Archives of Biochemistry and Biophysics Vol. 375, No. 2, March 15, pp. 304 –314, 2000 doi:10.1006/abbi.1999.1592, available online at http://www.idealibrary.com on
Ectonucleotide Diphosphohydrolase Activities in Entamoeba histolytica Fernanda S. Barros,* Lu´cia F. De Menezes,† Ana A. S. Pinheiro,* Edward F. Silva,‡ Angela H. C. S. Lopes,§ Wanderley De Souza,† and Jose´ R. Meyer-Fernandes* ,1 *Departamento de Bioquı´mica Me´dica, Instituto de Cieˆncias Biome´dicas, †Instituto de Biofı´sica Carlos Chagas Filho, §Instituto de Microbiologia Professor Paulo de Go´es, Universidade Federal do Rio de Janeiro, CCS, Bloco H, Cidade Universita´ria, Ilha do Funda˜o, 21541-590, Rio de Janeiro, RJ, Brazil; and ‡Departamento de Parasitologia, Universidade Federal de Minas Gerais, MG, Brazil
Received July 21, 1999, and in revised form October 15, 1999
In this work, we describe the ability of living cells of Entamoeba histolytica to hydrolyze extracellular ATP. In these intact parasites, whose viability was determined by motility and by the eosin method, ATP hydrolysis was low in the absence of any divalent metal (78 nmol P i/h/10 5 cells). Interestingly, in the presence of 5 mM MgCl 2 an ecto-ATPase activity of 300 nmol P i/h/10 5 cells was observed. The addition of MgCl 2 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 1.23 mM MgCl 2. Both activities were linear with cell density and with time for at least 1 h. The ecto-ATPase activity was also stimulated by MnCl 2 and CaCl 2 but not by SrCl 2, ZnCl 2, or FeCl 3. In fact, FeCl 3 inhibited both Mg 2ⴙ-dependent and Mg 2ⴙ-independent ecto-ATPase activities. The Mg 2ⴙ-independent ATPase activity was unaffected by pH in the range between 6.4 and 8.4, in which the cells were viable. However, the Mg 2ⴙ-dependent ATPase activity was enhanced concomitantly with the increase in pH. In order to discard the possibility that the ATP hydrolysis observed was due to phosphatase or 5ⴕ-nucleotidase activities, several inhibitors for these enzymes were tested. Sodium orthovanadate, sodium fluoride, levamizole, and ammonium molybdate had no effect on the ATPase activities. In the absence of Mg 2ⴙ (basal activity), the apparent K m for ATP 4ⴚ was 0.053 ⴞ 0.008 mM, whereas at saturating MgCl 2 concentrations, the corresponding apparent K m for Mg-ATP 2ⴚ for Mg 2ⴙ-dependent ecto-ATPase activity (difference between total and basal ecto-ATPase activity) was 0.503 mM ⴞ 0.062. Both ecto-ATPase activities were highly specific for ATP and were also able to hydrolyze ADP less efficiently. To identify the observed hydrolytic activiTo whom correspondence should be addressed. Fax: ⫹55-21-2708647. E-mail:
[email protected]. 1
304
ties as those of an ecto-ATPase, we used suramin, a competitive antagonist of P 2 purinoreceptors and an inhibitor of some ecto-ATPases, as well as the impermeant agent 4ⴕ-4ⴕ-diisothiocyanostylbenzene-2ⴕ-2ⴕ-disulfonic acid. These two reagents inhibited the Mg 2ⴙindependent and the Mg 2ⴙ-dependent ATPase activities to different extents, and the inhibition by both agents was prevented by ATP. A comparison among the ecto-ATPase activities of three amoeba species showed that the noninvasive E. histolytica and the free-living E. moshkovskii were less efficient than the pathogenic E. histolytica in hydrolyzing ATP. As E. histolytica is known to have a galactose-specific lectin on its surface, which is related to the pathogenesis of amebiasis, galactose was tested for an effect on ectoATPase activities. It stimulated the Mg 2ⴙ-dependent ecto-ATPase but not the Mg 2ⴙ-independent ATPase activity. © 2000 Academic Press Key Words: Entamoeba histolytica; ecto-ATPase; ecto-phosphatase.
Entamoeba histolytica is an enteric protozoan that infects 10% of the world’s human population. This parasite causes 50 to 100 million cases of invasive colitis or liver abscess and up to 100,000 deaths per year (1, 2). Amebiasis is one of the great parasitic diseases of humankind, which predominantly affects individuals of lower socioeconomic status, who live in developing countries. During its life cycle, E. histolytica exists in either trophozoite or cyst form. Infection is acquired by ingestion of the cyst form, which is resistant to the acid pH of the stomach, and excystation occurs in the small bowel, in which the cysts divide and differentiate into trophozoites (1). The pathogenic, invasive E. histolytica and the most prevalent, nonpathogenic E. dispar are two morphologically indistinguishable species that ex0003-9861/00 $35.00 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved.
ECTO-ATPases IN Entamoeba histolytica
hibit several antigenic, genetic, and isoenzymatic differences (1, 3). Cell– cell recognition and adherence are processes central to many fundamental areas of biology. One common theme for many pathogenic microorganisms is the use of microbial lectins for colonization and invasion. It is not surprising that lectin-based protein– carbohydrate interactions are important in adherence, as the extracellular surfaces of eukaryotic cells are rich in complex carbohydrates that are attached to glycoproteins and glycolipids. Lectins make up a class of proteins that bind carbohydrates specifically and noncovalently (4). Parasite recognition of glycoconjugates plays an important role in the pathogenesis of amebiasis. Killing of host cells by E. histolytica trophozoites in vitro occurs only upon direct contact, which is mediated by an amebic adhesin that recognizes N- and O-linked oligosaccharides (5). This adhesin is specifically inhibited by millimolar concentrations of galactose (Gal) 2 and N-acetyl-D-galactosamine (GalNAc) and avoidance of membrane lysis by the complement complex attack is also mediated by this lectin (6). During the last two decades, considerable progress has been achieved in the study of ectonucleotidases in general (7) and ecto-ATPases in particular (8, 9). This progress is related to the finding that, in contrast to the established view, nucleotides can be found in significant concentrations outside cells (9, 10). Nucleotides released to the extracellular medium may exert their effects on other cells in the vicinity of the secretion site and modulate biological processes by binding to specific cell surface receptors (9). ATP may be one of the signaling molecules involved in cell-mediated cytotoxicity (11). It has been demonstrated that extracellular ATP has profound effects on cellular functions (11, 12), causing plasma membrane depolarization, Ca 2⫹ influx, and cell death (11, 12). Fillipini et al. (12) have shown that ATP can kill various types of cells, with the exception of those that express a high level of ATP-breakdown activity on their surface. Ecto-ATPases are glycoproteins present in the plasma membrane and have their active sites facing the external medium rather than the cytoplasm (8). The activities of these enzymes can be measured using viable cells and can be modulated by divalent cations, such as Mg 2⫹ and Ca 2⫹ (13). A previous study on nucleotidases of E. histolytica (14) described the activity of a Ca 2⫹-dependent nucleoside diphosphatase associated with the particulate fraction. This enzyme was shown to be responsible for the hydrolysis of ADP, UDP, GDP, and thiamine pyrophosphate. Subsequently, McCaul and Bird (15) showed that thiamine pyrophosphatase 2
Abbreviations used: Gal, galactose; GalNAc, N-acetyl-D-galactosamine; p-NPP, p-nitrophenylphosphate; DIDS, 4⬘-4⬘-diisothiocyanostylbenzene-2⬘-2⬘-disulfonic acid; NTPase, nucleoside triphosphate hydrolase.
