Skip to main content

Wild Egyptian medicinal plants show in vitro and in vivo cytotoxicity and antimalarial activities



Medicinal plants have been successfully used as an alternative source of drugs for the treatment of microbial diseases. Finding a novel treatment for malaria is still challenging, and various extracts from different wild desert plants have been reported to have multiple medicinal uses for human public health, this study evaluated the antimalarial efficacy of several Egyptian plant extracts.


We assessed the cytotoxic potential of 13 plant extracts and their abilities to inhibit the in vitro growth of Plasmodium falciparum (3D7), and to treat infection with non-lethal Plasmodium yoelii 17XNL in an in vivo malaria model in BALB/c mice.


In vitro screening identified four promising candidates, Trichodesma africanum, Artemisia judaica, Cleome droserifolia, and Vachellia tortilis, with weak-to-moderate activity against P. falciparum erythrocytic blood stages with mean half-maximal inhibitory concentration 50 (IC50) of 11.7 μg/ml, 20.0 μg/ml, 32.1 μg/ml, and 40.0 μg/ml, respectively. Their selectivity index values were 35.2, 15.8, 11.5, and 13.8, respectively. Among these four candidates, T. africanum crude extract exhibited the highest parasite suppression in a murine malaria model against P. yoelii.


Our study identified novel natural antimalarial agents of plant origin that have potential for development into therapeutics for treating malaria.

Peer Review reports


Malaria is caused by parasites belonging to the phylum Apicomplexa, genus Plasmodium. In 2019, approximately 229 million cases of malaria and 409,000 associated deaths were reported across 87 malaria-endemic countries [1]. Human malaria cases are caused by four different Plasmodium species—Plasmodium ovale, Plasmodium vivax, Plasmodium malariae, Plasmodium falciparum—of which P. falciparum is considered to be the most lethal [2, 3]. Plasmodium parasites are typically transmitted by the bite of an infected female Anopheles mosquito, although malaria can also be transmitted through exposure to blood products from an infected individual (transfusion malaria) or congenitally [4]. Multidrug-resistant Plasmodium parasites are the biggest challenge to health care in most malaria-endemic areas. Thus, research to develop new antimalarial drugs is critical [5].

The two main malaria species that are responsible for most human malaria cases, P. falciparum and P. vivax, have developed resistance against chloroquine. This drug resistance first emerged in the late 1950s in both Colombia and at the Cambodia–Thailand border and may stem from the great success of chloroquine and its multiple large-scale usages over the decades [6]. Great efforts have been made to develop the novel active agent, artemisinin, as an alternative drug to chloroquine. However, presently, there is no single drug effective for treating multi-drug resistant malaria, and effective combination treatment includes artemisinin derivatives, such as artesunate, or mixtures with previously developed drugs, such as an atovaquone-proguanil combination [7].

From ancient times, medical plants have been used for various pharmacological purposes because they contain many useful biological compounds [8]. More than 1277 plant species have been traditionally used for the treatment of malaria [9, 10]. Natural products still have an effective role in disease treatment. Finding anti-parasitic compounds produced by natural products, especially traditional medicinal plants from Asia, Africa, or the Americas, which have been reported as being successfully used to treat many diseases, could be an initial step toward controlling an array of diseases [11]. Currently, the WHO recommends widespread use of the RTS,S/AS01 (RTS,S) malaria vaccine among children at risk in sub-Saharan Africa [12]. Egypt has multiple aromatic and medicinal plants owing to its favorable geographical position, climate, and soil condition; thus, it is a useful site for exploring herbal and medicinal plants [13]. Previous studies illustrated the use of plant extracts in inhibition of P. falciparum in vitro [14,15,16,17] and in vivo using various doses of Ficus platyphylla plant extract ranging from 100 to 300 mg/kg/day against Plasmodium berghei infection [18]. In addition, other studies evaluated the effect of plant extracts by oral treatment in a murine model by chemotherapeutic test against different murine Plasmodium species in BALB/c mice [19,20,21]. Furthermore, the combination of the plant extracts with the reference drug artemisinin in treatment of Plasmodium yoelii was also reported [22]. Other reported study evaluated febrifugine and isofebrifugine mixture prepared from the dried leaves of H. macrophylla var. Otaksa against three rodent Plasmodia species; P. yoelii 17XL, P. berghei NK65, and P. chabaudi AS in Institute of Cancer Research (ICR) mice [23]. The previous reported experimental models support the use of our in vitro assay and in vivo model in treatment of P. falciparum in vitro and P. yoelii-infected mice in murine malaria model. Therefore, the present study aimed to evaluate the effectiveness of extracts from Egyptian medicinal plants randomly selected from the desert roads against human malaria, first via an in vitro assay and then with a murine malaria model.


Ethical statement

This study was performed in strict accordance with the recommendations of the Guide for the Care and Use of Laboratory Animals of the Ministry of Education, Culture, Sports, Science and Technology, Japan. The protocol was approved by the Committee on the Ethics of Animal Experiments at Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Japan (permit numbers 19–185, 20–157, 21–32). Mouse work, such as injection with parasites or extracts, and euthanasia was implemented under general inhalation anesthesia induced with isoflurane (2%) to minimize animal suffering. Mice were euthanized by cervical dislocation at 30 days after parasite infection.

P. Falciparum culture and maintenance

Plasmodium falciparum parasites were transferred to previously washed human O+ red blood cells (RBCs) obtained from Hokkaido Red Cross Blood Center maintained in fresh complete RPMI-1640 medium (Sigma, St Louis, MO, USA), which was supplemented with a mixture of 6 g of HEPES (Sigma), 2 g of NaHCO3, 25 mg of hypoxanthine, 5 g of albumax II (Gibco, Carlsbad, CA, USA), and 250 μl of gentamicin (stock concentration, 50 mg/ml) in dissolved in MilliQ water. The final prepared complete medium was filtered with a 0.20-μm membrane filter (IWAKI, Saitama, Japan). The parasite cultures were maintained at 37 °C in a 5% CO2 atmosphere.

Plant material collection and extraction

The plants used in this study were obtained from a field survey conducted in two locations in Qena governorate (Latitude: 26° 09′ 51.05“ N, Longitude: 32° 43’ 36.16” E), which is in the southern region of Egypt: Qena-Sohag and Qena-Safaga (after Km 85) desert roads, Eastern desert, Egypt. A map marking the collection sites is shown in Fig. S1. Plants collection sites coordinates were shown in Table S1. Material was collected from 13 different plant species in May 2019 (plant flowering season). Samples were collected between 4:00 AM and 12:00 PM. The plant samples were collected under the approval of South Valley University, Qena, Egypt and were microscopically identified by Dr. Mohamed Owis Badry, in the herbarium of South Valley University at Faculty of Science, South Valley University, Egypt and voucher specimens were deposited in the same herbarium. Identification was performed according to the available literature [24,25,26,27], and an official identification letter was obtained. Images of the herbarium sheets of the identified plant species from which samples were collected are shown in Fig. S2. Plant taxonomy and species were further updated in accordance with information from Plants of the World Online [28].

For plant collection from the study areas, although that there are no specific licenses were required for the field studies, permission for collection of plants was obtained from Faculty of Veterinary Medicine, South Valley University, Qena, Egypt. Collection was performed under the guidelines and rules of South Valley University, Qena. The surveyed locations were not protected or privately-owned in any way and the field studies did not include any protected or endangered Egyptian plant species. The Latin binomial names of all plant extracts studied in this study were shown in Table 1.

