Aspartyl protease inhibition interferes with Plasmodium falciparum asexual blood-stage and early gametocyte development

Abstract

Background

Plasmodium falciparum is the most influential species of malaria parasites, capable of causing severe illness and mortality, especially in pregnant women and children under the age of 5. Global distribution of disease impacted on billions of endemic people and travellers. Asexual stage and gametocyte cause harmful manifestations, impacting the patients and contributing to the spread of the disease in the community, respectively. Moreover, most recent therapeutic drugs did not affect the gametocyte. The discovery of a new drug with dual actions on both stages could elucidate a cost-effective way to combat malaria. Within a human host, the parasite possesses many activities for its survival, such as invasion, egress, haemoglobin degradation, and protein trafficking, many of which are related to aspartyl protease, revealing the potential for antimalarial drug targets.

Methods

Pepstatin A, the representative of the board-spectrum aspartyl protease inhibitor, was utilized to investigate the effects of aspartyl protease inhibition on parasite development. The experiments were separately performed in vitro for different developmental stages of parasites, including the asexual blood-stage, early gametocytes, late gametocytes, and gamete. To demonstrate the effects of pepstatin A, the number of intact parasites and their stage distribution were counted under the microscope and calculated as a percentage of inhibition compared to the control. Additionally, the morphology of pepstatin A-treated parasites was observed to identify cellular alterations in the parasites.

Results

Pepstatin A at 100 µM inhibited the asexual stage and early-stage gametocyte development by 47% and 73%, respectively. They exhibited morphological defects, including chromatin condensation, vacuolization and haemozoin clumping in both asexual blood-stage and early-stage gametocyte. However, it could not influence the late-stage gametocyte development and gamete formation.

Conclusion

The inhibition of aspartyl protease by pepstatin A moderately affected both asexual blood-stage and early-stage gametocyte development. Morphological changes on treated parasites implied the effect of pepstatin A on haemoglobin degradation process, suggesting its potential for reducing the severity of the disease and minimizing malaria transmission. However, further research and development are required to use aspartyl protease as a drug target, focusing on identifying and modifying the drug to be more sensitive and effective.

Background

Malaria is an infectious tropical and subtropical disease caused by the Plasmodium parasite. Plasmodium falciparum, the causative agent of human malaria, can manipulate and develop through several stages following red blood cell (RBC) invasion, including ring, trophozoite, schizont, and five stages of gametocyte [1]. Significantly, the intraerythrocytic stage was the most hazardous, causing a coma and death in approximately 500,000 people each year [2]. Moreover, the mature stage V gametocyte was a precursor stage that mediate malaria transmission [3].

Although there are various strategies to counteract malaria, such as developing new pharmaceuticals for treatment and prevention, inventing new innovations for malaria management, and educating the local community in endemic areas, there are no 100% effective malaria eradication strategies. Furthermore, the majority of antimalarial medicines had no effect on gametocytes. Malaria transmission to different communities continued for weeks after asexual stage parasites had been cleared [4]. As a result, the development of medications that target the key mechanisms in both the asexual stage parasite and the gametocyte may be the most cost-effective malaria eradication strategy.

The important source of nutrients for the parasite growth in the intraerythrocytic stage was the amino acid derived from the haemoglobin degradation process [5]. Previous research showed that this mechanism was essential for both asexual and gametocyte development. In this process, various protease enzymes worked together in a cascade to degrade the large molecules of haemoglobin into small molecules of amino acids [6–9]. This function in the parasite cellular mechanism suggested that inhibiting enzymes involved in this process could impact on the development of the intraerythrocytic parasites.

One type of protease involved in the haemoglobin breakdown process was aspartyl protease [5, 10]. Aspartyl proteases in P. falciparum were made up of ten members, plasmepsin I to X, and they played a crucial role in both the asexual stage parasite and the gametocyte. These proteases contribute to the parasite's survival and growth within the erythrocyte. They were not only participated in haemoglobin breakdown but also contributed to other critical activities such as protein export, and they may be involved in parasite invasion and egress [11–16]. Thus, the aspartyl protease could be a powerful enzyme critical to the parasite's survival, indicating its potential as a useful target for blocking parasite development and transmission.

Pepstatin A was a broad-spectrum aspartyl protease inhibitor that has been demonstrated to block haemoglobin degradation enzymes and to reduce the growth of the Babesia parasite [17], which belonged to the same Apicomplexa phylum as the Plasmodium parasite. Moreover, its effect on the asexual stage of the P. falciparum 3D7 strain had been demonstrated [18, 19]. For this reason, it can be hypothesized that the use of pepstatin A could help elucidate the effects of aspartyl protease inhibition on the development of asexual blood-stage parasites, gametocytes, and gametes, especially in transmission stages that have not yet been investigated.

