JAHP_13_3_688-694

INTRODUCTION

Gastrointestinal helminthiasis remains a notable health issue among free-range and backyard poultry production worldwide (Shifaw et al., 2021). This challenge is particularly pronounced across Asia, where the warm and humid tropical climate favors helminth development and survival (Dube et al., 2010; Shifaw et al., 2021). This has been well-documented in several countries including Bangladesh, Thailand and the Philippines, where high prevalence of helminth infections in small-scale and free-range chickens has been reported (Ybañez et al., 2018; Wuthijaree et al., 2019; Abdullah et al., 2021; Wuthijaree et al., 2024). Consequently, this ecological advantage boosts the dynamics of parasite populations as parasite eggs may build up and potentially develop infectivity in the soil, litter, pasture, or via animal vectors (Thapa et al., 2015; Wongrak et al., 2015). In such systems, native chickens in semi-scavenging systems are highly susceptible to soil-transmitted helminths such as Ascaridia spp. and Heterakis spp. (Mubarokah et al., 2019; Zalizar et al., 2021; Ritu et al., 2024). These parasites can lead to stunted growth, poor nutrient absorption, inefficient feed conversion, reduced egg production, and in severe cases, result in mortality (Zalizar et al., 2021; Zirintunda et al., 2022; Hazarika et al., 2023). While synthetic anthelmintics are routinely used, their frequent use has raised concerns regarding anthelmintic resistance and accumulation of drug residues (Kabir et al., 2004; Kaplan, 2004; Goetting et al., 2011; Escosteguy, 2014; Sattar et al., 2014; Mund et al., 2016; Owusu-Doubreh et al., 2023; Mesfin et al., 2024). In rural settings, limited access to veterinary services and high costs of commercial dewormers further hinder effective parasite control (Permin and Hansen, 1998; Jamil et al., 2022).

Over the past decades, there has been a growing scientific interest on investigating the anthelmintic potential of medicinal plants traditionally used in ethnoveterinary practices (Mali and Mehta, 2008; Raza et al., 2015; Kuralkar and Kuralkar, 2021; Ahmad et al., 2023). In various parts of the Philippines, native chicken growers commonly administer crude aqueous extracts of indigenous plants such as Azadirachta indica (Neem), Areca catechu (Betel Nut), Tinospora rumphii (Makabuhay), Basella alba (Alugbati) and Piper betel (Betel Leaf) to combat worm infections; however, despite their widespread use, farmers largely remain unaware of the specific bioactive compounds responsible for their therapeutic effects (Dela Vega and Aban-Gonzales, 2019; Nadela, 2020). Recent studies have begun to elucidate the pharmacological basis of these practices. For instance, A. catechu extract has demonstrated significant in vitro and in vivo activity against A. galli, with its anthelmintic properties potentially attributed to the presence of alkaloids and tannins and saponins which can cause morphological damage in parasites (Mubarokah et al., 2019).

Moreover, A. indica which are found to contain tannins, alkaloids, and flavonoids, has demonstrated significant anthelmintic activity against A. galli in both in vitro and in vivo studies (Raza et al., 2015). These metabolites exert their effects by damaging the parasite’s cuticle, inhibition of neuromuscular and mitochondrial functions (Stephen et al., 2022; Gasaliyu et al., 2022; Tiuria et al., 2024).

Similarly, Plectranthus amboinicus (Indian borage) has demonstrated significant in vitro anthelmintic activity against Pheretima posthuma and A. galli (Wadekar et al., 2011). This therapeutic effect is primarily attributed to tannins, which bind to glycoproteins on the parasite cuticle and disrupt oxidative phosphorylation, leading to structural damage, energy depletion, and eventual paralysis and death (Wadekar et al., 2011; Raza et al., 2015; Stephen et al., 2022; Gasaliyu et al., 2022; Tiuria et al., 2024). Given these pharmacological insights, the present study conducted a preliminary phytochemical screening of medicinal plants traditionally used by native chicken farmers in Agusan del Norte, Philippines. By identifying the presence of key secondary metabolites, this study provides foundational evidence to support the scientific validation and future development of these ethnoveterinary botanicals as accessible, safe, and sustainable alternatives to conventional anthelmintics.