Table 1 Latin binomial name of all plant extracts used in this study

Plant samples were dried in the shade for 3–10 days, then a fine powder was obtained from the dried leaves, flowers, fruit, or seed parts by using a kitchen blender. The powdered plant material from each plant was dissolved at a 1:10 ratio in 80% methanol, 70% ethanol, or distilled water (100 g of plant powder/1 L of solvent) for a minimum of 1–3 days. The plant supernatant was further collected and filtrated by glass filtration apparatus and was collected in wide conical flask, and then it was dissolved in a wide petri dish at room temperature for 1–3 days. The final crude extract was collected in centrifuge tubes and stored in − 30 °C until use. To test the antimalarial potential of the various plant extracts, they were solubilized individually in the solvent dimethyl sulfoxide (DMSO) to prepare stock solutions (100 mg/ml).

Determination of cytotoxicity of plant extracts

To determine the cytotoxic potential of the plant extracts, their cytotoxicity against human foreskin fibroblast (HFF) cells was evaluated. Cell suspensions (1 × 105 cells/ml) in Dulbecco’s Modified Eagle medium (DMEM, Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% fetal bovine serum (FBS) (Nichirei Bioscience, Tokyo, Japan) were plated at 100 μl/well in 96-well plates and incubated at 37 °C in a 5% CO2 atmosphere for 48 h. The plant extracts were added to the cells at final concentrations of a two-fold serial dilution starting from 1000 μg/ml. To evaluate cell viability, cell proliferation inhibition (%) was calculated as described previously [29, 30].

In vitro anti-plasmodial activity

Plasmodium falciparum (3D7 strain) was maintained in O+ human erythrocytes (1% hematocrit) in complete RPMI medium (Sigma-Aldrich). P. falciparum was further synchronized to the ring stage with 5% sorbitol (>90%, as verified by light microscopy on Giemsa-stained blood smears [Giemsa stain for microscopy, Merck, Darmstadt, Germany]). Parasite solutions were prepared at 0.5% parasitemia and 2% hematocrit in complete RPMI medium. A 50-μl sample of the infected erythrocytes was added to each well of 96-well plates containing 50 μl of plant extract (concentrations ranging from 0.25–100 μg/ml). Medium only was used as a negative control, while chloroquine was used as a positive control. The plates were then incubated in an atmosphere of 5% CO2, 5% O2 at 37 °C for 72 h.

Parasite growth inhibition was determined by adding 100 μl of 0.02% of Syber Green I stain (SYBR® Green I Nucleic acid stain 10,000×, Lonza, Rockland, ME, USA) in lysis buffer (25 mM Tris, pH 7.5, containing 10 mM ethylenediamine tetraacetic acid, 0.01% saponin, and 0.1% Triton X-100) to each well, mixing gently, and incubating the plates for 1–2 h in the dark [31, 32]. The relative fluorescent inhibition values were determined by using a fluorescent plate reader Fluoroskan Ascent (Thermo Labsystems, Waltham, MA, USA) with excitation and emission wavelengths of 485 nm and 518 nm, respectively [29, 31, 33]. Parasite morphology was observed by examining Giemsa-stained blood smears with an all-in-one microscope BioRevo BZ-9000 (Keyence BioRevo, Tokyo, Japan). Parasite growth inhibition percentages were calculated as described previously [31, 33]. The antiplasmodial activities of the natural plant extracts used in this study were classified as follows: IC50 < 0.1 μg/ml: very good activity; IC50 between 0.1–1 μg/ml: good activity; IC50 between 1.1–10 μg/ml: good to moderate activity; IC50 between 11 and 50 μg/ml weak activity; IC50 > 100 μg/ml: inactive according to the classification mentioned in the reported study [34].

In vivo antimalarial efficacy of plant extracts

BALB/c mice, originally purchased from Clea Japan (Tokyo, Japan), were bred under specific pathogen-free conditions in the animal facility of the National Research Center for Protozoan Diseases at Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Japan. The animals were treated in accordance with the guiding principles for the care and use of research animals published by the Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Japan. The animals were kept under standard laboratory conditions on a 12/12-h light/dark cycle at 21 °C under 40% relative humidity and fed with commercial food and water ad libitum.

The non-lethal strain Plasmodium yoelii 17XNL was recovered from a stock of frozen parasitized RBCs (pRBC) via passage in donor mice intraperitoneally inoculated. Parasitemia was monitored daily. When the parasitemia level reached 20–30%, the donor mice were anesthetized, and blood was collected by cardiac puncture into a syringe containing 0.1 ml of ethylene diamine-N, N, N′, N′- tetraacetic acid disodium salt (EDTA) (Djindo Kumamoto, Japan).

Two male BALB/c mice aged 8–10 weeks and weighing 25–30 g, were infected with approximately 1 × 107P. yoelii-infected erythrocytes in total volume of 0.5 ml of phosphate-buffered saline (PBS). For each independent experiment, the mice were randomly divided into three groups of five according to previous published studies. AMA was aware of the group allocation at the different stages of the experiment. Total 17 mice were used for one trial. When the level of parasitemia reached 1%, oral treatment using 100 mg/kg/day of plant extracts was begun and continued for 1 week from day 0 (3 h post-challenge) until day 6 post-infection. The negative-control animals received only PBS. The parasitemia was assessed daily until 30 days post-infection by examining thin blood films made from mouse tail blood and stained with 10% Giemsa solution. The films were examined using a microscope to determine the parasite suppression activity of each extract. To measure the hematocrit percentage, 10 μl of blood was collected from the tail vein every other day until 30 days post-infection and measured by Celltac-α MEK-6550 (Nihon Kohden, Tokyo, Japan). A parasitemia suppression test (chemotherapeutic test) was performed daily for 1 week from challenge (day 0) until 6 days post-infection; the percentage of parasite growth suppression was calculated by using the following previously reported eq. [35], which is slightly modified from the study that originally reported it [36].

$$\%\ of\ parasite\ growth\ suppression=\left(A-B\right)/A\times 100$$

Where A is the mean parasitemia of the untreated group and B is parasitemia of each individual mouse in the treated groups.

The parasitemia percentage, bodyweight, and survival rates were monitored daily, and the hematocrit was monitored every other day. The percentage of parasitemia of each mouse was calculated by counting the number of parasite-infected erythrocytes per 600–1000 erythrocytes visible under a light microscope in 4–5 randomly selected fields of methanol-fixed thin blood smears slides stained with 10% Giemsa solution.


Order of treatment starts from the control then the treated groups. Order of challenge infection starts from the control then the treated groups. Measurements of body weight, hematocrit, and parasitemia were randomly done in group starting from control then the treated groups. Cage location was not changed from the start of the experiment until the end.

Statistical analysis

Graph Pad Prism 8.4.3 software (Graph Pad Software Inc. La Jolla, CA, USA) was used for all statistical tests. For the in vitro data, the IC50 values for the inhibition percentage of parasites and host cells were determined. The final mean IC50 of anti-P. falciparum (3D7) activity was calculated based on three independent experiments, and mean IC50 values against HFF cells were calculated based on three independent experiments. For the in vivo data (mean parasitemia %, mouse bodyweight and hematocrit changes), statistical analyses were performed using a two-way analysis of variance (ANOVA). Survival curves were generated with the Kaplan–Meier method, and survival rates were analyzed by a χ2 test. Statistically significant differences (those with a p-value of < 0.05) are marked in the figures by asterisks and defined in each figure legend. There were no any criteria used for including and excluding animals.


