©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Inositol Trisphosphate-dependent and -independent Ca Mobilization Pathways at the Vacuolar Membrane of Candida albicans (*)

(Received for publication, December 7, 1994)

Caroline M. Calvert Dale Sanders (§)

From the Biology Department, University of York, York YO1 5DD, United Kingdom

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Vacuolar membrane vesicles were isolated from Candida albicans protoplasts, and marker enzyme assays were employed to identify the membranes as vacuolar in origin. The mechanisms of Ca uptake and Ca release at the vacuolar membrane were investigated. Ca accumulation by vacuolar membrane vesicles can be generated via H/Ca antiport. The inside-acid pH is in turn generated by a vacuolar-type H-ATPase, as demonstrated by the sensitivity of Ca uptake to ionophores and the vacuolar H-ATPase inhibitor bafilomycin A(1). Vacuolar membrane vesicles exhibit two Ca release pathways: one induced by inositol 1,4,5-trisphosphate (InsP(3)) and the other by inside-positive voltage. These two pathways are distinct with respect to the amount of Ca released, the nature of response to successive stimuli, and their respective pharmacological profiles. The InsP(3)-gated pathway exhibits a K(0.5) for InsP(3) of 2.4 µM but is not activated by inositol 4,5-bisphosphate or inositol 1,3,4,5-tetrakisphosphate at concentrations up to 50 µM. Ca release by InsP(3) is blocked partially by low molecular weight heparin. Ca released by the voltage-sensitive pathway occurs at membrane potentials estimated to be over a physiological range from 0 to 80 mV. The voltage-sensitive Ca release pathway can be blocked by lanthanide ions and organic channel blockers such as ruthenium red and verapamil. Furthermore, the voltage-sensitive Ca release pathway exhibits Ca-induced Ca release. These findings are discussed in relation to the mechanism of Ca-mediated cellular signaling in C. albicans and other fungi.


INTRODUCTION

Candida albicans is a dimorphic yeast, which grows either as an ellipsoidal bud (often referred to as the blastophore or the yeast form) or in a filamentous fashion producing pseudohyphae or true septate hyphae (Odds, 1988). C. albicans is an opportunistic pathogen which generally produces mild superficial infections. Although, inspection of infected tissues reveals a mixture of budding, mycelial and pseudomycelial C. albicans cells (Odds, 1988), the pathogenicity of C. albicans is often linked to structural dimorphism. Thus the ability to grow filamentously may be advantageous during tissue invasion, and hyphal formation may be an escape mechanism from a phagocytosing host cell.

The transition from yeast to hyphal growth in C. albicans can be initiated by several factors (reviewed by Shepherd et al., 1985; Odds, 1988). Significantly, Ca is one of the factors that is able to regulate the dimorphic potential in C. albicans. Roy and Datta(1987) demonstrated inhibition of germ tube formation by Ca ionophores and calmodulin inhibitors. Exogenous Ca can also induce the dimorphic transition (Sabie and Gadd, 1989), and germ tube forming cells have more active calmodulin (Paranjape et al., 1990). The importance of Ca in dimorphism has parallels in other filamentous fungi (Muthukumar and Nickerson, 1984; Gadd and Brunton, 1992). Furthermore, cytosolic free Ca is thought to play a crucial regulatory role in hyphal tip growth in a diverse range of fungi (Jackson and Heath, 1993).

The fungal vacuole acts as a Ca buffering system, maintaining low cytosolic Ca concentrations. Halachmi and Eilam(1989) estimated the cytosolic free Ca in Saccharomyces cerevisiae with the Ca-sensitive fluorescent dye indo-1 as 346 nM while the vacuolar concentration was calculated as 1.3 mM. A lower estimate of 116 ± 90 nM for cytoplasmic Ca concentration was obtained by Iida et al.(1990), and in Neurospora crassa cytosolic free Ca measured with ion-selective microelectrodes is 92 ± 15 nM (Miller et al., 1990). One mechanism for vacuolar accumulation of Ca is an exchange of Ca for nH via a H/Ca antiporter (Ohsumi and Anraku 1983; Okorokov et al. 1985), although there is also evidence for a Ca-ATPase at the same membrane in yeast (Cunningham and Fink, 1994). Conclusive evidence for the Ca homeostatic function of the vacuole in S. cerevisiae arises from work on the mutant Deltavma4, which is deficient in functional vacuolar-type H-ATPase (Ohya et al., 1991) and hence in Ca uptake by Ca/H exchange: this mutant is unable to control cytosolic Ca concentration.

Little is known concerning the pathways of Ca release from fungal vacuoles, and, hence of the likely mechanisms of intracellular Ca mobilization during cell signaling. Patch clamp studies on yeast vacuoles have revealed the presence of voltage-sensitive ion channels which conduct Ca (Bertl and Slayman, 1990). In addition, InsP(3), (^1)a second messenger widely involved in intracellular Ca mobilization in animal cells (Berridge and Irvine, 1989), elicits Ca release from vacuoles of N. crassa (Cornelius et al., 1989) and S. cerevisiae (Belde et al., 1993). The physiological relevance of InsP(3)-elicited Ca release is supported by findings that the elements of a phosphoinositide cycle are present in both yeast and some filamentous fungi (Kaibuchi et al., 1986; Kato et al., 1989; Robson et al., 1991). It is not yet known whether both voltage- and InsP(3)-sensitive pathways for Ca release from fungal vacuoles occur in the same species, as is the case in plants (Johannes et al., 1992).

The ubiquity of cytosolic Ca as a second messenger in eukaryotic cells, coupled with the likelihood that Ca plays a specific role in C. albicans dimorphism and clear indications that the vacuole contains the major mobilizable intracellular Ca store in fungi, suggested the need to characterize the Ca transport pathways at the vacuolar membrane of C. albicans. We demonstrate here the presence of two such pathways which appear to be discrete. The first is gated open by InsP(3), whereas the second is gated open by cytosol-negative transmembrane voltages in the physiological range.


EXPERIMENTAL PROCEDURES

Culture Conditions and Spheroplast Formation

C.albicans (strain C316) from the Glaxo (Greenford, United Kingdom) culture collection was used throughout. Liquid cultures of the yeast form were grown in 1% (w/v) glucose-supplemented yeast nitrogen base medium (Difco, Detroit, MI) in an orbital incubator at 37 °C and a rotation speed of 200 revolutions/min. Cells were harvested by filtration in the mid-exponential phase of growth, washed, and resuspended in buffer (1.2 M sorbitol in 30 mM BTP adjusted to pH 7.5 with MES) to a concentration which gave an absorbance of 6.0 at 800 nm.

