1 Program in Molecular and Cell Biology, Oklahoma Medical Research Foundation,
825 N.E. 13th Street, Oklahoma City, OK 73104, USA
2 Zoologisches Institut, Ludwig-Maximilians-Universität, Munich,
Germany
* Author for correspondence (e-mail: clarkem{at}omrf.ouhsc.edu )
Accepted 16 February 2002
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Summary |
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Key words: Dictyostelium, V-ATPase, Endocytosis, pH regulation
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Introduction |
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Using the soil amoeba Dictyostelium discoideum, an organism in
which genetic as well as biochemical tools can be used to explore protein
function, we are analyzing the large (100 kDa) transmembrane subunit of
the V0 domain of the V-ATPase
(Liu and Clarke, 1996
).
Although this subunit has been the subject of study in both yeast and
mammalian cells, its structure and function are not well understood. Three
genes encoding isoforms with different tissue distributions have been
demonstrated in vertebrates; one isoform may be specific to osteoclasts and
other cells that target proton pumps to plasma membranes
(Nishi and Forgac, 2000
;
Mattsson et al., 2000
). C.
elegans has four isoforms that are strongly expressed in distinct cells
(Oka et al., 2001
). In S.
cerevisiae, there are two isoforms that are localized to different
endomembranes (Manolson et al.,
1992
; Manolson et al.,
1994
). Initial results suggested that the yeast 100 kDa subunits
may act to target the enzyme complex to particular endomembranes and/or
regulate its function in different compartments
(Manolson et al., 1994
).
Site-directed mutagenesis of one of the yeast isoforms (VPH1)
confirmed that this subunit is important in enzyme organization
(Leng et al., 1998
) and showed
that it is involved in proton translocation, which suggested that it is
functionally related to the a subunit of the F-ATPase
(Leng et al., 1996
). Other
data indicate that this subunit may provide a fixed structural link between
the V1 and V0 domains, thereby acting as a stator for
the rotary motor (Landolt-Marticorena et
al., 1999
; Landolt-Marticorena
et al., 2000
).
In Dictyostelium, vacuolar proton pumps are concentrated primarily
in membranes of the contractile vacuole complex, an osmoregulatory organelle
(Heuser et al., 1993;
Fok et al., 1993
;
Bush et al., 1994
); they are
also present in lesser abundance in membranes of the endo/lysosomal
compartment (Rodriguez-Paris et al.,
1993
; Temesvari et al.,
1994
; Bush et al.,
1994
; Clarke and Heuser,
1997
). Our studies of the 100 kDa V-ATPase subunit in
Dictyostelium have detected only a single gene (vatM)
encoding this subunit and have shown that the product of vatM is
greatly enriched in contractile vacuole membranes
(Liu and Clarke, 1996
).
Biochemical fractionation experiments have shown that the same protein is also
present in endosomal membranes (Adessi et
al., 1995
). Our attempts to inactivate vatM by homologous
recombination yielded no strains in which the gene had been disrupted.
Instead, when transformants were screened by immunoblot to detect clones
lacking the 100 kDa protein, only regulatory mutants with low vatM
mRNA levels and a transient reduction in protein levels were obtained, and the
protein soon returned to normal (Liu and
Clarke, 1996
). These results suggest that the level of the 100 kDa
protein is tightly regulated and that a cell in which the gene has been fully
inactivated is probably not viable.
To examine the function of this protein, a means of conditionally blocking
expression of the vatM gene would be helpful. One possible means of
accomplishing this would be through expression of an antisense transcript.
Although this approach has been effective for several genes in
Dictyostelium, for vatM an antisense-mediated reduction in
mRNA levels had only a temporary effect on protein levels
(Liu and Clarke, 1996).
Accordingly, we developed an alternative strategy; we altered the expression
of vatM by inactivating its endogenous promoter and substituting
another promoter with a different pattern of regulation. We describe here the
consequences of the altered expression of vatM in growing
Dictyostelium cells.
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Materials and Methods |
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Construction and use of vector for promoter replacement
The vector for promoter replacement was prepared in three steps that are
summarized here. (1) From plasmid pVM2
(Liu and Clarke, 1996) we
isolated a fragment that contained 0.6 kb of vatM upstream DNA plus
10 vatM codons (i.e. up to the BglII site in the gene). This
fragment was cloned into pBluescriptSK- (Stratagene) between the
XbaI and BamHI sites. (2) The complete vatM cDNA
was recovered as a polymerase chain reaction (PCR) product from the phagemid
pVM1 (Liu and Clarke, 1996
),
using a forward primer 5'-GGATCCATGAGCTTTTTAAGACC-3' (which added
a BamHI site immediately before the ATG translation initiation
codon); the reverse primer was the M13 sequence that flanked vatM in
the phagemid. The PCR product was cloned into the BamHI site of
pDNeo67 (Da Silva and Klein,
1990
), immediately downstream of the act6 promoter. A
fragment containing the act6 promoter plus 1 kb of vatM
coding sequence (to the XhoI site in vatM) was isolated and
subcloned into the plasmid created in step 1, just downstream of the
vatM promoter. (3) The pyr5-6 gene was recovered by PCR as a
1.8 kb fragment from DIV2 (Kuspa and
Loomis, 1994
) and the gene was moved into the plasmid created in
step 2, positioned between the vatM and act6 promoters. The
final construct, pVATM-act6, is shown schematically in
Fig. 1. DH1 cells were
transformed with this plasmid using the electroporation protocol described
previously (Kuspa and Loomis,
1994
). The pVATM-act6 plasmid was cut with SacII and
XhoI to free the vatM-containing segment from the
pBluescript backbone and 40 µg of the linear vatM-containing
segment was used to transform DH1 cells. Following transformation, the cells
were plated in ten 96-well plates at a density expected to yield single clones
(approximately 200 cells/well) and cultured axenically in FM medium.