In vitro antimalarial efficacy of plant extracts

The in vitro activities against P. falciparum 3D7 growth of 13 different types of Egyptian plant extracts were evaluated. Among the 13 tested plant extracts, four (extracts of Trichodesma africanum, Artemisia judaica, Cleome droserifolia, and Vachellia tortilis) showed low-to-moderate activity against P. falciparum 3D7; their mean IC50 values were 11.7 μg/ml, 20.0 μg/ml, 32.1 μg/ml, and 40.0 μg/ml, respectively, and their mean selectivity index values were 35.2, 15.8, 11.5, and 13.8, respectively, (Table 2). Despite the low-to-moderate activities of the previously mentioned plant extracts, they possess good selectivity index values (Table 2). The ethanolic extract of Pulicaria undulata and both the ethanolic and methanolic extracts of Citrullus colocynthis showed weak activity against P. falciparum 3D7 growth (mean IC50 values: 18.9 μg/ml, 51.7 μg/ml, 45.9 μg/ml, respectively) and had mean selectivity index values of 2.9, 1.7, and 1.4, respectively (Table 2).

Table 2 Mean IC50 of Egyptian plant extracts against Plasmodium falciparum (3D7) and HFF cells in vitro

All extracts of Aerva javanica and Anabasis setifera, both the aqueous and methanolic extracts of Calotropis procera, Carthamus tinctorius, Forsskaolea tenacissima, Ochradenus baccatus, and Ocimum basilicum, and the methanolic extract of P. undulata showed no efficacy against the growth of P. falciparum (3D7) in vitro, with IC50 values of > 100 μg/ml (Table 2).

In vitro effect of plant extracts on P. falciparum growth stages and morphology

To confirm the in vitro antimalarial efficacy of the tested plant extracts, we observed thin blood smears from 72-h parasite culture (Fig. 1). Treatment with plant extracts for 72 h caused dose-dependent suppression of the parasite growth in the percentage of parasites (Fig. 1A). Among the four tested plant extracts, T. africanum crude extract showed the highest level of parasite growth inhibition at all parasite stages (Fig. 1A). Morphological alterations, such as cell shrinkage, and parasite fragmentation were observed following treatment with plant extract concentration of 50 μg/ml as well as after treatment with the positive control drug, chloroquine, in comparison with untreated parasites (Fig. 1B).

Fig. 1
figure 1

Effect of plant extracts on stage-specific P. falciparum (3D7) morphology in vitro. A Percentage of parasites at each stage (i.e., ring, trophozoite, or schizont) after treatment with 10, 25, 50, or 100 μg/ml of extracts of Artemisia judaica, Trichodesma africanum, Cleome droserifolia, or Vachellia toritilis plants, chloroquine, or medium alone. The number of parasites at each stage was determined from a total of 600–900 erythrocytes. Data are representative of two independent experiments with similar results. P. falciparum parasites were treated with 50 μg/ml of an extract of A. judaica, T. africanum, C. droserifolia, or V. tortilis. Chloroquine (0.025 μg/ml) was used as a positive control, and medium alone was used as a negative control. Three wells were used for each plant or drug concentration. After 72 h, the parasite morphology was observed via microscopy (× 100 magnification) on Giemsa-stained thin blood smears. Line arrow indicate the fragmented parasites, while arrow head indicate the shrinkage parasites. Data shown here are representative of two independent experiments that produced similar results

Cytotoxicity of plant extracts

The cytotoxic potential of all included plant extracts at concentrations ranging from 1000 to 7.8 μg/ml (two-fold serial dilutions) were determined, and the mean IC50 values against HFF cells were calculated (Table 2). The methanolic and aqueous extracts from C. procera, the ethanolic extract from P. undulata, both the methanolic and ethanolic extracts from C. colocynthis, the methanolic extract from P. undulata, and the ethanolic extract from O. basilicum showed the highest cytotoxicity against HFF cells with mean IC50s of 2.9, 41.5, 55.5, 65.6, 88.0, 197.5, and 252.6 μg/ml, respectively. Extracts from plants A. judaica, C. droserifolia, A. javanica, T. africanum, F. tenacissima, and V. tortilis showed moderate-to-weak toxicity against HFF cells with mean IC50s of 316.8, 370.9, 378.1, 413.0, 519.0, and 554.5 μg/ml, respectively. Lastly, Extracts from A. setifera and O. baccatus were nontoxic or safe for HFF cells as cytotoxicity was not observed and their mean IC50s were > 1000 μg/ml of 1263.6 μg/ml, and 1179.0 μg/ml, respectively (Table 2).

In vivo antimalarial activity of plant extracts

Their in-vitro results suggested that plant extracts from T. africanum, C. droserifolia, A. judaica, and V. tortilis have low or no cytotoxicity and might have activity against Plasmodium parasites. Therefore, we decided to evaluate their efficacy against P. yoelii in a murine malaria model. A chemotherapeutic test of each plant extract was performed beginning at the treatment start time of each extract (3 h post-challenge; day 0) and continuing through 6 days post-infection (the end of the course of the treatment). The four tested extracts each showed a time-dependent suppression of parasitemia through 7 days post-infection (Fig. 2). The mean level of parasite growth suppression observed after treatment with extracts of A. judaica, C. droserifolia, T. africanum, or V. tortilis ranged from 13.5–60.6%, 17.1–61.9, 35.2–65.5%, and 36.3–72.5%, respectively (Fig. 2, Table S2).

Fig. 2
figure 2

Suppression percentage of Plasmodium yoelii in mice induced by plant extracts at 1-week post-infection. Five mice were used per group. Artemisia judaica and Cleome droserifolia plant extracts were tested in one independent experiment that shared the same control, and Trichodesma africanum and Vachellia tortilis plant extracts were tested in another independent experiment that shared the same control. All mice were challenged by an intraperitoneal injection of approximately 1 × 107P. yoelii-infected erythrocytes and then were treated orally with 100 mg/kg/day of each plant extract for 1 week. The mean and standard deviation of the parasite growth inhibition percentages were calculated against the untreated group mice from 24 h after challenge (dpi = 1) until 7 days post-infection (24 h after end course of treatment) (A–D) Parasite growth inhibition percentages in mice treated with A. judaica (A), C. droserifolia (B), T. africanum (C), or V. tortilis (D) plant extract. N.D. not detected

In the murine malaria model, a significantly reduced level of parasitemia was observed from 6 to 15 days post-infection after oral treatment with T. africanum extract (Fig. 3C), whereas there was no significant difference in the hematocrit, bodyweight change, or survival rate of T. africanum extract-treated mice as compared with mice in the untreated group (Figs. S3C, S4C, and S5C). Parasite growth suppression by C. droserifolia extract was not observed, except at day 14 post-infection (Fig. 3B). The hematocrit, bodyweight change, and survival rate of C. droserifolia extract-treated mice did not show any significant difference compared with untreated animals (Figs. S3B, S4B, and S5B). Although some parasite suppression efficacy was observed for A. judaica extract following treatment initiation through 6 days post-infection (Table S2; Fig. 2A), the level of parasite suppression at the peak of parasitemia was not significant, and A. judaica extract-treated mice took a similar length of time to recover as compared with the untreated mice (Fig. 3A); furthermore, the hematocrit, bodyweight percentage, and survival rate of these mice were not different from those of the untreated animals (Figs. S3A, S4A, and S5A). The in vivo parasite suppression induced by V. tortilis extract was only partial, but it was significant during the peak of parasitemia at days 10, 11, and 12 post-infection (Table S2, Fig. 3D); however, the hematocrit, bodyweight change, and survival rate of V. tortilis extract-treated mice were not significantly different from those of the untreated mice (Figs. S3D, S4D, and S5D).