The enzymes (all supplied by Sigma-Poole, U.K.) used for spheroplast formation were lyticase from Arthrobacter luteus, chitinase from Serratia marcescens, and glucuronidase type H2 from Helix pomatia. Cell suspension (200 ml) was preincubated for 15 min with 2 mM dithiothreitol and then incubated with 50 ml of lyticase (100 units/ml), 16 units of chitinase, and 200,000 units of glucuronidase for 3 h at 30 °C with shaking (150 revolutions/min). Spheroplasts were harvested by centrifugation for 15 min at 2200 times g using a Beckman Ti-35 rotor, washed in spheroplast buffer (1 M sorbitol, 30 mM BTP adjusted to pH 8.0 with MES) and reharvested.

Membrane Preparation

Vacuolar membrane vesicles were prepared using a method modified from Uchida et al.(1988). Briefly, each pellet was resuspended in Buffer A (10 mM Tris adjusted to pH 6.9 with MES, 0.1 mM MgCl(2) and 12% (w/v) Ficoll 400), and homogenized with a loose fitting glass homogenizer in order to lyse the spheroplasts. Unlysed spheroplasts and cell debris were removed by centrifugation at 4,500 times g in a SW 27 rotor for 10 min, and the supernatant was retained for vacuole isolation. All subsequent manipulations were carried out at 4 °C. To isolate vacuoles, 20-ml volumes of supernatant were transferred to SW 27 rotor tubes, and 10 ml of Buffer A was layered on top. This was then centrifuged at 51,900 times g for 30 min. The white wafer floating on top of the Ficoll was carefully removed using a spoon-shaped spatula (which had been prewetted with Buffer A) and resuspended in Buffer A. To further purify this vacuolar suspension, 15-ml volumes were overlaid with 15 ml of Buffer B (10 mM Tris-MES, pH 6.9, 0.5 mM MgCl(2), 8% (w/v) Ficoll 400) and recentrifuged under the same conditions.

The vacuoles recovered from the top of Buffer B were then converted into vesicles by diluting them first with an equal volume of Buffer C (10 mM Tris-MES, pH 6.9, 0.5 mM MgCl(2), 25 mM KCl), and then two volumes of Buffer D (20 mM Tris-MES, pH 6.9, 1.0 mM MgCl(2), 50 mM KCl). The vesicles were pelleted at 37,000 times g for 20 min. The pellet was resuspended in 35 mM BTP-MES, pH 8.0, 0.3 M glycerol, 2 mM dithiothreitol, and 1% (w/v) bovine serum albumin (fraction V, protease-free). Protein concentration of the membrane vesicle preparation was determined using a Bio-Rad (Hemel Hempstead, U.K.) assay kit with bovine serum albumin (fraction V, essentially fatty acid-free) as the standard. Typically the protein concentration of the lysed protoplasts was 360 mg which yielded 7.5 mg of membrane vesicles.

Marker Enzyme Assays

Marker enzyme assays were employed to determine the purity of the vesicle preparation. Plasma membranes were identified using vanadate-sensitive ATPase activity, mitochondrial membranes using azide-sensitive ATPase activity (Widell and Larsson, 1990), and vacuolar membranes using bafilomycin A(1)-sensitive ATPase activity (Bowman et al., 1988). ATPase activity was measured by estimating total P(i) release (Ames, 1966). Bafilomycin A(1) stock solutions were prepared in Me(2)SO and stored at -20 °C. IDPase was used as a marker for Golgi membranes and was measured using the method of Green(1983). The reaction medium (final volume 1.0 ml) comprised 3 mM IDP, 1 mM MgCl(2), and 50 mM Tris buffered to pH 7.0 with HCl. The IDPase assay was carried out at 25 °C and was initiated by the addition of membrane vesicles (10-50 µg of protein). The reaction was stopped after 1 h by the addition of ice-cold 20% (w/v) trichloroacetic acid. The precipitated protein was removed by centrifugation, and the supernatant was assayed for P(i) using the method of Ames(1966). Nonspecific phosphatase activity was measured by the method of Leigh and Walker(1980). Membrane vesicles were incubated at 37 °C with reaction medium (30 mM BTP-MES, pH 6.0 or 8.0, ±30 mM molybdate), and the reaction was initiated by the addition of 3 mM disodium p-nitrophenol phosphate. The reaction was stopped after 1 h by the addition of 1 ml of 10% (w/v) Na(2)CO(3) and the absorbance read at 400 nm. The assay was calibrated using p-nitrophenol as a standard. NADPH cytochrome c reductase was used as a marker for endoplasmic reticulum membranes (Schekman, 1982). The activity of cytochrome c reductase was measured as described by Hodges and Leonard(1974). The reaction was initiated by the addition of 3 mM NADPH to reaction medium containing membrane protein (10-50 µg), 0.45 mM cytochrome c (oxidized), 50 mM potassium phosphate buffer, pH 7.7, and two inhibitors of respiratory chain complexes (50 mM NaCN and 0.4 µM antimycin A). The increase in absorbance (as a result of cytochrome c reduction) was monitored continuously at 550 nm.

Fluorescence Assays

Transmembrane pH gradients generated in response to activation of the H-ATPase were assayed using the pH gradient-sensitive dye quinacrine (Rottenberg, 1979). Fluorescence measurements were made in a Perkin-Elmer LS-5 luminescence spectrometer at 22 °C. Data were collected on-line and subsequently analyzed using the program LSR (Jennings et al., 1988) on an IBM PC.

Membrane vesicles were incubated with 5 µM quinacrine in 1 ml of reaction medium comprising 4.0 mM ATP, 50 mM KCl, 3.5 mM BTP-MES, pH 8.0, 2 mM NaN(3), 0.1 mM VO(4) and 0.3 M glycerol. All solutions were stirred constantly. The transport-related percent quench in fluorescence on addition of 10 mM MgCl(2) was estimated from the percent quench recovered on the addition of an uncoupler such as NH(4)Cl (5 mM).

