Mg-Dependent Ecto-ATPase Activity inLeishmania tropica

ARCHIVES OF BIOCHEMISTRY AND BIOPHYSICS

Vol. 341, No. 1, May 1, pp. 40–46, 1997 Article No. BB979933

Mg-Dependent Ecto-ATPase Activity in Leishmania tropica Jose´ R. Meyer-Fernandes,*,1 Patrı´cia M. L. Dutra,† Claudia O. Rodrigues,† Jorge Saad-Nehme,* and Angela H. C. S. Lopes† *Departamento de Bioquı´mica Me´dica, Instituto de Cieˆncias Biome´dicas, Universidade Federal do Rio de Janeiro, CCS, Bloco H, Cidade Universita´ria, Ilha do Funda˜o, 21541-590, Rio de Janeiro, Brasil; and †Instituto de Microbiologia, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brasil

Received November 4, 1996, and in revised form February 3, 1997

ATPase activity has been located on the external surface of Leishmania tropica. Since Leishmania is known to have an ecto-acid phosphatase, in order to discard the possibility that the ATP hydrolysis observed was due to the acid phosphatase activity, the effect of pH in both activities was examined. In the pH range from 6.8 to 8.4, in which the cells were viable, the phosphatase activity decreased, while the ecto-ATPase activity increased. To confirm that the observed ATP hydrolysis was promoted by neither phosphatase nor 5*-nucleotidase activities, a few inhibitors for these enzymes were tested. Vanadate and NaF strongly inhibited the phosphatase activity; however, no effect was observed on ATPase activity. Neither levamizole nor tetramizole, two specific inhibitors of alkaline phosphatases, inhibited this activity. The lack of response to ammonium molybdate indicated that 5*-nucleotidase did not contribute to the ATP hydrolysis. Also, the lack of inhibition of the ATP hydrolysis by high concentrations of ADP at nonsaturating concentrations of ATP discarded the possibility of any ATP diphosphohydrolase activity. The ATPase here described was stimulated by MgCl2 but not by CaCl2 . In the absence of divalent metal, a low level of ATP hydrolysis was observed, and CaCl2 varying from 0.1 to 10 mM did not increase the ATPase activity. At 5 mM ATP, half-maximal stimulation of ATP hydrolysis was obtained with 0.29 { 0.02 mM MgCl2 . The apparent Km for Mg-ATP20 was 0.13 { 0.01 mM and free Mg2/ did not increase the ATPase activity. ATP was the best substrate for this enzyme. Other nucleotides such as ITP, CTP, GTP, UTP, and ADP produced lower reaction rates. To confirm that this Mg-dependent ATPase was an ecto-ATPase, an impermeant inhibitor, 4,4*-diisothiocyanostylbene-2,2 *-disulfonic acid was 1 To whom correspondence should be addressed at present address: Department of Biochemistry, Biological Sciences West Building, University of Arizona, Tucson, Arizona 85721. Fax: (520) 621-9288.

used. This amino/sulfhydryl-reactive reagent did inhibit the Mg-ecto-ATPase activity in a dose-dependent manner (I0.5 Å 27.5 { 1.8 mM). q 1997 Academic Press Key Words: Leishmania; Mg-ecto-ATPase.

Surface membrane interactions between Leishmania and their hosts are of critical importance for the survival of the parasite, from both the immunological and physiological viewpoints. The immunological interactions between leishmanial surface membrane constituents and the mammalian host immune system have been well documented (1, 2). Of special interest, the role of parasite membrane components in the uptake of these organisms by macrophages has been considered (3, 4). Less well studied, but equally important, is the role that specific parasite surface membrane constituents play in the protection from the cytolytic effects of extracellular molecules. Plasma membrane of cells contain enzymes whose active sites face the external medium rather than the cytoplasm. The activities of these enzymes, referred to as ecto-enzymes, can be measured using intact cells (5, 6). Leishmania donovani promastigotes and amastigotes and promastigotes of Leishmania mexicana mexicana have been shown to possess three distinct phosphomonoesterase activities, all of which are localized, at least in part, on the external surface of the plasma membrane (7–10). Acid phosphatases may play a crucial role in enabling the parasite to avoid the microbicidal activity of macrophages (11). It was recently shown that Leishmania acid phosphatases can regulate parasite binding to macrophages (12, 13). 5*-Nucleotidase has been detected in promastigotes of several Leishmania species (8–10, 14, 15); it is possible that this enzyme plays a key role in processing exogenously available nucleotides to a suitable form to be transported into the cell. 3*-Nucleotidase, an enzyme that

40

AID

0003-9861/97 $25.00 Copyright q 1997 by Academic Press All rights of reproduction in any form reserved.

