred to nitrocellulose membrane, and
immunoprobed with anti-plant PMA2
antibodies.
Figure 5:
pH dependence of ATP hydrolysis by the
plant PMA2 and the yeast PMA1. Crude membranes (the pellet resulting
from a 15,000 g
40 min centrifugation) were
prepared from the YAKpma2 strain (expressing only the plant pma2) and the YPS14-4 strain (expressing only the yeast PMA1) grown in the rich YGlu medium at pH 6.5 to 120
10
cells/ml and incubated in the presence of glucose or
sorbitol as described under ``Materials and Methods.'' The
total ATP and Mg
concentrations in the assay were
calculated so as to give 6 mM [MgATP]
and 1 mM free Mg
. The assay mixture
further contained 10 mM sodium azide, 0.1 mM sodium
molybdate, 100 mM potassium nitrate, 50 mM MES, 50
mM MOPS, 50 mM Tris adjusted to the indicated pH with
HCl or KOH.
The
kinetics of the ATPases was studied in the presence of an
ATP-regenerating system, 1 mM free Mg, and a
concentration of MgATP
varying from 10.3 µM to 4.05 mM. The Michaelis-Menten curves obtained after
incubation with glucose or sorbitol were similar. The Eadie-Hofstee
plots (Fig. 6, A and B) revealed a similar K
(23.3 and 23.8 µM) and V
(0.265 and 0.232 µmol of P
min
mg
) in
both cases. Interestingly, the first two points of these plots,
corresponding to the highest MgATP
concentrations,
did not align with the others, suggesting that the kinetics of the
enzyme may be different at high ATP concentration.
Figure 6:
Kinetics of the plant PMA2. Crude
membranes were prepared from the YAKpma2 strain (expressing only the
plant pma2) grown in the rich YGlu medium at pH 6.5 to 120
10
cells/ml. and incubated in the presence of
glucose or sorbitol as described under ``Materials and
Methods.'' The ATP hydrolysis rate was measured in the presence of
[MgATP]
varying from 0.0103 to 4.05
mM, 1 mM Mg
, 10 mM sodium
azide, 50 mM MES/KOH, pH 6.8. An ATP-regenerating system (10
mg/ml pyruvate kinase and 5 mM phosphoenolpyruvate) was added
to the mixture. Data are presented as Eadie-Hofstee plots. The slope
corresponds to -K
and the intercept
on the y axis corresponds to the V
. The
linear regression is the mean of three independent assays (the values
at 1.66 and 4.05 mM [MgATP]
were
excluded from calculation), the points are the means of the three
assays, and the dashed line gives a confidence interval of
95%.
Vanadate, a
specific inhibitor of cation-transporting ATPases forming a
phosphorylated intermediate during their catalytic cycle, inhibited the
ATPase activity of the PMA2-expressing strain, with a K of 15.2 µM (sorbitol-incubated cells) or 18
µM (glucose-incubated cells) (results not shown). On the
other hand, this ATPase activity was stimulated to 169% ± 15%
(results not shown) by 300 µg/ml lysophosphatidylcholine, a
detergent-like phospholipid known to activate plant
H
ATPases in native plant membranes (Palmgren et
al., 1988) and when expressed in yeast (Palmgren and Christensen,
1994).
In order to provide final proof that the N.
plumbaginifolia PMA2 is indeed a proton pump, we studied
ATP-dependent proton translocation using purified plasma membranes
(Dufour et al., 1982) resealed by a lecithin/deoxycholate
treatment (Venema et al., 1993). The plasma membranes
collected after acid precipitation exhibited no ATP-dependent
proton-pumping activity (data not shown), but when dispersed with the
lysolecithin-deoxycholate mix, sealed plasma membrane vesicles were
obtained after centrifugation. Interestingly, the ATPase activity lost
after acid precipitation of membranes prepared from YAKpma2 (Table 3) was recovered and even increased after reconstitution
(0.51 µmol of P
min
mg
). The proton-pumping activity of the plant PMA2,
illustrated in Fig. 7, was estimated by measuring the rate of
ATP-dependent quenching of ACMA fluorescence, a well known
pH
probe. ACMA quenching was released by the proton-specific ionophore
carbonyl cyanide p-trifluoromethoxyphenylhydrazone (trace
A) and fully inhibited by vanadate before addition of MgATP (trace B). The initial rate of ATP-dependent proton pumping
was proportional to the ATPase activity measured at the optimal pH of
6.0 or 6.9 of the corresponding yeast PMA1 or plant PMA2 containing
membranes. The ratio of specific ATPase activity (5.6 µmol of
P
min
mg
for control strain and 0.51 µmol of P
min
mg
for
YAKpma2 strain) to proton pumping (initial rate of quenching, 1, 100%
quenching
min
mg
for control strain and 79% quenching
min
mg
for YAKpma2 strain) is about the
same for both enzymes, 0.0065 and 0.0051, respectively.
