Open Access
Research Article
Issue
Parasite
Volume 23, 2016
Article Number 24
Number of page(s) 6
DOI https://doi.org/10.1051/parasite/2016024
Published online 14 June 2016

© A.R. Williams et al., published by EDP Sciences, 2016

Licence Creative Commons
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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).

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.

thumbnail 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.

thumbnail 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.

Acknowledgments

The authors are grateful to Katrine Krydefelt for assistance with extraction of plant material. ARW was supported by the Danish Council for Independent Research (Technology and Production Sciences, Grant No. 12-126630.

References

  1. Agyepong N, Agyare C, Adarkwa-Yiadom M, Gbedema SY. 2014. Phytochemical investigation and anti-microbial activity of Clausena Anisata (Willd), Hook. African Journal of Traditional, Complementary, and Alternative Medicines, 11(3), 200–209. [CrossRef] [Google Scholar]
  2. Al-Mathal EM, Alsalem AA. 2013. Pomegranate (Punica granatum) peel is effective in a murine model of experimental Cryptosporidium parvum ultrastructural studies of the ileum. Experimental Parasitology, 134(4), 482–494. [CrossRef] [PubMed] [Google Scholar]
  3. Ali N, Jamil A, Shah S, Shah I, Ahmed G, Junaid M, Ahmed Z. 2015. Parasiticidal and brine shrimp cytotoxicity potential of crude methanolic extract of rind of Punica granatum Linn against round worms and tape worms. Pakistan Journal of Pharmacuetical Sciences, 28(3), 959–962. [Google Scholar]
  4. Azando EVB, Hounzangbé-Adoté MS, Olounladé PA, Brunet S, Fabre N, Valentin A, Hoste H. 2011. Involvement of tannins and flavonoids in the in vitro effects of Newbouldia laevis and Zanthoxylum zanthoxyloïdes extracts on the exsheathment of third-stage infective larvae of gastrointestinal nematodes. Veterinary Parasitology, 180(3–4), 292–297. [CrossRef] [PubMed] [Google Scholar]
  5. Buckingham SD, Partridge FA, Sattelle DB. 2014. Automated, high-throughput, motility analysis in Caenorhabditis elegans and parasitic nematodes: applications in the search for new anthelmintics. International Journal for Parasitology: Drugs and Drug Resistance, 4(3), 226–232. [CrossRef] [Google Scholar]
  6. Cooper PJ, Chico ME, Sandoval C, Espinel I, Guevara A, Kennedy MW, Urban JF, Griffin GE, Nutman TB. 2000. Human infection with Ascaris lumbricoides is associated with a polarized cytokine response. Journal of Infectious Diseases, 182(4), 1207–1213. [CrossRef] [Google Scholar]
  7. Dkhil MA. 2013. Anti-coccidial, anthelmintic and antioxidant activities of pomegranate (Punica granatum) peel extract. Parasitology Research, 112(7), 2639–2646. [CrossRef] [PubMed] [Google Scholar]
  8. Githiori JB, Höglund J, Waller PJ. 2005. Ethnoveterinary plant preparations as livestock dewormers: practices, popular beliefs, pitfalls and prospects for the future. Animal Health Research Reviews, 6(01), 91–103. [CrossRef] [Google Scholar]
  9. Holden-Dye L, Walker RJ. 2014. Anthelmintic drugs and nematicides: studies in Caenorhabditis elegans, in WormBook, Editor. C. elegans Research Community, WormBook, DOI: 10.1895/wormbook.1.143.2, http://www.wormbook.org [Google Scholar]
  10. Hounzangbe-Adote S, Fouraste I, Moutairou K, Hoste H. 2005. In vitro effects of four tropical plants on the activity and development of the parasitic nematode, Trichostrongylus colubriformis. Journal of Helminthology, 79(01), 29–33. [CrossRef] [PubMed] [Google Scholar]
  11. Jia T-W, Melville S, Utzinger J, King CH, Zhou X-N. 2012. Soil-transmitted helminth reinfection after drug treatment: a systematic review and meta-analysis. PLoS Neglected Tropical Diseases, 6(5), e1621. [CrossRef] [PubMed] [Google Scholar]
  12. Keiser J, Utzinger J. 2008. Efficacy of current drugs against soil-transmitted helminth infections: systematic review and meta-analysis. Journal of the American Medical Association, 299(16), 1937–1948. [PubMed] [Google Scholar]