ABB 9933

/

6b32$$$241

03-26-97 22:21:25

arca

ECTO-ATPase ACTIVITY IN Leishmania tropica

does not have a mammalian counterpart, was also detected on the surface membrane fraction of L. donovani promastigotes and L. mexicana mexicana (8–10). Although ATP is known to be an important extracellular signaling molecule which is supposed to increase the plasma membrane permeability and has important effects on cell membrane properties (16–24), none of these ectophosphomonoesterases, however, are able to promote high levels of ATP hydrolysis (25–27). Extracellular ATP may be one of the signaling molecules involved in cell-mediated cytotoxicity (22–24). It has been demonstrated that extracellular ATP has profound effects on cellular functions (17, 22, 24), causing plasma membrane depolarization, Ca2/ influx, and cell death (22, 24). Filippini et al. (22) have shown that ATP can kill different cells, with the exception of cells that express a high level of ATP-breakdown activity on their surface. Here we show the presence of a Mgdependent ecto-ATPase activity on the cell surface of living L. tropica and characterize the properties of this enzyme. MATERIAL AND METHODS Microorganisms. L. tropica stock IOC-L 571, from a WHO collection, was provided by Dr. G. Grimaldi Jr. (Fundac¸a˜o Oswaldo Cruz, Rio de Janeiro, RJ, Brazil). The parasites were cultured as promastigotes in Schneider’s Drosophila medium (GIBCO, Grand Island NY), supplemented with 10% fetal bovine serum (GIBCO) at 247C (28), and were harvested at the stationary phase of growth. Five days after inoculation, cells were collected by centrifugation, washed twice, and kept in 50 mM Tris–HCl, pH 7.2, 20 mM KCl, 100 mM sucrose, and 20 mM glucose. Cellular viability was assessed, before and after incubations, by mobility and trypan blue methods (29). The viability was not affected under the conditions employed here. Protein concentration was determined by the method of Lowry et al. (30) using bovine serum albumin as standard. Ecto-ATPase activity measurements. Intact cells were incubated for 1 h at 307C with gentle shaking (40 oscillations/min) in 0.5 ml of a mixture containing, unless otherwise specified, 50 mM Tris–HCl, pH 7.2, 5 mM ATP, 5 mM MgCl2 , and 1 mg/ml of protein which corresponds to 2.2 1 108 cells/ml. The ATPase activity was determined by measuring the hydrolysis of [g-32P]ATP (104 Bq/nmol ATP) (31). The experiments were started by the addition of living cells and terminated by the addition of 1 ml of a cold mixture containing 0.2 g charcoal in 1 M HCl. The tubes were then centrifuged at 1500g for 20 min at 47C, after which 400 ml of the supernatant was added to 9 ml of scintillation liquid (2 g PPO2 in 1 liter toluene) and counted in a liquid scintillation counter. The ATPase activity was calculated by subtracting the nonspecific ATP hydrolysis measured in the absence of cells. The ATP hydrolysis was linear with time under the assay conditions used and was proportional to the protein concentration. In the experiments in which other nucleotides were used, the hydrolytic activity measured under the same conditions described above was assayed spectrophotometrically by measuring the release of Pi from the nucleotides (32).

41

FIG. 1. Time course of ATPase activity by intact cells of Leishmania tropica. The reaction medium containing 50 mM Tris–HCl buffer, pH 7.2, 5 mM Tris–ATP [g-32P]ATP [sp act Å 104 Bq/nmol ATP]), 5 mM MgCl2 and 1 mg/ml of protein which correspond to 2.2 1 108 cells/ml. Reactions were at 307C. Data are means { SE of three determinations with different cell suspensions. Phosphatase activity measurements. In addition to the measurements of ecto-ATPase activity, the ecto-p-nitrophenylphosphatase activity was determined in the same medium for ATP hydrolysis except that ATP was replaced by 5 mM p-nitrophenylphosphate (p-NPP). The reaction was terminated by the addition of 1 ml of 1 N NaOH. The released p-nitrophenol was determined spectrophotometrically at 410 nm using an extinction coefficient of 14.3 1 1003 M01 cm01. Reagents. All reagents were purchased from Merck S.A. (Sao Paulo, S.P.) or Sigma Chemical Co. (St. Louis, MO). [g-32P]ATP was prepared as described by Glynn and Chappel (33). Distilled water deionized by the MilliQ system of resins (Millipore Corp., Bedford, MA) was used in the preparation of all solutions. Concentrations of free and complexed species (Mg2/, Ca2/, Mg-ATP20, Ca-ATP20, and ATP40) at equilibrium were calculated by using an iterative computer program described by Fabiato and Fabiato (34). Statistical analysis. All experiments were performed in triplicate, with similar results obtained at least in three separate cell suspensions. Apparent Km for Mg-ATP20 was calculated using a computerized nonlinear regression analysis of the data to the Michaelis– Menten equation (35). I50 for DIDS was determined by the equation vi/v0 Å Ki/(Ki / [DIDS]n),

[1]

where vi and v0 are the velocities of ATP hydrolysis in the presence and absence of DIDS, respectively, n is an index of cooperativity for the inhibitory effects of DIDS, and I0.5 Å K1/n is the concentration of i DIDS that gives half-maximal inhibition of ATP hydrolysis. Differences were evaluated for statistical significance by using Student’s t test for paired or unpaired data, as required.

