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Article

Biochemical Properties of the Acid Ectophosphatase Activity of Phytomonas serpens Involved in Cell Proliferation

by
Luiz Fernando Carvalho-Kelly
1,2,
Anita Leocadio Freitas-Mesquita
1,2,
Thaís Souza Silveira Majerowicz
3 and
José Roberto Meyer-Fernandes
1,2,*
1
Instituto de Bioquímica Médica Leopoldo de Meis, Centro de Ciências da Saúde, Universidade Federal do Rio de Janeiro, Rio de Janeiro 21941-590, RJ, Brazil
2
Instituto Nacional de Ciência e Tecnologia em Biologia Estrutural e Bioimagem, Rio de Janeiro 21941-590, RJ, Brazil
3
Instituto Federal de Educação, Ciência e Tecnologia do Rio de Janeiro, Rua Senador Furtado, 121. Maracanã, Rio de Janeiro 20270-021, RJ, Brazil
*
Author to whom correspondence should be addressed.
Kinases Phosphatases 2024, 2(4), 379-390; https://doi.org/10.3390/kinasesphosphatases2040024
Submission received: 25 November 2024 / Revised: 11 December 2024 / Accepted: 12 December 2024 / Published: 15 December 2024
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Abstract

Phytomonas is the only kinetoplastid that can parasitize plants, causing economically relevant issues. Phytomonas serpens share similarities with pathogenic trypanosomatids, including surface enzymes that are involved in adhesion to the salivary gland of their experimental host, the insect Oncopeltus fasciatus. Ectophosphatases are cell surface enzymes involved in host–parasite interactions that are widely distributed among microorganisms. This work aimed to perform the biochemical characterization of P. serpens ectophosphatase activity, investigating and discussing its possible physiological role. This activity presented an acidic profile, and its kinetic parameters Km and Vmax were calculated as 1.57 ± 0.08 mM p-NPP and 10.11 ± 0.14 nmol p-NP/(h × 108 flagellates), respectively. It was stimulated by cobalt, inhibited by zinc, and insensitive to EDTA, a divalent metal chelator. The inhibitor sodium orthovanadate was able to decrease P. serpens ectophosphatase activity and growth, suggesting its involvement in cell proliferation. Given that P. serpens can uptake inorganic phosphate (Pi) from the extracellular medium, it is likely that its ectophosphatase activity acts together with the transport systems in the Pi acquisition process. The elucidation of the molecular mechanisms involved in this process emerges as a relevant perspective, providing new strategies for controlling Phytomonas infection.