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Expression of vatM under its own promoter
From plasmid pVM2 an 0.6-kb XbaI/BglII fragment was
isolated consisting of genomic DNA upstream of vatM plus the first
few nucleotides of coding sequence (as described above). This fragment was
used to replace the corresponding XbaI/BglII fragment of the
vatM cDNA in pVM1. Finally, a 2.1 kb XbaI fragment
containing the G418 resistance gene TN903 flanked by the act15
promoter and terminator was recovered from pDNeo2
(Witke et al., 1987) and
cloned into the XbaI site of this plasmid, yielding pVMop, a
G418-selectable transformation vector containing vatM driven by its
own promoter. This vector was used to transform VatMpr cells, and
transformants were selected in HL5 containing 5 µg/ml G418 (Geneticin,
Sigma).
Construction and use of the A15-pHluorin vector
Ratiometric pHluorin (Miesenböck
et al., 1998) (a gift of Dino De Angelis, Memorial Sloan-Kettering
Institute, New York, NY) was received as coding sequence in the bacterial
expression vector pGEX-2T. The downstream EcoRI site was converted to
a XhoI site by insertion of the oligonucleotide AATTTTAACTCGAGTTAA,
which also created a stop codon in the pHluorin reading frame. The pHluorin
gene was then excised with BamHI and XhoI, and combined with
the A15-promoter from A15-UGus (Zaccaria
et al., 1998
) and the A8-terminator from pDdgal-17
(Harwood and Drury, 1990
) in
the V18Tn5-DRE Dictyostelium selection vector
(Wetterauer et al., 1996
;
Deichsel et al., 1999
). In the
predicted protein, there are 13 N-terminal amino acids (MDGEDVQARSTG) in place
of the wild-type-GFP initiation methionine, most of which arise from A15
coding sequence, while four amino acids (TGEF) at the C-terminus are derived
from pGEX2T, and four further C-terminal amino acids present in the De Angelis
construct are deleted. AX2, DH1 and VatMpr cells were transformed with this
vector as described above, and transformants were selected as described
[(Wetterauer et al., 1996
)
agar method] using living K. aerogenes and a G418 concentration of
100 µg/ml. The selection plates were examined directly with fluorescent
illumination, and transformants with uniform GFP expression were chosen. The
cells were cultured in HL5 with 20 µg/ml G418 until required for
experiments.
Detection of vatM DNA, mRNA and protein
Methods for preparing and analyzing DNA were as previous described
(Liu and Clarke, 1996), except
that the probe was the 1 kb EcoRI/XhoI fragment from pVM1
described in that report. For analysis of protein in cell lysates, the methods
of sample preparation, electrophoresis and transfer were as previously
described (Liu and Clarke,
1996
), except that nitrocellulose membrane (Schleicher and
Schuell) was used instead of PVDF. After transfer, the membrane was cut in
half using prestained molecular weight markers as guides. The upper portion
was stained with N2 and sometimes also N4 hybridoma culture supernatants
(1:100 and 1:30 dilutions, respectively) and the lower portion with anti-actin
antibodies (C-4, 1:2000), as previously described
(Liu and Clarke, 1996
). The
secondary antibody was alkaline phosphatase-conjugated anti-mouse IgG
(Promega, 1:1000 dilution), used as directed by the manufacturer.
Indirect immunofluorescence
Dictyostelium cells of strains DH1, VatMpr, and VMop were grown in
suspension in association with K. aerogenes for 2 or 3 days before
being examined. Samples were taken from low density, exponentially growing
cultures and washed free of bacteria by three cycles of centrifugation in 17
mM Na2H/KH2PO4 buffer (pH 6.4). After
agaroverlay, the cells were fixed in the same buffer containing 2%
formaldehyde (5 minutes, room temperature), followed by 1% formaldehyde in
methanol (5 minutes, -15°C). They were stained with N2 or N4 hybridoma
culture supernatants (Fok et al.,
1993) diluted 1:30 and 1:20, respectively, followed by
FITC-conjugated donkey anti-mouse IgG (1:500, Jackson ImmunoResearch
Laboratories) (for details, see Clarke et
al., 1987
).
Assays for phagocytosis
Phagocytosis of fluorescent yeast particles was measured essentially as
described (Maniak et al.,
1995). Fluorescent yeast particles were prepared by suspending 5 g
of yeast (Sigma YSC-2) in 50 ml PBS (150 mM NaCl in 20 mM
Na2H/KH2PO4, pH 7.2); the suspension was
stirred for 30 minutes in a boiling water bath, then washed five times in PBS
and twice in KP. A pellet of 2x1010 yeast particles was
resuspended in 20 ml Na2HPO4 (50 mM, pH 9.2) containing
2 mg tetramethylrhodamine isothiocyanate (TRITC, Sigma T3163, Isomer R) and
incubated for 30 minutes at 37°C. The yeast particles were washed twice in
the pH 9.2 phosphate buffer, then four times in KP buffer, and were stored at
-20°C. The uptake of yeast particles was measured using cells that had
been growing on a suspension of K. aerogenes for 2-3 days prior to
the assay. The cells were collected from log phase cultures, washed three
times in KP, and resuspended at 2x106 cells/ml in KP
containing 10% LF medium. The cells were then incubated on a rotary shaker for
30 minutes, and fluorescent yeast particles were added
(1.2x107 yeast particles/ml). Duplicate 1 ml samples were
taken at 20 minute intervals; each was added to a tube containing 0.1 ml of
trypan blue (2 mg/ml), which quenched the fluorescence of uningested particles
(Maniak et al., 1995
). The
samples were mixed by inversion for 1 minute, then centrifuged (500
g, 4 minutes). The pellets were resuspended in 1 ml KP, and emission
spectra were determined immediately in an SLM 8000C spectrofluorimeter
(
ex=544 nm,
em=576 nm).