Fig. 3
figure 3

Effect of wild plant extracts on Plasmodium yoelii growth in mice through 30 days post-infection. Five mice were used in each group. Artemisia judaica and Cleome droserifolia plant extracts were tested in one independent experiment that shared the same control, and Trichodesma africanum and Vachellia tortilis plant extracts were tested in another independent experiment that shared the same control. Mean parasitemia % was monitored daily from day 0 (challenge day) until 30 days post-infection. All mice were challenged by an intraperitoneal injection of approximately 1 × 107Plasmodium yoelii-infected erythrocytes and then treated orally with 100 mg/kg/day of plant extract for 1 week. The untreated group received only PBS. A–D The mean parasitemia % of P. yoelii-infected mice treated with A. judaica (A), C. droserifolia (B), T. africanum (C), or V. tortilis (D) extract. Data were analyzed by a two-way ANOVA followed by a Bonferroni test against the untreated group (*p < 0.05)


Trichodesma africanum has been reported to have multiple medicinal uses (Table S1). The antibacterial efficacy of an oil extract of T. africanum was evaluated against the growth of three bacterial strains obtained from the American type culture collection (ATCC) (Staphylococcus aureus [ATCC 25923], Escherichia coli [ATCC 25922], and Pseudomonas aeruginosa [ATCC 27853]) as well as against the growth of methicillin-resistant S. aureus (MRSA) isolates, and its antifungal activity was tested against Candida albicans [37]. The chemical constituents of T. africanum include essential oils, steroids, coumarins, flavonoids, phenolics, alkaloids, and glycosides [38]. Its chemical constituents might be involved in its biological activity, as flavonoids had been reported to have antiprotozoal activity, specifically anti-leishmanial and anti-trypanosomal activities [39].

Trichodesma africanum collected from Saudi Arabia was reported to have weak antimalarial efficacy in vitro against a chloroquine-sensitive strain of P. falciparum with an IC50 value of 32.0 μg/ml (SI of 2) [40]. Here, T. africanum from an Egyptian desert was found to possess a moderate-to-weak activity in vitro against P. falciparum with a mean IC50 value of 11.7 μg/ml (SI of 35.2; Table 2). These results suggest that the efficacy of plant extracts can vary owing to differences in the area, time of day, and season of plant collection, extraction procedures used in collection, and cell lines used for the determination of its cytotoxic potential. Although the antimalarial activity of T. africanum was previously examined in vitro, our study is the first to illustrate the antimalarial efficacy of a T. africanum extract in a mouse model of malaria.

Cleome droserifolia, which is facing extinction, is found in tropical and subtropical areas, such as North Africa and India [24, 41, 42]. C. droserifolia is an important plant species owing to its historical use in traditional medicine in Egypt [43,44,45]. Regarding its medicinal uses, it has an immediate effect on abdominal and rheumatic pain, is anti-inflammatory, and is also effective for improving wound healing and treating snake bites and scorpion stings [43,44,45,46]. These effects are attributed to the rubefacient, antimicrobial, analgesic, antipyretic, antioxidant, and anti-inflammatory activities of its components [45,46,47,48,49], which include flavonoids, glycosides, carbohydrates, buchariol, teucladiol, daucosterol, cardenolides, saponins, sterols, tannins, catechins, triterpenes and sesquiterpenes as well as a newly described alkaloid found in its aerial parts [50,51,52,53]. C. droserifolia was previously reported to have an antibacterial effect [54], but little is known about its antiprotozoal efficacy. Although there is no literature on the effect of C. droserifolia extract on malaria, another species in the Cleome genus (e.g., Cleome rutidosperma) was reported to have moderate anti-plasmodial activity against P. falciparum CQS D10 strain in vitro (IC50 value: 34.4 μg/ml) [55]. In the present study, C. droserifolia was evaluated with both in vitro and in vivo assays. It had a moderate-to-weak inhibitory effect, with an IC50 value of 32.1 μg/ml and an SI of 12.9 in vitro against P. falciparum, and partially inhibited murine malaria in a short-term treatment of 100 mg/kg/day.

Recently, A. tortilis plant name was changed and become well known as V. tortilis according to [28], therefore, all information about V. tortilis in this discussion section is reported under its old name, A. tortilis. This plant is one of 1200 species of Acacia. It grows in tropical and subtropical areas with temperatures in the range of 40–45 °C in summer and < 5 °C in winter, such as locations in African, Australian, and Arabian countries [56]. A. tortilis possesses multiple medicinal and pharmacological properties, e.g., antidiabetic, antifungal, antidiarrheal, antitussive, and anti-inflammatory [57], but little is known about its antiprotozoal potential. Despite of the potent effectiveness of the methanolic extract of A. tortilis collected from Kenya as an anti-parasitic treatment, there are conflicting reports regarding its antimalarial activity. Some studies found that it had moderate efficacy against P. falciparum [57, 58]. In contrast, although the first in a pair of studies initially showed that A. tortilis root bark showed antimalarial activity, its follow-up study did not consider it to be an antimalarial candidate [57, 59]. In the present work, the methanolic extract from A. tortilis seeds collected from an Egyptian desert showed moderate-to-weak activity in vitro against P. falciparum (3D7). These results suggest that the efficacy of V. tortilis extract against malaria may be correlated to its medicinal uses (i.e., as an antimicrobial or other treatment) or to its chemical constituents and may vary depending on the extracted plant part and area from which the plant was collected. In vivo parasite suppression against P. yoelii in mice was observed until 6 days post-infection, and this extract significantly suppressed the parasitemia during its peak from days 10–12 post-infection, suggesting that it has partial efficacy against murine malaria.

Artemisia judaica belongs to the family Asteraceae, which is one of the largest families of angiosperms and contains 1600 to 1700 genera and about 24,000 species distributed worldwide [60]. Known as shih in the Middle-East, A. judaica is an aromatic shrub found mainly in the deserts of the Middle-East, Egypt, and serval North African countries and is traditionally used as an anthelmintic drug [61]. Although A. judaica has not been previously reported to have antimalarial efficacy, numerous other Artemisia species have been found to have antimalarial activity, including Artemisia nilagirica [62], Artemisia maciverae (chloroform extract) [63], Artemisia maritima (ethanolic and petroleum extracts), Artemisia nilegarica, Artemisia japonica [64], Artemisia ciniformis, Artemisia biennis, and Artemisia turanica [65]. Artemisinin, the well-known conventional drug discovered by Chinese scientists obtained from Artemisia annua [66], has also been found in several other species of Artemisia, including Artemisia lancea, Artemisia apiacea [67], Artemisia vulgaris [68], A. japonica [69], Artemisia sieberi [70], Artemisia absinthium [71], Artemisia dubia, and Artemisia indica [72]. There is no available information about the presence of artemisinin in A. judaica.