1. Introduction

Phytomonas is a monophyletic genus with a global distribution composed of trypanosomatid flagellates with the unique ability to infect plants [1]. Plant diseases, such as phloem necrosis in coffee and Phytomonas wilt in coconut trees, represent a significant problem to agriculture, leading to high economic loss [2]. In the early 2000s, 43% of coconut plantations in the state of Amazonas in Brazil, were affected by Phytomonas wilt, leading to the destruction of several plantations [2,3].
Even infections considered relatively harmless to plants can be economically relevant since they can reduce the market value of the vegetable product [4]. Despite not causing severe systemic disease, Phytomonas serpens can be found in tomato sap, causing the appearance of yellow spots on the parasitized fruits [4,5]. This species is one of the most studied representatives of the genus because, unlike other pathogenic species, P. serpens is readily isolated and cultivated in the laboratory, enabling in vitro studies in a wide range of areas [6]. In addition, it presents natural characteristics similar to other pathogenic members belonging to the Trypanosomatidae family, such as Trypanosoma cruzi, the etiological agent of Chagas disease. Chagasic patients have been shown to display antibodies that can recognize P. serpens antigens and the inoculation of this phytopathogen in mice led to protection against T. cruzi infection [7,8,9]. From these studies, P. serpens has also gained importance in medical research [10].
Although little is known about P. serpens in terms of how the species has adapted to life inside plants, parasites may be spread between plant hosts by a broad range of different insects [1,10]. When a phytophagous insect feeds on an infected plant, the flagellates are drawn into the digestive system, migrate to the hemolymph through the intestinal barrier and invade the salivary gland. During the subsequent feed, the insect injects saliva carrying flagellates into the plant. The P. serpens biological cycle can be reproduced in the laboratory using the insect Oncopeltus fasciatus [11]. Some surface enzymes similar to those from Trypanosoma and Leishmania have already been studied in P. serpens. A cysteine protease similar to T. cruzi cruzipain and a metalloprotease similar to gp-63 from Leishmania spp. are both involved in adhesion to the salivary gland of the experimental host O. fasciatus [12,13]. Ectophosphatases are cell surface enzymes reported to be involved in the interaction between trypanosomatid parasite and the salivary gland of its host in at least two other systems: Trypanosoma rangeli–Rhodnius prolixus [14] and Strigomonas culicis–Aedes aegypti [15].
Ectophosphatases are widespread among different microorganisms including fungi [16,17,18,19,20,21,22], trypanosomatids [15,23,24,25,26,27,28,29,30], and other protists [31,32,33,34,35]. Several physiological roles have been attributed to ectophosphatases beyond parasite–host interaction, like their involvement in cell differentiation, proliferation, and nutrition [36]. The presence of ectophosphatase activities has already been identified in two different species of Phytomonas, namely, Phytomonas françai and Phytomonas mcgheei [24]. In this context, we hypothesize that P. serpens could also present an ectophosphatase activity. This work aimed to perform the biochemical characterization of this ectophosphatase activity, identifying a possible physiological role in cell proliferation, thus helping provide new strategies for controlling Phytomonas infection. Moreover, we analyzed in detail the main biochemical properties of the ectophosphatases of different microorganisms, comparing them with the data obtained in the present study.

2. Results

Phytomonas serpens ectophosphatase activity assays were carried out using intact cells obtained from a three-day-old culture when the protists reached the early exponential phase. The ectophosphatase activity measured in the presence of 1 × 108 flagellates/mL was linear up to 1 h (Figure 1A). Cell densities from 1 to 10 × 108 flagellates/mL also showed a linear increase in ectophosphatase activity (Figure 1B).
To confirm the ecto-localization of the phosphatase activity, we ruled out the possibility that the observed p-NPP substrate hydrolysis was due to secreted soluble enzymes. P. serpens cells were incubated for 60 min in the reaction medium in the absence of the substrate. Then, the suspension was centrifuged to remove the cells, and the supernatant was assayed for phosphatase activity. The rate of p-NP released from the supernatant was approximately 2% of the total activity observed in intact cells (Figure 2).
Ectophosphatase assays performed in reaction media buffered with different pH values revealed acidic activity, which was almost three times greater at pH 5.0 than at pH 7.0 (Figure 3). The ectophosphatase activity follows Michaelis–Menten kinetics, with apparent parameters Km and Vmax calculated as 1.575 ± 0.077 mM p-NPP and 10.11 ± 0.137 nmol p-NP/(h × 108 flagellates), respectively (Figure 4).
Several metals were tested for effect on P. serpens ectophosphatase activity, among which cobalt stimulated this activity (Figure 5). The alkaline phosphatase inhibitor Ethylenediaminetetraacetic acid (EDTA) did not exert any modulation, following the acidic character of the ectophosphatase activity (Figure 3). Zinc was the only metal showing an inhibitory effect on this ectophosphatase activity (Figure 5).
The addition of sodium orthovanadate (1 mM) led to a decrease of approximately 50% of the ectophosphatase activity of P. serpens (Figure 6A). Sodium orthovanadate also inhibited the protist growth. The presence of 1 mM of this inhibitor in the culture medium significantly decreased the number of parasites from the second up to the fifth day of growth (Figure 6B).