In this study, pepstatin A was co-cultured with the intraerythrocytic P. falciparum AMB47 Thai isolate to examine aspartyl protease inhibitory effects on various developmental stages, including asexual blood stage, early, late gametocyte, and gamete. Notably, the morphology of the treated parasites was illustrated for describing the possible cellular process interfered by pepstatin A.

Methods

In vitro culture of Plasmodium falciparum

Plasmodium falciparum AMB47, a gametocyte-productive strain from Thailand, was cultivated using the method described by Read with some modifications [20]. In brief, the parasite’s asexual stage was constantly cultured at 5% haematocrit (HCT, O-type blood cells purchased from Thai Red Cross, Bangkok, Thailand) under mixed gas conditions (90% N₂, 5% O₂ and 5% CO₂) at 37 °C. The complete culture medium for the asexual stage of P. falciparum consists of RPMI 1640 powdered medium with L-Glutamine (Gibco), 2 g/L sodium bicarbonate (Mallinckrodt), 25 mM HEPES (Sigma), 2 g/L dextrose (M&B), 50 mg/L hypoxanthine (Sigma), 10% heat-inactivated human serum type A positive and 10 mg/L gentamicin (Siam pharmaceutical). The parasite was synchronized with 5% sorbitol (PanReac AppliChem ITW Reagents) to convert the mixed stage culture to ring stage [21].

The gametocyte culture followed the method outlined by Bounkeua [22] method with some changes. It was maintained under the same conditions as the asexual stage culture, excluding gentamicin. Initially, the parasitaemia of the asexual stage culture, ranging from 5–8%, was diluted to achieve a parasitaemia of 0.4–0.6%. It was then cultured at 6% HCT until the parasitaemia was up to more than 5%. At this point, the haematocrit culture percentage was reduced to 3.5% HCT (Day1). To prevent undesired gametogenesis, all gametocyte culture processes were carried out on a 37 °C hot plate. Mature stage V gametocytes were typically observed in the culture on Days 12–14.

Drug and inhibitor preparation

Pepstatin A (Sigma), an aspartyl protease inhibitor, was prepared at a concentration of 1 mM in 5% Dimethyl sulfoxide (DMSO). The positive control for the asexual stage inhibition experiment was 100 nM Artesunate (ATS), and the control for the gametocyte development experiment was 100 nM Chloroquine (CQ). They were stored at − 20 °C before the experiment. The negative control used in this investigation was 0.5% DMSO.

Development inhibition for asexual stage development

In a 96-well plate, the synchronized late ring stage at 1% parasitaemia was co-cultured with pepstatin A concentrations ranging from 0.1 to 100 µM, along with negative and positive controls, for 26–28 h. After incubation, the treated parasites were harvested, and Giemsa-stained thin blood smears were prepared to investigate the defects of the parasite under light microscope.

Early-stage gametocyte development inhibition (EGDI) and late-stage gametocyte development inhibition (LGDI)

The experiments were carried out using the modified method of Adjalley [22]. Various concentrations of pepstatin A were treated in the 2% HCT of gametocyte culture daily for 3 consecutive days. The investigation of the early-stage gametocyte development inhibition (EGDI) began on Day 3 of gametocyte development when the parasites were in stage I and II gametocytes. Late-stage gametocyte development inhibition (LGDI) was initiated on Day 8 when the parasites were in stage III and IV gametocytes. Giemsa-stained thin blood smears were prepared for light microscopic examination after 3 days of treatment.

Gamete formation inhibition

The interval from day 12 to 14 of gametocyte culture is an adequate period to generate gamete. The gametocyte was pre-incubated with 10 µM or 100 µM of pepstatin A for 15 min at 37 °C [23]. Afterward, they were spun down and immediately mixed with 21 °C ookinete medium in a 2:1 ratio for 15 min at room temperature. The ookinete medium contains incomplete medium, 20% non-heat inactivated serum type A positive, and 100 µM of xanthurenic acid to induce gamete formation. The number of exflagellation centers was counted using a haemocytometer, and Giemsa-stained thin blood smears were prepared to examine the number of macrogamete formations.

Light microscopic observation

Under a light microscope, the percentage of parasitaemia and morphological changes were examined after treating the parasites with pepstatin A. The number of normal parasites in 10,000 total red blood cells (total RBCs) was counted and calculated as a percentage of inhibition relative to the untreated control.

