Purification and characterization of a specific late-larval esterase from two species of the Drosophila repleta group: contributions to understand its evolution
© Lopes et al.; licensee Springer. 2014
Received: 28 September 2013
Accepted: 9 January 2014
Published: 24 January 2014
After duplication, one copy of an original gene can become redundant and decay toward a pseudogene status or functionally diverge. Here, we performed the purification and biochemical characterization of EST-4 (a late larval β-esterase) from two Drosophila repleta group species, Drosophila mulleri and Drosophila arizonae, in order to establish comparative parameters between these enzymes in these species and to contribute to better understand their evolution.
In D. mulleri, EST-4 had an optimal activity in temperatures ranging from 40° to 45°C and at pH 7.5, maintaining stability in alkaline pH (8.0 to 10.0). It was classified as serine esterase as its activity was inhibited by PMSF. No ion negatively modulated EST-4 activity, and iron had the most positive modulating effect. In D. arizonae, it showed similar optimum temperature (40°C), pH (8.0), and was also classified as a serine esterase, but the enzymatic stability was maintained in an acidic pH (5.5 to 6.5). Fe+2 had the opposite effect found in D. mulleri, that is, negative modulation. Al+3 almost totally inhibited the EST-4 activity, and Na+ and Cu+2 had a positive modulation effect. Kinetic studies, using ρ-nitrophenyl acetate as substrate, showed that EST-4 from D. mulleri had higher affinity, while in D. arizonae, it showed higher V max and catalytic efficiency in optimal reaction conditions.
EST-4 from D. mulleri and D. arizonae are very closely related and still maintain several similar features; however, they show some degree of differentiation. Considering that EST-4 from D. mulleri has more conspicuous gel mobility difference among all EST-4 studied so far and a lower catalytic efficiency was observed here, we proposed that after duplication, this new copy of the original gene became redundant and started to decay toward a pseudogene status in this species, which probably is not occurring in D. arizonae.
Keywordsβ-esterase EST-4 D. mulleri D. arizonae Evolutionary biochemistry
Several Drosophila species hold a class of β-esterase isoenzymes that have been reported to be encoded by a cluster of genes that are products of gene duplications (Zouros et al.1982; Collet et al.1990; East et al.1990; Brady and Richmond1992; Karotam et al.1993; Oakeshott et al.1999; Balakirev et al.2005; Robin et al.2009). These enzymes have been studied regarding their biochemistry and genetic and evolutionary aspects, including some differences in enzymatic and biochemical properties (Zouros and van Delden1982; Pen et al.1984,1986a,1990; Korochkin et al.1990; Brady and Richmond1990; Richmond et al.1990; Mateus et al.2009,2011), amino acid (Pen et al.1986b, Pen et al.1990) and gene sequencing (Balakirev et al.2003; Robin et al.2009), and gene localization in the chromosome (Gomes and Hasson2003). In general, with some exceptions, two enzymes are detected in most species that hold this gene cluster, one expressed during all insect's life cycle and present mostly in the hemolymph, and other expressed in the late larvae and early pupae in the integument.
The more intensively β-esterase genes studied are Est6 and Est7 of Drosophila melanogaster subgroup (Oakeshott et al.1995,1999; Balakirev et al.2006). They are tightly linked, and Est6 has acquired a novel function in this set of species (Oakeshott et al.2000). In D. melanogaster, it is highly expressed in the sperm ejaculatory duct of the adult male instead of in the hemolymph, and the enzyme is transferred to the female during mating and modifies her subsequent egg-laying and remating behaviors (Meikle et al.1990; Richmond et al.1990; Saad et al.1994). Est7 is predominantly expressed in the integumental tissue of late larvae and early pupae (Dumancic et al.1997), and its function there still remains unknown.