3. Discussion

Phytomonas is the only genus in the Kinetoplastid family that can parasitize plants, causing economically relevant issues by reducing the market value of the vegetable product. Pharmacological studies for treatment are scarce, especially due to the limitation of in vitro multiplication of the species that cause the most serious diseases; however, P. serpens can be easily cultivated in laboratory conditions, making it a promising model [1,6]. This phytopathogen shares some surface enzymes with other protists, including the ectophosphatase activity that was identified and characterized in the present work. An effective technique for identifying ectophosphatases is the phosphatase activity assay using intact and living cells. The cell viability during our experiments was confirmed by a Trypan blue exclusion assay. Moreover, p-NPP hydrolysis was linear with time (Figure 1A), which suggests that possible cell breakage during incubation did not contribute to the phosphatase activity measured in our assays [37]. The ectophosphatase activity also has a linear relationship with the cell density (Figure 1B), and no significant secreted phosphatase activity was detected (Figure 2), which suggests a negligible contribution of internal enzymes during the reaction [37].
The biochemical characterization of the ectophosphatase activity on P. serpens allow us to discuss its similarities and differences from those previously described in other microorganisms. The comparison between different studies of ectophosphatase activities is hampered by variations in available protocols. Experimental conditions such as the type of substrate and its concentration, pH, buffer, and temperature affect the measurement of enzymatic activity [38]. Among these parameters, using p-NPP as the substrate is the most constant and has prevailed since the end of the 1960s. Due to its ease and convenience, p-NPP is routinely used in quantitative measurements of phosphatase activity in soil, plants, animals, and microorganisms [39,40]. To analyze and compare the biochemical properties of ectophosphatase activities of P. serpens and other microorganisms, we selected studies in which phosphatase activity was performed using the same approach; therefore, no significant variations were expected. Table 1 reports the optimal pH range, Km and Vmax values, and inhibitory and stimulatory metals of ectophosphatase activities of twenty-one species of microorganisms, including seven fungi [16,17,18,19,20,21,22], nine trypanosomatids [15,23,24,25,26,27,28,29,30], and five other protists [31,32,33,34,35].
During their lifecycle, P. serpens lives in at least one acidic environment, the tomato fruit [4]. As ectophosphatase activity peaks in the acid pH range (Figure 3), it is possible that this activity contributes to the success of the protist’s infection in tomato fruit. As observed in Table 1, most microorganisms studied present greater ectophosphatase activity at acidic pHs, including the Phytomonas species P. françai and P mcgheei [24]. With respect to the kinetic parameters of P. serpens ectophosphatase activity (Figure 4), its substrate affinity lies within the range found for other trypanosomatids, such as Herpetomonas muscarum [23], Leishmania amazonensis [27], Trypanosoma cruzi (Colombiana strain) [28] and Trypanosoma rangeli (macias strain) [29]. Compared with that of other Phytomonas species, the Km of P. serpens ectophosphatase activity is closer to that of P. mcgheei and significantly lower than that of P. françai [24] (Table 1).
The observed stimulation of P. serpens ectophosphatase activity by cobalt (Figure 5) has been described for the fungal Rhinocladiella aquaspersa [20], and the protists Trypanosoma brucei [26], and Acanthamoeba castellanii [34]. The diversified use of cobalt in different branches of industry is leading to an excessive accumulation of this metal in the environment. Although being toxic when exceeding the threshold levels, it is known that maintaining its optimal content in microorganism cells can assist growth processes [41]. The inhibition of the ectophosphatase activity by zinc is quite common, being observed in practically all microorganisms tested (Table 1). As expected, zinc also inhibited the ectophosphatase activity of P. serpens, being the only one among the metals tested that showed an inhibitory effect (Figure 5). EDTA’s inability to modulate the ectophosphatase activity (Figure 5) suggests that contaminant metals in the reaction media had no effect on catalysis and that metalloenzymes are not a significant component of the ectophosphatase pool.