The formula for calculating the percentage of inhibition;

$$\%\text{Inhibition}=100-\left(\frac{\%\text{parasitaemia}\text{of}\text{treated}\text{group}}{\%\text{parasitaemia}\text{of}\text{untreated}\text{group}}\times 100\right)$$

The morphological investigation

The morphological changes of the parasite were observed and investigated as shown in Tables 1 and 2. Shape, nucleus, haemozoin, and vacuolization were the key constituents studied.

Table 1.

The criteria for morphological characterization on asexual blood stage parasite

Composition	Normal			Defect
	Ring	Trophozoite	Schizont
Nucleus	1 or 2 nuclei	≤ 2 nuclei	> 2 nuclei	Chromatin condensation, pyknosis, karyorrhexis and karyolysis
Haemozoin	No haemozoin formation	Brown pigment	Dark brown pigment	Clumped black haemozoin
Vacuolization	No vacuole formation	Tiny size, ≤ 3 vacuole formation		Enlarged or increased the number (with/without the content)

Table 2.

The criteria for morphological characterization on gametocyte

Composition	Normal					Defect
	I	II	III	IV	V
Shape	Difficult to differentiate from trophozoite	D-shape	Elongated D-shape with blunt-ends	Elongated shape with pointed ends	Crescent-shape with blunt-ends	Shrinkage or swelling
Nucleus		Elongate nucleus which located in the terminal site		Male gametocyte has the nucleus larger than female		Chromatin condensation, pyknosis, karyorrhexis and karyolysis
Haemozoin	Distribute throughout the cytoplasm of gametocyte					Clumped haemozoin or accumulation in the same food vacuole
Vacuolization	Tiny size, difficult to observe in the gametocyte by light microscope					Enlarged or increased the number (with/without the content)

Statistical analysis

The statistical analysis to compare the significant difference was performed by one-way ANOVA. The half-maximal inhibitory concentration (IC₅₀) was determined through Probit analysis from IBM® SPSS® statistics 24.

Results

The effect of aspartyl protease inhibition on asexual blood-stage

From the past until now, several aspartyl protease inhibitors have been synthesized and tested for their ability to inhibit this influential enzyme. Among these inhibitors, pepstatin A has shown the ability to inhibit all plasmepsins (Table 3). Therefore, pepstatin A was selected to investigate the inhibition of aspartyl proteases that could interfere with the development of the asexual blood-stage parasite.

Table 3.

Summary of plasmepsin inhibitor testing in Plasmodium spp

Inhibitor	Target plasmepsin	Experiment	References	Note	Inhibitor	Target plasmepsin	Experiment	References	Note
Azacyclic plasmepsin inhibitors	Plasmepsin II	In vitro enzyme activity	[57]	Fluorescence‐based proteolysis assay	KNI-764	Plasmepsin IV	In vitro enzyme activity	[58]	Pm
		Crystallization				Plasmepsin IV	Crystallization		Pm
	Plasmepsin IV	In vitro enzyme activity		Fluorescence‐based proteolysis assay	KNI-10006	Plasmepsin I	Crystallization	[59]
		Crystallization				Plasmepsin I	Molecular Docking	[60]
Azole-based inhibitor	Plasmepsin II	Molecular Docking	[61]			Plasmepsin II	Molecular Docking	[60]
		In vitro enzyme activity				Plasmepsin IV	Molecular Docking
	Plasmepsin IX	Molecular Docking			KNI-10395	HAP	Crystallization	[62]
		In vitro enzyme activity			Pepstatin A	All plasmepsins	Molecular Docking	[63]
	Plasmepsin X	Molecular Docking				Haemoglobin degradation enzyme	In vitro enzyme activity	[6]
		In vitro  enzyme activity				Plasmepsin II	Crystallization	[64]
Canavanine	Plasmepsin V	In vitro  enzyme activity	[65]	Fluorescence‐based proteolysis assay			Crystallization	[66]
		Crystallization					Crystallization	[67]
		Molecular Docking				HAP	Crystallization	[68]
Hydroxyethylamine derivative	Plasmepsin I	In vitro  enzyme activity	[69]			Plasmepsin IV	Dimerization	[67]	Pf,Pv
	Plasmepsin II	In vitro  enzyme activity	[70]				Crystallization	[71]	Pv
		Molecular Docking				Plasmepsin V	In vitro enzyme activity	[14]	Partially inhibited
		Crystallization	[72]				In vitro enzyme activity	[73]
	Plasmepsin IV	In vitro  enzyme activity	[70]			Plasmepsin IX	MD simulations	[74]
		Molecular Docking				Plasmepsin X	MD simulations
		In vitro  enzyme activity	[75]		Pepstatin A analogue	Plasmepsin II	In vitro enzyme activity	[76]
		Molecular Docking				Plasmepsin II	Crystallization	[76]
	Plasmepsin V	In vitro  enzyme activity	[77]		PEXEL peptidomimetic inhibitors	Plasmepsin V	In vitro enzyme activity	[78]	Fluorescence‐based proteolysis assay
	Plasmepsin IX	Parasite egress inhibition	[79]	Pb			Crystallization
		Parasite invasion inhibition		Pb			Molecular Docking
		In vitro  enzyme activity		Pb	RS367	Plasmepsin II	Crystallization	[80]
	Plasmepsin X	Parasite egress inhibition		Pb	RS370	Plasmepsin II	Crystallization	[80]
		Parasite invasion inhibition		Pb	HIV-1 inhibitors	Plasmepsin II	Crystallization In vitro  enzyme activity	[81]
	Plasmepsin X	In vitro enzyme activity		Pb		Plasmepsin X	In vitro  enzyme activity	[82]