MATERIALS AND METHODS

Geographical Sampling and Ethnoveterinary Survey

This study adopted a purposive geographical strategy to identify focal areas with the highest populations of native chicken raisers in Agusan del Norte (Figure 1). Preliminary data on chicken population densities were obtained from the Provincial Veterinary Office, while the Municipal Agriculture Offices provided barangay-level inventory of farmers actively engaged in native chicken production. Barangays exhibiting the highest flock densities were selected as study sites. Structured, face-to-face interviews were conducted using primarily close-ended questionnaires, supplemented with open-ended items, and delivered in the local dialect to accurately capture comprehensive ethnoveterinary knowledge related to medicinal plant use for deworming. Inclusion criteria for respondents included active management of native chickens under a semi-scavenging system and personal experience in the use of plant-based anthelmintics. To ensure methodological rigor, the required sample size was calculated using Cochran’s formula for large populations (adopted from Cochran, 1963; Singh and Masuku, 2014).

Where, n₀ is the required sample size, Z is the z-score corresponding to the desired confidence level, p is the estimated proportion of an attribute present in the population, q = 1 − p, and e is the desired level of precision. The methodological approach adheres to standards established in ethnobotanical and ethnoveterinary studies (Giday and Ameni, 2003; Lans et al., 2007; Bekalo et al., 2009). A total sample size of 189 respondents was selected to ensure representative coverage of native chicken farmers across major production areas in Agusan del Norte.

Selection and Identification of Plant Materials

The selection of plant species for phytochemical screening was based on data obtained through ethnoveterinary interviews. These species were shortlisted based on the frequency of mention, consensus among informants, and prior corroboration with peer-reviewed ethnobotanical and ethnoveterinary literature. A total of eight plant species and their traditionally used parts were identified and collected from multiple locations in six municipalities: Cabadbaran City, Tubay, Kitcharao, Las Nieves, Buenavista, and Carmen. Collection per species ranged from 3-5 replicates per site with a total of 24-30 samples. These included the leaves and vines of B. alba (Alugbati), leaves of C. papaya (Papaya), bulbs of A. sativum (Garlic), stems of T. rumphii (Panyawan), leaves of A. indica (Neem), leaves of P. amboinicus (Indian Borage), leaves of P. betel (Betel Leaf), and the fruit (nut) of A. catechu (Betel Nut).

Voucher specimens of the identified plant species were prepared and subjected to taxonomic verification by a qualified botanist using standard morphological descriptors. The specific plant parts reportedly used were documented during the interviews and substantiated with photographic records to ensure scientific validation and reproducibility.

Sample Preparation

Fresh plant materials were thoroughly washed with clean water to remove soil and debris, then air-dried in a well-ventilated shaded area for 5 to 7 days to prevent degradation of heat-sensitive phytochemicals. Once fully dried, the samples were ground into a fine powder using a mechanical grinder and stored in airtight, labeled containers under dry conditions to avoid moisture-induced degradation.

Aqueous extraction was performed by soaking 10 g of each powdered plant material in 100 mL of distilled water (1:10 w/v) at room temperature (~25°C) for 24 hours with occasional stirring. This maceration procedure was designed to replicate traditional decoction practices commonly used by rural poultry farmers in the preparation of herbal anthelmintics. After extraction, the mixtures were filtered through Whatman No. 1 filter paper to obtain clear filtrates, which were subsequently used for phytochemical screening. The use of water as an extraction solvent was intended to mimic traditional ethnoveterinary practices, where water-based preparations are most preferred.

Phytochemical Screening Methods

A qualitative phytochemical screening was carried out to detect major classes of secondary metabolites present in the aqueous extracts of the selected medicinal plants. The screening focused on the identification of alkaloids, anthraquinones, flavonoids, cyanogenic glycosides, saponins, steroids, and tannins, based on their established veterinary ethnopharmacological relevance in animal health (Harborne, 1998; Trease and Evans, 2002). Each test was selected based on its diagnostic specificity and was performed under controlled laboratory conditions. Observable color changes or precipitate formations served as qualitative indicators of compound presence.

All assays were conducted in duplicate to ensure consistency and reliability of observations, following methodological standards adopted in veterinary phytochemical and ethnomedicinal research (Ayoola et al., 2008; Doughari, 2012). Table 1 summarizes the phytochemical groups evaluated, the specific tests employed, and the diagnostic criteria used for interpretation.

Table 1: Preliminary phytochemical screening and diagnostic observations.

Phytochemical Group	Test Applied	Observation
Alkaloids	Culvenor–Fitzgerald	Formation of a precipitate or turbidity (orange-brown complex).
Anthraquinones	Modified Borntrager’s	Pink to red coloration in ammoniacal layer.
Flavonoids	Bate-Smith and Metcalf	Red or violet coloration upon acid hydrolysis.
Cyanogenic glycosides	Guignard’s picrate paper	Sodium picrate paper turns from yellow to brick-red.
Saponins	Froth (foam)	Persistent froth formation (>1 cm)
Steroids	Keller–Kiliani	Reddish-brown ring at interface with bluish-green upper layer.
Tannins	Ferric chloride	Blue-black or green coloration upon addition of FeCl₃.