Phagocytosis of the fluorescent yeast particles and endocytosis of
fluorescent dextrans were also monitored by confocal laser scanning microscopy
using a Zeiss LSM510 equipped with a 100x 1.4 NA Plan-Neofluar
objective. The 543 nm line of the HeNe laser was used for excitation of
TRITC-labeled yeast or dextran, and the 488 nm line of the Argon laser for
excitation of FITC-dextran. Cells that had been growing in LF were plated in
one-third strength LF in dishes with coverslip glass bottoms (WillCo-Dish,
WillCo Wells, The Netherlands). Bacterially grown cells were washed as
described above, then plated in one-tenth strength LF (to maintain osmotic
conditions similar to those in which the cells had been growing and to avoid
triggering development) and incubated for 30 minutes at room temperature. For
measuring endosomal transit time, a mixture of TRITC- and FITC-labeled
dextrans (2 mg/ml and 0.2 mg/ml, respectively, average Mr
70,000; Sigma) was added (Jenne et al.,
1998; Maniak,
1999
). For phagocytosis experiments, TRITC-yeast particles were
added in one-tenth strength LF thirty minutes after the cells had been plated.
The cells were then covered with a thin layer of agarose, excess fluid was
removed, and the cells were observed using low laser power (10%).
Confirmation of the cytosolic location of pHluorin
Cells of AX2/A15-pHluorin, grown overnight in LF medium, were allowed to
settle on coverslips and examined from below at 100x magnification using
epifluorescent illumination (ex=405 or 480 nm,
em=520 nm). At both excitation wavelengths the cell edge
was sharply defined with conspicuous bright filopodia. Agar-overlayed cells
presented a `Swiss cheese' appearance with abundant dark vesicles. Ratio
images showed an essentially uniform intracellular pH, with the only
variations attributable to cellular motion between blue and UV exposures (data
not shown).
Spectral acquisition with axenically growing cells
Cells were cultured overnight in LF medium that had been supplemented with
0.7 g/liter yeast extract. Cells grew with a normal doubling time in this
medium, but since the saturation density was reduced to about
3x106 cells/ml from the normal value of
2x107 in HL5, cells were harvested at 1x106
or below. For measurement the cells were resuspended in LF medium without
either peptone or yeast extract; this medium still contains 50 mM glucose, so
the cells would not be expected to exit the growth phase into development for
several hours (Marin et al.,
1980). Fluorescence spectra were taken using
ex=350 to 490 nm,
em=510 nm. The cells
were again collected by centrifugation, and a second spectrum recorded from
the supernatant; this was then subtracted from the spectrum of the cell
suspension.
Spectral acquisition with bacterially grown cells
Cells were harvested after culture in bacteria for at least 48 hours,
during which time the cell density was not allowed to exceed
1x106 cells/ml. The cells were collected by centrifugation
and washed three times in PB6 (20 mM Na/K phosphate, pH 6.0, containing 10
µM CaCl2 and 200 µM MgCl2), then suspended in the
buffer described above. (We were not able to work with cells directly in
bacterial suspension because of light absorption and scattering by the
bacteria and the presence of fluorescent bacterial secretion products.) The
fluorescence spectrum was recorded and the supernatant spectrum subtracted as
above.
Spectral analysis
The pHluorin excitation spectrum has two peaks: one at 400 nm and the other
at 475 nm, whose ratio depends on pH
(Miesenböck et al.,
1998). To obtain pH values from experimental spectra, the ratio of
peak heights was compared to a calibration table. To suppress noise in the
experimental spectra these were first fitted as the sum of two Gaussian
curves, using the `Solver' feature of Microsoft Excel (97) to fit values of
the mean, standard deviation and height with the sum of squared deviations
between measurement and model as the goodness-of-fit criterion. The peak
heights used for pH measurements were then determined from the fitted spectra
using a calibration table. To create this table, we first prepared calibration
spectra using cell extracts from AX2/A15-pHluorin cells that had been grown in
LF medium and developed overnight in PB; comparison with similarly treated,
untransformed AX2 showed that cellular autofluorescence is a negligible
component of the total fluorescence in this material. The cell extracts were
prepared in 20 mM Na/K phosphate buffer and titrated to round values of pH.
The spectra differed slightly from the published curves, perhaps reflecting
differences in the N- and C-termini of our construct; in particular, the 400
nM excitation peak at pH 7 is more nearly intermediate between the peaks at pH
6.5 and pH 7.5. We fitted our calibration spectra with paired Gaussian curves
as above, and the ratios of the fitted curve heights were expressed as a
function of pH. Finally, the last-mentioned curve was itself fitted with a
sigmoid curve, whose value was calculated at 1000 points to construct the
calibration table.
In bacterially grown cells, cellular autofluorescence was evident as a peak
or shoulder on the pHluorin excitation spectrum centered at
ex=360 nm,
em=510 nm. We corrected for
this by subtracting the fluorescence of a matched suspension of identically
treated untransformed AX2 cells. In some experiments an emission spectrum was
recorded at
ex=390 nm,
em=410-550 nm; in
this case cellular autofluorescence was evident as a peak that appeared to be
centered below 410 nm. When emission spectra were available, they were used to
estimate autofluorescence directly, and the result was used to correct the
excitation spectra. The cytosolic pH was determined in all cases from
corrected spectra.