The life cycle of P. falciparum required around 48 h for its development, progressing from the ring to the trophozoite and finally the schizont stage. In this study, the late ring stage (10–16 h) was co-cultured with various concentrations of pepstatin A. After 26–28 h of incubation, the parasite showed 8.2 ± 6.4% inhibition in 0.5% DMSO treatment, 2.2 ± 8.6%, 3.3 ± 10.8%, 25.5 ± 5.6%, 46.7 ± 13.4% inhibition for pepstatin A treatments ranging from 0.1, 1, 10, to 100 µM, and 94.8 ± 4.0% inhibition for ATS treatment, in comparison to the untreated control (Fig. 1A). The parasite in the negative group developed into a typical trophozoite and schizont, characterized by ≥ one nucleus with the brown pigment of haemozoin.

Fig. 1.

Pepstatin A treatment inhibits the development of asexual blood-stages. The bar graph represents the percentage of development inhibition after treatment compared to the untreated control (A). The parasites were staged under a light microscope both before (0 h) and after treatment. The distribution of the asexual stage and % parasitaemia in each condition is shown by the 100% stacked column (B). The conversion rate graph of parasites from ring to trophozoite and schizont (C). The signification was indicated by an asterisk [n = 6 biological replicates, F_{(6, 35)} = 58.26, *P = 0.0494, ***P < 0.0001 by ordinary one-way ANOVA]. ATS artesunate, DMSO dimethylsulfoxide

The treatment with 100 µM of pepstatin A inhibited development by 46.7% at the IC₅₀ greater than 100 µM (Fig. 1A). Although the IC₅₀ for pepstatin A was higher than expected, it exhibited a trend of development inhibition, as demonstrated by the reducing proportion and % parasitaemia of developing parasites compared to controls (Fig. 1B). Furthermore, the conversion rate of parasite from ring to trophozoite and schizont stage, indicative of normal parasite development, was revealed to be significantly reduced in 100 µM of pepstatin A when compared to untreated parasite (Fig. 1C).

Pepstatin A treatment caused chromatin condensation, vacuolization, and haemozoin clumping in asexual stage development

The morphological investigation was a simple yet advantageous study that helped to discover or predict the relationship between the parasite's defect after treatment and the underlying mechanism. In this study, the morphological defects of the asexual blood-stage parasite were investigated under a light microscope following drug treatment. The normal ring stage before treatment represented 1–2 nuclei without haemozoin formation (0 h). After 26–28 h, the parasite in the untreated control and 0.5% DMSO treatment group developed into trophozoite and schizont, characterized by ≥ two nuclei with normal dark brown haemozoin pigment. Following 100 µM pepstatin A treatment, three primary morphological defects—chromatin condensation, vacuolization, and haemozoin clumping—were identified in the asexual blood-stage. In 100 nM ATS treatment, dead parasites were recognized by the dark blue colour of the pyknotic nucleus (Fig. 2).

Fig. 2.

The morphological changes of the asexual blood-stage after treatment. The Giemsa-stained thin smears were observed under a light microscope to investigate the parasite's defect after treatment. The parasites were photographed before (0 h) and after treatment (26–28 h), including untreated control, 0.5% DMSO, 100 µM pepstatin A, and 100 nM ATS. Arrow indicated abnormal vacuole formation, red arrowhead denoted haemozoin clumping, and black arrowhead specified chromatin condensation. A micron bar indicated 10 µm. ATS Artesunate, DMSO dimethylsulfoxide

In general, the asexual blood-stage haemozoin pigment was brown to dark brown in colour and accumulated in the same location. In this investigation, a high dose of pepstatin A treatment (100 µM) caused haemozoin clumping and turned the malaria pigment into black colour (Fig. 2). The normal food vacuoles in parasites were usually found in a tiny size. However, this study demonstrated that pepstatin A treatment increased both size and the number of vacuole formations in the parasites (Fig. 2). Importantly, the sign of cell death also was observed in pepstatin A-treated parasites as show the chromatin condensation starting from the edge of nucleus or forming ring condensation (Fig. 2).

Targeting aspartyl protease by pepstatin A affected early-stage gametocyte development but not late-stage gametocyte development

Aspartyl protease was involved not only in the asexual blood-stage but also in the gametocyte. The effect of pepstatin A treatment on gametocyte development was examined. P. falciparum undergoes five stages of gametocyte development, stage I through V, each with a unique metabolism. In this study, investigations on gametocyte development were divided into early-stage gametocyte (stage I to III) and late-stage gametocyte (stage IV and V).

Pepstatin A was co-cultured in tenfold dilutions ranging from 100 to 0.1 µM, as in the asexual stage experiment. After 3 days of treatment, the gametocytes in the negative control developed normally from stage I or II to stage II or III in the EGDI test and from stage III or IV to stage IV or V in the LGDI assay. Pepstatin A treatment highlighted the importance of aspartyl protease in gametocyte development, demonstrating that early-stage gametocyte development was inhibited by 100 µM pepstatin A (IC₅₀ = 53.9 µM). In comparison to the untreated control, the early-stage gametocyte showed 1.6 ± 13.8% inhibition in 0.5% DMSO treatment, 10.9 ± 10.2%, 15.5 ± 10%, 42.5 ± 5.0% and 72.6 ± 7.1% inhibition with pepstatin A treatment ranging from 0.1, 1, 10, to 100 µM, respectively. CQ treatment exhibited 89.09 ± 8.83% inhibition (Fig. 3A). Moreover, the percentage of early-stage gametocyte inhibition was also related to the reduction of % parasitaemia of stage II and III gametocyte (Fig. 3B) and conversion rate of stage I&II to stage II&III gametocyte (Fig. 3C).

Fig. 3.

The gametocyte development inhibition after drug treatment. Early-stage gametocyte treatment (A–C). Late-stage gametocyte treatment (D–F). The bar graphs represented the early-stage gametocyte development inhibition (EGDI) relative to untreated control (A). For staging, gametocytes were examined under a light microscope both before (0 h) and after treatment. The distribution of the early-stage gametocyte and %gametocytemia in each condition is shown by the 100% stacked column (B). The bar graph shows the conversion rate of gametocyte from stage I&II to stage II&III (C) (n = 6 biological replicates, F _{(6, 30)} = 47.22, ***P < 0.0001 by one-way ANOVA). The bar graph depicts late-stage gametocyte development inhibition (LGDI) relative to the untreated control (D). The distribution of gametocyte stages and %gametocytemia before treatment (0 h) and after treatment in the LGDI assay (E). The bar graph illustrates the conversion rate of gametocytes from stage III&IV to stage IV&V when compared to the untreated control (F) [n = 6 biological replicates, F_{(6, 35)} = 17.33, **P = 0.0017, ***P < 0.0001 by ordinary one-way ANOVA]. CQ chloroquine, DMSO dimethylsulfoxide

On the other hand, the pepstatin A treatment had no effect on late-stage gametocyte development (IC₅₀ > 100 µM). After 3 days of treatment, the late-stage gametocyte showed 0 ± 19.5% inhibition in DMSO treatment, 6.9 ± 3.6%, 14.7 ± 11.7%, 17.8 ± 10.3%, and 31.1 ± 13.5% inhibition after pepstatin A treatment ranging from 0.1, 1, 10, to 100 µM, respectively. The positive control, CQ, exhibited 63.1 ± 10.4% inhibition (Fig. 3D). Although development inhibition increased in a dose-dependent manner (Fig. 3D) and the conversion of gametocyte stage III &IV to stage IV and V showed the significant reduction compared to untreated control (Fig. 3F), it had no effect on gametocyte development to stage V (Fig. 3E).

Pepstatin A treatment induced morphological changes exclusively in the early stages of gametocyte development

Morphological defects in both early and late-stage gametocytes were examined under a light microscope. The normal morphology of a stage II gametocyte characterized by a D-shape with blunt-ends and representing the normal distribution of haemozoin, was observed before treatment (0 h). After 3 days of continuous treatment, the untreated control and 0.5% DMSO-treated parasites progressed to stage II and III gametocytes, exhibiting an elongated D-shape with blunt ends and a normal haemozoin distribution. The defective stage III gametocyte could be found in 100 nM CQ and 100 µM pepstatin A treatment group. The gametocyte from the 100 nM CQ treatment group represented the haemozoin clumping. Chromatin condensation, haemozoin clumping and vacuole formation were distinctly evident in the pepstatin A treatment (Fig. 4A).

Fig. 4.

The morphological defects investigated in pepstatin A-treated gametocytes. A thin smear of Giemsa's stain was examined under a light microscope to investigate the parasite's defect after treatment. The parasite was photographed before (0 h) and after treatment, including untreated control, 0.5% DMSO, 100 µM pepstatin A, and 100 nM CQ. The morphology of gametocyte in the early-stage development inhibition (EGDI) experiment (A). The morphology of gametocyte in the late-stage development inhibition (LGDI) experiment (B). Haemozoin clumping is indicated by the red arrowhead, black arrowhead indicated chromatin condensation and vacuole formation was specified by arrow. The micron bar indicated 10 µm. CQ chloroquine, DMSO dimethylsulfoxide

In late-stage gametocytes, 0-h stage III gametocytes displayed an elongated D-shape with blunt ends and normal haemozoin distribution. After 3 days of continuous treatment under all conditions, parasites developed into normal stage IV and V gametocytes. Stage IV gametocytes had an elongated shape with pointed ends and a normal distribution of haemozoin pigment, while mature stage V gametocytes exhibited a crescent shape with normal haemozoin distribution at the center (Fig. 4B) Notably, abnormalities were exclusively observed in early-stage gametocytes following treatment with a high dose of pepstatin A (100 µM); no abnormalities were found in late-stage gametocytes.

The haemozoin pigment distribution in gametocytes differs from that in asexual stage parasites, as it is distributed throughout the cytoplasm of gametocytes rather than collecting in a specific area. In this work, a high dose of pepstatin A treatment (100 µM) induced haemozoin aggregation in the same vacuole (Fig. 4A). Additionally, similar to the asexual stage treatment, chromatin condensation indicated by dark purple colour concentrated at the inner surface of the nuclear membrane and vacuole formation characterized by increase in both size and number were observed in early-stage gametocytes.

The effect of aspartyl protease inhibition by pepstatin A during gamete formation

The transmission of the mature gametocyte from the patient to another human requires the critical process of gamete formation inside a female Anopheles mosquito. This process involved in the transformation of male and female gametocytes into microgametes and macrogametes [24]. It occurred after the gametocytes were activated by a change in pH, a drop in temperature, or exposure to xanthurenic acid within the mosquito [25, 26]. The process of microgamete formation was called exflagellation. The parasite replicates its genome to create an octoploid and develops eight flagella to find the macrogamete. The macrogamete was differentiated from the female gametocyte. The female gametocyte started to round up and emerged from the host red blood cell after being activated.

To determine whether pepstatin A could inhibit gamete formation, mature gametocytes were pre-incubated with pepstatin A for 15 min. After the activation of gametocyte, the results showed that pepstatin A could not inhibit the process of gamete formation. There were no significant differences between the pepstatin A treatments and the untreated control (Fig. 5A and B). Although there was a significant difference in the number between male gametocytes and male gamete formation, there was no significant difference in the number of male gamete formation between untreated control and pepstatin A treatment. Moreover, the morphology of macrogametes was normal as represented by no identification of RBC membrane surrounding parasites (Fig. 3C). The macrogamete normally exhibited a circular shape, dispersed haemozoin, a cytoplasm stained blue, and a single mass of chromatin similar to previously identified [27].

Fig. 5.

Gamete formation after treatment. The number of male gametocytes and exflagellation events was represented per 10,000 total RBCs in each group from the exflagellation assay (A). The number of female gametocytes and macrogamete formations was represented per 10,000 total RBCs in each group from the Giemsa-stained blood smear (B). Gametocyte morphology prior to activation and gamete formation in each condition (C). n = 3 biological replicates for gametocyte, and 6 biological replicates for gamete, F_{(3, 17)} = 692.6, ***P < 0.0001 by ordinary one-way ANOVA

Discussion

The malaria parasite has numerous mechanisms for survival in the human host. Each stage of parasite development has a unique mechanism that is important for that stage. For example, the invasion mechanism is for the parasite's invasive stages (merozoite, ookinete, and sporozoite), the egress mechanism is for schizont, mature gametocyte, and oocyst, and the haemoglobin degradation process is for parasite growth in asexual blood-stage and early-stage gametocyte. Although each stage of the parasite has its own mechanism, these mechanisms may share a key molecule.