Plasmodium often develops drug resistance, and there is now evidence of resistance to artemisinin drugs [73]. Therefore, there is an urgent need to find novel candidates for the development of drugs to treat Plasmodium; other Artemisia plants, such as A. judaica, and other plant species may be useful sources. Several Artemisia species have been evaluated for their antimalarial activity in rodent malaria models. A. vulgaris showed potent activity without toxicity when administrated orally to mice infected with P. yoelii, according to the results of a 4-day suppressive test performed following treatment with high doses (500 mg/kg and 1000 mg/kg) of A. vulgaris extract [74]. Furthermore, the efficacy of an ethanolic extract of A. vulgaris leaves was confirmed against Plasmodium berghei ANKA strain in ICR mice. Treatment with doses of 500, 750, and 1000 mg/kg of this extract significantly reduced parasitemia by 79.3, 79.6, and 87.3%, respectively [75]. A. sieberi from Iran showed antimalarial efficacy against P. berghei in NMRI mice, reducing some pathophysiological signs of malaria [76]. An infusion of A. annua (tea) failed to cause any reduction to the parasitemia caused by Plasmodium chabaudi in OF1 mice [77], whereas the oral administration of dried whole A. annua leaves killed these parasites more effectively than did a comparable dose of the pure drug artemisinin in C57BL/6 mice [78].

Here, a methanolic leaf extract of A. judaica was found to possess moderate-to-weak antimalarial activity against P. falciparum with no apparent cytotoxicity along with a moderate efficacy against murine malaria at a lower dose (100 mg/kg/day) than used in previous studies. However, it did not cause a significant reduction in the parasitemia of P. yoelii during the peak of infection in mice.

Calotropis procera has been reported to have multiple biological and medicinal uses [79]. An ethanolic extract of this plant was reported to have a schizonticidal effect in vitro [80]. Furthermore, fractions from the leaf extract show anti-plasmodial activity [81]. However, in our study, neither the methanolic nor the aqueous extract of C. procera flowers showed any efficacy in vitro against P. falciparum (3D7) when administered at a dose of 100 μg/ml.

Three rodent-specific Plasmodium species, P. berghei, P. yoelii, and P. chabaudi, are commonly used in animal models of malaria; these models exhibit different manifestations of the human disease. In vitro cultures of these parasites are not well established; thus, they require maintenance in mice [82]. Here, we used a P. yoelii mouse model to evaluate the antimalarial efficacy of four plant extracts. In previous plant extract treatment trials of P. yoelii in Swiss albino mice, treatment with aqueous or ethanolic extracts of Phyllanthus amarus at doses of 200, 400, 800, and 1600 mg/kg/day was performed until 6 days post-infection; respectively, the aqueous extract induced 56.0, 68.0, 77.9, and 81.2% parasite suppression, and the ethanolic extract induced 51.7, 67.9, 74.2, and 52.3% parasite suppression [83]. Another study used 1.25 g/kg of methanolic extract from Nigella sativa seeds. Parasite suppression of 84.6, 89.2, and 94% was observed at days 6, 7, and 8 post-infection with P. yoelii nigeriensis, respectively [84]. Furthermore, the efficacies of methanolic-chloroform (MC) and methanolic-aqueous (MA) extracts from Brucei mollis collected from India were evaluated against P. yoelii N-67 (chloroquine-resistant strain [CQR]); they had respective median effective doses 50 (ED50s) of 30 mg/kg/day and 72 mg/kg/day at 4 days post-infection and of 66 mg/kg/day and 79 mg/kg/day at 6 days post-infection [85]. In the present study, treatment with 100 mg/kg/day methanolic extracts of A. judaica, C. droserifolia, T. africanum, or V. tortilis from 0 to 6 days post-infection each caused significant parasite suppression, with mean suppression percentages ranges of 13.5–60.6%, 17.1–61.9, 35.2–65.5%, and 36.3–72.5%, respectively, despite the extract dose being lower compared with previously reported studies (Table S2).


This study showed that crude extracts of four wild plants collected from Egypt had antimalarial efficacy against the human malaria-causing parasite P. falciparum in vitro and against the murine malaria-causing parasite P. yoelii in a mouse model. Although the administration of these extracts at a dose of 100 mg/kg/day for a 7-day course of treatment did not achieve 100% inhibition of P. yoelii growth in BALB/c mice, the parasite suppression data suggests that these extracts may have potent antimalarial activity. Their efficacies are likely correlated with their multiple medicinal uses and their chemical constituents. Among the four tested candidates, the T. africanum crude extract possessed the highest parasite suppression ability in a short-term treatment course in vivo and had the highest IC50 in vitro against the human malaria-causing parasite P. falciparum, whereas V. tortilis extract showed moderate-to-weak effect against P. falciparum in vitro and induced partial inhibition against P. yoelii in vivo. These data support the use of these extracts in the future development of an antimalarial therapeutic. Further study will be needed to understand the mechanism of action and identify the main biological components of these crude extracts.

Availability of data and materials

All data generated or analysed during this study are included in this published manuscript and its supplementary information file.


P. falciparum:

Plasmodium falciparum

P. yoelii :

Plasmodium yoelii



IC50 :

Half maximal inhibitory concentration 50


days post infection


Human foreskin fibroblast


American type culture collection


Selectivity index


Methicillin-resistant staphylococcus aureus


Methanol 80%


Ethanol 70%




Phosphate-buffered saline




Chloroquine resistant





ED50 :

median effective dose 50


  1. World Health Organization (WHO). World Malaria Report 2020 Medicines for Malaria Venture. 2020.

    Google Scholar 

  2. White NJ, Pukrittayakamee S, Hien TT, Faiz MA, Mokuolu OA, Dondorp AM. Malaria. Lancet. 2014;383:723–35.

    Article  PubMed  Google Scholar 

  3. Garrido-Cardenas JA, González-Cerón L, Manzano-Agugliaro F, Mesa-Valle C. Plasmodium genomics: an approach for learning about and ending human malaria. Parasitol Res. 2019;118:1–27.

    Article  PubMed  Google Scholar 

  4. Trampuz A, Jereb M, Muzlovic I, Prabhu RM. Clinical review: severe malaria. Crit Care. 2003;7:1–9.

    Article  Google Scholar 

  5. Kremsner PG, Krishna S. Antimalarial combinations. Lancet. 2004;364:285–94.

    Article  CAS  PubMed  Google Scholar 

  6. Payne D. Spread of chloroquine resistance in plasmodium falciparum. Parasitol Today. 1987;3:241–6.

    Article  CAS  PubMed  Google Scholar 

  7. Taylor WR, White NJ. Antimalarial drug toxicity. Drug Saf. 2004;27:25–61.

    Article  CAS  PubMed  Google Scholar 

  8. Dias DA, Urban S, Roessner U. A historical overview of natural products in drug discovery. Metabolites. 2012;2:303–36.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  9. Willcox M, Benoit-Vical F, Fowler D, Bourdy G, Burford G, Giani S, et al. Do ethnobotanical and laboratory data predict clinical safety and efficacy of anti-malarial plants? Malar J. 2011;10:1–9.

    Article  Google Scholar 

  10. Rasoanaivo P, Wright CW, Willcox ML, Gilbert B. Whole plant extracts versus single compounds for the treatment of malaria: synergy and positive interactions. Malar J. 2011;10:1–2.

    Article  Google Scholar 

  11. Wink M. Medicinal plants: a source of anti-parasitic secondary metabolites. Molecules. 2012;17:12771–91.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. World Health Organization (WHO). 2021. Who-recommends-groundbreaking-malaria-vaccine-for-children-at-risk. Accessed 24 Feb 2022.