In other species of the genus, the β-esterase gene cluster seems to be more complex in composition and function. In Drosophila pseudoobscura (obscura group, also D. melanogaster's subgenus Sophophora), three in tandem genes are found, and they show evidences of gene conversion or reciprocal recombination (Brady and Richmond1992; King1998). One of these three encodes the major adult hemolymph β-esterase (as D. melanogaster's Est6 above), but no function was detected for the other two genes (Brady and Richmond1990; Tamarina et al.1997). In Drosophila virilis (virilis group) and several cactophilic species (repleta group) of the subgenus Drosophila, the basic adult hemolymph and preadult integument β-esterases have been detected, and at least one and at most three β-esterases are present in the male ejaculatory bulb (Oakeshott et al.1990,1993). In Drosophila mojavensis, Robin et al. (2009) identified six paralog genes belonging to the β-esterase cluster and annotated, based on amino acid (Pen et al.1986b) and gene sequence comparisons, that the isoenzymes, named EST-4 and EST-5 (Zouros et al.1982), are probably the products of Est2c and Est2a genes, respectively. These enzymes have the same substrate preference for β-naphtyl esters (Zouros and van Delden1982; Pen et al.1984; Mateus et al.2011), but as expected by default, they are highly differentiated about their temporal and tissue expression patterns. EST-4 is found only during the late larval stage and manly in the larvae cuticle, and EST-5 is present during throughout the insect's life cycle and predominantly in the hemolymph and fat body (Zouros et al.1982; Pen et al.1984). Drosophila buzzatii, a South American cactophilic species, also has very closely linked loci in chromosome 2 encoding esterase isozymes with the typical hemolymph and late larval/early pupal cuticle expression profiles (East et al.1990; Gomes and Hasson2003).
In a more specific analysis in the repleta group, Zouros et al. (1982) detected EST-5 activity in all the 13 species they analyzed, but apparently, this was not true for EST-4. The activity of β-esterase in the larval carcass varied considerably among the species. It was abundant in most species, but it was barely detected in three (Drosophila aldrichi, D. repleta, and Drosophila peninsularis) and totally missing in other three (Drosophila tira, Drosophila hydei, and Drosophila eohydei). Mateus et al. (2011) studied six species of the same group of species and detected EST-4 and EST-5 activity in all the species: D. mojavensis cluster species (D. mojavensis, D. arizonae, and Drosophila navojoa) showed fainter bands than D. mulleri cluster species (D. mulleri, D. aldrichi, and Drosophila wheeleri).
A number of other results have shown that EST-4 and EST-5 are closely related, but with differences other than those already presented above. They have similar isoelectric points (between 6.0 and 7.0; Mateus et al.2011), exhibit 82% identity in N-terminals of amino acid sequences (Pen et al.1986b), and form an interloci heterodimer (Zouros et al.1982; Mateus et al.2011). However, they have different molar masses (Pen et al.1984,1986b; Mateus et al.2009,2011) and exhibit differential inhibition profiles, with EST-4 being inhibited by phenylmethanesulfonyl fluoride (PMSF) and not affected by malathion, and EST-5 being inhibited by malathion and not affected by PMSF (Mateus et al.2011).
After duplication, a new copy of an original gene can take two possible pathways. It can become redundant and decay toward a pseudogene status or it functionally diverges. Gene duplication followed by functional divergence has been long considered the primary mechanism of molecular evolution (Lewis1951; Ohno1970). Balakirev and Ayala (1996) have detected high frequencies of null alleles (more than 60%) for Est7 gene, which encodes the integumental tissue of late larvae and early pupae enzyme in D. melanogaster. The two explanations above have been suggested for this result. Balakirev and Ayala (2003) and Balakirev et al. (2006) proposed that the EST-7 protein has become redundant, and the gene is decaying toward a pseudogene status. Alternatively, Balakirev and Ayala (2004) suggested that the Est7 gene maintains a function that is not disabled by the stop codons or frame-shifting mutations detected.
According to Lima-Catelani et al. (2004), differences in esterase synthesis during the insect life cycle are probably due to differences in the regulatory mechanisms acting accordingly with metabolic function requirements of a variable number of processes in which esterases are involved during development. On the other hand, Robin et al. (2009) detected contrasting examples of the types and stages of loss of gene function for β-esterases as they inferred missing orthologs, pseudogenes, and null alleles, and a minimum of nine gene gain-loss events in the 12 species genome analyzed. They speculated that their results are probably related to fluctuation in the requirements for the functions of these genes over evolutionary time, possibly in response to changes in environmental niches. However, the reproductive functions of some β-esterases suggest that the copy number could have been changed by sexual competition or conflict.
Thus, this work aimed to investigate differences and compare several biochemical and enzymatic properties of EST-4 in two Drosophila species from the repleta group, D. mulleri and D. arizonae, in order to contribute to better understand the differentiation and evolution of this enzyme. Preliminary characterization of this enzyme showed, in D. mulleri, that it has the most different electrophoretic pattern from all other species analyzed so far, aside from some other distinct enzymatic features such as having the highest isoelectric point and molecular weight (Mateus et al.2011). According to Harms and Thornton (2013), an integration of evolution with biochemistry is indispensable to achieve a more complete understanding of why biological molecules have the properties that they do. Our results are in agreement to previous achievements, and the lower V max and catalytic efficiency detected for D. mulleri lead us to propose that after duplication, this copy of the original gene became redundant and started to decay toward a pseudogene status in this species, which probably is not occurring in D. arizonae.
Multifemale lineages of the two species, D. mulleri and D. arizonae, were obtained from Prof. Dr. Carlos Roberto Ceron (Department of Chemistry and Environmental Sciences, IBILCE/UNESP, São José do Rio Preto, Brazil). They were maintained as mass cultures in 250-mL culture vials with standard banana agar medium in constant temperature of 25°C ± 1°C and 12-h photoperiod.
Late-third instar larvae of both species were obtained directly from the maintenance vials, and in order to maximize sample attainment, intraspecific crosses were performed. Virgin males and females were separated, and after 7 days, ten vials containing standard culture medium were prepared. Five couples were crossed for 21 days, transferring them into new vials every 7 days. All vials were daily checked for tracking larval development, separating those larvae that were at the desired stage.
The larvae collected were immediately frozen in liquid nitrogen and stored at -80°C. The enzyme extracts were obtained by macerating 400 late-third instar larvae in 0.1 M phosphate buffer at pH 6.2 and centrifuging at 10,000 × g in 4°C for 10 min. A sample of each supernatant was electrophoresed in 10% non-denaturing polyacrylamide gel, as described by Mateus et al. (2011) and adapted in ‘Molar mass exclusion and ion exchange chromatographies’ section below, in order to detect the presence of EST-4. An adult was used as a comparative sample.
Molar mass exclusion and ion exchange chromatographies
The purification in gel filtration through molecular mass exclusion chromatography (MMEC) was performed using Sephadex G-75 resin (GE Healthcare, São Paulo, Brazil), which was packed into a 4 × 100-cm (diameter × height) column. Fractionation was performed at 4°C using 0.1 M phosphate buffer at pH 6.2 and a 0.6-mL/min flow. Fractions of 5 mL were collected and individually analyzed for larval esterase activity in polyacrylamide gel electrophoresis (PAGE), mixing 20 μL of the fraction with 5 μL sample buffer (25% Tris–HCl buffer (0.05 M) pH 6.8, 20% glycerol, and 0.02% bromophenol blue). The electrophoresis was performed in 10% PAGE as described by Mateus et al. (2011) at constant voltage of 110 V in room temperature. To test the substrate specificity, the gels were soaked with usual α- and β-naphthyl acetate solution (Mateus et al.2011), and their products were stained for 2 h using Fast Blue RR salt (Sigma-Aldrich, São Paulo, Brazil).
The fractions that showed EST-4 activity were joined and dialyzed in 20 mM N-[tris(hydroxymethyl)methyl]-3-aminopropanesulfonic acid (TAPS) buffer at pH 8.5 for 24 h at 4°C and with three exchanges. The dialyzed material was submitted to ion exchange chromatography (IEC) in a Q-Sepharose resin (GE Healthcare, São Paulo, Brazil) with the same buffer above. The sample was washed with elution buffer (20 mM TAPS buffer of pH 8.5 without NaCl) to remove unbound material. The elution of proteins was initiated with linear salt gradient ranging from 0 to 2 M NaCl in the same buffer. Fractionation was performed at 4°C with a 1.0-mL/min flow, and 5-mL fractions were collected.
After each chromatography, EST-4 purity was certified through denaturing gel electrophoresis (10% sodium dodecyl sulfate (SDS)-PAGE) according to Laemmli (1970). The samples (20 μL) were mixed with 10 μL sample buffer (25% Tris–HCl buffer (0.05 M), pH 6.8, 3.1% DTT (w/v), 0.02% Bromophenol Blue, 20% glycerol, and 4% SDS (w/v)). This mixture was boiled for 5 min at 96°C, and after electrophoresis, the gel was stained with silver nitrate (See and Jackowski1989).
EST-4 activity test
where, Abs means absorbance at 410 nm, V R is the volume of reaction which is 500 μL, e is the molar extinction coefficient of ρ-nitrophenyl which is 18.5 μmol mL-1 cm-1, t is time in minutes, and V E is the volume of enzyme which is 25 μL.
The esterase quantification was determined according to the method described by Bradford (1976), using standard curve constructed with bovine serum albumin (BSA).
Biochemical characterization of EST-4
pH effect on the EST-4 activity and stability
Optimum pH and pH stability characterization of both enzymes were carried out with pH ranging from 4.5 to 10.5, varying 0.5 U. The following buffers were used: acetate (pH 4.5 and 5.0), 2-[N-morpholino]ethanesulfonic acid (MES; pH 5.5, 6.0, and 6.5), HEPES (pH 7.0, 7.5, and 8.0); N,N-bis(2-hydroxyethyl)glycine (BICINE; pH 8.5 and 9.0), and 3-(cyclohexylamino)-1-propanesulfonic acid (CAPS; pH 9.5, 10.0, and 10.5). All the buffers were prepared with 0.05 M of concentration. The reaction mixes contained 25 μL of purified enzyme, 25 μL of buffer (ranging from pH 4.5 up to 10.5, described above), and 450 μL of ρ-nitrophenyl acetate substrate. Thus, thirteen different mixes were set up, each one with different final pH, and they were incubated at 40°C for 30 min. After this period, the esterase activity was measured accordingly to the method described above (‘EST-4 activity test’ section).
The pH stability was determined by incubating the enzymes (25 μL) for 1 h at 25°C at different pH values (using the buffers described above), subsequently adding 450 μL of ρ-nitrophenyl acetate substrate, 13 μL of optimum pH buffer (pH 6.5 to D. mulleri and pH 7.5 to D. arizonae), and determining the activity as described above at 40°C for 30 min.
Temperature effect on the EST-4 activity
The influence of temperature on the activity of EST-4 was performed in optimal pH (7.0 for D. mulleri and 7.5 for D. arizonae) and temperature of 25°C to 55°C, with variations of 5°C. The pure enzymes (25 μL) were mixed with 25 μL of optimum pH buffer and 450 μL of ρ-nitrophenyl acetate substrate, as described in the ‘EST-4 activity test’ section. The enzyme activities were evaluated proceeding incubation for 30 min in the respective temperatures.
Chemical effect on the EST-4 activity
The determination of active site constitution of both enzymes was performed according to the protocol described by Dunn (1989) with modifications. The following reagents were used in a final concentration of 5 mM: PMSF, EDTA (ethylenediamine tetraacetic acid), and pepstatin.
The pure enzymes (25 μL) were premixed with 2.5 μL of each inhibitor, incubated at 40°C for 5 min, and after that period, 22.5 μL of optimum pH buffer and 450 μL of ρ-nitrophenyl acetate substrate were added. The enzyme activities were checked by incubating them for 30 min at 40°C. The control tube was made with addition of 25 μL pure enzyme, 450 μL of ρ-nitrophenyl acetate, and 25 μL of optimum pH buffer.
The effects of metal ions on the esterase activities were investigated by adding monovalent (Li+, Na+, and K+) and divalent ions (Ba2+, Ca2+, Mg2+, Mn2+, Fe2+, Ni2+, Cu2+, and Zn2+) to a final 10 mM concentration. The pure enzymes (25 μL) were preincubated with each ion (2.5 μL) at 40°C for 5 min. Subsequently, the enzyme activities were evaluated at 40°C for 30 min and added with 450 μL of ρ-nitrophenyl acetate substrate and 22.5 μL of optimum pH buffer. The control tube was made with the addition of 25 μL pure enzyme, 450 μL of ρ-nitrophenyl acetate, and 25 μL of optimum pH buffer.
EST-4 enzymatic kinetics
The enzyme kinetics was obtained for both EST-4 by adding increasing concentrations of ρ-nitrophenyl acetate substrate, from 0.1 to 1.0 mM. The experiments were performed in the optimum pH and at 40°C, and the results were read in a spectrophotometer with 410-nm absorbance.