The key to studying N. plumbaginifolia pma2 expression in yeast was to avoid any yeast
H-ATPase expression while enabling the cells to grow.
First, the recipient strain was completely deprived of its own two PMA genes to avoid the reconstitution of a functional gene by
recombination. Next, since the plant H
-ATPase was able
to sustain yeast growth, the plasmid-borne yeast PMA1 gene
could be removed as well.
Expression of the plant PMA2 correlated with the appearance of PMA2-containing karmellae structures. Such membranes were originally observed in yeast cells overexpressing hydroxymethylglutaryl-CoA reductase by Wright et al.(1988), who hypothesized that they derive from the ER. Osmium/potassium ferricyanide fixation, which labels the ER and Golgi membranes (Rambourg et al., 1993), also stained the karmellae structures (data not shown), and the KAR2 protein, an ER marker, co-sedimented with the lower density PMA2-containing membranes proliferating in the stationary phase in a minimal medium. This led us to suggest that this low density fraction contains the karmellae.
The appearance of karmellae does not
seem to be linked directly to the level of overexpression of the N.
plumbaginifolia PMA2. For instance, we found approximately the
same PMA2 level in strain YAK2 harboring 2µp(PMA1)pma2,
whether it expressed the yeast PMA1 (galactose medium, stationary
phase) or not (glucose medium, stationary phase), yet karmellae were more abundant in the latter case by an order of magnitude. An
even greater difference was seen with the strain containing only the
plant pma2 gene; although the amount of PMA2 was similar in
exponentially growing and stationary phase cells, the karmellae contents were very different: <1% (exponential phase) and 25%
(stationary phase). The latter observation concerned cells grown in a
minimal medium; growth in a rich medium did not cause proliferation of karmellae. We therefore conclude that overexpression of the
plant ATPase is a necessary but not a sufficient condition for karmellae proliferation. When we compare the various
situations in which karmellae were observed (Table 2),
it clearly appears that these structures became abundant when the yeast PMA1 was no longer expressed (after a shift to glucose medium
or after removal of the plasmid-borne gene) and, in addition, when the
cells were in a minimal medium and not dividing (glucose medium at a
nonpermissive pH or stationary phase at a permissive pH). It seems that
the karmellae develop when or because the cells are prevented
from growing. Although not pointed out by the authors, this conclusion
can also be drawn from the results of Villalba et al.(1992)
and Palmgren and Christensen(1994) concerning the expression of A.
thaliana H-ATPases in yeast which was reported to
be located in ER-like structures; in these studies, the cells were
harvested in a minimal medium, did not express the yeast PMA1,
and, in contrast to our observation with the N. plumbaginifolia
pma2 gene, the Arabidopsis H
-ATPase
failed to complement the nonexpression of the yeast PMA1 gene.
PMA1-null strain cells whose defect was complemented by the N.
plumbaginifolia pma2 did not recover a wild-type phenotype. Growth
was prevented below pH 5.5, and at a permissive pH, the doubling time
remained longer (2.7 or 1.6 times the wild-type generation time in rich
or minimal medium, respectively). These observations probably reflect
the comparatively poorer performance of the plant PMA2 expressed in
yeast compared with the native S. cerevisiae H-ATPase. It is harder to establish a potential
difference across the plasma membrane when the external pH is more
acid, as shown by the sensitivity to low pH of yeast strains with a
partially defective H
-ATPase (Mc Cusker et
al., 1987). The analogy between these mutants and the strain
analyzed in this work is further evidenced by their common resistance
to hygromycin, interpreted in the former case as a consequence of a
decreased electrochemical proton gradient (Mc Cusker et al.,
1987).
What might cause the lower performance of the plant
H-ATPase expressed in yeast? Coupling between
H
pumping and ATPase activity seems unaltered, since
in sealed vesicles the ratio of the H
pumping activity
(as reflected by the initial rate of ACMA quenching) to the ATPase
activity is similar for the strain expressing the yeast PMA1 and the
strain expressing the plant PMA2. On the other hand, we could readily
compare the amounts of plant PMA2 and yeast PMA1 by staining the plasma
membrane proteins of cells expressing one or the other enzyme. The
plant PMA2 amounted to approximately 25% of the quantity of yeast PMA1,
the other proteins being in similar amount in preparations from both
strains. The ATPase activity ratio between the plant and yeast enzymes
was more difficult to estimate, because although the purified plasma
membrane fraction obtained after acid precipitation was richer in
immunodetectable PMA2, it exhibited a lower specific ATPase activity
than the crude membrane preparation, possibly because of the acid
treatment. Why the PMA2 content is lower is a difficult question. Any
level of expression from transcription to protein stability might be
involved. Retention of the Arabidopsis H
-ATPase in the ER was the proposed explanation
for the absence of complementation (Palmgren and Christensen, 1994),
but as discussed above, retention of the N. plumbaginifolia PMA2 in ER-derived membranes varies according to the growth stage
and growth medium, indicating that retention is rather a secondary
effect than due to an intrinsic inability of PMA2 to be brought to the
plasma membrane. Alternatively, the fraction of PMA2 that is not
correctly targeted might be degraded rapidly in active cells but
accumulate in karmellae in cells with a low catabolic
activity.