  13. Mostafa N, Singab A. Prospective of herbal medicine in Egypt. Med Chem (Los Angeles). 2018;8:116–7.

    Article  Google Scholar 

  14. Hout S, Chea A, Bun SS, Elias R, Gasquet M, Timon-David P, et al. Screening of selected indigenous plants of Cambodia for antiplasmodial activity. J Ethnopharmacol. 2006;107:12–8.

    Article  PubMed  Google Scholar 

  15. Bagavan A, Rahuman AA, Kaushik NK, Sahal D. In vitro antimalarial activity of medicinal plant extracts against plasmodium falciparum. Parasitol Res. 2011;108:15–22.

    Article  PubMed  Google Scholar 

  16. Rufin Marie TK, Mbetyoumoun Mfouapon H, Madiesse Kemgne EA, Jiatsa Mbouna CD, Tsouh Fokou PV, Sahal D, et al. Anti-plasmodium falciparum activity of extracts from 10 Cameroonian medicinal plants. Medicines. 2018;5:115.

    Article  PubMed Central  CAS  Google Scholar 

  17. Kwansa-Bentum B, Agyeman K, Larbi-Akor J, Anyigba C, Appiah-Opong R. In Vitro Assessment of Antiplasmodial Activity and Cytotoxicity of Polyalthia longifolia Leaf Extracts on Plasmodium falciparum Strain NF54, Malaria Research and Treatment. 2019;2019:9. Article ID 6976298.

  18. Shittu I, Emmanuel A, Nok AJ. Antimalaria Effect of the Ethanolic Stem Bark Extracts of Ficus platyphylla Del. J Parasitology Res. 2011;2011:5. Article ID 618209.

  19. Tepongning RN, Yerbanga SR, Dori GU, Lucantoni L, Lupidi G, Habluetzel A. In vivo efficacy and toxicity studies on Erythrina senegalensis and Khaya ivorensis used as herbal remedies for malaria prevention in Cameroon. Eur J Med Plants. 2013;3:454–64.

    Article  Google Scholar 

  20. Chandel S, Bagai U, Vashishat N. Antiplasmodial activity of Xanthium strumarium against plasmodium berghei-infected BALB/c mice. Parasitol Res. 2012;110:1179–83.

    Article  PubMed  Google Scholar 

  21. Chutoam P, Klongthalay S, Somsak V. Effect of crude leaf extract of Bauhinia strychnifolia in BALB/c mice infected with plasmodium berghei. Malar Cont Elimination. 2015;4:S1–002.

    Google Scholar 

  22. Mohd Ridzuan MAR, Sow A, Noor Rain A, Mohd Ilham A, Zakiah I. Eurycoma longifolia extract-artemisinin combination: parasitemia suppression of plasmodium yoelii-infected mice. Trop Biomed. 2007;24:111–8.

    CAS  PubMed  Google Scholar 

  23. Ishih A, Miyase T, Ohori K, Terada M. Different responses of three rodent plasmodia species, plasmodium yoelii 17XL, P. berghei NK65 and P. chabaudi AS on treatment with febrifugine and isofebrifugine mixture from Hydrangea macrophylla var. Otaksa leaf in ICR mice. Phytother Res. 2003;17:650–6.

    Article  CAS  PubMed  Google Scholar 

  24. Boulos L. Flora of Egypt, Volume 1: Azollaceae - Oxalidaceae. Cairo, Egypt: Al Hadara Publishing; 1999. p. 419.

    Google Scholar 

  25. Boulos L. Flora of Egypt, volume 2: Geraniaceae–Boraginaceae. Cairo, Egypt: Al Hadara Publishing; 1999. p. 352.

    Google Scholar 

  26. Boulos L. Flora of Egypt, volume 3: Verbenaceae-Compositae. Cairo, Egypt: Al Hadara Publishing; 2002. p. 373.

    Google Scholar 

  27. Boulos L. Flora of Egypt checklist - revised Annotated Edition. Cairo, Egypt: Al Hadara Publishing; 2009. p. 410.

    Google Scholar 

  28. POWO. Plants of the world online. Facilitated by the Royal Botanic Gardens, Kew. 2019. Available from: Accessed June 2021.

  29. Leesombun A, Boonmasawai S, Nishikawa Y. Ethanol extracts from Thai plants have anti-plasmodium and anti-toxoplasma activities in vitro. Acta Parasitol. 2019;64:257–61.

    Article  CAS  PubMed  Google Scholar 

  30. Pagmadulam B, Tserendulam D, Rentsenkhand T, Igarashi M, Sawa R, Nihei CI, et al. Isolation and characterization of antiprotozoal compound-producing Streptomyces species from Mongolian soils. Parasitol Int. 2020;74:101961.

    Article  CAS  PubMed  Google Scholar 

  31. Leesombun A, Iijima M, Pagmadulam B, Orkhon B, Doi H, Issiki K, et al. Metacytofilin has potent anti-malarial activity. Parasitol Int. 2021;81:102267.

    Article  CAS  PubMed  Google Scholar 

  32. Johnson JD, Dennull RA, Gerena L, Lopez-Sanchez M, Roncal NE, Waters NC. Assessment and continued validation of the malaria SYBR green I-based fluorescence assay for use in malaria drug screening. Antimicrob Agents Chemother. 2007;51:1926–33.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Ariefta NR, Koseki T, Nishikawa Y, Shiono Y. Spirocollequins a and B, new alkaloids featuring a spirocyclic isoindolinone core, from Colletotrichum boninense AM-12-2. Tetrahedron Lett. 2021;64:152736.

    Article  CAS  Google Scholar 

  34. Rasoanaivo P, Deharo E, Ratsimanga-Urverg S, Frappier F. Guidelines for the Non-clinical Evaluation of the Efficacy of Traditional Antimalarials. In: Willcox M, Rasoanaivo P, Bodeker G, editors. Traditional medicine plants and malaria. London: CRC Press LLC Boca Raton; 2004. p. 255–70.

    Google Scholar 

  35. Kweyamba PA, Zofou D, Efange N, Assob JC, Kitau J, Nyindo M. In vitro and in vivo studies on anti-malarial activity of Commiphora africana and Dichrostachys cinerea used by the Maasai in Arusha region, Tanzania. Malar J. 2019;18:119.

    Article  PubMed  PubMed Central  Google Scholar 

  36. Peters W. The four-day suppressive in vivo antimalarial test. Ann Trop Med Parasitol. 1975;69:155–71.

    Article  CAS  PubMed  Google Scholar 

  37. Jaradat NA, Zaid AN, Abuzant A, Shawahna R. Investigation the efficiency of various methods of volatile oil extraction from Trichodesma africanum and their impact on the antioxidant and antimicrobial activities. J Intercult Ethnopharmacol. 2016;5:250–6.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  38. El-Moaty, A. Active Constituents and antimicrobial activity of Trichodesma africanum (L.) R. Br. var. heterotrichum Bornm. & Kneuck. Egyptian J Agricultural Sciences. 2009;60(4):357-65.

  39. Tasdemir D, Kaiser M, Brun R, Yardley V, Schmidt TJ, Tosun F, et al. Antitrypanosomal and antileishmanial activities of flavonoids and their analogues: in vitro, in vivo, structure-activity relationship, and quantitative structure-activity relationship studies. Antimicrob Agents Chemother. 2006;50:1352–64.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. Abdel-Sattar E, Maes L, Salama MM. In vitro activities of plant extracts from Saudi Arabia against malaria, leishmaniasis, sleeping sickness and Chagas disease. Phytother Res. 2010;24:1322–8.