RESULTS 2

Abbreviations used: AMPPCP, b,g-methyleneadenosine 5*-triphosphate; DIDS, 4,4*-diisothiocyanostylbene-2,2*-disulfonic acid; Mops, 4-morpholinepropane sulfonic acid; Pi , orthophosphate; PPO, 2,5-diphenyloxazole; p-NPP, p-nitrophenylphosphate.

AID

ABB 9933

/

6b32$$$242

03-26-97 22:21:25

The time course of ATPase activity present on the external surface of L. tropica is linear up to 1 h (Fig. 1). Living promastigotes of L. tropica at physiological

arca

42

MEYER-FERNANDES ET AL.

FIG. 2. Effect of pH on the ATPase and p-nitrophenylphosphatase activities of intact cells of L. tropica. Cells were incubated for 1 h at 307C in a medium containing 5 mM Tris-ATP [g-32P]ATP (sp act Å 104 Bq/nmol ATP)), or 5 mM p-NPP, 5 mM MgCl2 , 1 mg/ml of protein which correspond to 2.2 1 108 cells/ml, and 50 mM Mops–Tris buffer, adjusted to pH values between 6.8 and 8.4 with HCl or Tris. In this pH range (6.8–8.4) the cells were viable during all the course of the experiments. ATPase activity (l); p-nitrophenylphosphatase activity (j). Data are means { SE of three determinations with different cell suspensions.

pH (pH 7.2) were able to hydrolyze 380.0 { 15.4 nmol/ mg/h. To check the possibility of the observed phenomena being the result of secreted soluble enzymes, we prepared a reaction mixture with cells and it was incubated in the absence of ATP. Subsequently, it was centrifuged to remove the cells and the supernatant was checked for ATPase activity. This supernatant that at pH 5.0 presents phosphatase activity (p-NPP as substrate; 12); at pH 7.2 it failed to demonstrate ATP hydrolysis (data not shown). The presence of an acid phosphatase on the external surface of Leishmania has been described (7, 10, 36–38). Therefore, to discard the possibility that ATP hydrolysis was due to acid phosphatase activity, the effect of pH in both activities was examined in more detail. As shown in Fig. 2, at the pH range from 6.8 to 8.4, in which the cells were alive, while the phosphatase activity decreased, the ecto-ATPase activity increased. To confirm that the ATP hydrolysis was not due to phosphatase activity, different inhibitors of phosphatase activities were tested. In Table I it is shown that NaF and vanadate, two potent inhibitors of acid phosphatase activity (36–38), strongly inhibited the phosphatase activity, whereas no effect was observed on ATPase activity. Levamizole and tetramizole, two specific inhibitors for alkaline phosphatases

AID

ABB 9933

/

6b32$$$242

03-26-97 22:21:25

(39, 40), also failed to inhibit the ATPase activity. Another possible explanation for the ATP hydrolysis was that 5*-nucleotidase, another enzyme present on the external surface of Leishmania (8, 10, 14, 15) might be responsible for this hydrolysis. The lack of response to ammonium molybdate (Table I), a potent inhibitor of 5*-nucleotidase activity (8) and to its substrate, AMP (Table I), indicated that this enzyme did not contribute for the observed ATP hydrolysis. In L. donovani, 5*nucleotidase activity is inhibited by an increase in the pH (8), while the ATPase activity measured here was stimulated by an increase in the pH (Fig. 2). In human hepatoma cells it has been reported the presence of an ecto-Ca2/-ATPase activity that is also stimulated by an increase in the pH (41). In invertebrates an ATP diphosphohydrolase that hydrolyzes ATP to AMP, releasing 2 mol Pi/mol nucleotide has been described (42– 44). ATP hydrolysis catalyzed by this enzyme is inhibited by ADP with a Ki of 0.2 mM (45). The lack of inhibition by high concentrations of ADP at nonsaturating concentrations of ATP (Table I) caused the possibility that ATP hydrolysis was catalyzed by an ATP diphosphohydrolase activity to be discarded. In Table I it is shown that this Mg-activated ecto-ATPase activity was insensitive to oligomycin and sodium azide, two known inhibitors of mitochondrial Mg-ATPase. Recently it has been shown that Mg2/ is an important extracellular signal on the regulation of Salmonella virulence (46). Using intact cells in the absence of