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Results |
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A change in restriction fragment size was expected if homologous recombination between chromosomal vatM and the transforming DNA led to the insertion of the uracil marker plus the act6 promoter between vatM and its own promoter (Fig. 1B). Twenty-two tranformants were obtained and analyzed by Southern blot. A single transformant manifested the larger BclI fragment size consistent with the desired double crossover (Fig. 1C). This clone was designated VatMpr (for promoter replacement).
Effect of growth conditions on the level of VatM protein in VatMpr
cells
The amount of the 100 kDa V-ATPase subunit, the protein product of
vatM, was compared in VatMpr cells grown in liquid medium (HL5) and
on bacteria. Axenically growing VatMpr cells were harvested during mid log
phase growth. Cells growing in association with K. aerogenes on
nutrient agar plates were also collected during exponential growth, prior to
any visible clearing of the bacterial lawn, and the cells were washed free of
bacteria. The lysate of an equal number of cells from each growth condition
was electrophoresed on a polyacrylamide gel, and the level of the 100 kDa
V-ATPase subunit was examined by immunoblot. The upper portion of the blot was
stained with N2 monoclonal antibodies, which specifically recognize the 100
kDa subunit of the V-ATPase (Fok et al.,
1993). The lower portion of the blot was stained with anti-actin
antibodies, as a control for the amount of protein loaded in each lane. The
results are shown in Fig. 2A
for VatMpr and DH1. In the parental DH1 cells, growth conditions had no
significant effect on the level of the 100 kDa protein. However, in VatMpr
cells, the level of this V-ATPase subunit was much lower in cells grown on
bacteria than those grown axenically. Phosphorimager analysis indicated that
the amount of the 100 kDa subunit present in bacterially grown VatMpr cells
(normalized for actin levels) was 29% of that present in axenically grown
cells.
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Time course experiments using suspension cultures of cells shifted from axenic to bacterial growth conditions showed that approximately 2 days of growth on bacteria were required to produce the maximal reduction in VatM protein levels. After 2 days, the level of VatM protein in VatMpr cells was typically one-third that of control levels (range 29-37%).
Growth properties of VatMpr cells correlate with the level of
expression of vatM
We examined the consequences of the reduced level of the 100 kDa subunit on
growth rate and other properties of VatMpr cells. When cultured axenically,
DH1 and VatMpr cells grew at the same rate, with doubling times of
approximately 12 hours. However, when cultured with K. aerogenes on
nutrient agar plates, VatMpr cells cleared the bacterial lawn much more slowly
than did DH1 (Fig. 3). A
similar result was observed for cells grown on a suspension of bacteria. Under
these conditions, the doubling time was about 8 hours for VatMpr compared with
3 hours for DH1 cells. To determine whether the reduced level of the 100 kDa
subunit was responsible for the poor growth of VatMpr cells on bacteria, we
tested whether restoration of normal levels of this subunit would restore
normal growth. We constructed a plasmid that contained the vatM
coding region plus 0.6-kb of genomic DNA upstream of vatM,
presumed to include the endogenous promoter for this gene. After addition of a
G418 drug resistance cassette, this plasmid was transformed into VatMpr cells.
Several transformants were obtained that grew at normal rates on bacteria. In
Fig. 3, the growth of one of
these (VMop-11) on a bacterial lawn is compared with that of VatMpr and DH1.
This figure shows the size of plaques formed by each strain on a lawn of
K. aerogenes after 5 days and 6 days of growth. The very slow growth
rate of VatMpr is evident from its tiny plaque size
(Fig. 3A,B); addition of
vatM driven by its own promoter was sufficient to restore normal
growth (compare VMop-11 in Fig.
3E,F to DH1 in Fig.
3C,D). Immunoblot analysis confirmed that the level of the 100 kDa
subunit had been restored to normal in VMop-11 cells
(Fig. 2B). These data indicate
that the reduced level of the 100 kDa subunit in VatMpr cells was sufficient
to account for the slow growth of these cells on bacteria.
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The effect of reduced VatM levels on the distribution and
accumulation of VatA
Monoclonal antibodies specific for two different subunits of the
Dictyostelium V-ATPase were used to examine the effects of the VatMpr
mutation on the V-ATPase enzyme complex. N2 antibodies recognizing VatM and N4
antibodies recognizing VatA (subunit A of the peripheral V1 domain,
responsible for ATP hydrolysis), were used to visualize V-ATPase distribution
in bacterially grown VatMpr and DH1 cells
(Fig. 4). In normal cells, the
staining patterns of these two antibodies are indistinguishable; they both
stain membranes of the contractile vacuole complex
(Fok et al., 1993). The
VatM-specific N2 antibodies gave only faint staining of VatMpr cells, in
agreement with the immunoblot results shown in
Fig. 2A. The cell population
was somewhat heterogeneous with respect to staining intensity, but all VatMpr
cells were very weakly labeled compared with DH1
(Fig. 4B versus 4A). Where
staining was visible, N2 antibodies were associated with contractile vacuole
membranes.
|
In contrast, N4 antibodies revealed a striking difference in the distribution of the A subunit in mutant versus control cells. In DH1 cells, these antibodies labeled membranes of the contractile vacuole complex in the usual manner (Fig. 4G). However, in bacterially grown VatMpr cells, N4 staining was mostly diffuse and cytoplasmic (Fig. 4H). Labeling of contractile vacuole membranes could still be detected in some cells (as expected owing to residual VatM in VatMpr cells), but this pattern was largely obscured by bright cytoplasmic fluorescence. Thus, in the absence of normal amounts of the 100 kDa subunit, much of the A subunit became mislocalized to the cytosol.