305
was associated with the cytoplasmic vacuole of this parasite. Moreover, Serrano et al. (16) reported the presence of a Mg 2⫹-dependent ATPase activity in the plasma membrane of this parasite. Recently, upon addition of ATP to intact cells of E. histolytica, it was demonstrated that a Ca 2⫹–ATPase activity of this parasite is exposed to the medium and is inhibited by extracellular addition of the impermeant reagent diazotized sulfanilic acid (17). It has been proposed that this E. histolytica ecto-ATPase could participate in a chain of nucleosidase activities on the surface of the parasite and that this pathway could provide the parasites with a source of extracellular adenosine (17), which is known to be part of the dietary requirement of E. histolytica and is taken up by these cells (18). Here, we show the presence of Mg 2⫹-independent and Mg 2⫹-dependent ecto-ATPase activities on the cell surface of living E. histolytica trophozoites. We characterize the properties of these enzymes and demonstrate the effects of galactose. We also compare ATP hydrolysis by the invasive E. histolytica with that catalyzed by the noninvasive E. histolytica and the freeliving E. moshkovskii. MATERIAL AND METHODS Reagents. All reagents were purchased from E. Merck (D-6100 Darmstadt, Germany) or Sigma Chemical Co. (St. Louis, MO). [␥ 32P]ATP was prepared as described by Glynn and Chappell (19). Distilled water was deionized using a MilliQ system of resins (Millipore Corp., Bedford, MA) and was used in the preparation of all solutions. Concentrations of free and complexed species at equilibrium were calculated by using an iterative computer program that was modified (20) from that described by Fabiato and Fabiato (21). Microorganisms. Invasive E. histolytica (strain HM1:IMSS), noninvasive E. histolytica (strain RPS) (22), and free-living E. moshkovskii (strain FIC) trophozoites were cultured axenically in TYIS-33 medium (22, 23), supplemented with 10% adult bovine serum and 3% vitamin (Diamond Vitamin, Tween 80; JRH Biosciences) at 37°C, and were harvested at the stationary phase of growth, 48 h after inoculation. The parasites were then collected by centrifugation, washed twice, and kept in 116.0 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, 50.0 mM Hepes, pH 7.2. Cellular viability was assessed, before and after incubations, by motility and the eosin method (23). The viability was not affected by the conditions employed. Ecto-ATPase activity measurements. Intact cells were incubated for 1 h at 30°C in 0.5 ml of a mixture containing, unless otherwise specified, 116.0 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, 50.0 mM Hepes–Tris buffer, pH 7.2, 5.0 mM ATP, and 1.0 ⫻ 10 6 cells/ml, in the presence or in the absence of 5.0 mM MgCl 2. The ATPase activity was determined by measuring the hydrolysis of [␥- 32P]ATP (10 4 Bq/nmol ATP) (24). The experiments were started by the addition of living cells and terminated by the addition of 1.0 ml of a cold mixture containing 0.2 g charcoal in 1.0 M HCl. The tubes were then centrifuged at 2000 rpm for 10 min at 4°C. Aliquots (0.5 ml) of the supernatant containing the released 32P i were transferred to scintillation vials containing 9.0 ml of scintillation fluid (2.0 g PPO in 1 liter of toluene). 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 cell number. In the experiments in which other nucleotides were used, the hydrolytic activity measured under the same conditions described above was assayed spectrophotometri-
306
BARROS ET AL.
cally by measuring the release of P i from the nucleotides (25). The values obtained for ATPase activities measured using both methods (colorimetric and radioactive) are exactly the same. In the experiments in which high concentrations of Ca 2⫹, Mn 2⫹, and Fe 3⫹ were tested, possible precipitates formed were checked as previously described (26). Under the conditions employed, in the reaction medium containing 50 mM Hepes, pH 7.2, 116 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, and 5 mM ATP, no phosphate precipitates were observed in the presence of these cations. Phosphatase measurements. In addition to the measurements of ecto-ATPase activity, the ecto-p-nitrophenylphosphatase activity was determined in the same medium as that for ATP hydrolysis, except that ATP was replaced by 5.0 mM p-nitrophenylphosphate (p-NPP). The reaction was determined spectrophotometrically at 425.0 nm using an extinction coefficient of 14.3 ⫻ 10 3 M ⫺1 cm ⫺1. Statistical analysis. All experiments were performed in triplicate, with similar results obtained in at least three separate cell suspensions. Apparent K m and V max values were calculated using a computerized nonlinear regression analysis of the data to the Michaelis–Menten equation (24). Statistical significance was determined by Student’s t test. Significance was considered as P ⬍ 0.05.