    Article  CAS  PubMed  Google Scholar 

  41. Batanouny KH, Aboutabl E, Shabana M, Soliman F. Wild medicinal plants in Egypt. Cairo: The Palm press; 1999.

    Google Scholar 

  42. Kamel WM, Abd El-Ghani MM, El-Bous M. Cleomaceae as a distinct family in the flora of Egypt. Afr J Plant Sci Biotechnol. 2010;4:11–6.

    Google Scholar 

  43. Aparadh VT, Mahamuni RJ, Karadge BA. Taxonomy and physiological studies in spider flower (Cleome species): a critical review. Plant Sci Feed. 2012;2:25–46.

    Google Scholar 

  44. Rahman MA, Mossa JS, Al-Said MS, Al-Yahya MA. Medicinal plant diversity in the flora of Saudi Arabia 1: a report on seven plant families. Fitoterapia. 2004;75:149–61.

    Article  PubMed  Google Scholar 

  45. Moustafa A, Sarah R, Qiqa S, Mansour S, Alotaibi M. Cleome droserifolia: An Egyptian natural heritage facing extinction. Asian J Plant Sci Res. 2019;9:14–21.

    CAS  Google Scholar 

  46. Sarhan WA, Azzazy HM, El-Sherbiny IM. Honey/chitosan nanofiber wound dressing enriched with Allium sativum and Cleome droserifolia: enhanced antimicrobial and wound healing activity. ACS Appl Mater Interfaces. 2016;8:6379–90.

    Article  CAS  PubMed  Google Scholar 

  47. El-Ghazali GE, Al-Khalifa KS, Saleem GA, Abdallah EM. Traditional medicinal plants indigenous to Al-Rass province, Saudi Arabia. J Med Plant Res. 2010;4:2680–3.

    Article  Google Scholar 

  48. Abd El-Gawad AM, El-Amier YA, Bonanomi G. Essential oil composition, antioxidant and allelopathic activities of Cleome droserifolia (Forssk). Delile Chem Biodiversity. 2018;15:e1800392.

    Article  CAS  Google Scholar 

  49. Panicker NG, Balhamar SO, Akhlaq S, Qureshi MM, Rehman NU, Al-Harrasi A, et al. Organic extracts from Cleome droserifolia exhibit effective caspase-dependent anticancer activity. BMC Complement Med Ther. 2020;20:1–3.

    Article  Google Scholar 

  50. Aboushoer MI, Fathy HM, Abdel-Kader MS, Goetz G, Omar AA. Terpenes and flavonoids from an Egyptian collection of Cleome droserifolia. Nat Prod Res. 2010;24:687–96.

    Article  CAS  PubMed  Google Scholar 

  51. Hussain J, Khan H, Ali L, Latif Khan A, Ur Rehman N, Jahangir S, et al. A new indole alkaloid from cleome droserifolia. Helv Chim Acta. 2015;98:719–23.

    Article  CAS  Google Scholar 

  52. Abdullah W, Elsayed WM, Abdelshafeek KA, Nazif NM, Singab AN. Chemical constituents and biological activities of Cleome genus: a brief review. Int J Pharmacogn Phytochem Res. 2016;8:777–87.

    Google Scholar 

  53. Singh H, Mishra A, Mishra AK. The chemistry and pharmacology of Cleome genus: a review. Biomed. 2018;101:37–48.

    CAS  Google Scholar 

  54. Muhaidat R, Al-Qudah MA, Samir O, Jacob JH, Hussein E, Al-Tarawneh IN, et al. Phytochemical investigation and in vitro antibacterial activity of essential oils from Cleome droserifolia (Forssk.) Delile and C. trinervia Fresen. (Cleomaceae). S Afr J Bot. 2015;99:21–8.

    Article  CAS  Google Scholar 

  55. Bose A, Smith PJ, Lategan CA, Gupta JK, Si S. Studies on in vitro antiplasmodial activity of Cleome rutidosperma. Acta Pol Pharm Drug Res. 2010;67:315–8.

    Google Scholar 

  56. Ibrahim AA, Aref IM. Host status of thirteen Acacia species to Meloidogyne javanica. J Nematol. 2000;32:609.

    CAS  PubMed  PubMed Central  Google Scholar 

  57. Yadav P, Kant R, Kothiyal P. A review on Acacia tortilis. Int J Pharm Phytopharm Res. 2013;3:93–6.

    Google Scholar 

  58. Kigondu EV, Rukunga GM, Keriko JM, Tonui WK, Gathirwa JW, Kirira PG, et al. Anti-parasitic activity and cytotoxicity of selected medicinal plants from Kenya. J Ethnopharmacol. 2009;123:504–9.

    Article  PubMed  Google Scholar 

  59. Nguta JM, Mbaria JM. Brine shrimp toxicity and antimalarial activity of some plants traditionally used in treatment of malaria in Msambweni district of Kenya. J Ethnopharmacol. 2013;148:988–92.

    Article  CAS  PubMed  Google Scholar 

  60. Hussain A, Hayat MQ, Sahreen S, Ain QU, Bokhari SA. Pharmacological promises of genus Artemisia (Asteraceae): a review. Proc Pakistan Acad Sci: B Life Environ Sci. 2017;54:265–87.

    Google Scholar 

  61. Wyk BEV, Wink M. Medicinal plants of the world: An illustrated scientific guide to important medicinal plants and their uses. Pretoria, South Africa: CABI, Briza Publications; 2004.

    Google Scholar 

  62. Panda S, Rout JR, Pati P, Ranjit M, Sahoo SL. Antimalarial activity of Artemisia nilagirica against Plasmodium falciparum. J Parasit Dis. 2018;42:22–27.

    Article  PubMed  Google Scholar 

  63. Ene AC, Atawodi SE, Ameh DA, Ndukwe GI, Kwanashie HO. Bioassay-guided fractionation and in vivo antiplasmodial effect of fractions of chloroform extract of Artemisia maciverae Linn. Acta Trop. 2009;112:288–94.

    Article  CAS  PubMed  Google Scholar 

  64. Valecha NE, Biswas S, Badoni V, Bhandari KS, Sati OP. Antimalarial activity of Artemisia japonica, Artemisia maritima and Artemisia nilegarica. Indian J Pharm. 1994;26:144.

    Google Scholar 

  65. Mojarrab M, Naderi R, Afshar FH. Screening of different extracts from Artemisia species for their potential antimalarial activity. Iran J Pharm Sci. 2015;14:603.

    Google Scholar 

  66. Covello PS. Making artemisinin. Phytochemistry. 2008;69:2881–5.

    Article  CAS  PubMed  Google Scholar 

  67. Qian GP, Yang YW, Ren QL. Determination of artemisinin in Artemisia annua L. by reversed phase HPLC. J Liq Chromatogr Relat Technol. 2005;28:705–12.

    Article  CAS  Google Scholar 

  68. Numonov S, Sharopov F, Salimov A, Sukhrobov P, Atolikshoeva S, Safarzoda R, et al. Assessment of artemisinin contents in selected Artemisia species from Tajikistan (Central Asia). Medicines. 2019;6:23.

    Article  CAS  PubMed Central  Google Scholar 

  69. Rashmi TR, Francis MS, Murali S. Determination of Artemisinin in selected Artemisia L. species by HPLC. Indo Am J Pharm. 2014;4:2637–44.

    Google Scholar 

  70. Arab HA, Rahbari S, Rassouli A, Moslemi MH, Khosravirad F. Determination of artemisinin in Artemisia sieberi and anticoccidial effects of the plant extract in broiler chickens. Trop Anim Health Prod. 2006;38:497–503.