VatMpr cells were also examined under conditions in which normal levels of the 100 kDa subunit were present, that is, when growing axenically or after being transformed with the plasmid carrying vatM under the control of its own promoter. Cells of VatMpr and its transformant VMop-11, both harvested after growth on bacteria, were stained with N2 and N4 antibodies. In bacterially grown VMop-11 (Fig. 4I) and in axenically grown VatMpr (not shown), N2 and N4 antibodies stained contractile vacuole membranes, just as in wild-type cells. Thus, when approximately normal levels of the 100 kDa subunit were restored, mislocalization of the A subunit did not occur.
The degree of N4 staining in bacterially grown VatMpr cells (Fig. 4H) suggested that A subunit levels were not reduced, and, indeed, might be elevated, as a consequence of the 100 kDa subunit deficiency. To examine this possibility, we grew DH1 and VatMpr cells either axenically or for 2 days on a suspension of K. aerogenes, then subjected equal aliquots of the four cultures to electrophoresis and immunoblotting. The blot was stained to detect VatM, VatA and actin (Fig. 5). Analysis of staining intensity using Scion Image software showed that the total amount of A subunit (normalized to the value of actin) was one-third greater in bacterially grown VatMpr than in DH1 or axenically grown VatMpr cells.
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The contractile vacuole system in VatMpr cells
Vacuolar proton pumps are heavily concentrated in membranes of the
contractile vacuole complex of Dictyostelium, where they are
responsible for the accumulation of water by this osmoregulatory organelle
(Heuser et al., 1993;
Bush et al., 1994
;
Clarke and Heuser, 1997
).
Electron microscopy of freeze-dried cells shows the V1 domains of
the V-ATPase as a dense array of `studs' on the cytoplasmic surface of
contractile vacuole membranes (Heuser et
al., 1993
). Such images are shown in
Fig. 6 for elements of the
contractile vacuole system from DH1 and VatMpr cells examined after two days
of growth on bacteria. In agreement with the immunostaining data reported
above, the density of proton pumps in contractile vacuole membranes of VatMpr
cells was found to be substantially reduced relative to that in control
cells.
|
The possibility of impaired osmoregulatory function in VatMpr cells was
tested by subjecting these cells to osmotic shock conditions that have been
shown to kill Dictyostelium mutants defective in osmoregulation
(Schuster et al., 1996).
Mutant and parental cells were grown on a suspension of K. aerogenes
for 2 days prior to osmotic challenge. Exponentially growing cultures at a
density of 1x106 cells/ml were split in half, and sorbitol
(final concentration 0.4 M) was added to one-half of each culture. After 2
hours, samples were taken from each culture and diluted 100-fold and 1000-fold
with KP buffer; 10 minutes later, aliquots were plated to determine viable
titer. No significant differences in viable titer were found between mutant
and parental cells or between cells that had and had not been subjected to
osmotic shock. Thus, the reduced levels of the 100 kDa subunit present in
mutant cells did not significantly impair their ability to survive osmotic
shock.
Contractile vacuole filling and emptying was also monitored in living cells
by microscopy. Bacterially grown VatMpr cells, observed under hypotonic
conditions using either interference reflection microscopy or styryl dyes
(Heuser et al., 1993),
revealed an apparently normal, albeit somewhat sluggish, contractile vacuole
cycle (J. Heuser, personal communication). Thus, VatMpr cells manifested no
obvious impairment in osmoregulatory function.
Endo/lysosomal function in VatMpr cells
The slow growth of VatMpr cells on bacteria, evident in
Fig. 3, raised the possibility
that endo/lysosomal function might be affected. Accordingly, the endocytic and
phagocytic capabilities of mutant and control cells were examined. For these
experiments, cells were grown either axenically or for two days on bacteria
prior to analysis. AX3 cells were used as controls in addition to (or instead
of) DH1 cells in these experiments, so that wild-type and mutant cells could
be analyzed in the same growth medium (i.e. without the 100 µM uracil
required by DH1). During assays for both endocytosis and phagocytosis, care
was taken to keep the cells in osmotic conditions similar to those in which
they had been growing. We found that if cells were shifted from growth on
bacteria (suspended in 17 mM phosphate buffer) to the much greater osmotic
strength of HL5 or LF medium, endocytosis and phagocytosis rates were
significantly depressed for an hour or more, until the cells had time to adapt
to the new conditions (data not shown).
Endosomal transit time was monitored using an assay developed by Maniak and
co-workers (Jenne et al.,
1998; Maniak,
1999
). In this assay, cells are fed a mixture of TRITC and
FITC-labeled dextrans (10:1 ratio) and monitored by fluorescence microscopy.
FITC fluorescence is quenched at acidic pH, so endo/lysosomes appear red
throughout most of their cycle. However, late in the pathway, prior to
exocytosis of indigestible material, the pH of an endosome rises, restoring
the FITC fluorescence and changing the color of the endosome from red to
yellow. Thus, the time of appearance of yellow endosomes in a cell population
is a convenient indicator of endosomal transit time.
Cells from exponentially growing axenic or bacterial Dictyostelium cultures were plated on glass coverslips in low fluorescence (LF) nutrient medium (see Materials and Methods for details). At T0, they were fed a FITC/TRITC-dextran mixture and observed by confocal laser scanning microscopy. For all axenically grown cells (AX3, DH1 and VatMpr), yellow endosomes first appeared after about 1 hour (range 52-68 minutes in four experiments). For bacterially grown AX3 and DH1 cells, similar timing was observed (range 65-75 minutes). However, for bacterially grown VatMpr cells, orange or yellow endosomes first appeared after about 2 hours (range 110-140 minutes in five experiments). Thus, the endosomal transit time for bacterially grown VatMpr cells was approximately twice as long as that of control cells or axenically grown VatMpr cells.