The membrane potential of the vacuolar membrane vesicles was monitored by loading membrane vesicles with 5 µM oxonol V (Molecular Probes, Junction City, OR). On attainment of a steady fluorescence reading, 10 mM TPMP was added to generate an inside positive-membrane potential. Delta was estimated using the Nernst equation, after preloading the membrane vesicles for 5 min in reaction medium containing 1 mM TPMP.

Calcium Transport Assays

Calcium transport was assayed using a radiometric-filtration technique. Membrane vesicles (20 µg/ml) were incubated at 22 °C in reaction medium (3.5 mM BTP-MES, pH 8.0, 0.3 M glycerol, 50 mM KCl, 4.5 mM MgSO(4), 2 mM NaN(3) 0.1, mM VO(4), and 4.0 mM BTP-ATP) to generate a steady transmembrane H gradient. After 5 min Ca (Amersham International, Amersham, U.K.: 2.02 mCi ml on arrival) diluted with cold CaCl(2) was added at the desired concentration. Samples (50 µl) were removed from the medium at specified times, pipetted onto prewetted filters (Whatman cellulose nitrate membrane filters, pore diameter 0.45 µm), and washed rapidly under vacuum using a Millipore filtration unit, with 6 ml ice-cold wash medium. Wash medium contained 3.5 mM BTP-MES, pH 8.0, 0.3 M glycerol, and CaCl(2) at a concentration 20 times that used to initiate uptake. The filtration rate was approximately 0.7 ml of wash medium s. Radioactivity on the filters was then counted by standard liquid scintillation techniques. Ca uptake is expressed as nmol Ca/mg protein. Controls were performed in the absence of ATP to account for any ``passive'' uptake and have been subtracted from the results except when the minus ATP results are clearly depicted on the same graph. The amount of Ca binding to the filter was estimated by repeating the experiment without membranes and did not exceed 0.03% of the total Ca normally retained by membrane vesicles.

Ca-Release Assays

After Ca had been accumulated to a steady state within the vesicles, potential agonists of Ca release (InsP(3) and TPMP) were added from buffered stock solution with rapid mixing. Potential inhibitors of Ca release were added 1 min prior to the initiation of the H/Ca antiport. Ca release is expressed as a percentage in relation to the total ionophore (A23187)-sensitive Ca accumulation.


RESULTS

Membrane Characterization

Bafilomycin was used as a V-type ATPase inhibitor. Of the total ATPase activity in the membrane vesicle preparation, at least 70% was bafilomycin sensitive (Fig. 1). This suggests that the dominant membrane fraction in the preparation was vacuolar. The I for inhibition of ATPase activity in membrane vesicles by bafilomycin was 0.6 µM (Fig. 1), a value comparable to the I values for other V-type ATPases (Bowman et al., 1988). The preparation was contaminated predominantly with mitochondrial and plasma membranes. Thus, ATPase activity was inhibited 13% by azide and 8% by vanadate, with the residual activity (about 10%) accounted for by the action of nonspecific phosphatases. Activities of marker enzymes for Golgi and endoplasmic reticulum were barely detectable indicating neglible contamination by these membranes.


Figure 1: Bafilomycin inhibits V-type H-ATPase activity in C. albicans. Vacuolar membrane vesicles (10 µg) were assayed for H-ATPase activity in reaction medium comprising 4.0 mM ATP, 2.0 mM MgSO(4), 50 mM KCl, and 50 mM BTP-MES, pH 8.5, ± bafilomycin A(1). Results are the mean of three separate experiments and are fitted to a rectangular hyperbola (solid line) using nonlinear least-squares.



The ATP-dependent proton pumping, as monitored by fluorescence quenching of quinacrine, was completely inhibited by bafilomycin A(1) (Fig. 2). This result is significant as it demonstrates that intravesicular acidification is generated by the V-type H-ATPase alone. In total, these marker enzyme results suggest that this membrane vesicle preparation is suitable for the study of ion transport at the vacuolar membrane.


Figure 2: Inhibition of ATP-dependent H pumping by bafilomycin A(1). H pumping was assayed as quenching of quinacrine fluorescence, as described under ``Experimental Procedures.'' A, 100 µg of membrane protein was preincubated for 5 min in the presence of 3 µM bafilomycin. B, relaxation of steady state pH gradient by addition of 3 µM bafilomycin.



H/Ca Antiport at the Vacuolar Membrane

Ca uptake at the vacuolar membrane could be driven by ATP (Fig. 3). A steady state accumulation was achieved after 6 min and could be reversed to the level of ATP-independent uptake by the addition of the Ca ionophore A23187. In principle, Ca uptake could be driven either by a Ca-ATPase or by the proton motive force set up by the primary H-ATPase. To discriminate between these possibilities, 10 µM FCCP was added to the reaction medium to dissipate the H gradient. Fig. 4shows that ATP-dependent Ca uptake is largely abolished by FCCP. Moreover, Ca uptake is completely eliminated by 3 µM bafilomycin A(1). These results confirm that Ca accumulation is driven by a H gradient generated by the V-type ATPase.


Figure 3: Ca uptake by vacuolar membrane vesicles. 50 µg of membrane protein was preincubated for 5 min with reaction medium as detailed under ``Experimental Procedures,'' with (bullet) or without (circle) 4.0 mM ATP. 10 µMCaCl(2) was then added at time = 0 and mixed rapidly. The time scale refers to the time when aliquots of reaction medium were filtered. 10 µM A23187 was added at the time shown to release the Ca accumulated. Each point represents the mean of at least three independent experiments ± S.E.




Figure 4: Effect of FCCP and bafilomycin on vacuolar Ca accumulation. 40 µg of membrane protein was preincubated in reaction medium in the absence (bullet) or presence of either 10 µM FCCP (circle) or 3 µM bafilomycin ().



The initial rate of H/Ca antiport (assayed after 15 s) displayed saturation kinetics with respect to Ca concentration, and possesses a K(m) of 7.3 ± 1.5 µM (Fig. 5).


Figure 5: Concentration-dependence of ATP-dependent Ca uptake into vacuolar vesicles. 20 µg of membrane protein were preincubated in 0.5 ml of reaction medium with or without 4.0 mM ATP. The initial influx was calculated as ATP-dependent Ca uptake after 15 s. Each point is the mean of three determinations ± S.E. The solid line is a nonlinear least-squares fit (Marquardt, 1963) to the Michaelis-Menten equation.