Although these are distinct processes, the key molecules involved can overlap. Aspartyl proteases are a good example of a molecules involved in many important processes. They participate in the haemoglobin degradation, protein exportation and may be involved in invasion and egress of the parasites [11–15, 28]. Thus, the aspartyl proteases represent a group of critical enzymes essential for parasite survival. Notably, a previous study demonstrated that inhibiting two specific aspartyl enzymes inhibited the parasite at multiple stages [29].

This study explored the effects of pepstatin A, a broad-spectrum aspartyl protease inhibitor. Previous studies demonstrated the effect of pepstatin A in P. falciparum, confirming the impact of aspartyl protease inhibition on asexual blood-stage development [18, 19]. In this study, in addition to influencing the asexual blood stage of the parasite, pepstatin A also induced morphological changes in early gametocyte development. However, no effect was observed on late gametocyte development and gamete formation. Previous research supported this finding by highlighting the dependency of the parasite's development in the asexual stage and early-stage gametocyte on the haemoglobin degradation process, which was not required for late gametocyte development [8, 9].

Previous studies have indicated that pepstatin A itself poorly penetrates the parasite [30, 31]. However, it was later shown that commercial pepstatin A preparations often contain pepstatin butyl ester as a contaminant, which can enter the parasite more efficiently. Once inside, esterases cleave the ester, releasing active pepstatin to inhibit aspartyl proteases intracellularly [31]. The observed activity of pepstatin A on asexual blood stages and early gametocytes suggests that the preparation may have contained active ester forms, although their levels were not directly quantified.

Nevertheless, pepstatin A did not inhibit gametocyte maturation or gamete formation, including exflagellation, even at 100 μM. A previous study of 1,2-Epoxy-3-(p-nitrophenoxy) propane (EPNP), another aspartyl protease inhibitor, also reported limited effect on exflagellation at concentrations below 200 μM, with an IC₅₀ of 225.8 ± 20.5 μM [32]. These findings suggest that the inhibition of exflagellation by aspartyl protease inhibitors may only occur at high concentrations, possibly through off-target effects rather than specific aspartyl protease inhibition. Moreover, the EPNP study did not examine early-stage gametocytes; therefore, the current findings extend the understanding of stage-specific responses to aspartyl protease inhibition. Although pepstatin A did not inhibit gametocyte maturation or gamete formation under the conditions tested, the data overall reinforce the potential role of aspartyl proteases during early parasite development, including the asexual stage and early gametocytogenesis. Therefore, it is implied that inhibition of aspartyl proteases by pepstatin A can help reduce parasite burden and potentially limit transmission by targeting earlier developmental stages (Fig. 6).

Fig. 6.

The effect of aspartyl protease inhibition by pepstatin A treatment. Within 48 h, the asexual blood-stage developed from the ring stage to the trophozoite and schizont stages

Although dihydroartemisinin (DHA) is a frontline antimalarial drug known to affect early gametocyte development, it was not included in this study, as the focus was on characterizing pepstatin A as an aspartyl protease inhibitor. A previous study reported that DHA inhibits early-stage gametocytes with high potency, with IC₅₀ values in the low nanomolar range (< 120 nM) [22]. In contrast, pepstatin A showed only moderate activity at micromolar levels. While a direct comparison would be informative, chloroquine was selected as a reference due to its well-characterized morphological effects on early gametocytes and its utility in validating stage-specific responses in the assay system. Given the variability in ester content that influences pepstatin A’s uptake and activity, future comparative studies using standardized preparations would more clearly define the relative efficacy of pepstatin A compared to other antimalarials, such as DHA.

The first morphological defect visible in parasites treated with pepstatin A was vacuole formation. It was identified as cytoplasmic vacuolization in asexual blood-stage and early-stage gametocytes after treatment with pepstatin A. Vacuolization was usually associated with autophagy [33–35]. Generally, autophagy occurred to remove the damaged or aged organelles [36]. It was also related to cell death and became more prevalent as the cells lose nourishment [37–39]. Previous research demonstrated that vacuolization occurred in P. falciparum after antimalarial drug incubation [40, 41]. In this study, cytoplasmic vacuolization was observed in asexual and early-stage gametocytes following pepstatin A treatment. While vacuolization is often associated with autophagy in other systems, pepstatin A has not been shown to inhibit canonical autophagic pathways in P. falciparum [42]. Instead, the vacuole formation may result from nutrient stress due to disrupted haemoglobin digestion, a process heavily reliant on aspartyl and cysteine proteases. Inhibiting these enzymes likely reduces amino acid availability, impairing the parasite’s ability to sustain growth and potentially leading to death via starvation-related mechanisms, rather than via classical autophagy inhibition.