To monitor phagocytosis, we labeled heat-killed S. cerevisiae with
TRITC and fed the TRITC-yeast to Dictyostelium cells, as described
(Maniak et al., 1995). For
cells growing in suspension, uptake was monitored by harvesting aliquots at 20
minute intervals after yeast were added, quenching the fluorescence of
undigested particles and determining the cell-associated fluorescence
spectrofluorimetrically (see Materials and Methods for details). For
bacterially grown cells, the rate of uptake by VatMpr cells was one-half that
of AX3 cells (Fig. 7).
|
Uptake of TRITC-yeast was also monitored by fluorescence microscopy of cells plated on coverslips. The behavior of AX3 cells and axenically grown VatMpr cells was similar; after 30-40 minutes, most cells in the population had ingested one or two yeast particles (Fig. 8A,C). For bacterially grown VatMpr cells, only rare cells had ingested more than a single yeast particle. Even after 70 minutes, many VatMpr cells contained no yeast, although yeast particles were enriched at the surface of the cells (Fig. 8B), suggesting that binding was not impaired.
|
Cytosolic pH in VatMpr cells
Ratiometric pHluorin, a pH-sensitive variant of GFP
(Miesenböck et al.,
1998), was used to examine VatMpr for possible alterations of
cytosolic pH or its regulation. For axenically growing cells, cytosolic pH was
7.55±0.07 (n=10) for AX2/A15-pHluorin and 7.55±0.08
(n=10) for VatMpr/A15-pHluorin. For cells grown on bacteria, we
obtained values suggesting that the cytosolic pH is unaltered in AX2
(7.6±0.24//0.11) but slightly reduced in VatMpr (7.3±0.18//0.06)
(mean±s.d. //s.d.m.). Although this difference is formally significant
at the 1% level, we view it with caution, since the expression of pHluorin was
much reduced in cells growing at low density on bacteria, presumably owing to
the lowered activity of the A15 promoter
(Cohen et al., 1986
).
Autofluorescence and bacterial fluorescence, which were negligible in
comparison with the signal from axenic cells, made significant contributions
to the spectra of cells cultured on bacteria. Corrections for these were
necessarily imperfect, resulting in significantly larger standard deviations
for measurements made on bacterially grown cells. Thus, our results suggest
but do not conclusively establish a decrease in cytosolic pH on the order of a
few tenths of a pH unit for bacterially grown VatMpr cells.
We next explored the possibility that VatMpr cells might be maintaining
their cytosolic pH with the help of mechanisms normally used only under acid
growth conditions. We probed for this situation in the two strains by
measuring the dynamic responses of the cytosolic pH to increased acid loading.
Bacterially grown cells were washed and suspended in PB6, and the fluorescence
at the `acid' pHluorin peak (ex=475 nm,
em=510 nm) was monitored continuously while acetate was
added to the medium without a change in extracellular pH. In bacterially grown
AX2, the fluorescence increased smoothly with time, indicating a decrease in
pH with roughly inverse-exponential kinetics and an estimated half-time of 70
seconds. Bacterially grown VatMpr initially acidified in a manner similar to
AX2, but the curve abruptly flattened, reaching a plateau at 30-45 seconds
(Fig. 9A).
|
This type of very rapid pH stabilization is characteristic of cells that have been growing under conditions of acid stress, as illustrated in Fig. 9B. In this experiment we analyzed AX2 cells that had been grown overnight in LF medium at pH 7.0, then either left in this medium or transferred to LF medium buffered to pH 5.0. Four hours later, the two cultures were subjected to acetate challenge as described above. AX2 cells that had been grown at pH 5.0 exhibited very rapid pH regulation, closely resembling the behavior of bacterially grown VatMpr.
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Discussion |
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Several laboratories have attempted antisense mutagenesis of
Dictyostelium V-ATPase subunits using an inducible promoter,
discoidin-I, to drive expression of the antisense transcripts
(Liu and Clarke, 1996
;
Xie et al., 1996
;
Davies, L. et al., 1996
). This
approach was ineffective in altering VatM protein levels
(Liu and Clarke, 1996
) but
reductions in Ca2+ transport and acidic vesicle content were
detected in vatP antisense cells
(Xie et al., 1996
). Further,
poor development was reported for antisense strains of vatP
(Xie et al., 1996
) and
vatB (Xie et al.,
1996
); Davies, L. et al.,
1996
). However, each group found that the ability to induce the
antisense effects was lost over time. Moreover, even initially, a mixture of
affected and unaffected cells is to be expected, since transcripts regulated
by the discoidin-I
promoter are not uniformly expressed in a cell
population (Clarke and Gomer,
1995
).
As an alternative means of exploring the function of the 100 kDa
transmembrane subunit of the Dictyostelium V-ATPase, we altered the
regulation of the chromosomal copy of vatM by replacing its promoter.
Our choice of the act6 promoter for this purpose was based on the
observations of Knecht and Loomis, who generated myosin-deficient
Dictyostelium cells by expressing an antisense transcript of the
myosin heavy chain gene under the control of the act6 promoter
(Knecht and Loomis, 1987).