Inositol 1,4,5-Trisphosphate Releases Ca from Vacuolar Membrane Vesicles of C. albicans

On addition of 20 µM InsP(3) to vacuolar membrane vesicles which had been allowed to accumulate Ca to a steady state, approximately 24% of the A23187-sensitive Ca pool was released (Fig. 6). This reproducible Ca release was rapid, and no reuptake was observed. The time course for InsP(3)-induced release was not discernible within the time resolution of the filtration assay (the fastest sampling time is approximately 15 s). The specificity of the InsP(3) response was examined with respect to InsP(2) and InsP(4): neither of these compounds elicited any Ca release at concentrations up to 50 µM.


Figure 6: InsP(3)-induced Ca release from vacuolar membrane vesicles. 40 µg of membrane protein were preincubated for 5 min with Ca uptake reaction medium, as detailed under ``Experimental Procedures.'' Ca uptake was initiated at t = 0 by addition of 10 µMCaCl(2), followed by rapid mixing. 10 µM InsP(3) and 10 µM A23187 were added to the reaction medium at the times shown and mixed rapidly. Each point represents the mean of at least three independent experiments.



The concentration dependence of InsP(3)-elicited Ca release is shown in Fig. 7. These data exhibit monophasic saturation kinetics and yield a K(m) for InsP(3)-induced Ca release of 2.4 ± 0.2 µM, with maximal release at saturating InsP(3) concentrations amounting to 24%. After an initial dose of 20 µM InsP(3), no further release of Ca was observed on subsequent application of InsP(3) (data not shown).


Figure 7: InsP(3) release of Ca is dependent on InsP(3) concentration. The amount of Ca released from the A23187-sensitive Ca pool was measured in varying concentrations of InsP(3). The InsP(3) stock solution was diluted appropriately so that the same volume of InsP(3) was added each time. Other experimental details are described in the legend to Fig. 3. Data (the mean ± S.E. of three separate experiments) were fitted by nonlinear least-squares to the Michaelis-Menten equation.



Inhibitor Studies on InsP(3)-stimulated Ca Release

Several inhibitors known to block Ca channels in animals and plants were tested for their ability to block InsP(3)-induced Ca release. The results are summarized in Table 1(left-hand column). Low molecular weight heparin, a potent antagonist of InsP(3)-induced Ca release in animals (Koybayashi et al., 1988) and plants (Brosnan and Sanders, 1990), inhibited Ca release, but only by 40% at 10 µM (0.05 mg ml). Other established InsP(3)-gated Ca release blockers, dantrolene and TMB-8, were without effect at concentrations of 200 µM.



Other Ca channel blockers were examined as potential inhibitors. The lanthanides Gd and La both blocked Ca release when applied at a concentration of 100 µM. 1 mM Mn had no effect on Ca release. The endomembrane calcium channel blocker ruthenium red completely inhibited Ca release at 100 µM. Verapamil (100 µM) did not exert any inhibitory effects on InsP(3)-induced Ca release.

The possibility that InsP(3)-elicited Ca release is regulated by cytosolic free Ca was investigated by addition of 200 µM EGTA 2 min prior to the addition of InsP(3). Although EGTA itself had no effect on preaccumulated Ca, this chelation of Ca in the medium resulted in a 60% enhancement of InsP(3)-elicited Ca release.

Membrane Potential-driven Ca Release

Addition of the lipophilic cation TPMP to membrane vesicles generated a physiological inside-positive membrane potential indicated by the quenching of oxonol V fluorescence (Fig. 8, inset). The quench was sustained and could be recovered by the addition of detergent. The imposition of a Delta with 10 mM TPMP elicited Ca release from Ca-loaded membrane vesicles (Fig. 8). Ca release was dependent on the concentration of TPMP applied (Fig. 9). Thus, half-maximal release of Ca was attained at 5.9 ± 0.8 mM TPMP, with maximal release accounting for 82% of the A23187-sensitive Ca pool. In order to test whether Ca release was complete after a single application of TPMP, two near-saturating doses of TPMP (20 mM) were administered to the vesicles. Fig. 10shows that two successive Ca release responses of equivalent magnitude were elicited.


Figure 8: TPMP releases Ca from vacuolar membrane vesicles. Vacuolar membrane vesicles were preloaded with Ca, as detailed in the legend to Fig. 6. 10 mM TPMP was added with rapid mixing at the time shown. Each point represents the mean of four separate experiments. Inset, TPMP-induced quenching of oxonol V fluorescence. 20 µg of membrane protein was incubated in 0.5 ml of Ca uptake reaction medium containing 5 µM oxonol V. 10 mM TPMP was added as indicated to impose an inside-positive membrane potential (and hence induce the fluorescence quench), and then 0.02% (v/v) Triton X-100 was added to disrupt the vesicles. The trace shown is representative of two experiments.




Figure 9: Release of Ca is dependent on TPMP concentration. Vacuolar membrane vesicles were allowed to accumulate Ca as described in the legend to Fig. 6. After 6 min TPMP was added with rapid mixing. Aliquots were then removed from the reaction medium 1 min after TPMP addition and assayed for Ca retained by the vesicles. The data are fitted to the Michaelis-Menten equation by nonlinear least-squares. Each data point represents the mean of three experiments.




Figure 10: Two near-saturating doses of TPMP elicit two responses of equal size. After Ca loading and accumulation to a steady state (as in Fig. 6), 20 mM TPMP was added to the reaction medium. Two aliquots were removed, and the amount of Ca retained was determined. A further dose of TPMP was then added, and the amount of Ca retained was measured. The results are the mean ± S.E. of three experiments.



The Delta dependence of Ca release was quantified by preincubating vesicles in the presence of 1 mM TPMP. The Delta resulting on subsequent addition of various concentrations of TPMP was then calculated by application of the Nernst equation. The results are shown in Fig. 11, and demonstrate measurable Ca release over a range of intravesicular Delta between 0 and 80 mV. Since the membrane potential of intact yeast vacuoles is also thought to reside around positive potentials, when referenced to lumen cytosol (Bertl et al., 1992), the results in Fig. 11are in accord with activation of Ca release over a physiological range of membrane potentials.


Figure 11: Ca release increases with membrane potential. 40 µg of membrane protein were preincubated with Ca uptake reaction medium containing 1 mM TPMP. After subsequent uptake of Ca for 6 min, TPMP was added at the desired concentration, and the amount of Ca released after 1 min was measured. The results are expressed as Ca release as a percent of the amount of Ca accumulated in the steady state. Results shown are the data from three independent determinations ± S.E.