Why could N. plumbaginifolia PMA2, contrary to
the A. thaliana AHA1, AHA2, or AHA3 H-ATPases
(Palmgren and Christensen, 1994), sustain growth of a yeast strain
lacking its own PMA genes? Possibly, the pH of the minimal
medium used to test AHA complementation was too low. Growth at various
pH values has not been reported for these AHA enzymes. Two additional
properties, however, distinguish the H
-ATPases of the
two plant species. One is that N. plumbaginifolia PMA2 has a
lower K
(23 µM) than AHA1 (150
µM), AHA2 (150 µM), and AHA3 (1.5
mM) (Palmgren and Christensen, 1994), but it is unclear
whether this higher apparent affinity of PMA2 for ATP has any
physiological consequences in a cell where the cytosolic ATP
concentration is usually well above the K
. A
second difference is that the pH optimum of PMA2 is pH 6.8. This means
that at the physiological pH, this enzyme is more active than the A. thaliana AHA or S. cerevisiae PMA1 enzymes, whose
pH optima are between 6.0 and 6.5. Interestingly, a similar pH optimum
was found and complementation was observed in yeast when the A.
thaliana AHA2 was truncated of 92 residues at the carboxyl
terminus (Palmgren and Christensen, 1993). We checked that no mutation
occurred in the plant pma2 borne by the yeast strain depleted
of its own PMA1. These results might therefore indicate that
the two plant enzymes, which belong to two distinct subfamilies, have
different kinetics. As a matter of fact, reporter gene analysis
indicates that plant pma genes are differentially expressed
according to the cell type and might thus be involved in activating
different secondary transport systems (Michelet and Boutry, 1995). (
)It would be interesting in this regard to test PMA4 of N. plumbaginifolia, for which a cDNA is available and whose
close resemblance to A. thaliana AHA1, AHA2, and AHA3 might
prevent it from functionally replacing an absent yeast
H
-ATPase in yeast.
An interesting observation
concerning PMA2 kinetics is the unusual behavior of the enzyme at high
ATP concentration; the velocity clearly departs from linearity in the
Eadie-Hofstee plots, suggesting another kinetic mode at high ATP
concentration. This should be further explored. It will be necessary to
examine whether a single PMA2 enzyme displays double kinetics or
whether two populations of PMA2 with distinct kinetics exist in yeast.
The material used in this experiment came from cells grown in a rich
medium and displayed a single ATPase peak upon subcellular
fractionation on a sucrose gradient. If two populations of PMA2 exist,
they must therefore belong to membranes with identical density. Some
observations do support the existence of different states of the plasma
membrane H-ATPase. For instance, yeast PMA1 is
converted to a more active form upon incubation in the presence of
glucose (Serrano, 1983), a process probably involving phosphorylation
by protein kinases (Chang and Slayman, 1991). The sarcoplasmic
reticulum Ca
-ATPase, moreover, exists in two states
with different affinities for ATP, and conversion from one state to the
other also involves phosphorylation (Chambeil et al., 1988).
H
-ATPases from plants such as oat or maize also
exhibit complex kinetics (Roberts et al., 1991; Ramos et
al., 1994), but these results cannot be unambiguously interpreted
as they were obtained with membranes prepared from whole organs which
probably express various isoforms. The lowest K
observed in this case was 11-16 µM (Roberts et al., 1991), a figure close to that observed for PMA2 in
yeast.
In conclusion, we have obtained a S. cerevisiae strain whose plasma membrane H-ATPase is replaced
by the N. plumbaginifolia PMA2. This has enabled us to
characterize biochemically, a single plant H
-ATPase
whose expression permits cell growth. We have shown, moreover, that the
trafficking dysfunction revealed in certain circumstances by the
proliferation of karmellae structures in this strain depends
on the growth medium and growth stage. These observations should enable
us to design experiments for testing which growth-related or
growth-influencing parameters induce karmellae formation.