TABLE I

Influence of Various Agents on ATPase and p-Nitrophenylphosphatase Activities Additions Levamizole (1 mM) Tetramizole (1 mM) NaF (1 mM) Vanadate (1 mM) Molybdate (0.1 mM) Oligomicine (1 mM) Azide (1 mM) AMP (1 mM) AMP (10 mM) ADP (1 mM) ADP (10 mM)

PNPase 108.1 79.9 11.1 5.3 7.8 93.7 106.6 101.5 100.1 98.4 92.2

{ { { { { { { { { { {

8.4 9.1 1.3 0.6 0.8 8.3 11.1 6.9 8.6 11.9 10.4

ATPase 107.5 97.3 96.6 98.3 94.6 91.2 93.3 93.1 99.7 99.1 94.6

{ { { { { { { { { { {

9.4 7.6 8.2 8.7 7.4 9.5 8.9 10.2 9.3 9.7 8.6

Note. ATPase and phosphatase activities were measured in the standard assay with 0.1 mM ATP and 5 mM p-NPP. Activities are expressed as a percentage of that measured under control conditions, i.e., without other additions. The ATPase (366.3 { 27.4 nmol/mg/h) and phosphatase (119.0 { 7.9 nmol/mg/h) activities were taken as 100%. The standard errors were calculated from the absolute activity values of four experiments with different cell suspensions and converted to percentage of the control value. The unpaired t test showed, in all cases with respect to ATPase activity, that there were not statistical differences (P ú 0.05) with respect to values found for each compound.

arca

ECTO-ATPase ACTIVITY IN Leishmania tropica

43

high level of hydrolysis of this nucleotide. Intact cells hydrolyzed AMPPCP at a much slower rate (4.2 { 0.5% of the rate for ATP, Table II). This Mg-dependent ectoATPase present on the external surface of L. tropica did not have its activity modified when cells were in log or stationary phase (data not shown). The fact that we have employed intact cells for these experiments indicated that this Mg-dependent ATPase was an ectoATPase. In order to confirm this, we applied the criterion that an authentic ectoenzyme should be inhibited by an added extracellular impermeant reagent (6, 41, 47) such as 2*,2*-diisothiocyanostylbene disulfonic acid (DIDS) (31, 41, 47–49). As shown in Fig. 5, this amino/ sulfhydryl-reactive compound did inhibit the Mg-ectoATPase activity in a dose-dependent manner (I0.5 Å 27.5 { 1.8 mM; n Å 0.7 { 0.06). DISCUSSION

This paper reports the presence of Mg-dependent ecto-ATPase present on the external surface of L. tropFIG. 3. Dependence of MgCl2 or CaCl2 concentration on the ATPase and p-nitrophenylphosphatase activities. Cells were incubated for 1 h at 307C in a medium containing 50 mM Tris–HCl, pH 7.2, 5 mM Tris–ATP ([g-32P]ATP (sp act Å 104 Bq/nmol ATP)), or 5 mM pNPP, 1 mg/ml of protein which corresponds to 2.2 1 108 cells/ml and increasing concentration of MgCl2 (j; l) or CaCl2 (h; s) as shown on the abscissa. Curve for Mg-ATPase activity represents the fit of experimental data by linear regression using the Michaelis–Menten equation as described under Materials and Methods. ATPase activity (l; s); p-nitrophenylphosphatase activity (j; h). Data are means { SE of four determinations with different cell suspensions.

MgCl2 , a low level of ATP hydrolysis was observed (Fig. 3), and the addition of MgCl2 to the extracellular medium increased the ecto-ATPase activity in a dose-dependent manner. At 5 mM ATP, half-maximal stimulation of ATP hydrolysis was obtained with 0.29 { 0.02 mM MgCl2 , while the phosphatase activity was not stimulated by MgCl2 (Fig. 3). In human hepatoma cells it has been shown that the presence of the Mg2/- or Ca2/-stimulated ecto-ATPase activities (41). CaCl2 (0.1 to 10 mM) did not increase the phosphatase or the ATPase activities (Fig. 3). At 5 mM ATP, in the absence of Ca2/ and Mg2/, the concentration of ATP40 was 3.1 mM. In the presence of 10 mM CaCl2 , the concentrations of ATP40 and Ca-ATP20 were 0.1 and 4.8 mM, respectively. The changes in ATP40 and Ca-ATP20 levels had no effect on ATPase activity (Fig. 3). These data indicated that Mg-ATP20 was the substrate for this enzyme. The apparent Km for Mg-ATP20 was 0.13 { 0.01 mM (Fig. 4) and free Mg2/ did not increase the ATPase activity (Fig. 4, inset). In Table II it is shown that ATP was the best substrate for this enzyme. Other nucleotides such as ITP, CTP, GTP, UTP, and ADP produced lower reaction rates. Then, the lack of inhibition by ADP showed in Table I could not be attributed to a