Effect of Inhibitors on Delta-sensitive Ca Release

Several Ca channel antagonists were examined for their effects on TPMP-induced Ca release (Table 1). The first group included two lanthanides (Gd and La) and two divalent cations (Zn and Mn). Gd and La fully blocked, and Mn (1 mM) partially (up to 80%) blocked Ca release, whereas Zn (100 µM) had no effect on Ca release. The dose-response relationships for inhibition of TPMP-elicited Ca release by La and Gd are shown in Fig. 12. The derived values for half-maximal inhibition are 0.87 ± 0.38 and 8.4 ± 3.6 µM, respectively.


Figure 12: Dose-response curves for La and Gd inhibition. Various concentrations of La (A) or Gd (B) were added to the Ca uptake reaction medium. Vesicles were then loaded with CaCl(2) for 6 min prior to the addition 10 mM TPMP. The data (mean ± S.E. of three independent experiments) are fitted to the Michaelis-Menten equation using nonlinear least-squares.



The endomembrane channel blocker ruthenium red (Lee and Tsien, 1983) exerted a complete blockade on Ca release at 100 µM. Verapamil (an inhibitor of animal plasma membrane Ca channels: Biden et al., 1984) also significantly reduced the amount of Ca released (7% released).

In contrast to its effects on InsP(3)-elicited Ca release, EGTA (200 µM) considerably reduced TPMP-generated Ca release to only 5% of the A23187-sensitive pool. Thus, it appears likely that voltage-sensitive Ca release requires the presence of cytosolic free Ca for full activity.

One possible mode of TPMP-induced Ca release might be that the shift in membrane potential induced by TPMP reverses the antiport mechanism. However, this is very unlikely since we detected no effect of La on Ca uptake via H/Ca antiport.

InsP(3)- and Delta-gated Ca Release Appear to be Mediated by Separate Pathways

When TPMP and a saturating dose of InsP(3) are added sequentially (in either order), Ca release to approximately the same level is observed (Fig. 13). This suggests that Ca release by TPMP and InsP(3) is independent and represents different release pathways. The disparate pharmacological profile of these pathways (summarized in Table 2) provides further evidence for separate pathways.


Figure 13: Sequential release by TPMP and InsP(3). Vacuolar membrane vesicles were preloaded with Ca as described in the legend to Fig. 6. When Ca uptake was at a steady state, 10 mM TPMP or 10 µM InsP(3) was added, and two aliquots were removed from the reaction medium to estimate the amount of Ca retained. Following this a subsequent dose of InsP(3) or TPMP was added (the reduction in reaction volume was accounted for), and the amount of Ca retained was estimated. Results are the means of two independent experiments ± standard deviation.






DISCUSSION

The H/Ca Antiporter

The present results demonstrate that uptake of Ca at the vacuolar membrane of C. albicans can be fuelled by H/Ca antiport. The possibility of a parallel uptake pathway involving Ca-ATPase activity, and analogous to that postulated for the vacuolar membrane of S. cereviseae (Cunningham and Fink, 1994), cannot be discounted as an additional in vivo mechanism, although were it to exist, it would be rendered inoperative by the presence of VO(4) in the uptake media. This protocol served to select for Ca uptake by vacuolar membrane vesicles.

The K(m) of 7 µM for H-coupled Ca transport into vacuolar vesicles of C. albicans is in close agreement with the K(m) for Ca uptake in some other vacuolar vesicles. In plant cells, Schumaker and Sze(1986) reported a K(m) of 10 µM Ca for H/Ca exchange in oat root vacuoles, and Bush and Sze(1986) obtained a K(m) of 21 µM for Ca uptake in tonoplast vesicles from cultured carrot cells. Previous estimates of the K(m) for vacuolar Ca uptake in yeasts are, however, somewhat higher, and range from 60 µM in Saccharomyces carlsbergensis to 100 µM in S. cerevisiae (Okorokov et al., 1985; Ohsumi and Anraku, 1983). In all cases the K(m) for Ca transport was calculated on the basis of Ca added to the reaction medium, so the actual K(m) for free calcium may be lower than these values indicate if there is significant Ca chelation. Nevertheless, since cytosolic free Ca in fungi resides normally at submicromolar levels (see Introduction), it seems likely that one major function of vacuolar H/Ca antiport would be to clear cytosolic Ca when it is abnormally high. Such conditions might apply locally, and especially in the vicinity of the vacuolar membrane, during stimulus-evoked Ca mobilization.

InsP(3)-gated Ca Release Pathway

At 24%, the proportion of Ca released by a saturating dose of InsP(3) from vacuolar membrane vesicles is similar to that reported for other non-animal systems (e.g. 26% in oat roots: Schumaker and Sze, 1987; 20% in red beet: Brosnan and Sanders, 1990). This limited release has been quantitatively accounted for as resulting from the low native density of InsP(3) regulated channels in vacuoles, which results in the formation of many vesicles which lack the channels (Brosnan, 1990).

A similar explanation appears likely to account for the limited InsP(3)-gated Ca release from C.albicans vesicles. Furthermore, it is possible that not all of the small vacuoles present in the blastospore of C. albicans possess an InsP(3) receptor. This could provide a mechanism for short bursts of localized Ca elevation in the cytosol, possibly advantageous for actin localization at the apex of the developing hyphae in C. albicans (Lasker and Riggsby, 1992). The loss of responsiveness of membranes to a second saturating dose of InsP(3) is characteristic and can be attributed to saturation of the InsP(3) receptor (Prentki et al., 1984).

The K(m) of 2.4 µM for InsP(3)-induced Ca release in C. albicans also compares favorably with values reported for other systems. In Neurospora vacuoles, the K(m) is 5.2 µM (Cornelius et al., 1989), and in Saccharomyces the K(m) is 0.4 µM (Belde et al., 1993). In plants the K(m) for InsP(3) mobilization of Ca varies from 8 µM in corn coleoptile microsomes (Reddy and Poovaiah, 1987) to as little as 0.2 µM in Acer vacuoles (Ranjeva et al., 1988) and 0.5 µM in red beet microsomes (Brosnan and Sanders, 1990). In pancreatic acinar cells, Ca is released from non-mitochondrial stores by InsP(3) with a K(m) of 1.1 µM (Streb et al., 1983).