Classical phosphatase inhibitors are commonly used to investigate the biological role of ectophosphatase activities [14,42,43]. Sodium orthovanadate, a phosphotyrosine phosphatase inhibitor, can affect different processes; however, its main biological action is on the surface of living cells, as it is an anion and is not able to cross the plasma membrane [34,44]. Several reports associate the ectophosphatase activity with the ability of parasites to interact with their host cells due to the inhibitory effect promoted by sodium orthovanadate [15,16,19,21,34]. Some possible scenarios have been described to explain the role played by ectophosphatases in the host–parasite interaction, taking into account their peculiar location in the parasite plasma membrane. The removal of phosphate groups from surface proteins in host cells could attenuate electrostatic repulsion between parasite and host cells or even expose additional sites for the interaction to occur. Ectophosphatases themselves could contain adhesive domains that directly promote the attachment of parasite cells to their hosts, thus functioning similarly to well-characterized microbial adhesins [43]. Once P. serpens–O fasciatus interaction can be reproduced in the laboratory [11], a study investigating the role played by ectophosphatase activity in this process would be a great perspective to deepen the knowledge in this field.
In other approaches, the addition of sodium orthovanadate to the culture medium of different microorganisms impaired their growth, suggesting the involvement of ectophosphatase activity in cell proliferation [20,27,45]. Corroborating these previous data, we observed that sodium orthovanadate was able to inhibit both ectophosphatase activity and proliferation of P. serpens (Figure 6). Based on this result, we can hypothesize that ectophosphatase activity plays a significant role in this protist’s growth. Because of its extracellular active site, ectophosphatase activity allows the supply of inorganic phosphate (Pi) from organic phosphates found in the environment. Pi is an essential nutrient for the proliferation of all organisms, being required for the biosynthesis of nucleic acids, proteins, lipids, and sugars, and also for energy metabolism and signal transduction [36].
The first study describing a mechanism for Pi transport across the plasma membrane of trypanosomatids was performed with T. rangeli, identifying two systems involved in Pi acquisition during the parasite life cycle [46]. Posteriorly, several Pi-uptake systems were identified and characterized in different trypanosomatids, including Leishmania infantum [47], T. cruzi [48], T. brucei [49], L. amazonensis [50], and other protists such as Giardia duodenalis [51] and A. castellanii [52,53]. At least two high-affinity Pi-transport systems were identified in P. serpens, one transport coupled to Na+-ATPase and the other coupled to H+-ATPase [54]. Besides the biochemical studies, phylogenetic analysis of two hypothetical protein sequences found in the Phytomonas protein database showed similarities to the well-known high-affinity Pi transporters Pho84 and Pho89 of Saccharomyces cerevisiae [54]. Given that protists can uptake Pi from the extracellular medium, it is likely that ectophosphatases act together with the transport systems in the Pi acquisition process, by generating the extracellular Pi that can cross the microorganisms’ plasma membranes through a co-transport with Na+ or H+.
The transcription of genes encoding phosphatases and the Pi transporters can be modulated by the Pi availability. The phosphate sensing and acquisition pathway, known as the PHO pathway, has been extensively characterized in S. cerevisiae. The induction of approximately twenty genes involved in phosphate acquisition, transport, and storage is initiated by the nuclear accumulation of the transcription factor Pho4 in response to Pi starvation. These include secreted acid phosphatases, such as Pho5, Pho10, and Pho11, which are found in either the periplasmic space or the cell wall, being able to provide phosphate from many different substrates [55]. Although the occurrence of the PHO system in trypanosomatids remains to be elucidated, it has already been described that Pi starvation leads to an increase in both ectophosphatase activity and Pi uptake [36,46,48,51]. In this context, the elucidation of the molecular mechanisms involved in the phosphate acquisition process of P. serpens, such as the differential expression of phosphatases and Pi transporters in response to Pi availability, emerges as a relevant perspective in future research directions.
Further study of the biology of P. serpens may contribute to research into human diseases caused by pathogenic trypanosomatids, since ectophosphatase activity has been identified as potential therapeutic target [27]. Furthermore, the development of an efficient treatment against the damage caused to plants parasitized by Phytomonas would have a great impact from an economic point of view. The present work provides the biochemical characterization of the ectophosphatase activity of P. serpens, adding relevant information to the current knowledge about this phytopathogen. Moreover, we believe that the detailed comparison of the main biochemical properties of the ectophosphatases of different microorganisms brings an important contribution to the study of this remarkable enzymatic activity.

