Experimental Parasitology 124 (2010) 459–465
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Research Brief
Entamoeba histolytica: Identification and partial characterization of a-mannosidase activity Clara E. Santacruz-Tinoco, Julio C. Villagómez-Castro, Everardo López-Romero * Departamento de Biología, División de Ciencias Naturales y Exactas, Universidad de Guanajuato, Apartado Postal No. 187, Guanajuato, Gto. 36000, Mexico
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Article history: Received 18 March 2009 Received in revised form 16 October 2009 Accepted 21 December 2009 Available online 4 January 2010 Keywords: Entamoeba histolytica N-Protein glycosylation a-Mannosidases
a b s t r a c t Despite their well recognized importance in pathogenesis of Entamoeba histolytica there are few studies dealing with the assembly and secretion of glycoproteins that participate in the adhesion to target cells and in the dissemination of the parasite in infected tissues. Some of these studies refer to the identification and, in some cases, the characterization of glycosyl transferases and glycosidases involved in the biosynthesis of these macromolecules as well as to compartments involved in the amoeba dolichol-linked glycosylation pathway. While an N-glycan trimming a-mannosidase has been demonstrated in E. histolytica, little is known on its cellular distribution and properties. Here we describe the presence and partial biochemical characterization of soluble and MMF-associated forms of a-mannosidase and the separation of at least three internal membrane structures enriched with this glycosidase. Results are discussed in terms of the possible identity of a-mannosidase activity and the potential precursor–product relationship between the two enzyme forms. Ó 2009 Elsevier Inc. All rights reserved.
1. Introduction
a-Mannosidases participate in the processing of Asn-linked oligosaccharides providing trimmed N-glycans that serve as precursors of complex glycoproteins in a vast diversity of organisms. They also participate in the catabolism of N-oligosaccharides, a function that takes place mostly in the cytosol, vacuoles and lysosomes (Daniel et al., 1994; Herscovics, 1999b), as well as in the endoplasmic-reticulum associated degradation (ERAD) as part of the glycoprotein quality control process (Helenius and Aebi, 2004). Based on aminoacid sequence analysis and some biochemical properties, a-mannosidases from different species are grouped into glycosylhydrolase families 47 (EC 3.2.1.113) and 38 (EC 3.2.1.24/EC 3.2.1.114) (Henrissat and Davis, 1997). Members of family 47 are membrane-bound a1,2-mannosidases and can be found in two cellular locations: the endoplasmic reticulum (ER) of yeast and mammalian cells (Herscovics, 1999a), and the Golgi apparatus (Herscovics, 1999b; Tremblay and Herscovics, 2000), where they perform substrate-specific N-glycan processing functions. These a1,2-mannosidases are inhibited preferentially by 1deoxymannojirimycin (DMJ) and can be activated by calcium ions. Family 38 includes Golgi a-mannosidases II, IIx and III as well as lysosomal, vacuolar acidic and cytosolic/ER neutral a1,2-, a1,3and a1,6-mannosidases, which are preferentially inhibited by swainsonine (SWN) (Daniel et al., 1994; Herscovics, 1999a; Jordan * Corresponding author. Fax: +52 473 73 20006x8153 E-mail address:
[email protected] (E. López-Romero). 0014-4894/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.exppara.2009.12.014
et al., 2001; Bourne and Henrissat, 2001). Cytosolic a-mannosidases have been studied extensively in mammals (Tulsiani and Touster, 1987; Haeuw et al., 1991; De Gasperi et al., 1992), while their presence in lower eukaryotes is poorly documented. Recently, a putative Entamoeba histolytica a-mannosidase gene (Gene ID: 3408991) showing homology to members of glycosylhydrolase family 92, which mostly consists of bacterial enzymes, has been reported (Loftus et al., 2005; Magnelli et al., 2008). Pathogenesis of E. histolytica has been related to the presence of a number of glycoproteins that participate in adherence and the ensuing colonization of the host tissue by the parasite (Meza et al., 1990; McCoy et al., 1993; Stanley et al., 1995). Despite their unquestionable importance, the biosynthetic events leading to the assembly and secretion of glycoproteins in this protozoan are poorly understood. Previous studies from this laboratory allowed the identification and characterization of essential glycosyl transferases such as dolichol phosphate mannose synthase (DPMS) (Villagómez-Castro et al., 1998, 2000), the enzymes that catalyze the first two reactions of N-linked glycosylation (Vargas-Rodríguez et al., 1998) and soluble and MMF-bound a-glucosidases (Zamarripa-Morales et al., 1999; López-Romero et al., 2000; Bravo-Torres et al., 2003, 2004) thereby providing strong evidence of the presence of the machinery required for protein N-linked glycosylation in E. histolytica. Further support came later when results of subcellular fractionation indicated that some of the enzymes involved in the dolichol pathway and the ER molecular markers EhSec61 asubunit, EhDPMS and EhPDI (protein disulfide isomerase) are all contained in membrane compartments with similar density
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(Salgado et al., 2005). No evidence of genes or enzyme activities supporting further assembly steps was obtained at that time. However, work by other authors has documented the presence in the amoeba of other components of the N-glycosylation pathway such as oligosaccharyltransferase (OST) and UDP-Glc:glycoprotein glucosyltransferase (Samuelson et al., 2005; Kelleher et al., 2007). It has been described that E. histolytica contains a swainsonineinhibitable a-mannosidase with a proposed role in N-glycan trimming (Magnelli et al., 2008) but little is known about its cellular location and properties. Studies of amoeba a-mannosidases by our group have been seriously limited because of the low enzyme activity detected in cell extracts, particularly in internal membranes, and the difficulty to obtain large amounts of trophozoites. The present work was undertaken to gain an insight into the subcellular distribution, properties and possible type of amoeba a-mannosidases, by using a very sensitive fluorometric method. 2. Materials and methods 2.1. Organism, culture conditions and preparation of enzyme fractions Trophozoites of E. histolytica, strain HM1:IMSS, were maintained under axenic conditions in the TYI-S-33 medium formulated by Diamond et al. (1978). For propagation, 1-L Roux bottles, usually 16, containing 120 ml of medium were inoculated with trophozoites at a final density of 4–6 103 cells/ml an incubated at 37 °C. After 72 h, cells were collected by low speed centrifugation and washed twice with PBS buffer (15 mM potassium phosphate and 175 mM sodium chloride, pH 7.2). The PBS-washed cell pellet (3.5–4.5 108 cells) was mixed with a sufficient amount of E-64 [trans-epoxy-succinyl-L-leucylamido-(guanidine)butane] to give a final concentration of 10 lM and trophozoites were broken with glass beads (0.45–0.50 mm in diameter) by alternate periods of vortexing (20 s) and ice-cooling until 2 min of breakage were completed. The homogenate was aspirated with a Pasteur pipette, taken to 8–10 ml (depending on the amount of cells) with 5 mM citrate–phosphate buffer, pH 4.5, supplemented with 2 lM E-64 and centrifuged at 110,000g for 60 min. The resulting high-speed supernatant (soluble fraction, SF) was saved and the pellet, consisting of a mixed membrane fraction (MMF), was resuspended in a volume of 5 mM citrate–phosphate buffer, pH 4.5, containing 2 lM E-64, equal to that of the supernatant. Both the SF and MMF were used freshly to measure a-mannosidase activity as described below. 2.2. Fractionation of amoeba internal membranes in a sucrose density gradient Plasma membrane-free homogenates were obtained by the method of Aley et al. (1980) essentially as described previously (Salgado et al., 2005) except that 3.5–4.5 106 trophozoites were used. The top portion of the Aley gradient containing the cytosol and the internal membranes (10–30 mg protein) was collected and layered on top of a continuous 10–65% (p/v) sucrose density gradient prepared in 5 mM Tris/HCl buffer, pH 7.5, that was centrifuged at 167,000g for 4 h. Twenty-nine fractions (1.3 ml) were collected from the top to the bottom of the gradient and used to determine protein content and refractive index. Aliquots (1.2 ml) of fractions 9–29 were mixed with an equal volume of water and centrifuged at 333,000g for 25 min to eliminate sucrose. The membrane pellet was homogenized in 0.5 ml of 100 mM citrate–phosphate buffer, pH 4.5, containing 2 lM E-64, and used freshly to measure enzyme activities as described below. Top fractions 1–8 were diluted with 100 mM citrate–phosphate buffer, pH 4.5, and used directly to measure a-mannosidase activity.
2.3. Enzyme assays
a-Mannosidase activity was measured with 4-methylumbelliferyl-a-D-mannopyranoside (4-MU-Man) as substrate essentially as previously described (Zamarripa-Morales et al., 1999). Briefly and unless otherwise stated, reaction mixtures containing 0.1 mM 4MU-Man, the enzyme fraction, 100 mM citrate–phosphate buffer, pH 4.5, containing 2 lM E-64 in a total volume of 200 ll were incubated at 45 °C. After 1 h, the reaction was stopped by adding 2.3 ml of 0.5 M Na2CO3 in 0.1 N NaOH, pH 10.4, and the fluorescence of released 4-methylumbelliferone (4-MU) was monitored in a PerkinElmer luminescence spectrometer with excitation and emission set at 350 and 440 nm, respectively. Activity was expressed as pmoles of 4-MU released in one min. Specific activity was referred to one mg of protein. In some experiments, activity was determined with p-nitrophenyl a-D-mannopyranoside (p-NP-Man) as substrate. In this case, mixtures containing 2 mM p-NP-Man, the enzyme fraction, and 100 mM citrate–phosphate buffer, pH 4.5, containing 2 lM E-64 in a final volume of 200 ll were incubated at 45 °C. After 1 h, the reaction was stopped as described above and the amount of released p-NP was determined in a Beckman DU-650 spectrophotometer at 405 nm. Before reading, reaction mixtures prepared with the membrane fractions were centrifuged at 3000g for 10 min to remove turbidity. Activity was expressed as pmoles of p-nitrophenolate (p-NP) released in 1 min. Specific activity was referred to 1 mg of protein. With both methods, assays were carried out in duplicate and, in some cases, in triplicate. Values plotted in figures shown below are the average of at least three independent experiments. DPMS and acid phosphatase (AP) activities were measured in internal membrane fractions as described elsewhere (Salgado et al., 2005). 2.4. Protein assay Protein in enzyme preparations was quantified by the modified Lowry’s method (1951) using the Bio-Rad DC protein assay kit with bovine serum albumin as standard. 2.5. Chemicals 4-MU-Man, p-NP-Man, 1-deoxymannojirimycin (DMJ; 1,5-dideoxy-1,5-imino-D-mannitol), swainsonine (SWN; 8a-indolizidine1,2,8-triol) and E-64 were purchased from Sigma Chemical Co. (St. Louis, MO). Ingredients of the Diamond’s medium such as peptone biotryptase, dextrose and yeast extract were from Becton Dickinson de Mexico (Mexico City, Mexico). Adult bovine serum and vitamins were obtained from Laboratorios Microlab, S.A. de C.V. (Mexico City, Mexico). All other chemicals were of the highest purity commercially available. 3. Results 3.1. Subcellular distribution and some general properties of amannosidase activity When measured under optimal conditions using 4-MU-Man, a very sensitive fluorogenic substrate, results from several experiments revealed that 37% and 63% of total a-mannosidase activity was distributed in the MMF and SF, respectively (not shown). Both enzyme fractions linearly hydrolyzed 4-MU-Man up to 2 h of incubation at 45 °C and as a function of the amount of protein up to 536 and 752 lg per assay (200 ll), respectively (not shown). Hydrolysis of the fluorogen by the two enzyme preparations occurred maximally at pH 4.5 with 100 mM citrate–phosphate buffer although
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the SF a-mannosidase was slightly more active with sodium acetate buffer (not shown). The effect of cations is a useful parameter to distinguish among a-mannosidases. For instance, members of family 47 can be stimulated by Ca2+ ions whereas activity of class II and III a-mannosidases of family 38 is increased by Zn2+ and Co2+ or fully depend on Co2+, respectively (Herscovics, 1999a; Nakajima et al., 2003). It was therefore important to examine the effect of some common divalent cations on hydrolysis of 4-MU-Man by the amoeba a-mannosidases. As observed in Fig. 1 and 5 mM Co2+ or Mn2+ inhibited over 80% of the MMF-associated enzyme whereas at the same concentration they reduced activity of the SF by 67– 73%. Concentrations above 5 mM resulted in higher enzyme inhibition. On the other hand, Ca2+ and Mg2+ did not significantly affect activity. The effect of metal ions on hydrolysis of p-NP-Man (see below) by the MMF-associated enzyme was also measured. Up to 10 mM, Ca2+, Mg2+ and Mn2+ had no effect on activity whereas it was fully inhibited by 2.5 mM Co2+ (not shown). 3.2. Kinetics of hydrolysis of 4-MU-Man and p-NP-Man by amoeba amannosidases MMF and SF hydrolyzed 4-MU-Man in a manner proportional to substrate concentration up to 0.2 mM but the SF exhibited about a 3-fold higher affinity than the MMF as judged from the Km values of 0.24 and 0.72 mM, respectively. Corresponding Vmax values were 225 and 300 pmol of 4-MU/min/mg (Fig. 2A). It has been observed that some a-mannosidases of family 38 but not of family 47 can cleave other synthetic substrates such as p-NP-Man (Moremen et al., 1994; Daniel et al., 1994). Therefore, it was also important to determine whether amoeba a-mannosidases acted upon this substrate. Results revealed that both enzyme fractions cleaved pNP-Man but activity of the membrane-associated fraction was substantially higher than that of the soluble enzyme (Fig. 2B, inset).
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Hydrolysis of this substrate followed a typical Michaelis–Menten kinetics, yielding Km and Vmax values of 0.52 mM p-NP-Man and 0.48 pmol of p-NP/min/mg, respectively (Fig. 2B). These constants were not calculated for the SF a-mannosidase. 3.3. Effect of inhibitors of a-mannosidase on enzyme activity To have a further clue into the type of a-mannosidases present in the amoeba trophozoites, DMJ and SWN, two preferential inhibitors of family 47 and 38 a-mannosidases, respectively, were tested for their ability to inhibit the hydrolysis of 4-MU-Man and p-NP-Man. The effect of inhibitors was measured at two concentrations of substrate to determine the type of inhibition. While hydrolysis of the fluorogenic substrate by both enzyme fractions was not affected by DMJ (Fig. 3, left panels), this was strongly and comparably inhibited by SWN yielding IC50 values of 0.15–0.17 and 0.14 lM for the MMF and SF enzyme preparations, respectively (Fig. 3, right panels). On the other hand, both inhibitors failed to affect the hydrolysis of p-NP-Man by the membrane-associated enzyme (not shown). 3.4. Distribution of a-mannosidase activity in internal membranes Detection of up to 37% of total a-mannosidase activity in the amoeba MMF prompted us to separate the plasma membrane-free internal membranes by isopycnic centrifugation in a sucrose density gradient in an attempt to identify the amoeba compartments enriched with the enzyme. As depicted in Fig. 4, apart from the expected peak at the top of the gradient corresponding to the SF, amannosidase activity was distributed into at least three other peaks exhibiting sucrose densities of 1.121 (A), 1.135 (B) and 1.214 (C) g/ml. Minor peak B (observed here as a shoulder of peak A) and major peak C were enriched with ER- and lysosome-resident activities, respectively, as previously described (Salgado et al., 2005). No markers were used to identify the slower sedimenting peak A; however, according to its density (1.121 g/ml) it may very well correspond to Golgi structures whose presence in the amoeba has been suggested and/or demonstrated by us (Salgado et al., 2005) and others (Mazzuco et al., 1997; Ghosh et al., 1999; Bredeston et al., 2005; Teixeira and Huston, 2008). All three enzyme peaks were associated to membrane fractions containing barely detectable levels of protein. Membrane peaks located between the ER and lysosome markers (fractions 16–23) were not identified but they probably correspond to transit vesicles or lysosome precursors. 4. Discussion
Fig. 1. Effect of metal ions on a-mannosidase activity. Enzyme was assayed as a function of increasing concentrations of the indicated divalent cations. All other components of the reaction mixture remained unchanged. MMF, MMF-associated enzyme; SF, soluble fraction. Each point represents the mean of three independent determinations ±SD.
While a-mannosidases have been extensively studied in fungi and mammalian cells, their knowledge in E. histolytica and other protozoans is rather scarce. Accordingly, it has been described that E. histolytica contains an a-mannosidase that trims N-glycans to Man3GlcNAc2 and Man4GlcNAc2 oligosaccharides in reactions that are inhibited by SWN (Magnelli et al., 2008) but not much is known on its subcellular distribution and properties. Experimental conditions used here demonstrated that most of the enzyme is present in the soluble fraction, with a substantial amount (37%) remaining associated to the amoeba membranes. Soluble and particulate amannosidases have been described in other organisms such as Trypanosoma cruzi where a neutral a1,2-mannosidase converts Man9GlcNAc2 into Man7GlcNAc2 (Xavier et al., 1994; Bonay and Fresno, 1999), Saccharomyces cerevisiae (Jelinek-Kelly and Herscovics, 1988) and Candida albicans. In the latter, we demonstrated an ER membrane-located and two soluble a-mannosidases named as E-I and E-II (Vázquez-Reyna et al., 1993, 2000; Mora-Montes
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Fig. 2. Kinetics of hydrolysis of 4-MU-Man and p-NP-Man by the MMF-associated (closed circles) and soluble (open circles) enzyme fractions. a-Mannosidase activity was measured as a function of increasing concentrations of 4-MU-Man (A, inset) or p-NP-Man (B, inset). All other components of the reaction mixture remained unchanged. Data were plotted according to Lineweaver and Burk to calculate the Km and Vmax constants. Each point represents the mean of three independent determinations ±SD.
et al., 2006). Biochemical characterization in terms of maximum pH of activity, sensitivity to metal ions and kinetics of hydrolysis of 4-MU-Man revealed comparable properties between the two enzyme preparations except for a higher affinity of the soluble enzyme for 4-MU-Man and the ability of the MMF-associated but not of the soluble enzyme to readily hydrolyze p-NP-Man. Although we cannot fully explain the different 4-MU-Man affinities observed for the SF and MMF activities nor the ability of MMF but not of SF to efficiently hydrolyze p-NP-Man, it is conceivable that some catalytic properties of the enzyme are affected by its physical state. Moreover, the presence in the membranes of other enzymes active on p-NP-Man cannot be formally ruled out. Accordingly, pPNP-Man could be epimerized to p-PNP-Glc which could then be cleaved by an a-glucosidase. In fact, we have demonstrated the presence of a-glucosidase activity in the amoeba membranes (Bravo-Torres et al., 2004). However, if one considers epimerization, two questions arise: (a) would sugar epimerases act on PNP-bound mannose? and (b) a search in the literature reveals that E. histolytica possesses and aldolase 1-epimerase (mutarotase) (Villalobo et al., 2005) but not an aldolase 2-epimerase. Clearly, more studies will be required to discriminate among these possibilities and also to explain the lack of effect of SWN on hydrolysis of p-PNP-Man. In silico analysis of the E. histolytica genome predicts the presence of a single a-mannosidase gene (Gene ID: 3408991) which
encodes for a protein of 737 amino acids, with a calculated molecular weight of 85126 and a pI of 5.78. This gene exhibits a high sequence homology with a putative a-mannosidase belonging to glycosyl hydrolase family 92 which is composed mostly of bacterial enzymes (Loftus et al., 2005; Magnelli et al., 2008). Since no biochemical data are presently available for members of this family, enzyme characteristics observed here were compared with amannosidases belonging to families 38 and 47. Properties such as the lack of effect of Ca2+, the optimum pH of activity, the enzyme ability to cleave p-NP-Man, in particular the MMF-associated form (see below), and its sensitivity to SWN but not to DMJ are all consistent with an acidic a-mannosidase of the family 38 glycosylhydrolases. However, in contrast to the behavior expected for a member of this family, no stimulation was observed by Co2+ which, like Mn2+, strongly reduced activity on 4-MU-Man. On the contrary, Mn2+ and also Ca2+ and Mg2+ failed to affect hydrolysis of p-NP-Man suggesting that the effect of some ions such as Mn2+ may depend on the substrate used to determine activity. Family 38 includes the Golgi a-mannosidases II, IIx and III and lysosomal, vacuolar acidic and cytosolic/ERneutral a1,2-, a1,3- and a1,6-mannosidases (Daniel et al., 1994; Herscovics, 1999a; Jordan et al., 2001; Bourne and Henrissat, 2001). Centrifugation in a sucrose density gradient allowed separation of a-mannosidase activity into three internal membrane fractions
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Fig. 3. Effect of DMJ and SWN on hydrolysis of 4-MU-Man by MMF-associated (closed symbols) and SF (open symbols) enzyme fractions. a-Mannosidase activity was assayed in the presence of 100 (circles) or 200 (squares) mM 4-MU-Man as a function of increasing concentrations of DMJ (left panels) or SWN (right panels). All other components of the reaction mixture remained unchanged. Each point represents the mean of three independent determinations ±SD.
Fig. 4. Distribution profile of a-mannosidase activity in an isopycnic sucrose density gradient. A sample of plasma membrane-free homogenate of E. histolytica trophozoites was layered on the top of a sucrose gradient (10–65%, w/v) that was centrifuged to equilibrium and processed as described in the text to determine protein, a-mannosidase activity and sucrose density. The position of enzyme markers DPMS (ER) and AP (lysosomes) as well as the sucrose densities are indicated. Results are representative of more than 10 experiments.
corresponding to ER, lysosome and most likely Golgi compartments. ER equilibrated as a shoulder of the peak at 1.121 g/ml and contained about half of the enzyme activity detected in the other structures. This activity most likely corresponds to an a1,2mannosidase specifically involved in N-glycan processing, a function previously demonstrated in E. histolytica (Magnelli et al., 2008) and T. cruzi (Bonay and Fresno, 1999). In mammalian cells, N-oligosaccharides are further modified to complex glycans by Golgi glycosyl hydrolases, including a-mannosidases, and transferases (Herscovics, 1999a,b). Thus, it seems reasonable to expect a similar role for the Golgi a-mannosidases detected in this study. On the other hand, it has been described that degradation of N-oligosaccharides is carried out in the cytosol and acid compartments such as vacuoles and lysosomes (Daniel et al., 1994; Herscovics,
1999b). For instance, the acid a-mannosidase of T. cruzi is a broad-specificity hydrolase involved in the catabolism of glycoconjugates, presumably in the digestive vacuole (Vandersall-Nairn et al., 1998). This function could also be expected for the lysosomal a-mannosidase encountered here. If the prediction of a single a-mannosidase gene is confirmed, then an interesting question is why does the amoeba exhibit two enzyme locations, i.e., membrane-associated and cytosolic?. A feasible answer may be obtained from studies in C. albicans where amannosidase total activity distributes in the cytosol (80–85%) and in the cell membranes (15–20%) (Vázquez-Reyna et al., 1993, 2000). Recently, we demonstrated that in this organism, a 65 kDa a-mannosidase present in the ER and Golgi-derived vesicles is processed in the Golgi complex by Kex2 protease into a soluble,
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52 kDa a-mannosidase (E-I), which is transported into the cytosol where it converts soluble Man8GlcNAc2 into Man7GlcNAc2 (MoraMontes et al., 2008). We also demonstrated that E-I and the membrane-bound a-mannosidases can be converted into a soluble amannosidase E-II by a pepstatin A-sensitive protease (Mora-Montes et al., 2006). Similarly, a soluble ER a-mannosidase of 44.5 kDa purified from S. cerevisiae was obtained after pepstatin A-sensitive proteolysis of a 60 kDa membrane-bound precursor (Jelinek-Kelly and Herscovics, 1988). If a similar situation prevails in E. histolytica, one may expect some differences between the two a-mannosidase forms. For instance, conversion of the particulate enzyme into the soluble form may involve changes in the catalytic properties of a-mannosidase that would help to explain the much higher and lower affinities of the soluble enzyme for 4-MU-Man and p-NP-Man, respectively, and also the inability of SWN to affect hydrolysis of p-NP-Man by the membrane-associated enzyme. In support of this view, hydrolysis of p-NP-Man by E-I and E-II in C. albicans was slightly stimulated or not affected, respectively, by both DMJ and SWN (Vázquez-Reyna et al., 1999). In conclusion, results presented in this study provide useful insights into the properties and possible type of amoeba a-mannosidase activity. Although biochemical data reveal similarities with members of family 38, the amoeba putative a-mannosidase gene exhibits homology with members of glycosylhydrolase family 92 which includes prokaryotic and eukaryotic organisms whose amannosidase activity has not been yet characterized. Clearly, more biochemical and molecular studies will be required to propose a reliable classification of amoeba a-mannosidase. Acknowledgments This work was supported by Grants No. 2002-COI-39528/A-1 and 2002-COI-39529/A-1 from SEP-CONACyT, México, and by DINPO, Universidad de Guanajuato. CEST was a scholarship recipient from CONACyT, México. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.exppara.2009.12.014. References Aley, S.B., Scott, W.A., Cohn, Z.A., 1980. Plasma membrane of Entamoeba histolytica. Journal of Experimental Medicine 152, 391–404. Bonay, P., Fresno, M., 1999. Isolation and purification of a neutral a(1,2)mannosidase from Trypanosoma cruzi. Glycobiology 9, 423–433. Bourne, Y., Henrissat, B., 2001. Glycoside hydrolases and glycosyltransferases: families and functional modules. Current Opinion in Structural Biology 11, 593– 600. Bravo-Torres, J.C., Calvo-Méndez, C., Flores-Carreón, A., López-Romero, E., 2003. Purification and biochemical characterization of a soluble a-glucosidase from the parasite Entamoeba histolytica. Antonie van Leeuwenhoek 84, 169–178. Bravo-Torres, J.C., Villagómez-Castro, J.C., Calvo-Méndez, C., Flores-Carreón, A., López-Romero, E., 2004. Purification and biochemical characterization of a membrane-bound a-glucosidase from the parasite Entamoeba histolytica. International Journal for Parasitology 34, 455–462. Bredeston, L.M., Caffaro, C.E., Samuelson, J., Hirschberg, C.B., 2005. Golgi and endoplasmic reticulum functions take place in different subcellular compartments of Entamoeba histolytica. Journal of Biological Chemistry 280, 32168–32176. Daniel, P.F., Winchester, B., Warren, C.D., 1994. Mammalian a-mannosidasesmultiple forms but a common purpose? Glycobiology 4, 551–566. De Gasperi, R., Al Daher, S., Winchester, B., Warren, C.D., 1992. Substrate specificity of the bovine and feline neutral alpha-mannosidases. Biochemical Journal 286, 47–53. Diamond, L.S., Harlow, D.R., Cunnick, C.C., 1978. A new medium for the axenic cultivation of Entamoeba histolytica and other Entamoeba. Transactions of the Royal Society of Tropical Medicine and Hygiene 72, 431–432. Ghosh, S.K., Field, J., Frisardi, M., Rosenthal, B., Mai, Z., Rogers, R., Samuelson, J., 1999. Chitinase secretion by encysting Entamoeba invadens and transfected Entamoeba histolytica trophozoites: localization of secretory vesicles,
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