RESULTS
The invasive E. histolytica presented two ectoATPase activities on its external surface, a Mg 2⫹-independent ATPase (activity measured in the absence of any cation added or in the presence of 1 mM EDTA) and a Mg 2⫹-dependent ATPase. At pH 7.2, in the absence of MgCl 2, living trophozoites of invasive E. histolytica were able to hydrolyze ATP (78.0 nmol P i/h/10 5 cells), ADP (39.0 nmol P i/h/10 5 cells), and p-NPP (3.0 nmol P i/h/10 5 cells) (Fig. 1). As can be seen in Fig. 1, the Mg 2⫹-stimulated ecto-enzyme activities (calculated as total [measured in the presence of 5 mM MgCl 2] minus basal ecto-enzyme activities) present in these parasites hydrolyzed ATP (258.0 nmol P i/h/10 5 cells), ADP (82.0 nmol P i/h/10 5 cells) and p-NPP (1.2 nmol P i/h/10 5 cells). These intact cells were unable to hydrolyze 5⬘ AMP in the absence or in the presence of 5 mM MgCl 2 (Fig. 1). To check the possibility that the observed ATP hydrolysis was the result of secreted soluble enzymes, as seen in Tetrahymena thermophila (27), we prepared a reaction mixture with cells that were incubated in the absence of ATP. Subsequently, the suspension was centrifuged to remove the cells and the supernatant was checked for ATPase activity. This supernatant failed to show ATP hydrolysis either in the absence or in the presence of MgCl 2 (data not shown). The time course of hydrolysis by the ecto-ATPase present on the surface of E. histolytica was linear for at least 60 min for both activities (Fig. 2). Similarly, in assays to determine the influence of cell density, Mg 2⫹independent, as well as the Mg 2⫹-dependent activities, measured over 60 min, were linear over a nearly 10fold range of cell densities (Fig. 3). Recently, it was shown that Mg 2⫹ is an important extracellular signal in the regulation of Salmonella virulence (28). The addition of MgCl 2 to the extracellular medium increased the ecto-ATPase activity of E. histolytica in a dose-dependent manner (Fig. 4A). At 5
FIG. 1. Influence of MgCl 2 on the ecto-phosphohydrolase activities of intact cells of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the reaction medium (final volume: 0.5 ml) containing 50 mM Hepes, pH 7.2, 116 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, 1.0 ⫻ 10 6 cells/ml, and 5 mM of either ATP, ADP, 5⬘ AMP, or p-NPP. Hatched bars: Mg 2⫹-stimulated activity (total activity, measured in the presence of 5 mM MgCl 2, minus the basal activity). Black bars: basal activity (measured in the absence of MgCl 2). In these experiments, ATP hydrolysis was measured using the same colorimetric assay (described under Material and Methods) for P i release as that used for the other nucleotides. Data are means ⫾ SE of three determinations with different cell suspensions.
mM ATP, half-maximal stimulation of ATP hydrolysis was obtained with 1.23 mM MgCl 2. Very low concentrations of Fe 3⫹ inhibited the Mg 2⫹-dependent ATPase in a dose-dependent manner, whereas the Mg 2⫹-independent ecto-ATPase activity was less sensitive to Fe 3⫹, being inhibited only by concentrations higher than 3.0 mM Fe 3⫹ (Fig. 4B). Low concentrations of Ca 2⫹ and Mn 2⫹ also stimulated the ecto-ATPase activity, whereas high concentrations of these cations inhibited it (Figs. 4C and 4D, respectively). The presence of an acid phosphatase on the plasma membrane of E. histolytica has already been described (29). A possible contribution to ATP hydrolysis caused by acid phosphatase activity was examined by measuring pH dependence of both activities. As shown in Fig. 5, in the pH range from 6.4 to 8.4, in which the cells were alive throughout the time course of reaction, the Mg 2⫹-independent (A) and the Mg 2⫹-dependent (B) phosphatase activities decreased. On the other hand, no effect of pH was observed on the Mg 2⫹-independent ATPase activity (lower curve, Fig. 6). However, the Mg 2⫹-dependent ATPase activity in this pH range progressively increased to reach a maximal level at pH 8.4 (upper curve, Fig. 6). In Leishmania tropica, it has also been reported that the ecto-Mg 2⫹-ATPase activity is stimulated by an increase in pH (30). To confirm the conclusion that the ATP hydrolysis was not due to phosphatase or to some other type of ATPase activities, different inhibitors were used. In Table I, it is shown that NaF and sodium orthovana-
ECTO-ATPases IN Entamoeba histolytica
307
FIG. 2. Time course of the ecto-ATPase activities of intact cells of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the reaction medium (final volume: 0.5 ml) containing 50 mM Hepes, pH 7.2, 116 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, 1 ⫻ 10 6 cells/ml, and 5 mM Tris–ATP [␥- 32P]ATP [sp act ⫽ 10 4 Bq/nmol ATP], without (A) or with (B) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
date, two potent inhibitors of acid phosphatase activities (31, 32), strongly inhibited the Mg 2⫹-independent and the Mg 2⫹-dependent phosphatase activities, whereas no effect was observed on the ATPase activities. Levamizole, a specific inhibitor of alkaline phosphatases (33, 34), also failed to inhibit the ATPase activities and the phosphatase activities (Table I). Moreover, p-NPP, a substrate for phosphatases, did not inhibit the ATPase activities. Another possible explanation for the ATP hydrolysis might be a 5⬘-nucleotidase (35). However, as shown in Fig. 1, these intact cells were unable to hydrolyze 5⬘ AMP in the absence or in the presence of 5 mM MgCl 2. Moreover, the lack of response to ammonium molybdate (Table I), a potent inhibitor of 5⬘-nucleotidase activity (35), indicates that this enzyme did not contribute to the ATP hydrolysis. It is also shown in Table I that the ATPase and phos-
phatase activities were insensitive to oligomycin and sodium azide, two known inhibitors of mitochondrial Mg-ATPase, to bafilomycin A 1, a V-ATPase inhibitor, and to ouabain, a Na/K-ATPase inhibitor (36). The ATP hydrolysis described here may be due to an ATPdiphosphohydrolase activity, as it was inhibited by high concentrations of ADP and AMP, but not by adenosine (Table I). The Mg 2⫹-dependent and the Mg 2⫹-independent ecto-ATPase activities showed different kinetic patterns with respect to the ATP concentration (Fig. 7). The Mg 2⫹-independent activity had an apparent K m for ATP 4⫺ of 0.053 ⫾ 0.008 mM and a V max of 87.6 ⫾ 9.2 nmol P i/h/10 5 cells (Fig. 7A). For the Mg 2⫹-dependent ecto-ATPase activity, the apparent K m for MgATP 2⫺ was 0.503 ⫾ 0.062 mM and the V max was 276.6 ⫾ 30.6 nmol P i/h/10 5 cells (Fig. 7B). We also analyzed the
FIG. 3. Cell density dependence of the ecto-ATPase activity of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the same reaction medium (final volume: 0.5 ml) as that described in the legend of Fig. 2, without (A) or with (B) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
308
BARROS ET AL.