The K m and V max kinetic values were obtained using Michaelis-Menten equation calculated by non-linear regression of data from hydrolysis of the substrate using the software GraFit version 5.0 (Erithacus Software Ltd., Surrey, UK). K m, K cat, and K cat/K m were evaluated by determining the enzyme activities against ρ-nitrophenyl acetate substrate in ideal conditions.
Results and discussion
Purification of EST-4 from D. mulleri and D. arizonae
The molar mass of EST-4 was determined using protein molecular weight marker (low molecular weight SDS calibration kit for SDS electrophoresis, GE Healthcare) in the SDS-PAGE gel. As seen in Figure 4, the molar masses of the purified EST-4 (correspondent to the purified protein subunit (arrow)) of D. mulleri and D. arizonae are approximately 45 kDa. Pen et al. (1984) determined the molar mass of EST-4 of D. mojavensis also using denaturing gel electrophoresis (SDS-PAGE) and obtained values between 62 and 64 kDa for its subunits. Similarly, the molar mass of 64 to 66 kDa for the subunits of EST-5 of the same species was found by Pen et al. (1986a). As pointed out before, EST-4 and EST-5 are expressed by duplicated genes, Est2c and Est2a, respectively, in these species. Mateus et al. (2011) and Pen et al. (1984) have determined the molar mass of the dimeric EST-4 protein as being between 83 and 95 kDa. Therefore, our results showed a more congruent data for the molar mass of the subunit, as it appears to have half the mass of the dimeric protein, and not the anomalous behavior previously found by Pen et al. (1984,1986a).
EST-4 purification features from Drosophila mulleri and Drosophila arizonae after MMEC and IEC
Total protein (mg)
Total activity (U)
Specific activity (U/mg)
Biochemical characterization of EST-4
Effect of pH on the EST-4 activity and stability
Therefore, regarding pH, both EST-4 had similar optimum pH curve but different pH stability. Thomazine (2007) studied two enzymatic variants of EST-5, called fast (EST-5 F) and slow (EST-5S), and found that these allozymes presented pH profiles similar to that described here, i.e., optimum activity in alkaline pH and lower activity in acid pH. In a study of characterization of JHE in D. melanogaster, Campbell et al. (1992) found that below pH 6.0, considerable non-enzymic, acid hydrolysis of juvenile hormone (JH) occurred. Over the pH range from 6.0 to 8.6, the JHE activity almost doubled, increasing linearly with increasing pH. No further change in activity was observed at pH 9.0. Therefore, they showed that JHE from D. melanogaster also had a tendency of better activity on alkaline environment.
Effect of temperature on the EST-4 activity
These results provide evidence that both enzymes had similar optimum temperatures, with minor differences as EST-4 from D. mulleri operated better at higher temperatures (45°C) when compared to D. arizonae. Thomazine (2007) found even higher optimum temperature in experiments with variants of EST-5 (slow and fast) of D. mulleri, which showed higher activity at 50°C. However, when these variants were incubated for 10 min at 55°C, no activity was detected for EST-5 F.
Effect of chemicals on the EST-4 activity
These inhibition results corroborate the previously data of Mateus et al. (2011) who detected that the EST-4 of six species of the D. repleta group, including the two studied here, were all inhibited by PMSF. According to Dunn (1989), this type of inhibition probably occurs because this compound irreversibly binds to the hydroxyl side chain of the serine residue, impeding the enzymatic catalysis. This class of esterase is commonly found in insects (see Krejci et al.1991, Anthony et al.1995, Hinton and Hammock2003, Coutinho-Abreu et al.2007, Yu et al.2009, and Li et al.2010 as examples). However, in some insects, they were not detected. For example, in a study of biochemical identification and characterization of esterases in Tribolium castaneum, Gigliolli et al. (2011) observed that all enzymes were inhibited by eserine sulfate and/or malathion, and none by PMSF.
Other two compounds also affected the EST-4 activity in our experiments. EDTA was responsible for more than 70% decrease in EST-4 activity in D. arizonae and about 35% decrease in the D. mulleri, suggesting that these esterases probably have their activity modulated positively in the presence of metal ions. In the presence of this compound, the ions were possibly chelated, resulting in the observed inhibition. Pepstatin was responsible for reducing approximately 20% of the enzymatic activity of EST-4 in both species. Serine enzymes commonly present a catalytic triad composed of serine, histidine, and aspartate residues, which indirectly interact, enhancing enzyme activity (Zhou et al.1994). This inhibitor probably is connected to the aspartate residue causing the observed reduction in the EST-4 activity.