The specificity of the Ca release for InsP(3) suggests that the response is mediated by a defined receptor in the vacuolar membrane vesicles of C. albicans. Such specificity has previously been demonstrated in plants (Ranjeva et al., 1988; Schumaker and Sze, 1987) and Saccharomyces (Belde et al., 1993) but was not observed in Neurospora, as several inositol phosphates also elicited Ca release (Schultz et al., 1990).

Of the inhibitors tested, low molecular weight heparin is considered to be a good probe for the presence of an InsP(3) receptor (Ghosh et al., 1988) and is thought to interact directly with the InsP(3) receptor as it is able to displace bound InsP(3) (Cullen et al., 1988; Brosnan and Sanders, 1993). Heparin is not a very effective inhibitor in C. albicans (present work) or Neurospora (Cornelius et al., 1989), and this may reflect differences in receptor structure between fungi and other eukaryotes.

The potentiation of InsP(3)-dependent Ca release by EGTA suggests that cytosolic Ca exerts an effect on the InsP(3) response. This result is in agreement with previous work done on animal systems where extravesicular Ca has been demonstrated to inhibit Ca release by optimal doses of InsP(3) (Jean and Klee, 1986; Chueh and Gill, 1986).

Membrane Potential Sensitive Ca Release

The Delta-dependent Ca release elicited by TPMP raises the strong possibility that voltage-operated Ca release channels reside in the vacuolar membrane of C. albicans. There are clear parallels with vacuolar cation channels reported from S. cereviseae (Bertl and Slayman, 1990) in that both pathways are opened at cytosol-negative (&cjs0809;inside-positive) Delta thought to prevail in vivo as a result of the operation of the electrogenic H-ATPase.

Lanthanides are known to block stretch-activated channels in Xenopus oocytes (Yang and Sachs, 1989) as well as vacuolar voltage-sensitive Ca release channels in plants (Allen and Sanders, 1994). Gd is also an inhibitor of the mechanosensitive plasma membrane calcium channel in the fungus Uromyces appendiculatus (Zhou et al., 1991). The inhibitory effects on voltage-gated Ca release of the ions tested in the present study might be explained in terms of a physical blockade (Stein, 1990). The unhydrated ionic radius of Ca is 0.099 nm, and La and Gd are close to this with radii of 0.106 and 0.094 nm, respectively. The unhydrated ionic radius of Mn is 0.080 nm, which is considerably smaller than that of Ca, and smaller still is Zn at 0.074 nm. As Mn partially inhibits the TPMP-induced Ca release, and Zn has no effect, this may reflect a critical size of radius required for inhibition.

The inhibitory effects of EGTA suggest that Ca release is controlled by external Ca. Elevation of cytosolic Ca to micromolar levels is known to enhance the activity of the voltage-dependent slowly activating vacuolar (SV) channel at the plant vacuolar membrane (Hedrich and Neher, 1987) which has recently also been demonstrated to operate as a Ca release channel (Ward and Schroeder, 1994). Cation channels at the vacuolar membrane of Saccharomyces are also activated by Ca, albeit at high levels (1 mM) (Wada et al., 1987). Later work on yeast vacuolar cation channels which can conduct Ca reports that this unphysiologically high Ca requirement can be lowered to 1 µM in the presence of a reducing agent (1 mM dithiothreitol or 10 mM 2-mercaptoethanol; Bertl and Slayman, 1990).

Parallel Pathways of Ca Release

Table 2summarizes the evidence for the independence of the two Ca release pathways described in this article. The differences include: the size of the inducer-sensitive Ca pool, response to successive doses of inducer, and pharmacological differences. These differences may provide the answer to how the two pathways may operate in vivo. Voltage-sensitive release appears to be controlled by desensitisation, while InsP(3)-induced Ca release is controlled by the presence of InsP(3). Thus, successive doses of InsP(3) to membrane vesicles loaded with Ca prompt only one response, whereas successive changes in the membrane potential give two Ca release responses.

Another notable difference between the two pathways is the role of cytosolic Ca concentration. Voltage-sensitive Ca release requires the presence of Ca: if free Ca is substantially lowered by EGTA, then no Ca release is observed. For InsP(3)-induced Ca release, endogenous Ca may also be involved, as Ca removal by EGTA partially stimulates the release of Ca. These results hint at an interesting phenomenon; Ca-regulated Ca release pathways at the vacuolar membrane of C. albicans. This difference in the two pathways might mean that, functionally, the pathway for Ca release from the vacuole would depend on the prevailing cytoplasmic Ca concentration. The different responses of the two pathways to cytoplasmic Ca concentration may suggest a mechanism whereby limited Ca release elicited by InsP(3) could serve to trigger more substantial Ca-induced Ca release. The requirement for cytoplasmic Ca in voltage-sensitive release can be viewed as a positive feedback mechanism, as observed in Saccharomyces (Bertl and Slayman, 1992). This would ensure fast and effective release of Ca from the vacuole.

Physiological Relevance of the Ca Release Pathways

If these Ca release pathways are of physiological significance, there must be a signal which triggers the Ca release. One physiological signal for Ca release in C. albicans could be the cytosolic alkalinization observed after induction of the dimorphic transition (Stewart et al., 1988). This pH change during germ tube formation could stimulate voltage-sensitive Ca release from the vacuole either directly or indirectly through a change in Delta if the alkalinization were generated through activation of the V-type H-ATPase. It is therefore relevant that Bertl and Slayman (1992) report that the cation channel at the vacuolar membrane in Saccharomyces opens at alkaline pH. Activation of the InsP(3)-gated release pathway might arise through elevation of cAMP levels which have been observed on germination (Chattaway et al., 1981). In Saccharomyces cAMP is known to activate PI and PIP kinases which results in enhanced InsP(3) production (Kato et al., 1989).

Downstream signaling events ensuing a projected rise in cytosolic free Ca could follow a well established pattern. Intracellular mobilization of free Ca will result in activation of calmodulin, which is known to be present in C. albicans (Muthukumar and Nickerson, 1987) and which has been implicated in the dimorphic transition of C. albicans (Sabie and Gadd, 1989; Paranjape et al., 1990). Calmodulin could then activate various phosphodiesterases and protein kinases (Miyakawa et al., 1989), and it is therefore noteworthy that an increase in protein phosphorylation has been observed in germinating cells (Roy and Datta, 1987).

The wide range of signaling events with which Ca has been associated in C. albicans also includes regulation of chitin synthase activity (Datta, 1992) and clustering of actin granules at the tip of the germ tube (Schmid and Harold, 1988; Soll, 1986). The presence of discrete pathways for intracellular Ca mobilization potentially endows cells with the capacity for modulation in the spatial or temporal patterns of Ca release. Thus, despite the wide range of signaling events with which cytosolic free Ca is likely to be associated in C. albicans, elements of specificity in stimulus-response coupling have the potential to be attained.


FOOTNOTES

*
This work was supported by a Science and Engineering Research Council-CASE studentship (to C. M. C.) and by Glaxo Group Research. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed. Tel.: +44-1904-43-2825; Fax: +44-1904-43-2860.

(^1)
The abbreviations used are: InsP(3), inositol 1,4,5-trisphosphate; Delta, membrane potential; BTP, bis-tris propane; cAMP, cyclic adenosine 3`5`-monophosphate; dantrolene, (1-[(5-[p-nitrophenyl]fur-furylidene)amine]hydantoin); FCCP, carbonyl cyanide p-trifluoromethoxyphenyl-hydrazone; InsP(2), inositol 4,5-bisphosphate; InsP(4), inositol 1,3,4,5-tetrakisphosphate; Me(2)SO, dimethyl sulfoxide; MES, 2-[N-morpholino]ethanesulfonic acid; oxonol V, bis-(3-phenyl-5-oxoisooxazol-4-yl)pentamethine oxonol; PIP(2), phosphatidyl inositol 4,5-bisphosphate; Quinacrine, 6-chloro-9-{[diethyl-amino)-1-methylbutyl]amino}-2-methoxy-acridine hypochloride; TMB-8, 8-(N,N-diethylamino)-octyl-3,4,5-trimethylbenzoate; TPMP, methyltriphenylphosphonium ion (CH(18)P).


ACKNOWLEDGEMENTS

We are grateful to Prof. K. H. Altendorf (Osnabrück) for the gift of bafilomycin A(1) and to Dr R. Irvine (Cambridge) for the supply of pure InsP(3). We also thank Glaxo Group Research (in particular Dr. J. Houston) for scientific assistance.


REFERENCES

  1. Allen, G. J., and Sanders, D. (1994) Plant Cell 6, 685-694 [Abstract/Free Full Text]
  2. Ames, B. N. (1966) Methods Enzymol. 8, 115-118
  3. Belde, P. J. M., Vossen, J. H., Borst-Pauwels, G. W. F. H., and Theuvenet, A. P. R. (1993) FEBS Lett. 323, 113-118 [CrossRef][Medline] [Order article via Infotrieve]
  4. Berridge, M. J., and Irvine, R. (1989) Nature 341, 197-205 [CrossRef][Medline] [Order article via Infotrieve]
  5. Bertl, A., and Slayman, C. L. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 7824-7828 [Abstract]
  6. Bertl, A., and Slayman, C. L. (1992) J. Exp. Biol. 172, 271-287 [Abstract/Free Full Text]
  7. Bertl, A., Blumwald, E., Coronado, A, Eisenberg, R., Findlay, G., Gradmann, D., Hille, B., Kohler, K., Kolb, H. A., MacRobbie, E., Meissner, G., Miller, C., Neher, E., Palade, P., Pantoja, O., Sanders, D., Schroeder, J., Slayman, C., Spanswick, R., Walker, A., and Williams, A. (1992) Science 258, 873-874 [Medline] [Order article via Infotrieve]
  8. Biden, T. J., Prentki, M., Irvine, R. F., Berridge, M. J., and Wollheim, C. B. (1984) Biochem. J. 223, 467-473 [Medline] [Order article via Infotrieve]
  9. Bowman, E. J., Siebers, A., and Altendorf, K. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 7972-7976 [Abstract]
  10. Brosnan, J. M. (1990) Nature 344, 593
  11. Brosnan, J. M., and Sanders, D. (1990) FEBS Lett. 260, 70-72 [CrossRef]
  12. Brosnan, J. M., and Sanders, D. (1993) Plant Cell 5, 931-940 [Abstract/Free Full Text]
  13. Bush, D. R., and Sze, H. (1986) Plant Physiol. 80, 549-555
  14. Chattaway, F. W., Wheeler, P. R., and O'Reilly, J. (1981) J. Gen. Microbiol. 123, 233-240 [Medline] [Order article via Infotrieve]
  15. Chueh, S. H., and Gill, D. L. (1986) J. Biol. Chem. 261, 13883-13886 [Abstract/Free Full Text]
  16. Cornelius, G., Gebauer, G., and Techel, D. (1989) Biochem. Biophys. Res. Comm. 162, 852-856 [Medline] [Order article via Infotrieve]
  17. Cullen, P. J., Comerford, J. G., and Dawson, A. P. (1988) FEBS Lett. 228, 57-59 [CrossRef][Medline] [Order article via Infotrieve]
  18. Cunningham, K. W., and Fink, G. R. (1994) J. Cell Biol. 124, 351-363 [Abstract]
  19. Datta, A. (1992) Curr. Sci. 62, 400-404
  20. Gadd, G. M., and Brunton, A. H. (1992) J. Gen. Microbiol. 138, 21561-1571
  21. Ghosh, T. K., Eis, P. S., Mullaney, J. M., Ebert, C. L., and Gill, D. L. (1988) J. Biol. Chem. 263, 11075-11079 [Abstract/Free Full Text]
  22. Green, J. R. (1988) in Isolation of Membranes and Organelles from Plant Cells (Hall, J. L., and Moore, A. L., eds) pp. 135-152, Academic Press, London
  23. Halachmi, D., and Eilam, Y. (1989) FEBS Lett. 256, 55-61 [CrossRef][Medline] [Order article via Infotrieve]
  24. Hedrich, R., and Neher, E. (1987) Nature 329, 833-836 [CrossRef]
  25. Hodges, T. K., and Leonard, R. T. (1974) Methods Enzymol. 32, 397-398
  26. Iida, H., Sakaguchi, S., Yagawa, Y., and Anraku, Y. (1990) J. Biol. Chem. 265, 21216-21222 [Abstract/Free Full Text]
  27. Jackson, S. L., and Heath, I. B. (1993) Microbiol. Rev. 57, 367-382 [Abstract]
  28. Jean, T., and Klee, C. B. (1986) J. Biol. Chem. 261, 16414-16420 [Abstract/Free Full Text]
  29. Jennings, I. R., Rea, P. A., Leigh, R. A., and Sanders, D. (1988) Plant Physiol. 79, 1257-1263
  30. Johannes, E., Brosnan, J. M., and Sanders, D. (1992) Plant J. 2, 97-102
  31. Kaibuchi, K., Miyajima, A., Arai, K., and Matsumoto, K. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 8172-8176 [Abstract]
  32. Kato, H., Uno, I., Ishikawa, T., and Takenawa, T. (1989) J. Biol. Chem. 264, 3116-3121 [Abstract/Free Full Text]
  33. Kobayashi, S., Somlyo, A. V., and Somlyo, A. P. (1988) Biochem. Biophys. Res. Commun. 153, 625-631 [Medline] [Order article via Infotrieve]
  34. Lasker, B. A., and Riggsby, W. S. (1992) Exp. Mycol. 16, 155-162
  35. Lee, K. S., and Tsien, R. W. (1983) Nature 302, 790-794 [Medline] [Order article via Infotrieve]
  36. Leigh, R. A., and Walker, R. R. (1980) Planta 150, 222-229
  37. Marquardt, D. W. (1963) J. Soc. Indust. Appl. Math. 11, 431-441
  38. Miller, A. J., Vogg, G., and Sanders, D. (1990) Proc. Natl. Acad. Sci. 87, 9348-9352 [Abstract]
  39. Miyakawa, T., Oka, Y., Tsuchiya, E., and Fukui, S. (1989) J. Bacteriol. 171, 1417-1422 [Medline] [Order article via Infotrieve]
  40. Muthukumar, G., and Nickerson, K. W. (1984) J. Bacteriol. 159, 390-392 [Medline] [Order article via Infotrieve]
  41. Muthukumar, G., and Nickerson, K. W. (1987) FEMS Microbiol. Lett. 41, 253-255
  42. Odds, F. C. (1988) Candida and Candidosis , 2nd Ed., Balliere and Tindall, London
  43. Ohsumi, Y., and Anraku, Y. (1983) J. Biol. Chem. 258, 5614-5617 [Abstract/Free Full Text]
  44. Ohya, Y., Umemoto, N., Tanida, I., Ohta, A., Iida, H., and Anraku, Y. (1991) J. Biol. Chem. 266, 13971-13977 [Abstract/Free Full Text]
  45. Okorokov, L. A., Kulakovskaya, T. V., Lichko, L. P., and Polorotova, E. V. (1985) FEBS Lett. 192, 303-306 [CrossRef][Medline] [Order article via Infotrieve]
  46. Paranjape, V., Gupta-Roy, B., and Datta, A. (1990) J. Gen. Microbiol. 136, 2149-2154 [Medline] [Order article via Infotrieve]
  47. Prentki, M., Biden, T. J., Janjic, D., Irvine, R. F., Berridge, M. J., and Wollkeim, C. B. (1984) Nature 309, 562-564 [Medline] [Order article via Infotrieve]
  48. Ranjeva, R., Carrasco, A., and Boudet, A. M. (1988) FEBS Lett. 230, 137-141 [CrossRef]
  49. Reddy, A. S. N., and Poovaiah, B. W. (1987) J. Biochem. (Tokyo) 101, 569-573 [Abstract]
  50. Robson, G. D., Trinci, A. P. J., Wiebe, M. G., and Best, L. C. (1991) Mycol. Res. 95, 1082-1084
  51. Rottenberg, H. (1979) Methods Enzymol. LV, 547-569
  52. Roy, G. B., and Datta, A. (1987) FEMS Lett. 41, 327-329 [CrossRef]
  53. Sabie, F. T., and Gadd, G. M. (1989) Mycopathology 108, 47-54
  54. Schekman, R. (1982) in Molecular Biology of the Yeast Saccharomyces , pp. 651-652, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  55. Schmid, J., and Harold, F. M. (1988) J. Gen. Microbiol. 134, 2622-2631
  56. Schumaker, K. S., and Sze, H. (1986) J. Biol. Chem. 261, 12172-12178 [Abstract/Free Full Text]
  57. Schultz, C., Gebauer, G., Metschies, T., Rensing, L., and Jastorff, B. (1990) Biochem. Biophys. Res. Commun. 166, 1319-1327 [Medline] [Order article via Infotrieve]
  58. Schumaker, K. S., and Sze, H. (1987) J. Biol. Chem. 262, 3944-3946 [Abstract/Free Full Text]
  59. Shepherd, M. G., Poulter, R. T. M., and Sullivan, P. A. (1985) Ann. Rev. Microbiol. 39, 579-614 [CrossRef][Medline] [Order article via Infotrieve]
  60. Soll, D. R. (1986) Bioessays 5, 5-11 [Medline] [Order article via Infotrieve]
  61. Stein, W. D. (1990) Channels, Carriers and Pumps: an Introduction to Membrane Transport. pp. 221-269, Academic Press Inc., New York _
  62. Stewart, E., Gow, N. A. R., and Bowen, D. V. (1988) J. Gen. Microbiol. 134, 1079-1087 [Medline] [Order article via Infotrieve]
  63. Streb, H., Irvine, R. F., Berridge, M. J., and Schultz, I. (1983) Nature 306, 67-69 [Medline] [Order article via Infotrieve]
  64. Uchida, E., Oshumi, Y., and Anraku, Y. (1988) Methods Enzymol. 157, 544-562 [Medline] [Order article via Infotrieve]
  65. Wada, Y., Ohsumi, Y., Tanifuji, M., Kasai, M., and Anraku, Y. (1987) J. Biol. Chem. 262, 17260-17263 [Abstract/Free Full Text]
  66. Ward, J. M., and Schroeder, J. I. (1994) The Plant Cell 6, 669-683 [Abstract/Free Full Text]
  67. Widell, S., and Larson, C. (1990) In the plant plasma membrane (Larsson, C., and Moller, I. M. eds) pp. 16-43, Springer-Verlag
  68. Yang, X. C., and Sachs, F. (1989) Science 243, 1068-1071 [Medline] [Order article via Infotrieve]
  69. Zhou, X. L., Stumpf, M. A., Hoch, H. C., and Kung, C. (1991) Science 253, 1415-1417 [Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.