FIG. 4. Influence of different cation concentrations on the ecto-ATPase activities of intact cells of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the same reaction medium (final volume: 0.5 ml) as that described in the legend of Fig. 2, with the addition of increasing concentrations of Mg 2⫹ (A), Fe 3⫹ [with (F) or without (E) the addition of 5 mM MgCl 2] (B), Ca 2⫹ (C), and Mn 2⫹ (D). Data are means ⫾ SE of three determinations with different cell suspensions.
specificity of these ecto-ATPase activities for various other nucleotides. In Table II, it is shown that ATP was the best substrate for these enzymes, although they
were also able to hydrolyze ADP. Other nucleoside 5⬘-triphosphates (ITP, CTP, GTP, and UTP) produced very low reaction rates.
FIG. 5. Effect of pH on the p-nitrophenylphosphatase activities of intact cells of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the reaction medium (final volume: 0.5 ml) containing 116 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, 5 mM p-NPP, 1.0 ⫻ 10 6 cells/ml, and 50 mM Hepes–Tris buffer, adjusted to pH values between 6.4 and 8.4. In this pH range (6.4 – 8.4), the cells were viable thougout the course of the experiments. The assays were performed without (A) or with (B) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
ECTO-ATPases IN Entamoeba histolytica
309
much higher ATPase activities than the noninvasive E. histolytica and the free-living E. moshkovskii. It is well known that the pathogenesis of invasive amebiasis is due to an amebic Gal- and GalNAc-inhibitable adherence lectin that exclusively mediates the adherence of E. histolytica to mammalian target cells (6, 39, 40). The physiological role of the ecto-ATPases is still unknown, but a possible involvement in cellular adhesion has been suggested (41– 43). For these reasons, we examined the effect of D-galactose on the ectoATPase activities of this parasite (Fig. 10). The Mg 2⫹dependent ecto-ATPase was stimulated more than twofold by 50 mM D-galactose (Fig. 10B), although the Mg 2⫹-independent ecto-ATPase was not affected by this carbohydrate (Fig. 10A). D-Glucose had no effect on either of the hydrolytic activities tested. FIG. 6. Effects of pH on the ecto-ATPase activities of intact cells of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the reaction medium (final volume: 0.5 ml) containing 116 mM NaCl, 5.4 mM KCl, 5.5 mM D-glucose, 1 ⫻ 10 6 cells/ml, 5 mM Tris–ATP [␥- 32P]ATP [sp act ⫽ 10 4 Bq/nmol ATP], and 50 mM Hepes–Tris buffer, adjusted to pH values between 6.4 and 8.4. In this pH range (6.4 – 8.4) the cells were viable througout the course of the experiments. The assays were performed with (F) or without (E) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
The Mg 2⫹-dependent and the Mg 2⫹-independent ATPase activities present on the external surface of E. histolytica were the same when cells were in log phase as when they were in stationary phase (data not shown). Since we used intact cells for measuring the enzyme activities in all the experiments done in this work, it likely that the described activities are ectoATPases. To confirm this, we applied the criterion that an authentic ectoenzyme should be inhibited by an added extracellular impermeant reagent (13, 30, 37) such as 4⬘-4⬘-diisothiocyanostylbene-2⬘-2⬘-disulfonic acid (DIDS) (30, 37), and by an inhibitor of some ectoATPases, such as suramin, which is also a competitive antagonist of P 2-purinergic receptors (38). Figure 8 shows that 1 mM DIDS inhibited 100% of the Mg 2⫹independent ecto-ATPase activity (A) and 60% of the Mg 2⫹-dependent activity (B) and that 1 mM suramin promoted a 54% reduction of the E. histolytica Mg 2⫹independent ecto-ATPase activity (A) and a 64% reduction of the Mg 2⫹-dependent activity (B). The inhibition promoted by either DIDS or suramin was suppressed when the cells were preincubated in the presence of 5 mM ATP (Figs. 8A and 8B, insets). The noninvasive E. histolytica and the free-living E. moshkovskii were compared to the pathogenic E. histolytica concerning the presence of Mg 2⫹-dependent and Mg 2⫹-independent ecto-ATPase activities. As shown in Fig. 9, all three species express the Mg 2⫹independent (A) and the Mg 2⫹-dependent (B) ectoATPase activities, and pathogenic E. histolytica showed
DISCUSSION
In this paper, we report the characterization of Mg 2⫹independent and Mg 2⫹-dependent ecto-ATPase activities present on the external surface of E. histolytica (Fig. 1). Cellular viability was assessed, before and after the reactions, by motility and by the eosin method (23). The viability was not affected by any of the conditions used in the assays. The external localization of the ATP-hydrolyzing site is supported by its sensitivity to the impermeant reagent DIDS and to suramin, which is a noncompetitive inhibitor of ecto-ATPases and an antagonist of P 2 purinoreceptors, which mediate the physiological functions of extracellular ATP (38, 44). Also, the use of a battery of inhibitors for other ATPases that have intracellular ATP binding sites showed no change in the ATPase activities, either in the absence or in the presence of Mg 2⫹ (Table I). For the above reasons, we interpret our data according to the criteria for ectolocalization of the ATPase activities here described (8). These criteria include the use of nonpenetrating substrates, which have access to the enzyme in whole, intact cells, the demonstration of product formation extracellularly, and the demonstration that the activity can be inhibited by exposure of intact cells to impermeant reagents (13, 30, 45). Our data are consistent with observations showing that ecto-ATPases are widely distributed in nature, as they have been described in plants (46), several mammalian cell types (8, 47), insects (48), helminth endoparasites (49, 50), and protozoa, including T. thermophila (27), Leishmania tropica (30), Toxoplasma gondii (51), and E. histolytica (17), among others. The ATPase activities shown in this work cannot be attributed to a mitochondrial ATPase, since this activity was insensitive to oligomycin and sodium azide (Table I), two known F-type ATPase inhibitors (30). In Leishmania donovani, a H ⫹-ATPase responsible for intracellular pH maintenance has been identified in the plasma membrane (52). This H ⫹-ATPase was de-
310
BARROS ET AL. TABLE I
Influence of Various Agents on ATPase and p-Nitrophenylphosphatase Activities of E. histolytica Relative activity (%) p-NPPase
ATPase
Additions
Independent of MgCl 2
Dependent on MgCl 2
Independent of MgCl 2
Dependent on MgCl 2
Control Levamizole (1.0 mM) Vanadate (1.0 mM) Molybdate (1.0 mM) Tartrate (10.0 mM) NaF (10.0 mM) Ouabain (1.0 mM) Azide (10.0 mM) Bafilomycin (1 mM) Oligomycin (2 g/ml) Adenosine (10.0 mM) AMP (10.0 mM) ADP (10.0 mM) P i (10.0 mM) p-NPP (10.0 mM)
100.0 ⫾ 9.8 100.8 ⫾ 12.5 5.7 ⫾ 0.7 9.2 ⫾ 1.3 90.1 ⫾ 10.8 12.6 ⫾ 1.7 95.8 ⫾ 7.4 98.6 ⫾ 8.9 91.4 ⫾ 10.6 98.1 ⫾ 12.6 112.6 ⫾ 10.8 81.0 ⫾ 7.2 62.8 ⫾ 7.1 59.3 ⫾ 4.8 —
100.0 ⫾ 8.7 95.2 ⫾ 12.9 24.1 ⫾ 3.4 31.3 ⫾ 4.2 98.1 ⫾ 13.8 36.7 ⫾ 3.7 98.7 ⫾ 10.7 93.3 ⫾ 8.7 93.0 ⫾ 9.7 95.4 ⫾ 10.4 103.2 ⫾ 12.5 79.2 ⫾ 6.6 41.5 ⫾ 3.2 75.9 ⫾ 8.1 —
100.0 ⫾ 7.6 95.3 ⫾ 12.4 92.8 ⫾ 8.6 92.5 ⫾ 10.7 104.7 ⫾ 11.8 81.5 ⫾ 9.3 100.7 ⫾ 12.7 100.4 ⫾ 7.9 100.3 ⫾ 8.6 100.3 ⫾ 12.8 94.9 ⫾ 10.9 60.4 ⫾ 7.2 61.6 ⫾ 4.4 98.9 ⫾ 7.9 104.1 ⫾ 12.0
100.0 ⫾ 12.4 95.7 ⫾ 9.4 92.2 ⫾ 7.4 92.7 ⫾ 12.6 104.8 ⫾ 11.6 81.0 ⫾ 6.9 100.8 ⫾ 10.5 100.6 ⫾ 9.6 100.0 ⫾ 11.7 100.6 ⫾ 7.3 94.4 ⫾ 12.9 60.3 ⫾ 7.9 58.2 ⫾ 6.6 98.8 ⫾ 10.1 99.0 ⫾ 8.7
Note. ATPase and phosphatase activities were measured in the standard assay described under Material and Methods with 5 mM ATP or 5 mM p-NPP. Activities are expressed as a percentage of that measured under control conditions, i.e., without other additions. The Mg 2⫹-independent ATPase (78.2 ⫾ 5.9 nmol P i/h/10 5 cells), the Mg 2⫹-dependent ATPase (258.6 ⫾ 31.9 nmol P i/h/10 5 cells), the Mg 2⫹independent p-NPPase (2.8 ⫾ 0.2 nmol P i/h/10 5 cells), and the Mg 2⫹-dependent p-NPPase (1.3 ⫾ 0.1 nmol P i/h/10 5 cells) activities were 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 values. The unpaired t test showed, in all measurements of ATPase activities, that there were no statistical differences (P ⬎ 0.05) from the control.
scribed as a P-type cation transport ATPase, highly sensitive to orthovanadate (53). Since the Mg 2⫹-independent and the Mg 2⫹-dependent ecto-ATPases here described did not respond to vanadate (Table I), the possibility that these activities were due to a H ⫹-
ATPase present on the surface of the plasma membrane (30) was discarded. This ATP hydrolysis cannot be due to a phosphatase activity present on the external surface of Entamoeba membrane because, as shown in Fig. 5, the ecto-phosphatase activities present on the exter-
FIG. 7. Dependence on ATP 4⫺ (A) and Mg-ATP 2⫺ (B) concentrations of ecto-ATPase activities of intact cells of invasive E. histolytica. Cells were incubated at 36°C in the same reaction medium (final volume: 0.5 ml) as that described in the legend of Fig. 2, which corresponds to ATP 4⫺ (A) or Mg-ATP 2⫺ (B) concentrations varying as shown on the abscissa. Curves represent the fit of experimental data by nonlinear regression using the Michaelis–Menten equation as described under Material and Methods. The total amounts of ATP and MgCl 2 necessary to form the desired ATP 4⫺ (A) and Mg-ATP 2⫺ (B) concentrations were calculated as described under Material and Methods. Data are means ⫾ SE of three determinations with different cell suspensions.
ECTO-ATPases IN Entamoeba histolytica TABLE II
Substrate Specificity of Ecto-ATPase Activities Relative activity Nucleotides
Mg 2⫹-independent
Mg 2⫹-dependent
ATP CTP GTP ITP UTP ADP
100.0 ⫾ 7.6 1.3 ⫾ 0.1 1.0 ⫾ 0.1 9.3 ⫾ 0.6 3.6 ⫾ 0.3 50.6 ⫾ 5.7
100.0 ⫾ 12.4 4.2 ⫾ 0.3 1.1 ⫾ 0.1 3.2 ⫾ 0.4 1.0 ⫾ 0.1 32.1 ⫾ 3.6
Note. ATPase activities were measured in a standard assay described under Material and Methods with the nucleotides listed (5 mM), without or with the addition of 5 mM MgCl 2. Reactions were at 30°C. The Mg 2⫹-independent ATPase (78.2 ⫾ 5.9 nmol P i/h/10 5 cells) and the Mg 2⫹-dependent ATPase (258.6 ⫾ 31.9 nmol P i/h/10 5 cells) activities were 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 values. In these experiments, ATP hydrolysis was measured using the same colorimetric assay of P i release from other nucleotides as that described under Material and Methods.