4. Materials and Methods

Microorganism and growth curves: Phytomonas serpens (isolate 9T, CT-IOC-189) was axenically cultured in Warren medium supplemented with 10% heat-inactivated fetal bovine serum (Cripion, Rio de Janeiro, Brazil) at 28 °C [11]. Flagellates were collected on the third day, unless otherwise indicated, by centrifugation at 1500× g for 10 min. Prior to each experiment, the flagellates were washed three times in buffer containing 116 mM NaCl, 5.4 mM KCl, 5.5 mM glucose and 50 mM Hepes-Tris, pH 7.2. For growth curves, 2.0 × 106 flagellates/mL were inoculated in 15 mL of complete culture medium in triplicate. An aliquot was sampled from each flask daily, then fixed and counted using a Neubauer chamber.
Ectophosphatase activity assay: P. serpens were washed as described, and ectophosphatase activity was measured by incubating 1 × 108 flagellates/mL in a reaction medium, containing 5.0 mM p-nitrophenylphosphate (p-NPP) as the phosphatase substrate and buffered by the same buffer containing 116 mM NaCl, 5.4 mM KCl, 5.5 mM glucose and 50 mM Hepes-Tris, pH 7.2 for a final volume of 0.5 mL, for 1 h at 25 °C. When EDTA and divalent metals were tested, 1 mM of these compounds was added to the reaction medium. Other additions are explicitly stated in the respective figure’s legend. The reaction was interrupted with 1.0 mL of 1 M NaOH, and flagellates were pelleted as described. The product of the reaction, p-nitrophenolate (p-NP), was measured spectrophotometrically (λ = 425 nm). The phosphatase activity was expressed as the rate of p-NP production per hour and 108 flagellates. In the experiments where CuCl2, ZnCl2, CaCl2, and CoCl2 were tested, the possible phosphate precipitates formed were checked as described previously [56]. Under the conditions employed here, no phosphate precipitates were observed in the presence of these salts. To test different pH values, a buffer composed of 10 mM MES, 10 mM HEPES and 10 mM TRIS was used. Viability after the assays under all conditions was greater than 95% as measured by the Trypan blue method.
Statistical analysis: All experiments were performed in triplicate, with similar results obtained from at least three separate cell suspensions. The values presented in all experiments represent the mean ± SE. Differences were considered significant at p < 0.05, as determined by one-way analysis of variance using Turkey’s multiple comparisons test, unless otherwise specified in the figure legends. The kinetic parameters (apparent Km and Vmax values) were calculated using nonlinear regression analysis of data from the Michaelis–Menten equation. All statistical analyses were performed with GraphPad Prism 6.0 software (GraphPad Software).

Author Contributions

Conceptualization, L.F.C.-K., T.S.S.M. and J.R.M.-F.; Data curation, L.F.C.-K., A.L.F.-M. and T.S.S.M.; Formal analysis, L.F.C.-K., A.L.F.-M., T.S.S.M. and J.R.M.-F.; Funding acquisition, J.R.M.-F.; Investigation, L.F.C.-K., A.L.F.-M., T.S.S.M. and J.R.M.-F.; Methodology, L.F.C.-K.; Supervision, J.R.M.-F.; Validation, L.F.C.-K. and J.R.M.-F.; Writing—original draft, A.L.F.-M., T.S.S.M. and J.R.M.-F. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by grants from the Brazilian agencies Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq—Grant Number: 310670/2021-7), Coordenação de Aperfeiçoamento de Pessoal de Nível Superior Finance Code 001 (CAPES—Grant Number: 0012017) and Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ—Grant Number: E-26/201.136/2021).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available upon request from the corresponding author.