AID

ABB 9933

/

6b32$$$243

03-26-97 22:21:25

FIG. 4. Dependence on Mg-ATP20 concentration of Leishmania tropica ATPase activity. Cells were incubated at 307C in a medium containing 50 mM Tris–HCl, pH 7.2, 5 mM MgCl2 , 1 mg/ml of protein which corresponds to 2.2 1 108 cells/ml and Mg-ATP20 concentration varying as shown on the abscissa. Curve represents the fit of experimental data by nonlinear regression using the Michaelis–Menten equation as described under Materials and Methods. (Inset) Effects of free Mg2/ on the ATPase activity. Cells were incubated at 307C in a medium containing 50 mM Tris–HCl, pH 7.2, 1 mM EDTA, 10 mM Mg-ATP20 ([g-32P]ATP (sp act Å 104 Bq/nmol ATP)), and [Mg2/] varying as shown on the abscissa. The total amounts of ATP and MgCl2 necessary to form the desired Mg-ATP20 and free Mg2/ concentrations were calculated as described under Materials and Methods. Data are means { SE of three determinations with different cell suspensions.

arca

44

MEYER-FERNANDES ET AL. TABLE II

Substrate Specificity of Ecto-ATPase Activity Nucleotides

Relative activity

ATP ADP ITP CTP GTP UTP AMPPCP

100.0 21.9 41.3 39.9 25.4 15.8 4.2

{ { { { { { {

9.4 3.6 3.2 4.2 3.1 1.7 0.5

Note. The ATPase activity was measured in a medium containing nucleotides listed (5 mM), 5 mM MgCl2 , 50 mM Tris–HCl buffer, pH 7.2, and 1 mg/ml of protein which correspond to 2.2 1 108 cells/ml. Reactions were at 307C. The ATP hydrolysis (381.6 { 37.9 nmol/mg/ h) was taken as 100%. The standard errors were calculated from the absolute activity values of three experiments with different cell suspensions and converted to percentage of the control value.

ica. The external localization of the ATP-hydrolyzing site is supported by its sensitivity to an impermeant reagent (41, 47, 49) such as DIDS and the observation that inhibitors of other ATPases, with internal ATP binding sites did not change the activity of this enzyme. The Mg-activated ecto-ATPase activity shown here cannot be attributed to a mitochondrial Mg-ATPase, since this activity was insensitive to oligomycin and sodium azide (Table I). In L. donovani a H/-ATPase has been identified on the plasma membrane that is responsible for the intracellular pH maintenance (50). This H/-ATPase was described as a P-type ATPase highly sensitive to orthovanadate (Ki Å 7.5 mM) (51). Since this Mg-dependent ecto-ATPase did not respond to vanadate (Table I), the possibility that this activity was catalyzed by a H/-ATPase present in the surface of the plasma membrane was discarded. This ATP hydrolysis is not expected to be driven by phosphatase activity present on the external surface of Leishmania’s membrane because this enzyme has been described as an acid phosphatase (7, 10, 12, 36–38, 52) and the pH optimum for the Mg-dependent ecto-ATPase was shifted to alkaline range (Fig. 2). In addition, potent inhibitors for phosphatase activities were not capable of modifying the Mg-dependent ecto-ATPase activity (Table I). The Mg-dependent ecto-ATPase activity shown here cannot be attributed to a Leishmania 5*nucleotidase activity, since this enzyme is not stimulated by Mg2/ (8, 9), and the measured activity was insensitive to ammonium molybdate and AMP (Table I). The fact that it was effective toward ADP would be consistent with an ATP diphosphohydrolase-like activity. However, the observed properties do not conform with known ATP diphosphohydrolases (42–45). ADP was a much poorer substrate than ATP, the rate of ADP hydrolysis under saturating conditions being 21% of that of ATP (Table II), and ADP did not inhibit the