nal surface of E. histolytica are acid phosphatases, whereas the optimum pH for the Mg 2⫹-dependent ectoATPase lies in the alkaline range and the Mg 2⫹-independent activity was not affected by variation of the pH (Fig. 6). In addition, potent inhibitors for phosphatase activities were not capable of modifying the ectoATPase activities (Table I). Similar results were obtained for L. tropica Mg 2⫹-dependent ecto-ATPase, which also exhibits higher activity in alkaline pH and does not respond to phosphatase inhibitors (30). The Mg 2⫹-independent and the Mg 2⫹-dependent ectoATPase activities described here cannot be attributed to a 5⬘-nucleotidase, since AMP hydrolysis is not catalyzed by the intact cells (Fig. 1), and ammonium
311
molybdate does not inhibit the ecto-ATPase activities, although it does inhibit the phosphatase activities (Table I). These data are consistent with the previously described biochemical and ultrastructural localization of an ATPase, an ADPase, and an acid phosphatase and exclusion of a 5⬘-nucleotidase on the plasma membrane and on the inner surface of the cytoplasmic vacuoles in E. histolytica (16, 54). Here, we further characterize the E. histolytica ecto-ATPase activities, showing that the addition of MgCl 2 to the extracellular medium increased the ecto-ATPase activity in a dosedependent manner (Fig. 4A). Low concentrations of Ca 2⫹ and Mn 2⫹ also stimulated the ecto-ATPase activity, whereas high concentrations of these cations inhibited this activity (Figs. 4C and 4D, respectively). Very low concentrations of Fe 3⫹ (as low as 0.3 mM) were able to inhibit the Mg 2⫹-dependent ATPase but not the Mg 2⫹-independent ecto-ATPase activity (Fig. 4B). Differences in the response to pH variation observed for the two ecto-ATPases here described, as well as in sensitivity to the inhibition promoted by Fe 3⫹, suggest that these activities are due to different enzymes. As can be also seen in Figs. 7A and 7B, the Mg 2⫹-dependent ATPase activity was higher than the Mg 2⫹-independent ecto-ATPase activity, although with lower affinity for the corresponding substrates Mg-ATP 2⫺ and ATP 4⫺, respectively. It has been shown that the nucleoside triphosphate hydrolase (NTPase) purified from T. gondii is not a single enzyme but a mixture of two isozymes, termed NTPase I and NTPase II, and that a primary difference between these isozymes is that NTPase II hydrolyzes nucleoside triphosphate and diphosphate substrates at almost the same rate, whereas NTPase I hydrolyzes nucleoside diphosphate at a much slower rate (55). The Mg 2⫹-independent and the Mg 2⫹-dependent ectoATPases of E. histolytica characterized here demon-
FIG. 8. Effects of DIDS and suramin on the ecto-ATPase activities of intact cells of invasive E. histolytica. Cells were preincubated for 1 h at 36°C in the same reaction medium (final volume: 0.5 ml) as that described in the legend of Fig. 2, with or without the addition of 1 mM DIDS or suramin. After 1 h, the reaction was started by the addition of 5 mM Tris–ATP [␥- 32P]ATP [sp act ⫽ 10 4 Bq/nmol ATP]. (Insets) The cells were incubated in the same conditions described above in the presence of 5 mM ATP. The assays were performed without (A) or with (B) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
312
BARROS ET AL.
FIG. 9. Ecto-ATPase activities of intact cells of invasive E. histolytica (A), noninvasive E. histolytica (B), and free-living E. moshkovskii (C). Cells were incubated for 1 h at 36°C in the same reaction medium (final volume: 0.5 ml) as that described in the legend of Fig. 2, without (A) or with (B) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
strate high specificity for ATP, being much less active toward other nucleoside triphosphate substrates, such as ITP, CTP, UTP, and GTP (Table II), but are also able to hydrolyze ADP (Fig. 1, Table II). The ADP hydrolysis was 32 and 50% of the ATP hydrolysis for the Mg 2⫹-dependent and the Mg 2⫹-independent activities, respectively (Table II). Furthermore, ADP inhibited the two ATPase activities by about 40% (Table I). These data are highly suggestive that the ATP hydrolysis characterized here is due to an ATP-diphosphohydrolase activity, as already described in other organisms (9, 56 –59). In contrast, in L. tropica, a Mg 2⫹dependent ecto-ATPase that was not considered an ATP-diphosphohydrolase because of the lack of inhibition of the ATP hydrolysis by high concentrations of ADP at nonsaturating concentrations of ATP has been characterized (30). In T. gondii, it has been shown that different strains exhibit NTPases from two different groups, one group with ATPase/ADPase ratios of 4.0 or 5.0 and another group with ATPase/ADPase ratios of
approximately 1.0 (51). This ratio is similar to those shown by the ecto-ATPases of aortic and lung endothelial cells, erythrocytes, and other blood cells (8). The identity and function of ecto-ATPases have been reviewed and the nomenclature “E-type ATPases” was proposed for these enzymes (9), whose physiological role is still unknown. However, several hypotheses have been suggested, such as protection from cytolytic effects of extracellular ATP (11, 12), regulation of ectokinase substrate concentration (9), involvement with cellular adhesion and signal transduction (40, 60, 61), and regulation of NO release in smooth muscle cells (62). Considering the fact that E. histolytica is responsible for complicated infections, whose ulceration of the intestinal mucosa allows the parasite to penetrate into the host circulation, one could speculate that the presence of an ecto-ATPase activity might reflect some form of evasion of the parasite from host defense mechanisms in the circulation (1, 8). A highly active ectoATP-diphosphohydrolase was localized on the external
FIG. 10. Effects of D-glucose and D-galactose on the ecto-ATPase activities of intact cells of invasive E. histolytica. Cells were incubated for 1 h at 36°C in the same reaction medium (final volume: 0.5 ml) as that described in the legend of Fig. 2, in the absence (Control) or in the presence of 50 mM glucose (Glu) or 50 mM galactose (Gal) without (A) or with (B) the addition of 5 mM MgCl 2. Data are means ⫾ SE of three determinations with different cell suspensions.
ECTO-ATPases IN Entamoeba histolytica
surface of the tegument of Schistosoma mansoni (49, 50) and the authors suggested that this enzyme might be involved in an escape mechanism allowing the parasite to split ATP or ADP released in its vicinity by activated platelets or activated cytotoxic T lymphocytes (49, 50). The results presented in Fig. 9 demonstrate that the pathogenic amoeba E. histolytica has much higher ecto-ATPase activities than the noninvasive amoeba E. histolytica or the free-living amoeba E. moshkovskii. It is well known that coevolution of parasites and hosts is based upon gain or loss of certain characteristics that are acted on by natural selection, so that one organism becomes adapted to live as a parasite of the other, which becomes its host (63). An example is the acquisition of cytoplasmic virus-like particles by some strains of Endotrypanum spp., which has been attributed to the apparent faster rate of evolution and to the geographic distribution of these trypanosomatid parasites, as compared to other strains of the same genus (64, 65). Differences in the structure of glycoconjugates that are exposed on the parasites’ exterior surfaces are certainly among the most important acquisitions for intracellular parasitism (66 – 68). Since ecto-ATPases are glycoproteins themselves (8, 9), it is possible that the differences observed here in the ecto-ATPase activities of the free-living E. moshkovskii, the noninvasive E. histolytica, and the pathogenic E. histolytica (Fig. 9) were significant in the evolution of these species, a process occurring over a long period of time, during which they became so physiologically distinct that they remain taxonomically separate (2, 3). In T. gondii, it has been shown that there is no difference between the ATPase activities of virulent and those of avirulent strains, although it was suggested that these activities must be essential for its intracellular parasitism (51). If ecto-ATPases serve as adhesion molecules in Entamoeba, they could be considered pathogenesis markers for these cells. Surface membrane interactions between E. histolytica and its host are of critical importance, and understanding the biochemical basis for parasite adherence and cytolytic activities is crucial for vaccine development. Killing of host cells by E. histolytica trophozoites in vitro occurs only upon direct contact, which is mediated by an amebic adhesin, a lectin that recognizes N- and O-linked oligosaccharides (6, 39). This lectin is specifically inhibited by millimolar concentrations of galactose and N-acetyl-D-galactosamine (6). The essential role of the Gal/GalNAc lectin in amebic adherence to target cells has been demonstrated using carbohydrate inhibitors and using target cells deficient in glycosylation. In adherence assays in vitro, 50 mM Gal or GalNAc inhibited about 90% of the binding of E. histolytica trophozoites to Chinese hamster ovary cells, whereas other sugars had no effect (69, 70). When we investigated the possible effect of D-galactose on both ecto-
313
ATPases present in E. histolytica, 50 mM D-galactose stimulated the Mg 2⫹-dependent ecto-ATPase activity by more than twofold but did not interfere with the Mg 2⫹-independent ecto-ATPase activity (Fig. 10). Recently, it was shown that a 46-kDa lectin isolated from root extracts of the legume Doliclos biflorus is a Nod factor binding protein as well as a nucleoside di- and triphosphate hydrolase stimulated by carbohydrate ligands (71). As it has been suggested that ecto-ATPases may be involved with cellular adhesion (41, 60, 61), we intend to investigate whether the amebic lectin (6, 39) may be associated with this Mg 2⫹-dependent ecto-ATPase. ACKNOWLEDGMENTS We thank Dr. Martha M. Sorenson for critical reading of the manuscript. This work was partially supported by grants from the Brazilian agencies Conselho Nacional de Desenvolvimento Cientı´fico e Tecnolo´gico (CNPq), Financiadora de Estudos e Projetos (FINEP), Fundac¸a˜o de Amparo a` Pesquisa do Estado do Rio de Janeiro (FAPERJ), and Programa de Nu´cleos de Exceleˆncia (PRONEX, Grant 0885).
REFERENCES 1. Ravdin, J. (1989) J. Infect. Dis. 159, 420 – 429. 2. Aucott, J. N., and Ravdin, J. I. (1993) Infect. Dis. Clin. North Am. 7, 467– 485. 3. Jackson, T. F. H. G. (1998) Int. J. Parasitol. 28, 181–186. 4. Sharon, N., and Lis, H. (1989) Science 246, 227–234. 5. Guerrant, R. L, Brush, J., Ravdin, J. I., Sullivan, J. A., and Mandell G. L. (1981) J. Infect. Dis. 143, 83–93. 6. Ravdin, J. I., and Guerrant, R. L. (1981) J. Clin. Invest. 68, 1305–1313. 7. Zimmermann, H. (1996) Drug Dev. Res. 39, 337–352. 8. Plesner, L. (1995) Int. Rev. Cytol. 158, 141–214. 9. Dombrowski, K., Ke, Y., Brewer, K. A., and Kapp, J. A. (1998) Immunol. Rev. 161, 111–118. 10. Gordon, J. L. (1986) Biochem. J. 233, 309 –319. 11. Steinberg, T. H., and Di Virgilio, F. (1991) Curr. Opin. Immunol. 3, 71–75. 12. Filippini, A., Taffs, R. E., Agui, T., and Sitkovsky, M. V. (1990) J. Biol. Chem. 265, 334 –340. 13. De Pierre, J. W., and Karnovsky, M. L. (1973) J. Cell Biol. 56, 275–303. 14. McLaughlin, J., Lindmark, D. G., and Mu¨ller, M. (1978) Biochem. Biophys. Res. Commun. 82, 913–920. 15. McCaul, T. F., and Bird, R. G. (1978) Int. J. Parasitol. 8, 501– 506. 16. Serrano, R., Deas, J. E., and Swarren, L. G. (1977) Exp. Parasitol. 41, 370 –384. 17. Bakker-Grunwald, T., and Parduhn, H. (1993) Mol. Biochem. Parasitol. 57, 167–170. 18. Boonlayagoor, P., Albach, R. A., Stern, M. L., and Booden, T. (1978) Exp. Parasitol. 45, 225–233. 19. Glynn, I. M., and Chappell, J. B. (1946) Biochem. J. 90, 147–149. 20. Sorenson, M. M., Coelho, H. S. L., and Reuben, J. P. (1986) J. Membrane Biol. 90, 219 –230. 21. Fabiato, A., and Fabiato, F. (1979) J. Physiol. (Paris) 75, 423– 429.
314
BARROS ET AL.
22. Gomes, M. A., Silva, E. F., Macedo, M. A., Vago, A. R. and Melo, M. N. (1997) Parasitology 114, 517–520. 23. Diamond, L. S., Harlow, D. R., and Cunnick, C. C. (1978) Trans. R. Soc. Trop. Med. Hyg. 72, 431. 24. Guilherme, A., Meyer-Fernandes, J. R., and Vieyra, A. (1991) Biochemistry 90, 147–149. 25. Lowry, H. O., and Lopez, M. (1946) J. Biol. Chem. 162, 421– 428. 26. Meyer-Fernandes, J. R., and Vieyra, A. (1988) Arch. Biochem. Biophys. 266, 132–141. 27. Smith, T. M., Kirley, T. L., and Hennessey, T. M. (1997) Arch. Biochem. Biophys. 337, 351–359. 28. Garcia-Ve´scovi, Soncini, F. C., and Groisman, E. A. (1996) Cell 84, 165–174. 29. Anaya-Ruiz, M., Rosales-Encina, J. L., and Talamas-Rohama, P. (1997) Arch. Med. Res. 28, 182–183. 30. Meyer-Fernandes, J. R., Dutra, P. M. L., Rodrigues, C. O., SaadNehme, J., and Lopes, A. H. C. S. (1997) Arch. Biochem. Biophys. 341, 40 – 46. 31. Fernandes, E. C., Meyer-Fernandes, J. R., Silva-Neto, M. A. C., and Vercesi, A. E. (1997) Z. Naturforsch. 52c, 351–358. 32. Dutra, P. M. L., Rodrigues, C. O., Jesus, J. B., Lopes, A. H. C. S., Souto-Padro´n, T., and Meyer-Fernandes, J. R. (1998) Biochem. Biophys. Res. Commun. 253, 164 –169. 33. Van Belle, H. (1972) Biochim. Biophys. Acta 289, 158 –168. 34. Van Belle, H. (1976) Clin. Chem. 22, 972–976. 35. Gottlieb, M., and Dwyer, D. M. (1983) Mol. Biochem. Parasitol. 7, 303–317. 36. Cunningham, H. B., Yazak, P. J., Domingo, R. C., Oades, K. V., Bohlen, H., Sabbadini, R. A., and Dahms, A. S. (1993) Arch. Biochem. Biophys. 303, 32– 43. 37. Knowles, A. F. (1988) Arch. Biochem. Biophys. 263, 264 –271. 38. Hourani, S. M. O., and Chown, J. A. (1989) Gen. Pharmacol. 20, 413– 416. 39. Ravdin, J. I., Croft, B. Y., and Guerrant, R. L. (1980) J. Exp. Med. 152, 377–390. 40. Salata, R. A., Pearson, R. P., Murphy, C. F., and Ravdin, J. I. (1985) J. Clin. Invest. 76, 491– 499. 41. Stout, J. G., Strobel, R. S., and Kirley, T. L. (1995) J. Biol. Chem. 270, 11845–11850. 42. Cheung, P. H., Luo, W., Qui, Y., Zhang, X., Earley, K., Milirons, P., and Lin, S-H. (1993) J. Biol. Chem. 268, 24303–24310. 43. Aurivillus, M., Hansen, O. C., Lazrek, M. B. S., Bock, E., and Obrink, B. (1990) FEBS Lett. 264, 267–269. 44. Chen, B. C., Lee, C-M, and Lin, W-W. (1996) Br. J. Pharmacol. 119, 1628 –1634. 45. Trams, E. G., and Lauter, C. J. (1974) Biochim. Biophys. Acta 345, 180 –197. 46. Traverso-Cori, A., Chaimovich, H., and Cori, O. (1965) Arch. Biochem. Biophys. 109, 173–184. 47. Komoszynski, M., and Wojtczak, A. (1996) Biochim. Biophys. Acta 1310, 233–241.
48. Ribeiro, J. M. C., and Garcia, E. S. (1980) J. Insect Physiol. 26, 303–307. 49. Vasconcelos, E. G., Nascimento, P. S., Meirelles, M. N. L., Verjovski-Almeida, S., and Ferreira, S. T. (1993) Mol. Biochem. Parasitol. 58, 205–214. 50. 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, 267–269. 51. Asai, T., and Suzuki, Y. (1990) FEMS Microbiol. Lett. 72, 89 –92. 52. Zilberstein, D., and Dwyer, D. M. (1985) Proc. Natl. Acad. Sci. USA 82, 1716 –1720. 53. Zilberstein, D., and Dwyer, D. M. (1988) Biochem. J. 256, 13–21. 54. Kobayashi, S., Takeushi, T., Asami, K., and Fujiwara, T. (1981) Exp. Parasitol. 54, 202–212. 55. Asai, T., Miura, S., Sibley, L. D., Okabayashi, H., and Takeuchi, T. (1995) J. Biol. Chem. 270, 11391–11397. 56. Sarkis, J. J. F., Guimara˜es, J. A., and Ribeiro, J. M. C. (1986) Biochem. J. 233, 885– 890. 57. Handa, M., and Guidotti, G. (1996) Biochem. Biophys. Res. Commun. 218, 916 –923. 58. Wang, T-F., and Guidotti, G. (1996) J. Biol. Chem. 271, 9898 – 9901. 59. Zimmermann, H. (1999) Trends Pharmacol. Sci. 20, 231–236. 60. Margolis, R. N., Schell, M. J., Taylor, and Hubbard, A. L. (1990) Biochem. Biophys. Res. Commun. 166, 562–566. 61. Dubyak, G. R., and El-Motassin, C. (1993) Am. J. Physiol. 34, C577–C606. 62. Yagi, K., Nishiro, I., Eguchi, M., Kitagawa, M., Miura, Y., and Mizoguchi, T. (1994) Biochem. Biophys. Res. Commun. 203, 1237–1243. 63. Ridley, M. (1993) in Evolution, pp. 267–301, Blackwell Sci., Cambridge. 64. Lopes, A. H. C. S., Iovannisci, D., Petrillo-Peixoto, M. L., McMahon-Pratt, D., and Beverley, S. M. (1990) Mol. Biochem. Parasitol. 49, 151–162. 65. Fernandes, A. P., Nelson, K., and Beverley, S. M. (1993) Proc. Natl. Acad. Sci. USA 90, 11608 –11612. 66. Maue¨l, J. (1996) Adv. Parasitol. 38, 1–51. 67. Routier, F. H., Xavier da Silveira, E., Wait, R., Jones, C., Previato, J. O., and Mendonc¸a-Previato, L. (1995) Mol. Biochem. Parasitol. 69, 81–92. 68. Xavier da Silveira, E., Jones, C., Wait, R., Previato, J. O., and Mendonc¸a-Previato, L. (1998) Biochem. J. 329, 665– 673. 69. Saffer, L. D., and Petri, W. A., Jr. (1991) Exp. Parasitol. 72, 106 –108. 70. McCoy, J. J., Mann, B. J., and Petri, W. A., Jr. (1994) Infect. Immun. 62, 3045–3050. 71. Etzler, M. E., Kalsi, G., Ewing, N. N., Roberts, N. J., Day, R. B., and Murphy, J. B. (1999) Proc. Natl. Acad. Sci. USA 96, 5856 – 5861.