Acknowledgments

We would like to thank Fabiano Ferreira Esteves and Rosangela Rosa de Araújo for their excellent technical assistance.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Time course and flagellate density dependence of Phytomonas serpens ectophosphatase activity. Ectophosphatase activity is linear with both (A) time and (B) flagellate density ((A): r2 = 0.987, n = 3; (B): r2 = 0.991, n = 3), thus validating the use of intact flagellates to assess ectophosphatase activity.
Figure 1. Time course and flagellate density dependence of Phytomonas serpens ectophosphatase activity. Ectophosphatase activity is linear with both (A) time and (B) flagellate density ((A): r2 = 0.987, n = 3; (B): r2 = 0.991, n = 3), thus validating the use of intact flagellates to assess ectophosphatase activity.
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Figure 2. Phosphatase activity of intact cells and the supernatant of Phytomonas serpens. Intact cells were incubated for 60 min at 25 °C in the reaction buffer in the absence of the substrate p-NPP. After this time, the cells were centrifuged, and aliquots of the cells and supernatant were assayed for ectophosphatase activity (n = 3). The asterisk denotes significant difference (p < 0.05).
Figure 2. Phosphatase activity of intact cells and the supernatant of Phytomonas serpens. Intact cells were incubated for 60 min at 25 °C in the reaction buffer in the absence of the substrate p-NPP. After this time, the cells were centrifuged, and aliquots of the cells and supernatant were assayed for ectophosphatase activity (n = 3). The asterisk denotes significant difference (p < 0.05).
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Figure 3. Effect of pH on the ectophosphatase activity of Phytomonas serpens. All pH values were tested with the same buffering system (10 mM each MES/HEPES/TRIS) to guarantee both cell integrity and buffering capacity from pH 5.0 to 8.0. The ectophosphatase activity is acidic, although it was not possible to observe its optimum pH because of the need to keep microorganisms intact during all the assays. Black circles represent means ± standard errors (n = 3). In some cases, standard errors are graphically smaller than symbols.
Figure 3. Effect of pH on the ectophosphatase activity of Phytomonas serpens. All pH values were tested with the same buffering system (10 mM each MES/HEPES/TRIS) to guarantee both cell integrity and buffering capacity from pH 5.0 to 8.0. The ectophosphatase activity is acidic, although it was not possible to observe its optimum pH because of the need to keep microorganisms intact during all the assays. Black circles represent means ± standard errors (n = 3). In some cases, standard errors are graphically smaller than symbols.
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Figure 4. Effect of p-NPP concentration on the ectophosphatase activity of Phytomonas serpens. The first and second points of the ectophosphatase activities were measured within 30 min to ensure that p-NPP hydrolysis did not exceed 10% of the available substrate; other concentrations were assayed for 60 min. Black circles represent means ± standard errors (n = 3), and experimental data fitting shows a nonlinear regression using the Michaelis–Menten equation. Calculated apparent kinetic parameters were Km = 1.575 ± 0.077 mM p-NPP and Vmax = 10.11 ± 0.137 nmol p-NP/(h × 108 flagellates).