The next defect that was found in pepstatin A- treated parasite is a defect in nucleus. Previous studies demonstrated the parasite treated with pepstatin A exhibited pyknotic nuclei. This evidence supports the observed chromatin condensation. The chromatin condensation observed in pepstatin A-treated parasites, evident as dark blue staining along the inner nuclear membrane, may resemble early apoptotic-like morphology [43]. However, the presence and significance of classical apoptosis in P. falciparum remain controversial and poorly defined. Therefore, while the observed phenotype is consistent with apoptosis-like features, additional markers would be required to confirm cell death via this pathway.

The malaria pigment haemozoin originated from the haem detoxification process [44], transforming toxic haem, a byproduct of haemoglobin breakdown, into the non-toxic crystallized form of β-haematin or haemozoin [45–47]. The erythrocytic stage of the parasite could consume up to 80% of host haemoglobin [48], indicating substantial haem synthesis. Fitch’s findings suggested that a mere 0.1% of haem derived from erythrocyte haemoglobin could lyse the parasite in 10 min [49]. Free-form haem in a ferrous state caused membrane damage, block proteases, and led to parasite mortality [50]. Thus, the parasite needed the detoxification process to get rid of haem. Unfortunately, the parasite lacked haem oxygenase activity, which was the haem degradation enzyme found in the vertebrates [51]. However, the parasite could utilize another detoxification process called haemozoin synthesis [52]. In this study, asexual blood-stage and early gametocytes exhibited haemozoin clumping after pepstatin A treatment. This defect might relate to the haemozoin synthesis, as aspartyl protease contributes to 60–80% of the globin degradation process in the purified digestive vacuole [47, 53]. Additionally, plasmepsin II, histo aspartic protease or plasmepsin III and plasmepsin IV played a role in haemozoin synthesis by forming the protein complex with falcipain 2 and Haem Detoxification Protein (HDP) [53, 54]. Apart from the defect in haemozoin synthesis due to aspartyl protease inhibition, haemozoin clumping could be a result from a process known as haemozoin depolymerization, as demonstrated in 1997 studied by Pandey and colleagues. They showed that chloroquine could depolymerize purified haemozoin to haem, similarly representing the defect in haemozoin clumping [55]. Therefore, further study of the haemozoin clumping process was necessary, as the mechanism generating this defect remained unclear [52, 53, 56].

Some parasite populations grew into gametocytes, which further formed five stages of development in Plasmodium falciparum. The early-stage gametocyte was defined as the stage I through stage III gametocyte, while late-stage gametocytes were those that progressed from stage IV to mature stage V. Only mature stage V gametocytes could then egress from the host RBC and produced the gamete, which naturally occurred within the mosquito. This study depicted the dual effect of aspartyl protease inhibition by pepstatin A, mainly interfering with the asexual blood-stage and early gametocyte development. This suggested that aspartyl protease inhibition by pepstatin A could minimize the severity of disease caused by asexual stage development as well as reducing the chance of disease transmission to mosquitoes.

Conclusion

Although pepstatin A did not completely inhibit both asexual blood stage and gametocyte, it moderately affected the growth of asexual blood-stage and early-stage gametocytes development. Moreover, this study provided insights into the morphological changes of P. falciparum after aspartyl protease inhibition by pepstatin A. It exhibited chromatin condensation, vacuolization and haemozoin clumping in both asexual blood-stage and early-stage gametocyte, which are related to defects in the haemoglobin degradation process. Given its ability for aspartyl protease inhibition, it becomes a valuable target for an antimalarial drug. However, further studies on drug development or combination therapy are needed to increase the efficiency of aspartyl protease inhibition.

Abbreviations

ATS

Artesunate

CQ

Chloroquine

DMSO

Dimethyl sulfoxide

EGDI

Early-stage gametocyte development inhibition

HCT

Haematocrit

HDP

Haem detoxification protein

IC₅₀

The half-maximal inhibitory concentration

LGDI

Late-stage gametocyte development inhibition

RBC

Red blood cell

Author contributions

GN has designed the work, performed the experiments, interpreted data, drafted the manuscript, and revised the manuscript. CP is the colleague that also performed all experiments. RJ, WR, and JS have made contributions to the conception and advice on the experiments. VP and NK have major contributions to conceptualization, experimental design, data analysis, conclusion, and revised the manuscript. NK has approved the submitted version. All authors have contributed to the approval of the submitted version of manuscript.

Funding

Open access funding provided by Mahidol University. This research work was supported by Mahidol University (Fundamental Fund: fiscal year 2023 by National Science Research and Innovation Fund (NSRF)), and the CIF and CNI Grant, Faculty of Science, Mahidol University.

Data availability

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.