  13. Kotze AC, Hunt PW, Skuce P, von Samson-Himmelstjerna G, Martin RJ, Sager H, Krücken J, Hodgkinson J, Lespine A, Jex AR, Gilleard JS, Beech RN, Wolstenholme AJ, Demeler J, Robertson AP, Charvet CL, Neveu C, Kaminsky R, Rufener L, Alberich M, Menez C, Prichard RK. 2014. Recent advances in candidate-gene and whole-genome approaches to the discovery of anthelmintic resistance markers and the description of drug/receptor interactions. International Journal for Parasitology: Drugs and Drug Resistance, 4(3), 164–184. [CrossRef] [Google Scholar]
  14. Mena P, Ascacio-Valdés JA, Gironés-Vilaplana A, Del Rio D, Moreno DA, García-Viguera C. 2014. Assessment of pomegranate wine lees as a valuable source for the recovery of (poly)phenolic compounds. Food Chemistry, 145, 327–334. [CrossRef] [PubMed] [Google Scholar]
  15. Navarrete A, Hong E. 1996. Anthelmintic properties of α-sanshool from Zanthoxylum liebmannianum. Planta Medica, 62(03), 250–251. [CrossRef] [PubMed] [Google Scholar]
  16. Nejsum P, Betson M, Bendall RP, Thamsborg SM, Stothard JR. 2012. Assessing the zoonotic potential of Ascaris suum and Trichuris suis: looking to the future from an analysis of the past. Journal of Helminthology, 86(02), 148–155. [CrossRef] [PubMed] [Google Scholar]
  17. O’Grady J, Kotze AC. 2004. Haemonchus contortus: in vitro drug screening assays with the adult life stage. Experimental Parasitology, 106(3–4), 164–172. [CrossRef] [PubMed] [Google Scholar]
  18. Olounladé PA, Azando EVB, Hounzangbé-Adoté MS, Ha TBT, Leroy E, Moulis C, Fabre N, Magnaval JF, Hoste H, Valentin A. 2011. In vitro anthelmintic activity of the essential oils of Zanthoxylum zanthoxyloides and Newbouldia laevis against Strongyloides ratti. Parasitology Research, 110(4), 1427–1433. [CrossRef] [PubMed] [Google Scholar]
  19. Patino L, Prieto RJA, Cuca SLE. 2012. Zanthoxylum Genus as a potential source of bioactive compounds, in Bioactive compounds in phytomedicine. Rasooli I, Editor Intech: Rijeka, Croatia. p. 185–218. [Google Scholar]
  20. Pullan R, Smith J, Jasrasaria R, Brooker S. 2014. Global numbers of infection and disease burden of soil transmitted helminth infections in 2010. Parasites & Vectors, 7(1), 37. [CrossRef] [PubMed] [Google Scholar]
  21. Steenhard NR, Jungersen G, Kokotovic B, Beshah E, Dawson HD, Urban JF Jr, Roepstorff A, Thamsborg SM. 2009. Ascaris suum infection negatively affects the response to a Mycoplasma hyopneumoniae vaccination and subsequent challenge infection in pigs. Vaccine, 27(37), 5161–5169. [CrossRef] [PubMed] [Google Scholar]
  22. Tolossa K, Debela E, Athanasiadou S, Tolera A, Ganga G, Houdijk J. 2013. Ethno-medicinal study of plants used for treatment of human and livestock ailments by traditional healers in South Omo, Southern Ethiopia. Journal of Ethnobiology and Ethnomedicine, 9(1), 32. [CrossRef] [PubMed] [Google Scholar]
  23. Vercruysse J, Albonico M, Behnke JM, Kotze AC, Prichard RK, McCarthy JS, Montresor A, Levecke B. 2011. Is anthelmintic resistance a concern for the control of human soil-transmitted helminths? International Journal for Parasitology: Drugs and Drug Resistance, 1(1), 14–27. [CrossRef] [Google Scholar]
  24. WHO. 2012. Accelerating work to overcome the global impact of neglected tropical diseases – a roadmap for implementation. World Health Organization: Geneva. [Google Scholar]
  25. Williams AR, Fryganas C, Ramsay A, Mueller-Harvey I, Thamsborg SM. 2014. Direct anthelmintic effects of condensed tannins from diverse plant sources against Ascaris suum. PLoS One, 9(5), e97053. [CrossRef] [PubMed] [Google Scholar]

Cite this article as: Williams AR, Soelberg J & Jäger AK: Anthelmintic properties of traditional African and Caribbean medicinal plants: identification of extracts with potent activity against Ascaris suum in vitro. Parasite, 2016, 23, 24.

All Tables

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).

All Figures

thumbnail 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.

In the text
thumbnail 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.

In the text

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