Figure 4. Effect of p-NPP concentration on the ectophosphatase activity of Phytomonas serpens. The first and second points of the ectophosphatase activities were measured within 30 min to ensure that p-NPP hydrolysis did not exceed 10% of the available substrate; other concentrations were assayed for 60 min. Black circles represent means ± standard errors (n = 3), and experimental data fitting shows a nonlinear regression using the Michaelis–Menten equation. Calculated apparent kinetic parameters were Km = 1.575 ± 0.077 mM p-NPP and Vmax = 10.11 ± 0.137 nmol p-NP/(h × 108 flagellates).
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Figure 5. Effects of different metals on Phytomonas serpens ectophosphatase activity. The ectophosphate activity was inhibited by zinc and stimulated by cobalt; other metals tested could not modulate the activity. EDTA, a divalent cations chelator, had no effect on ectophosphatase activity, which suggests that this activity is not carried by a metalloenzyme (n = 3). EDTA and divalent metals were tested at a concentration of 1 mM. The asterisks denote significant differences (p < 0.05) compared to the control.
Figure 5. Effects of different metals on Phytomonas serpens ectophosphatase activity. The ectophosphate activity was inhibited by zinc and stimulated by cobalt; other metals tested could not modulate the activity. EDTA, a divalent cations chelator, had no effect on ectophosphatase activity, which suggests that this activity is not carried by a metalloenzyme (n = 3). EDTA and divalent metals were tested at a concentration of 1 mM. The asterisks denote significant differences (p < 0.05) compared to the control.
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Figure 6. Effects of sodium orthovanatade on Phytomonas serpens ectophosphatase activity and growth. The ectophosphate activity was inhibited by 1 mM sodium orthovanadate ((A), n = 3); P. serpens proliferation was inhibited by 1 mM sodium orthovanadate from the second up to the fifth day of growth ((B) (n = 3). The asterisks denote significant differences (p < 0.05) compared to the controls.
Figure 6. Effects of sodium orthovanatade on Phytomonas serpens ectophosphatase activity and growth. The ectophosphate activity was inhibited by 1 mM sodium orthovanadate ((A), n = 3); P. serpens proliferation was inhibited by 1 mM sodium orthovanadate from the second up to the fifth day of growth ((B) (n = 3). The asterisks denote significant differences (p < 0.05) compared to the controls.
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Table 1. Biochemical properties of ectophosphatase activities of different microorganisms.
Table 1. Biochemical properties of ectophosphatase activities of different microorganisms.
MicroorganismOptimal pH RangeKmVmaxInhibitory MetalsStimulant MetalsRef
Phytomonas françaiacid0.36 ± 0.06
(mM p-NPP)		5.34 ± 0.02
nmol Pi/(mg protein × min)		Zn2+ND[24]
Phytomonas mcgheeiacid1.33 ± 0.27
(mM p-NPP)		8.54 ± 0.68
nmol Pi/(mg protein × min)		Zn2+Cu2+[24]
Angomonas deaneiND5.35 ± 0.84
(mM p-NPP)		6.93 ± 0.42
nmol Pi/(h × 108 cells)		Zn2+ND[25]
Herpetomonas muscarumacid0.76 ± 0.04
(mM p-NPP)		4.90 ± 0.46
nmol Pi/(mg protein × min)		Zn2+Al3+[23]
Leishmania amazonensisND1.78 ± 0.32
(mM p-NPP)		31.93 ± 3.04
nmol Pi/(h × 107 cells)		Zn2+ND[27]
Strigomonas culicisacidNDNDZn2+Mg2+[15]
Trypanosoma bruceiacid0.66 ± 0.09
(mM p-NPP)		0.72 ± 0.07
nmol p-NP/(mg protein × min)		Zn2+Mg2+ Mn2+ Co2+ Cu2+[26]
Trypanosoma cruzi
(Y strain)		acid8.6
(mM p-NPP)		3.9
nmol p-NP/(mg protein × min)		Zn2+ND[28]
Trypanosoma cruzi
(Colombiana strain)		acid1.6
(mM p-NPP)		37.0
nmol p-NP/(mg protein × min)		NDND[28]
Trypanosoma rangeli
(Macias strain)		alkaline1.04 ± 0.16
(mM p-NPP)		8.94 ± 0.36
nmol p-NP/(h × 107 cells)		NDMg2+[30]
Trypanosoma rangeli
(H14 strain—short forms)		acid3.48 ± 0.62
(mM p-NPP)		26.27 ± 1.10
nmol p-NP/(h × 107 cells)		Zn2+ Mg2+[29]
Trypanosoma rangeli
(H14 strain—long forms)		acid0.22 ± 0.04
(mM p-NPP)		17.10 ± 1.07
nmol p-NP/(h × 107 cells)		Zn2+, Ca2+Mg2+ Mn2+, Sr2+[29]
Acanthamoeba castellaniiacid2.12 ± 0.54
(mM p-NPP)		26.12 ± 2.53
nmol Pi/(h × 106 cells)		NDMg2+ Co2+ Ni2+[34]
Euglena gracilisacid2.52 ± 0.12
(mM p-NPP)		3.62 ± 0.06
nmol Pi/(h × 106 cells)		Zn2+Cu2+[35]
Entamoeba histolyticaacid2.68 ± 0.25
(mM p-NPP)		8.00 ± 0.22
nmol Pi/(h × 105 cells)		Zn2+ND[32]
Giardia duodenalisacidNDNDNDND[33]
Trichomonas vaginalisacid0.24 ± 0.02
(mM p-NPP)		167.2 ± 19.8
nmol Pi/(h × 107 cells)		Zn2+ND[31]
Blastocladiella emersoniialkaline1.34 ± 0.18
(mM p-NPP)		5.29 ± 0.27
nmol Pi/(min × 107 cells)		NDND[22]
Candida parapsilosisacid0.47 ± 0.05
(mM p-NPP)		26.80 ± 1.13
nmol Pi/(h × 107 cells)		Zn2+Cu2+[17]
Cryptococcus neoformansacid29.2 ± 3.3
(mM p-NPP)		110.9 ± 8.1
nmol p-NP/(h × 108 cells)		Zn2+ND[16]
Fonsecae pedrosoiacid15.12 ± 3.55
(mM p-NPP)		18.07 ± 2.30
nmol p-NP/(h × 107 cells)		NDFe3+[19]
Metarhizium anisopliaeacidNDNDZn2+, Cu2+, Cd2+ND[21]
Pseudallescheria boydiialkaline6.66 ± 1.0
(mM p-NPP)		103.0 ± 8.6
nmol p-NP/(mg dry weight × min)		Cu2+, Cd2+Mg2+, Mn2+, Zn2+[18]
Rhinocladiella aquaspersaacid3.08 ± 0.38
(mM p-NPP)		24.46 ± 1.04
nmol p-NP/(h × 107 cells)		NDCo2+[20]
The metals added in the form of chloride or sulfide; ND = data not available.
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Carvalho-Kelly, L.F.; Freitas-Mesquita, A.L.; Majerowicz, T.S.S.; Meyer-Fernandes, J.R. Biochemical Properties of the Acid Ectophosphatase Activity of Phytomonas serpens Involved in Cell Proliferation. Kinases Phosphatases 2024, 2, 379-390. https://doi.org/10.3390/kinasesphosphatases2040024

AMA Style

Carvalho-Kelly LF, Freitas-Mesquita AL, Majerowicz TSS, Meyer-Fernandes JR. Biochemical Properties of the Acid Ectophosphatase Activity of Phytomonas serpens Involved in Cell Proliferation. Kinases and Phosphatases. 2024; 2(4):379-390. https://doi.org/10.3390/kinasesphosphatases2040024

Chicago/Turabian Style

Carvalho-Kelly, Luiz Fernando, Anita Leocadio Freitas-Mesquita, Thaís Souza Silveira Majerowicz, and José Roberto Meyer-Fernandes. 2024. "Biochemical Properties of the Acid Ectophosphatase Activity of Phytomonas serpens Involved in Cell Proliferation" Kinases and Phosphatases 2, no. 4: 379-390. https://doi.org/10.3390/kinasesphosphatases2040024

APA Style

Carvalho-Kelly, L. F., Freitas-Mesquita, A. L., Majerowicz, T. S. S., & Meyer-Fernandes, J. R. (2024). Biochemical Properties of the Acid Ectophosphatase Activity of Phytomonas serpens Involved in Cell Proliferation. Kinases and Phosphatases, 2(4), 379-390. https://doi.org/10.3390/kinasesphosphatases2040024

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