They observed high levels of the antisense transcript (and a mutant phenotype)
when the cells were grown axenically, but not when they were grown on
bacteria. Thus, expression of a transcript driven by the act6
promoter appeared to be dependent on growth conditions. Furthermore, although
an increase in act6-regulated transcription was detected at the onset
of development (Romans et al.,
1985
; Knecht and Loomis,
1987
), there was a steep decline in act6 mRNA in the
later stages of development (McKeown and
Firtel, 1981
; Romans et al.,
1985
), which suggested that a transcript controlled by the
act6 promoter would be absent in late development. Both of these
features seemed useful for our analysis of VatM.
Substitution of the act6 promoter for that of vatM in VatMpr cells was confirmed by the expected changes in the restriction map of the gene as assessed by Southern blot, and by the diminished abundance of the VatM protein in bacterially grown cells. Although not shown here, it was also confirmed by the pattern of vatM mRNA accumulation during development, which matched that of act6 rather than wild-type vatM (T.L. and M.C., unpublished). However, the correct double crossover leading to promoter replacement was not a frequent event; only one such mutant was found among 40 clones from two transformations that were analyzed by Southern blot.
Effect of reduced levels of VatM on proton pump assembly
In bacterially grown VatMpr cells, immunostaining revealed that the A
subunit of the V-ATPase was mostly cytoplasmic rather than membrane
associated, and electron microscopy showed that the density of proton pumps in
contractile vacuole membranes was reduced. These results indicate that
assembly of the V-ATPase enzyme complex is limited by availability of the 100
kDa transmembrane subunit. A related finding was reported by Manolson and
co-workers, who disrupted the VPH1 gene in S. cerevisiae and
demonstrated by cell fractionation that the peripheral nucleotide-binding
subunits of the V-ATPase became mislocalized
(Manolson et al., 1992).
Similarly, certain mutations introduced into VPH1 by site-directed
mutagenesis impaired assembly of the enzyme complex
(Leng et al., 1996
). We
conclude that in Dictyostelium, as in yeast, the 100 kDa subunit
plays an essential role in V-ATPase localization and assembly.
Our data also showed that an elevated amount of A subunit protein was
present in cells containing reduced levels of the 100 kDa subunit. Thus, the
synthesis and/or stability of V1 components does not depend upon
the availability of sufficient V0 domains to allow assembly of the
holoenzyme. Instead, our results suggest that Dictyostelium cells
have a means of monitoring the amount of functional V-ATPase present in the
cell and adjusting expression of the subunits accordingly. In VatMpr cells,
this homeostatic mechanism is deranged because vatM expression cannot
be properly coordinated with that of the other subunits. However, it is
noteworthy that VatM subunit levels remain as high as they do, since other
studies have shown that the act6 promoter has very little activity in
bacterially grown cells (Wetterauer et
al., 1996; Souza et al.,
1998
). Strong post-transcriptional regulation of VatM was also
evident on our earlier attempts to alter protein levels using anti-sense RNA
(Liu and Clarke, 1996
).
Use of a single isoform of VatM in contractile vacuoles and
endosomes
Multiple isoforms of the 100 kDa V-ATPase subunit have been detected in
organisms ranging from yeast to mammals and their localization in distinct
cell populations suggests that they possess specialized functions (see
Introduction). PCR-based efforts to detect additional isoforms in
Dictyostelium were not successful
(Liu and Clarke, 1996),
although the possibility that another isoform(s) exists cannot be ruled out
until the sequencing of the Dictyostelium genome has been completed.
However, the results of the present study are consistent with biochemical
evidence that the same VatM subunit is present in both contractile vacuole and
endosomal membranes (Adessi et al.,
1995
; Clarke and Liu, 1996). Furthermore, although the two
organelles are physically separated in the cell
(Gabriel et al., 1999
),
mutations in the clathrin heavy chain
(O'Halloran and Anderson,
1992
; Ruscetti et al.,
1994
) and in a Rab4-like GTPase
(Bush et al., 1996
) affect both
compartments in a manner suggestive of a membrane trafficking relationship
between them. Additional confirmation that the same VatM subunit is present in
both contractile vacuole and endosomal membranes has come from imaging of the
dynamics of VatM in living cells, visualized by expression of green
fluorescent protein fused to VatM (M.C., J. Köhler, Q.A., T.L. and G.
Gerisch, unpublished). The presence of this subunit in two compartments whose
membranes differ so greatly in proton pump density demonstrates that the
abundance of proton pumps in a particular endomembrane is not determined by
the isoform of the 100 kDa subunit that resides there.
Effect of reduced levels of VatM on cell growth and endocytic
function
VatMpr cells grew slowly on bacteria and manifested clear endocytic defects
under restrictive conditions. Both the rate of particle uptake by phagocytosis
and the transit time of fluorescent dextrans through the endo/lysosomal
pathway were affected by a factor of about two.
Light microscopy revealed that some cells in both wild-type and mutant cell populations did not take up particles (or fluid, in the case of fluorescent dextrans) during a given time interval, although a cell inactive during one time interval could be active during another. Such cell-to-cell variation within a population may reflect a dependence of endocytic activity on the cell cycle, although this possibility has not been explored. Whatever the basis of this variation, the bacterially grown VatMpr population contained a larger fraction of inactive cells (as illustrated in Fig. 8), contributing to the reduced rate of particle uptake. However, uptake was not the only activity affected. Endosomal transit time was also prolonged, a property measured not as a population average but as the earliest appearance of yellow endosomes in individual cells. Thus, the processing of endocytosed material was slower, even in the endocytically active fraction of the VatMpr population.