    Article  CAS  PubMed  Google Scholar 

  71. Zia M, Mannan A, Chaudhary MF. Effect of growth regulators and amino acids on artemisinin production in the callus of Artemisia absinthium. Pak J Bot (Pakistan). 2007;39:799–805.

    Google Scholar 

  72. Mannan A, Shaheen N, Arshad W, Qureshi RA, Zia M, Mirza B. Hairy roots induction and artemisinin analysis in Artemisia dubia and Artemisia indica. Afr J Biotechnol. 2008;7(18):3288-92. 17 September, 2008. Available online at ISSN 1684–5315 © 2008 Academic Journals.

  73. Dondorp AM, Nosten F, Yi P, Das D, Phyo AP, Tarning J, et al. Artemisinin Resistance in Plasmodium falciparum Malaria. N Engl J Med. 2009;361:455–67.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  74. Kodippili K, Daya Ratnasooriya W, Premakumara S, Udagama PV. An investigation of the antimalarial activity of Artemisia vulgaris leaf extract in a rodent malaria model. Int J Green Pharm. 2011;5(4). P-ISSN0973-8258 E-ISSN - 1998-4103.

  75. Bamunuarachchi GS, Ratnasooriya WD, Premakumara S, Udagama PV. Antimalarial properties of Artemisia vulgaris L. ethanolic leaf extract in a plasmodium berghei murine malaria model. J Vector Borne Dis. 2013;50:278–84.

    PubMed  Google Scholar 

  76. Nahrevanian H, Sheykhkanlooye Milan B, Kazemi M, Hajhosseini R, Soleymani Mashhadi S, Nahrevanian S. Antimalarial effects of Iranian flora Artemisia sieberi on Plasmodium berghei in vivo in mice and phytochemistry analysis of its herbal extracts. Malaria research and treatment. Hindawi Publishing Corporation Malaria Research and Treatment. 2012;2012:8. Article ID 727032.

  77. Atemnkeng MA, Chimanuka B, Dejaegher B, Vander Heyden Y, Plaizier-Vercammen J. Evaluation of Artemisia annua infusion efficacy for the treatment of malaria in plasmodium chabaudi chabaudi infected mice. Exp Parasitol. 2009;122:344–8.

    Article  PubMed  Google Scholar 

  78. Elfawal MA, Towler MJ, Reich NG, Golenbock D, Weathers PJ, Rich SM. Dried whole plant Artemisia annua as an antimalarial therapy. PLoS One. 2012;7:e52746.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Meena AK, Yadav AK, Niranjan US, Singh B, Nagariya AK, Sharma K, et al. A review on Calotropis procera Linn and its ethnobotany, phytochemical, pharmacological profile. Drug Invent Today. 2010;2:185–90.

    Google Scholar 

  80. Sharma P, Sharma JD. In-vitro schizonticidal screening of Calotropis procera. Fitoterapia. 2000;71:77–9.

    Article  CAS  PubMed  Google Scholar 

  81. Mudi SY, Bukar A. Anti-plasmodia activity of leaf extracts of Calotropis procera Linn. Biokemistri. 2011;23(1). Available online at

  82. Huang BW, Pearman E, Kim CC. Mouse models of uncomplicated and fatal malaria. Bio Protoc. 2015;5(13):e1514.

  83. Ajala TO, Igwilo CI, Oreagba IA, Odeku OA. The antiplasmodial effect of the extracts and formulated capsules of Phyllanthus amarus on plasmodium yoelii infection in mice. Asian Pac J Trop. 2011;4:283–7.

    Article  Google Scholar 

  84. Okeola VO, Adaramoye OA, Nneji CM, Falade CO, Farombi EO, Ademowo OG. Antimalarial and antioxidant activities of methanolic extract of Nigella sativa seeds (black cumin) in mice infected with plasmodium yoelli nigeriensis. Parasitol. 2011;108:1507–12.

    Article  Google Scholar 

  85. Prakash A, Sharma SK, Mohapatra PK, Bhattacharjee K, Gogoi K, Gogoi P, et al. In vitro and in vivo antiplasmodial activity of the root extracts of Brucea mollis wall. Ex Kurz. Parasitol Res. 2013;112:637–4.

    Article  PubMed  Google Scholar 

Download references


We thank Drs., Arpron Leesombun, Mingming Liu, Nanang Rudianto Ariefta and Ms. Iqra Zafar Wahla (Obihiro University of Agriculture and Veterinary Medicine) for their excellent technical assistance. We are also thankful for Dean of Faculty of Veterinary Medicine and Dean of Faculty of Science, South Valley University, Qena, Egypt for granting the official permit for plant taxa collection and identification. We are grateful to the great efforts from the South Valley University Workers, South Valley University, Qena for their help in plant specimen’s collection from the desert roads, we are also grateful to Prof. Dr. Shin-ichiro Kawazu (Obihiro University of Agriculture and Veterinary Medicine) for providing P. falciparum (3D7) parasites. We thank the Hokkaido Red Cross Blood Center for supplying human red blood cells. We thank Katie Oakley, Ph.D., from Edanz ( for editing a draft of this manuscript.


This research was supported by the Research Program on Emerging and Re-emerging Infectious Diseases (20fk0108137h [YN]) from the Agency for Medical Research and Development (AMED) and fund for the Promotion of Joint International Research (Fostering Joint International Research(B))” from the Ministry of Education, Culture, Sports, Science and Technology KAKENHI (20KK0152 [YN]).

Author information

Authors and Affiliations



A.M.A and Y.N designed the project and experiments. A.M.A, A. Sh. S., N.A and M.O.B conducted the experiments. A.M.A performed the statistical analysis, A.M.A wrote the manuscript, Y.N revised the manuscript. All authors have reviewed and approved the final draft of the manuscript.

Corresponding author

Correspondence to Yoshifumi Nishikawa.

Ethics declarations

Ethics approval and consent to participate

This study was performed in strict accordance with the recommendations of the Guide for the Care and Use of Laboratory Animals of the Ministry of Education, Culture, Sports, Science and Technology, Japan. The protocol was approved by the Committee on the Ethics of Animal Experiments at Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Japan (permit numbers 19–185, 20–157, 21–32). Plasmodium parasites preparation on human RBCs, was maintained based on the ethical review of Obihiro University of Agriculture and Veterinary Medicine, Obihiro, Japan (permit number 2011-06-3). For plant collection, no specific licenses were required for field studies. Collection was performed according to the approval, guidelines, and rules of South Valley University, Qena, Egypt. The surveyed locations were not protected or privately-owned and did not include any protected or endangered Egyptian plant species.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1: Table S1.

The plants used in this study and their reported medicinal uses. Table S2. Chemotherapeutic test of four plant extracts against the growth of Plasmodium yoelii in mice. Figure S1. Sampling map of the plant samples collected in Egypt. Figure S2. Images of the collected plant materials. Figure S3. Effect of wild plant extracts on the growth of plasmodium yoelii in male BALB/c mice. Figure S4. Effect of wild plant extracts on bodyweight change in Plasmodium-infected mice. Figure S5. Effect of wild plant extracts on survival rate of Plasmodium-infected mice.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Abdou, A.M., Seddek, Al.S., Abdelmageed, N. et al. Wild Egyptian medicinal plants show in vitro and in vivo cytotoxicity and antimalarial activities. BMC Complement Med Ther 22, 130 (2022).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: