Anthelmintic properties of traditional African and Caribbean medicinal plants: identification of extracts with potent activity against Ascaris suum in vitro.

Ascariasis affects more than 1 billion people worldwide, mainly in developing countries, causing substantial morbidity. Current treatments for Ascaris infection are based on mass drug administration (MDA) with synthetic anthelmintic drugs such as albendazole, however continual re-infection and the threat of drug resistance mean that complementary treatment options would be highly valuable. Here, we screened ethanolic extracts from 29 medicinal plants used in Africa (Ghana) and the Caribbean (US Virgin Islands) for in vitro anthelmintic properties against Ascaris suum, a swine parasite that is very closely related to the human A. lumbricoides. A wide variety of activities were seen in the extracts, from negligible to potent. Extracts from Clausena anisata, Zanthoxylum zanthoxyloides and Punica granatum were identified as the most potent with EC50 values of 74, 97 and 164 μg/mL, respectively. Our results encourage further investigation of their use as complementary treatment options for ascariasis, alongside MDA. © A.R. Williams et al., published by EDP Sciences, 2016.

Introduction

Soil-transmitted helminths remain one of the largest burdens on global health. Altogether, Ascaris lumbricoides (roundworm), Trichuris trichiura (whipworm) and Necator americanus (hookworm) infect more than a billion people, mainly in the developing world [20, 24]. Of these, the most prevalent is A. lumbricoides, which was estimated to infect around 800 million people in 2010, resulting in more than a million years lived with disability (YLD) [20]. Ascariasis can result in malnutrition and inhibit cognitive development and learning in children, and can also interfere with effective expression of immunity to other pathogens and vaccines due to polarisation of the immune system towards a regulatory/Th2-skewed state [6, 21].

At present, A. lumbricoides infections are treated through mass drug administration (MDA) programmes, involving annual or bi-annual treatment of school children, mainly with albendazole [12]. However, the sustainability of this approach has been questioned due to continual re-infection, arising from the hard-shelled eggs which survive for many years in the environment, and the ever-present threat of drug resistance, where cautionary tales can be drawn from the critical levels of resistance that have arisen in helminths of veterinary importance due to mass administration of synthetic drugs [11, 13, 23].

The use of indigenous medicinal plants for control of internal parasites has been practised for centuries, however scientific validation of these traditional practices has been lacking [8, 22]. Thus, a potentially vast resource of natural plant compounds with anthelmintic activity has been underutilised. The use of natural plant extracts, which can be prepared by decoction or other simple procedures, has several advantages such as low cost and easy integration into local community practices, if and where plants are locally available. The rational use of such an approach may complement current MDA programmes and be a useful tool to slow the onset of drug resistance.

Screening for anthelmintic activity is often accomplished by using the free-living nematode Caenorhabditis elegans [5, 9] due to low cost and amenability of laboratory culture methods. However, many differences exist between free-living nematodes and parasites such as Ascaris, as well as between free-living and parasitic stages of the same species [17]. Thus, anthelmintic activity is ideally assessed using as close as possible a model to the eventual target. Ascaris suum is a swine parasite and closely related parasite to A. lumbricoides, being morphologically indistinguishable, and indeed, for many years the human and pig forms of Ascaris were considered to be a single species [16]. This means that A. suum is an excellent model for studying possible new interventions such as vaccines or new drug candidates against A. lumbricoides. In particular, in vitro studies with A. suum are easily performed due to the ability to generate large numbers of infective, parasitic third-stage larvae (L3) from embryonated eggs recovered from the uteri of adult female worms. We have previously used a combination of motility and migration inhibition assays to test for in vitro anthelmintic effects of a number of compounds against A. suum L3 [25]. Here, we utilised local ethno-medical knowledge to compile a library of traditional medicinal plants from Ghana and the Caribbean, and used A. suum L3 to screen > 30 extracts for in vitro activity. Our results indicate that some of these plants may have value as a complementary treatment option for ascariasis.

Materials and methods

Plant material and extraction

Medicinal plants were collected during the period November 2013 to January 2014 in the Greater Accra region of Ghana or October to November 2014 on the US Virgin Islands (Table 1). The plants were dried at ambient temperature in the shade. The plants were identified and authenticated by an ethnobotanist (Jens Soelberg, University of Copenhagen, Denmark). Botanical voucher specimens were deposited at the Herbaria at the University of Copenhagen, Denmark; and at the University of Ghana, Ghana, and St. George Village Botanical Garden, respectively. Voucher numbers are given in Table 1. Ethanolic extracts were prepared by extracting 2 g of ground plant material with 20 mL of 96% ethanol in an ultrasonic bath for 30 min. Extracts were filtered through filter paper and taken to dryness at room temperature under nitrogen. For anthelmintic assays, extracts were dissolved in 100% DMSO.

Table 1.

Names and characteristics of plant species tested for anthelmintic activity against Ascaris suum. Plants were collected from either Ghana (G) or the US Virgin Islands (VI).

Species name	Location	Family	Voucher	Plant part
Aloe vera (L.) Burm. f.	VI	Xanthorrhoeaceae	JS 617	Leaves
Boerhavia diffusa L.	G	Nyctaginaceae	JS 281	Aerial parts
Boerhavia erecta L.	G	Nyctaginaceae	JS 282	Aerial parts
Clausena anisata (Willd.) Hook. f. ex Benth.	G	Rutaceae	JS 214	Roots
Deinbollia pinnata Schum. & Thonn.	G	Sapindaceae	JS 202	Aerial parts
Erythrina senegalensis DC.	G	Leguminosae	JS 231	Bark
Flacourtia flavescens Willd.	G	Salicaceae	JS 249	Leaves
Flueggea virosa (Roxb. ex Willd.) Royle	G	Phyllanthaceae	JS 252	Leaves
Gardenia ternifolia Schumach. & Thonn.	G	Rubiaceae	JS 246	Leaves
Gymnanthemum coloratum (Willd.) H. Rob. & B. Kahn	G	Asteraceae	JS 268	Leaves/flowers
Gymnanthemum coloratum (Willd.) H. Rob. & B. Kahn	G	Compositae	JS 268	Roots
Launaea taraxacifolia (Willd.) Amin ex C. Jeffrey	G	Compositae	JS 212	Leaves
Mallotus oppositifolius (Geiseler) Müll. Arg.	G	Euphorbiaceae	JS 208	Leaves
Mucuna pruriens (L.) DC.	VI	Leguminosae	JS 656	Fruit
Newbouldia laevis (P. Beauv.) Seem.	G	Bignoniaceae	JS 216	Leaves
Opuntia sp.	VI	Cactaceae	JS 672	Stem
Paullinia africana R.Br. ex Tedlie	G	Sapindaceae	JS 219	Aerial parts
Phyllanthus amarus Schumach. & Thonn.	G	Phyllanthaceae	JS 237	Aerial parts
Premna quadrifolia Schumach. & Thonn.	G	Lamiaceae	JS 283	Aerial parts
Psidium guajava L.	VI	Myrtaceae	JS 623	Leaves
Punica granatum L.	VI	Lythraceae	JS 615	Fruit peel
Pupalia lappacea (L.) Juss.	G	Amaranthaceae	JS 239	Aerial parts
Rivina humilis L.	VI	Phytolaccaceae	JS 608	Aerial parts
Senna occidentalis (L.) Link	G	Leguminosae	JS 234	Aerial parts
Senna occidentalis (L.) Link	G	Leguminosae	JS 234	Roots
Spathodea campanulata P. Beauv.	G	Bignoniaceae	JS 230	Bark
Spigelia anthelmia L.	VI	Loganiaceae	JS 651	Aerial parts
Stylosanthes erecta P. Beauv.	G	Leguminosae	JS 271	Aerial parts
Thonningia sanguinea Vahl	G	Balanophora	JS 296	Aerial parts
Triumfetta semitriloba Jacq.	VI	Malvaceae	JS 658	Aerial parts
Zanthoxylum zanthoxyloides (Lam.) Zepern. & Timler	G	Rutaceae	JS 243	Roots
Zanthoxylum zanthoxyloides (Lam.) Zepern. & Timler	G	Rutaceae	JS 243	Root bark

Parasite material

Gravid Ascaris suum worms were collected from fresh pig intestines at a local slaughterhouse (Danish Crown, Ringsted, Denmark). Eggs were isolated from the uteri of the worms and then subsequently embryonated at 25 °C for at least 60 days in 0.1 M H2SO4. Full embryonation was confirmed by light microscopy. To obtain the L3, eggs were washed, suspended in Hanks’ Buffered Salt Solution (HBSS) and then hatched by stirring together with 2 mm glass beads for 30 min at 37 °C. Viable L3 were then separated from unhatched eggs and debris by overnight migration into sterile HBSS using a Baermann apparatus equipped with 20 μm mesh. The L3 were then washed, counted and suspended in larval culture media (RPMI 1640 supplemented with 2 mM L-glutamine, 100 U/mL penicillin and 100 μg/mL streptomycin) for use in the migration inhibition assay.

Larval migration inhibition assay

The migration inhibition assay was conducted essentially as described previously [25]. Briefly, 100 L3 (in triplicate) were added to wells on a tissue culture plate and plant extracts in DMSO added (DMSO concentration never exceeded 1%). All assays included 1% DMSO in culture media as a negative control and 50 μg/mL ivermectin (Sigma-Aldrich) as a positive control. The plates were then incubated overnight at 37 °C in 5% CO2 in air. Then, an equal amount of 1.6% agar solution (45 °C) was added to each well and mixed thoroughly. The agar was allowed to solidify, before fresh culture media were added on top to cover the agar and the plates returned to the incubator overnight. The next day, the media were collected from each well and the number of larvae that had migrated from the setting agar was enumerated by light microscopy.

Data analysis and statistics

Migration inhibition was calculated relative to larvae incubated only in culture media (+1% DMSO) and expressed as percentage inhibition. Half-maximal effective concentration (EC50) values were calculated using non-linear regression. Analyses were performed in GraphPad Prism (v6.00, GraphPad Software, La Jolla, California, USA, www.graphpad.com).

Results and discussion

Screening of the plant extracts at a concentration of 1 mg/mL revealed a wide variety of potencies (Fig. 1). A number of extracts had no or negligible activity. However, 10 extracts inhibited larval migration by at least 50%. An arbitrary cut-off value of 90% migration inhibition was chosen to define plants that had potent activity, comparable to the inhibition achieved by 50 μg/mL ivermectin (positive control). Using this criterion, four extracts – Clausena anisata, Zanthoxylum zanthoxyloides (both the roots and root bark) and Punica granatum – were selected on the basis of potent activities for dose-dependent studies to confirm their activities and determine EC50 values. All four extracts displayed dose-dependent activity (Fig. 2). The EC50 (95% CI) values for C. anisata, Z. zanthoxyloides roots, Z. zanthoxyloides root bark and P. granatum were 74 (63.3–86.8), 97 (63.9–149.3), 132 (105.9–164.6) and 164 (124.7–271.1) μg/mL, respectively.

Figure 1.

Inhibition of migratory ability of Ascaris suum third-stage larvae after exposure to extracts of medicinal plants (1 mg/mL) or ivermectin (50 μg/mL). Results are the mean (± SEM) of three replicates from a single experiment. The vertical dashed line indicates 90% inhibition of larval migration.

Figure 2.

Dose-dependent inhibition of migration of Ascaris suum third-stage larvae after exposure to extracts of three different medicinal plants. Results are the mean of three independent experiments, each performed in triplicate, with the results expressed as the mean ± inter-experiment SEM.

We have thus confirmed that a number of plants that are traditionally used in medicinal form in Ascaris-endemic regions have direct anthelmintic activity against A. suum. The identification of three extracts with particularly potent activity is consistent with previous reports concerning their ethno-medical usage and anthelmintic properties. C. anisata is used as an anthelmintic by traditional healers in Kenya [24], and has also been noted to have in vitro activity against free-living larvae of the sheep nematode Haemonchus contortus [23]. Similarly, in vitro studies with Z. zanthoxyloides have demonstrated anthelmintic effects of crude extracts from this plant against H. contortus and another sheep nematode, Trichostrongylus colubriformis [4, 10]. Moreover, the distilled essential oils from Z. zanthoxyloides have been shown to have in vitro activity against the rat helminth Strongyloides ratti [18]. Punica granatum extracts also have in vitro activity against free-living worms such as Allolobophora caliginosa [7] and veterinary helminths such as the poultry roundworm Ascaridia galli [3]. Our data, which indicate a strong anti-Ascaris effect of these three plant extracts, extend on these previous studies by suggesting that they may have potential for treatment of A. lumbricoides in humans.

The active compounds of these extracts were not investigated here, but all three extracts (Z. zanthoxyloides, P. granatum and C. anisata) are known to be rich in secondary compounds such as tannins (with P. granatum being a particularly rich source of ellagitannins and gallic acid), flavonoids, terpenes and alkaloids, and these have been implicated in the apparent anti-microbial and ethno-medical properties of these plants [1, 14, 19]. Previous in vitro studies with H. contortus and S. ratti have suggested that the anthelmintic properties of Z. zanthoxyloides are partially dependent on flavonoids and related polymeric tannins, but that other bioactive compounds such as terpenes derived from the essential oil may contribute to the activity [4, 18]. Furthermore, a related plant, Z. liebmannianum, has also been shown to have activity against A. suum in vitro and here α-sanshool was isolated as a putative active compound [15]. Ellagitannins or related phenolic compounds found in P. granatum have been speculated to be responsible for activity that has been observed against protozoan parasites such as Cryptosporidium parvum [2]. Fractionation and identification of the active compounds in these extracts is ongoing in our laboratory.

The confirmation of anti-Ascaris activity of some of these traditional extracts encourages refinement of their use as treatments for roundworm infection in areas where these plants are found locally. Whilst our current experiments were performed with third-stage larvae, which are present in the intestinal tract for only a short period of time following egg ingestion, we have shown previously that activity of bioactive compounds against L3 correlates very closely with their activity against both fourth-stage larvae [25] and adult worms (A.R. Williams, unpublished data). Thus, there appears to be clear scope for these remedies to be used as therapeutic treatments against established infections in the small intestine. Having now confirmed the activity of these traditional extracts, further studies to investigate the pharmacokinetics in order to determine the best possible dosage and administration methods of these extracts are now warranted. Moreover, if activity can be solely or mainly ascribed to a single compound, then the possibility remains of synthesis and production of a new anthelmintic drug.

In conclusion, we have demonstrated that extracts of C. anisata, Z. zanthoxyloides and P. granatum have potent anthelmintic activity against A. suum in vitro, which encourages further investigation of their use as therapeutic agents against Ascaris infections in endemic regions.