AID

ABB 9933

/

6b32$$$243

03-26-97 22:21:25

Mg2/ ecto-ATPase activity (Table I). These data discarded the possibility that ATP hydrolysis was catalyzed by an ATP diphosphohydrolase-like activity and suggested that ADPase is distinct from the ATPase activity. In the parasitic protozoan Toxoplasma gondii a nucleoside triphosphate hydrolase (NTPase) able to hydrolyze different nucleotide triphosphates with almost the same rate has been described (53). Although originally described as a cytosolic enzyme, it has been demonstrated that it is located in dense granules and secretory vesicles of the parasite and is secreted into the vacuolar space of infected cells (54). Recently it has been shown that the purified enzyme is not a single enzyme but a mixture of two isozymes, termed NTPase I and NTPase II, and that the primary difference between these isozymes is that NTPase II hydrolyzes nucleoside triphosphate and diphosphate substrates at almost the same rate and NTPase I hydrolyzes nucleoside diphosphates at a much slower rate (55). The Mgdependent ecto-ATPase present on the external surface of L. tropica was less active toward other nucleotides triphosphates such as ITP, CTP, GTP, and UTP than ATP (Table II), and it is a not secreted enzyme. EctoATPases are enzymes that exhibit divalent cation-dependent ATPase activity on the extracellular side of the plasma membrane (6, 41, 56–59). These ATPases

FIG. 5. Effect of increasing concentration of DIDS on ATPase activity. Cells were incubated for 1 h at 307C in a medium containing 50 mM Tris–HCl, pH 7.2, 5 mM MgCl2 , 5 mM Tris–ATP ([g-32P]ATP (sp act Å 104 Bq/nmol ATP)), 1 mg/ml of protein which corresponds to 2.2 1 108 cells/ml and concentrations of DIDS shown on the abscissa. The line adjusted to the experimental points was fit by nonlinear regression using Eq. [1] (see Materials and Methods). Data are means { SE of three determinations with different cell suspensions.

arca

ECTO-ATPase ACTIVITY IN Leishmania tropica

have been characterized in intact cell assays with respect nucleotide and divalent cations requirements (41, 56 – 58). Ecto-ATPases stimulated by cations have been described in rat liver, pancreas (56 – 58), and Entamoeba histolitica (47). The substrate for the enzyme described here is the complex Mg-ATP20 (Km Å 0.13 { 0.01 mM, Fig. 4) and free Mg2/ was not able to increase the enzyme activity, as it was observed for the nucleoside triphosphate hydrolase from T. gondi (53). Most of the ecto-ATPases are Mg2/ or Ca2/ stimulated (41, 56, 57, 59); however, this enzyme was not stimulated by CaCl2 . These data suggest that CaATP20 is a not substrate for this enzyme. In living promastigotes of L. tropica the pH optimum for Mgdependent ecto-ATPase was also shifted to alkaline range as reported to the Ca2/-ecto-ATPase present in human hepatoma cells (41). ATP-diphosphohydrolase activities are present in a wide variety of organisms (42–45, 59–64). In animals cells, the enzyme is a membrane-bound ecto-ATP-diphosphohydrolase (59, 61, 62). A highly active ectoATP diphosphohydrolase was localized on the external surface of the tegument of Schistosoma mansoni (63, 64) and the authors suggested that this enzyme might be involved in escape mechanisms of the parasite by splitting ATP or ADP which could be released on its surface by activated platelet or activated cytotoxic T lymphocytes (63, 64). Protozoa of the genus Leishmania are obligate intracellular parasites of mammalian macrophages, surviving within the severe environment of phagolysosome (1). At least under the conditions employed here, ATP and its nonhydrolyzable analog (AMPPCP, Table II) did not affect the Leishmania viability (see Materials and Methods). The identity and function of ecto-ATPases have been reviewed and the nomenclature of ‘‘E-type ATPases’’ was proposed to describe these enzymes (59). The physiological role of these enzymes is still unknown. However, several hypotheses have been suggested, such as (i) protection from cytolytic effects of extracellular ATP (22–24), (ii) regulation of ectokinase substrate concentration (59), (iii) involvement in cellular adhesion (65–67), (iv) involvement in signal transduction (68, 69), and (v) regulation of NO release in smooth muscle cells (70). Recently, it has been suggested that signal transduction networks, involving ectoenzymes with tyrosine kinase and phosphatase activities may modulate crucial events during Leishmania infection (13). The physiological role of the Mg-dependent ecto-ATPase in L. tropica remains to be elucidated.

ACKNOWLEDGMENT This work was supported by grants from Conselho Nacional de Desenvolvimento Cientifico e tecnolo´gico (CNPq).