There are several ways in which the VatMpr mutation could slow endocytic
activity. The number of proton pumps delivered to endosomes might be reduced
as it is in the contractile vacuole system, or their rate of delivery might be
slower, with possible consequences to acidification kinetics. However, the
lumenal pH of organelles is not determined uniquely by pump number but rather
by a complex interplay of proton pumps with ion channels and transporters
(Futai et al., 1998;
Grabe and Oster, 2001
).
Consequently, there is no direct relationship between pump number and pH (as
the contractile vacuole system also makes evident). Another step that might be
affected is the delivery of lysosomal enzymes from the Golgi to
endo/lysosomes; this and other types of membrane trafficking also depend on
the V-ATPase (Gueze et al., 1983; Mellman,
1996
). Finally, there may be general effects on the efficiency of
energy utilization in mutant cells, as discussed below.
Effect of reduced levels of VatM on cytosolic pH regulation
Possible effects of altered VatM expression on cytosolic pH were examined
with `ratiometric pHluorin'
(Miesenböck et al.,
1998), a pH-sensitive GFP variant. We found that this probe gives
intracellular pH values similar to those estimated using null-point lysis
(Aerts et al., 1985
;
Coukell et al., 1997
) and NMR
(Satre et al., 1989
), and
allows excellent compartmental and temporal resolution. Under axenic growth
conditions, the cytosolic pH of VatMpr was indistinguishable from that of the
wild-type control. During growth on bacteria, VatMpr appeared to have a
slightly lowered cytosolic pH, while that of AX2 seemed to be unchanged. These
results, while statistically significant, should nevertheless be viewed with
caution owing to technical difficulties in collecting spectra from bacterially
grown cells. It is important to note that even a small drop in pH could have
important effects on growth, as a decrease of 0.2 pH units in the
Dictyostelium cell cycle is associated with a threefold decrease in
the rate of protein synthesis (Aerts et
al., 1985
), and lowering cytosolic pH by comparable amounts quite
generally inhibits cell cycle entry in cultured metazoan cells
(Grinstein et al., 1989
).
When we examined the dynamic regulation of cytosolic pH in bacterially
grown VatMpr and AX2 cells, we found clear evidence of changes in VatMpr. In
AX2 subjected to a sudden increase in acid loading, the cytosolic pH decreased
asymptotically with a half-time of about 70 seconds. The acidification curve
for bacterially grown VatMpr, although initially very similar, showed an
abrupt plateau at about 45 seconds. This suggests that the mutant cells employ
a mechanism for pH homeostasis that is not used by cells containing normal
levels of vacuolar proton pumps. In particular, it appears that bacterially
grown VatMpr cells make use of a system that is inactive in wild-type cells
under standard conditions, but can be induced in the latter cells when these
are grown in acid media. The system appears to require 30 seconds to 1 minute
after acid challenge to become fully active, so that the acidification curve
initially appears identical to that of uninduced cells, but then suddenly
flattens. In S. cerevisiae, the V-ATPase plays a major role in
cytosolic pH homeostasis (Nelson and
Harvey, 1999; Forgac,
2000
), but a P-type H+- ATPase in the yeast plasma
membrane assists in pH regulation (reviewed by
Portillo, 2000
). This enzyme,
Pma1p, is the major plasma membrane proton pump and is essential for
viability. It is activated, probably by phosphorylation
(Goosens et al., 2000
), when
cells are exposed to any of a number of environmental factors, the most
prominent of which are glucose and acid pH. At least in the response to
glucose stimulation, Pma1p seems to require about 1 minute to reach maximal
activity (Serrano, 1983
).
Yeast cells also contain a second plasma membrane H+-ATPase, Pma2p,
which is not essential for growth and is expressed at a much lower level; its
function has not been determined.
The Dictyostelium genome contains an open reading frame
(aa-numgf1149 in the genome sequencing database) that encodes a sequence 32%
identical to Pma1p. This predicted Pma1p homologue has not yet been
characterized. However, a Dictyostelium P-type H+-ATPase
homologous to Pma2p is induced under conditions of mild cytosolic
acidification (Coukell et al.,
1997), and our preliminary data suggest that the latter enzyme
(Pat2) helps to defend cytosolic pH in the presence of proton ionophores (C.M.
and H.M., unpublished). Our working hypothesis is that the V-ATPase is
normally responsible for pH regulation in Dictyostelium, but that
reduced expression of VatM leads to the induction of P-type proton ATPase(s)
to assist in pH regulation. This would account for the ability of VatMpr to
respond more rapidly than AX2 to acid challenge. It would also be consistent
with the report that, in Neurospora, inhibition of a V-type ATPase
led to the emergence of mutations in a P-type ATPase gene, which increased the
in vivo activity of that enzyme (Bowman et
al., 1997
).
This strategy raises certain questions of energy economy: V-type ATPases
pump up to two protons per ATP hydrolyzed
(Grabe et al., 2000), or twice
the yield of P-type enzymes (Davies et
al., 1994
; Briskin et al.,
1995
). Even under the most extreme conditions of our experiments
a gradient of about 2.5 pH units and a moderate membrane potential
(van Duijn et al., 1988
)
the energy of ATP hydrolysis should suffice for V-ATPase function
(Wieczorek et al., 1999
), and
the substitution of P-ATPases for V-ATPases would thus seem to squander ATP.
However, V-type ATPases are huge molecules with relatively low turnover
number, while P-type enzymes have about an eighth the mass and a much higher
throughput. It thus makes at least intuitive sense for a cell to employ V-type
enzymes for predictable base load applications but use P-type transporters to
handle load peaks. Since use of P-type ATPases entails additional ATP
consumption, activation of this mechanism could impose a brake on other cell
functions, thereby contributing to the growth phenotype of VatMpr. Studies to
test this hypothesis are in progress.
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Acknowledgments |
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