In the case of D. arizonae, Na+ and Ba+2, together with Cu+2 and Co+2, also activated more than 50% (around 60%) of the EST-4 activity (Figure 9B). However, different from D. mulleri, several ions decreased the esterase activity, such as Fe+2, Li+, Mg+2, Ca+2, K+, Mn+2, Zn+2, and Al+3. This last one inhibited nearly 100% of the enzyme activity, and in this case, it is possible that it bound to the enzyme, promoting negative modulatory effect and preventing it to perform its catalytic function. Negative modulation of serine protease activity in the presence of Al+3 was also observed by Silva (2011) in the fungus Aspergillus fumigatus Fresenius. They detected, among all ions tested, that the enzyme had its activity reduced about 80% only in the presence of this ion.
Therefore, the differences in the modulator effect of Fe+2 in both species are noteworthy. It was responsible for the activation of EST-4 in D. mulleri, and it had a negative modulatory effect in D. arizonae, decreasing approximately 40% of the esterase activity. Moreover, Al+3 showed an effect (positive or negative) over EST-4 of D. mulleri and was responsible for almost the complete inhibition of EST-4 in D arizonae. Thus, it can be suggested that although paralog genes with similar temporal and tissue expressions encode these enzymes, they are biochemically distinct in their catalysis regarding the presence of ions.
Kinetic parameters of EST-4
EST-4 kinetic parameters using ρ-nitrophenyl acetate as substrate
V max(mM min-1)
k cat/K m(mM-1 min-1)
Regarding the number of reaction cycles performed per unit of time, k cat, it was possible to infer that the EST-4 of D. arizonae can convert the substrate into a product with higher efficiency (1,933 min-1) as compared to the EST-4 of D. mulleri (94.54 min-1). The catalytic efficiency (k cat/K m) corroborated these results as the EST-4 of D. arizonae showed higher catalytic efficiency (2,583.8 mM-1 min-1) when compared to D. mulleri (532 mM-1 min-1).
Our results clearly showed that these enzymes are very closely related and still maintain some similar features, such as optimal temperature and pH. However, they already depict many other characteristics that show they have differentiated in the evolutionary time (effect of chemicals, pH stability, enzymatic affinity, V max, and catalytic efficiency). It seems that the EST-4 of D. arizonae is much better adjusted as an esterase enzyme than the EST-4 of D. mulleri because of its superior K cat and K cat/K m. Considering that this enzyme of D. mulleri has more conspicuous difference in gel mobility among all EST-4 studied so far (Mateus et al.2011) and its kinetic features observed here, it can be can propose that after duplication, one new copy of the original gene (in our case, the Est2c gene of EST-4) became redundant and started to decay toward a pseudogene status in this species, which probably is not occurring in D. arizonae. Balakirev and Ayala (1996) detected high frequencies of null alleles for the Est7 gene, which encodes the enzyme found in the integumental tissue of late larvae and early pupae in D. melanogaster. This seems like to be the possible explanation for the observations detected for the EST-4 of D. mulleri.
bovine serum albumin
molar extinction coefficient of ρ-nitrophenyl
ethylenediamine tetraacetic acid
ion exchange chromatography
juvenile hormone esterase
molecular mass exclusion chromatography
polyacrylamide gel electrophoresis
Sodium dodecyl sulfate
time in minutes
- VE :
volume of enzyme
- VR :
volume of reaction.
We are thankful to Prof. Dr. Carlos R. Ceron for providing the Drosophila strains, to Prof. Dr. Maura H. Manfrin for opening her laboratory to us in order to maintain and to perform the experiments with these strains to obtain all the samples for purification, and to Ronivaldo Rodrigues da Silva and Nathalia G. Rosa for technical assistance in the laboratory experiments. Funds were provided by the following: SETI/Fundação Araucária (V. F. Lopes Master's Fellowship), Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) (H. Cabral, grant number 2011/06986-0), Financiadora de Estudos e Projetos (FINEP), Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES), Faculdade de Ciências Farmacêuticas de Ribeirão Preto da Universidade de São Paulo (FCFRP/USP), and Universidade Estadual do Centro-Oeste (UNICENTRO).
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