AID

ABB 9933

/

6b32$$$243

03-26-97 22:21:25

45

REFERENCES 1. Alexander, J., and Russel, D. G. (1992) Adv. Parasitol. 31, 175– 254. 2. Handman, E., and Mitchell, G. F. (1985) Proc. Natl. Acad. Sci. USA 82, 5910–5914. 3. Handman, E., and Goding, J. W. (1985) EMBO J. 4, 329–336. 4. Russell, D. G., and Wilhem, H. (1986) J. Immunol. 136, 2613– 2620. 5. De Pierre, J. W., and Karnovsky, M. L. (1973) J. Cell. Biol. 56, 275–303. 6. De Pierre, J. W., and Karnovsky, M. L. (1974) J. Biol. Chem. 249, 7111–7120. 7. Gottlieb, M., and Dwyer, D. M. (1981) Exp. Parasitol. 52, 117– 128. 8. Gottlieb, M., and Dwyer, D. M. (1983) Mol. Biochem. Parasitol. 7, 303–317. 9. Dwyer, D. M., and Gottlieb, M. (1984) Mol. Biochem. Parasitol. 10, 139–150. 10. Hassan, H. F., and Coombs, G. H. (1987) Mol. Biochem. Parasitol. 23, 285–296. 11. Remaley, A. T., Kuhns, D. B., Basford, R. E., Glew, R. H., and Kaplan, S. S. (1984) J. Biol. Chem. 259, 11173–11175. 12. Vannier-Santos, M. A., Martiny, A., Meyer-Fernandes, J. R., and de Souza, W. (1995) Eur. J. Cell. Biol. 67, 112–119. 13. Martiny, A., Vannier-Santos, M. A., Borges, V. M., Meyer-Fernandes, J. R., Asseruy, J., Cunha e Silva, N. L., and de Souza, W. (1996) Eur. J. Cell. Biol. 71, 206–215. 14. Pereira, N. M., and Konigk, E. (1981) Tropenmed. Parasitol. 32, 209–214. 15. Hassan, H. F., and Coombs, G. H. (1985) Comp. Biochem. Physiol. 84B, 217–223. 16. Rozengurt, E., Heppel, L. A., and Friedberg, I. (1977) J. Biol. Chem. 252(13), 4584–4590. 17. Gordon, J. L. (1986) Biochem. J. 233, 309–319. 18. Steinberg, T. H., and Silverstein, S. C. (1987) J. Biol. Chem. 262(7), 3118–3122. 19. Steinberg, T. H., Newman, A. S., Swanson, J. A., and Silverstein, S. C. (1987) J. Biol. Chem. 262(18), 8884–8888. 20. Daniger, R. S., Raffaeli, S., Horeno-Sanches, R., Sakai, H., Capogrossi, M. C., Sporgeon, H. A., Hansford, R. G., and Lakatta, E. G. (1988) Cell Calcium 9, 193–199. 21. Greenberg, S., Di Virgilio, F., Steinberg, T. H., and Silverstein, S. C. (1988) J. Biol. Chem. 263(21), 10337–10343. 22. Filippini, A., Taffs, R. E., Agui, T., and Sitkovsky, M. V. (1990) J. Biol. Chem. 265, 334–340. 23. Zanovello, P., Bronte, V., Rosato, A., Pizzo, P., and Di Virgilio, F. (1990) J. Immunol. 145, 1545–1550. 24. Steinberg, T. H., and Di Virgilio, F. (1991) Curr. Opin. Immunol. 3, 71–75. 25. Remaley, A. T., Das, S., Campbell, P. I., LaRocca, G. M., Pope, G. M., and Glew, R. H., and Kaplan, S. S. (1985) J. Biol. Chem. 260, 880–885. 26. Neubert, T. A., and Gottlieb, M. (1990) J. Biol. Chem. 265, 7236– 7242. 27. Gbenle, G. O., and Dwyer, D. M. (1992) Biochem. J. 285, 41–46. 28. Lopes, A. H. C. S., and McMahon-Pratt, D. (1989) J. Protozool. 36, 354–361. 29. Barankiewicz, J., Dosch, H. M., and Cohen, A. (1988) J. Biol. Chem. 263, 7094–7098. 30. Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) J. Biol. Chem. 193, 265–275.

arca

46

MEYER-FERNANDES ET AL.

31. Guilherme, A., Meyer-Fernandes, J. R., and Vieyra, A. (1991) Biochemistry 30, 5700–5706. 32. Lowry, H. O., and Lopez (1946) J. Biol. Chem. 162, 421–428. 33. Glynn, I. M., and Chappel, J. B. (1964) Biochem. J. 90, 147–149. 34. Fabiato, A., and Fabiato, F. (1979) J. Physiol. (Paris) 75, 423– 429. 35. Noel, F., and Cumps, J. (1989) Braz. J. Med. Biol. Res. 22, 433– 445. 36. Gottlieb, M., and Dwyer, D. M. (1981) Science 212, 939–941. 37. Gottlieb, M., and Dwyer, D. M. (1982) Mol. Cell. Biol. 2, 76–81. 38. Glew, R. H., Czuczman, M. S., Diven, W. F., Berens, R. L., Tope, M. T., and Katsoulis, D. E. (1982) Comp. Biochem. Physiol. 72B, 581–590. 39. Van Belle, H. (1972) Biochim. Biophys. Acta 289, 158–168. 40. Van Belle, H. (1976) Clin. Chem. 22, 972–976. 41. Knowles, A. F. (1988) Arch. Biochem. Biophys. 263, 264–271. 42. Ribeiro, J. M. C., and Garcia, E. S. (1980) J. Insect. Physiol. 26, 303–307. 43. Ribeiro, J. M. C., and Garcia, E. S. (1981) Experientia 37, 384– 386. 44. Ribeiro, J. M. C., and Garcia, E. S. (1981) J. Exp. Biol. 94, 219– 230. 45. Sarkis, J. J. F., Guimara˜es, J. A., and Ribeiro, J. M. C. (1986) Biochem. J. 233, 885–890. 46. Garcia-Ve´scovi, Soncini, F. C., and Groisman, E. A. (1996) Cell 84, 165–174. 47. Barkker-Grunwald, T., and Parduhn, H. (1993) Mol. Biochem. Parasitol. 57, 167–170. 48. Romero, P. J., and Ortiz, C. (1988) J. Membr. Biol. 101, 237– 246. 49. Barbacci, E., Filippini, A., De Cesaris, P., and Ziparo, E. (1996) Biochem. Biophys. Res. Commun. 222, 273–279. 50. Zilberstein, D., and Dwyer, D. M. (1985) Proc. Natl. Acad. Sci. USA 82, 1716–1720. 51. Zilberstein, D., and Dwyer, D. M. (1988) Biochem. J. 256, 13– 21.

AID

ABB 9933

/

6b32$$$244

03-26-97 22:21:25

52. Glew, R. H., Saha, A. K., Das, S., and Remaley, A. T. (1988) Microbiol. Rev. 52, 412–432. 53. Asai, T., O’Sullivan, W. J., and Tatibana, M. (1983) J. Biol. Chem. 258, 6816–6822. 54. Bermudes, D., Peck, K. R., Afifi, M. A., Beckers, C. J. M., and Joiner, K. A. (1994) J. Biol. Chem. 269, 29252–29260. 55. Asai, T., Miura, S., Sibley, L. D., Okabayashi, H., and Takeuchi, T. (1995) J. Biol. Chem. 270, 11391–11397. 56. Lin, S-H. (1985) J. Biol. Chem. 260(13), 7850–7856. 57. Lin, S-H., and Russel, W. (1988) J. Biol. Chem. 263, 12253– 12258. 58. Meghji, P., Pearson, J. D., and Slakey, L. L. (1995) Biochem. J. 308, 725–731. 59. Plesner, L. (1995) Int. Rev. Cytol. 158, 1–61. 60. Handa, M., and Guidotti, G. (1996) Biochem. Biophys. Res. Commun. 218, 916–923. 61. Wang, T-F., and Guidotti, G. (1996) J. Biol. Chem. 271, 9898– 9901. 62. Strobel, R. S., Nagy, A. K., Knowles, A. F., Buegel, J., and Rosemberg, M. D. (1996) J. Biol. Chem. 271, 16323–16331. 63. Vasconcelos, E. G., Nascimento, P. S., Meirelles, M. N. L., Verjovski-Almeida, S., and Ferreira, S. T. (1993) Mol. Biochem. Parasitol. 58, 205–214. 64. Vasconcelos, E. G., Ferreira, S. T., Carvalho, T. M. U., de Souza, W., Kettlun, A. M., Mancilla, M., Valenzuela, M. A., and Verjovski-Almeida, S. (1996) J. Biol. Chem. 271, 22139–22145. 65. Aurivillus, M., Hansen, O. C., Lazrek, M. B. S., Bock, E., and Obrink, B. (1990) FEBS Lett. 264, 267–269. 66. Cheung, P. H., Luo, W., Qui, Y., Zhang, X., Earley, K., Millirons, P., and Lin, S-H. (1993) J. Biol. Chem. 268, 24303–24310. 67. Stout, J. G., Strobel, R. S., and Kirley, T. L. (1995) J. Biol. Chem. 270, 11845–11850. 68. Margolis, R. N., Schell, M. J., Taylor, and Hubbard, A. L. (1990) Biochem. Biophys. Res. Commun. 166, 562–566. 69. Dubyak, G. R., and El-Motassim, C. (1993) Am. J. Physiol. 34, C577–C606. 70. Yagi, K., Nishino, I., Eguchi, M., Kitagawa, M., Miura, Y., and Mizoguchi, T. (1994) Biochem. Biophys. Res. Commun. 203, 1237–1243.

arca