RESULTS AND DISCUSSION

Phytochemical Profiles and Ethnopharmacological Implications

The phytochemical screening revealed distinct qualitative profiles across eight indigenous plant species. Seven key phytochemical groups were evaluated such as alkaloids, anthraquinones, flavonoids, cyanogenic glycosides, saponins, steroids (deoxysugars), and tannins. The qualitative presence was categorized based on standard diagnostic observations as not detected (–), trace/weak (+), moderate (++), or strong (+++), as illustrated in Figure 2.

All plant samples exhibited the presence of at least five phytochemical groups. Notably, T. rumphii, P. betel, A. indica, and A. catechu emerged as the most phytochemically rich species. Flavonoids were strongly present (+++) in B. alba, C. papaya, A. sativum, T. rumphii, P. betel, and A. catechu, while tannins were also abundantly detected (+++) in T. rumphii, A. indica, P. betel, and A. catechu. Alkaloids were most prominent in C. papaya and T. rumphii, and moderately present in A. sativum, P. betel, and A. catechu. Saponins were strongly detected in B. alba, P. betel, and A. catechu, while steroids were consistently present at strong (+++) levels in all plants except P. amboinicus, which exhibited moderate levels (++). The phytochemical profiles observed in this study largely corroborate earlier findings on the abundant presence of alkaloids, flavonoids, tannins, and saponins in T. rumphii, P. betel, A. indica, and A. catechu (Mubarokah et al., 2019; Jamil et al., 2022; Ahmad et al., 2023). However, anthraquinones and cyanogenic glycosides were undetectable (–) in all tested extracts. This is possibly due to the differences in solvent polarity, where aqueous maceration excludes less polar metabolites which are often better extracted with methanol or chloroform (Ayoola et al., 2008). Variability in chemotypes, soil conditions, light exposure, and seasonality may further account for differences in phytochemical expression (Tchetan et al., 2022).

Structural and Digestive Disruption by Tannins and Saponins

High levels of tannins in T. rumphii, A. indica, P. betel, and A. catechu indicate potential for disrupting nematode cuticle integrity. Tannins bind to glycoproteins and digestive enzymes on the parasite surface, impairing nutrient uptake and energy metabolism. Mubarokah et al. (2019) and Stephen et al. (2022) demonstrated significant cuticle damage and death in Ascaridia galli following exposure to tannin-rich extracts. Similarly, saponins, detected strongly in B. alba, P. betel, and A. catechu, possess amphipathic properties that cause cytoplasmic leakage and parasite membrane rupture. These surfactant-like actions can potentiate synergistic interactions with other bioactives and have been linked to significant worm burden reduction in chickens treated with A. catechu (Mubarokah et al., 2019).

Neuromuscular and Metabolic Inhibition by Flavonoids and Alkaloids

Flavonoids which are abundant in C. papaya, T. rumphii, A. sativum, and P. betel reportedly interfere with ATPase function, calcium ion pumps, and mitochondrial metabolism, ultimately impairing motility and reproduction (Ahmad et al., 2023). A recent study of Tiuria et al. (2024) reported that flavonoid-rich P. betel extracts caused complete in vitro mortality of A. galli through paralysis and structural collapse. Moreover, alkaloids such as arecoline and piperine mimic or block neurotransmitters, leading to helminth paralysis. These alkaloids, found in T. rumphii, C. papaya, and A. sativum, mimic the mode of action of synthetic agents like levamisole due to their neurotoxicity and anti-reproductive properties (Jamil et al., 2022; Ahmad et al., 2023; Zirintunda et al., 2025).

Reproductive Interference and Immunomodulation by Steroidal Glycosides

Steroidal glycosides were consistently detected in all tested species except P. amboinicus. These compounds have been reported to interfere with mitochondrial membrane potential, which disrupts energy-dependent processes vital for helminth development. As a result, larval viability and egg hatching are significantly impaired, reflecting their ovicidal and larvicidal potential (Ahmad et al., 2023). In a study of Tchetan et al. (2022), extracts from Dioscorea spp., rich in steroidal glycosides, significantly suppressed egg production and larval development in H. contortus, which may be extrapolated to A. galli due to conserved metabolic pathways. Moreover, these compounds can enhance host immunity by promoting lymphocyte activity, boosting macrophage function, and increasing antibody production, which may improve resistance to reinfection (Kuralkar and Kuralkar, 2021).

CONCLUSIONS AND RECOMMENDATIONS

This present study validated the presence of key anthelmintic compounds in medicinal plants traditionally used for native chickens in Agusan del Norte. T. rumphii, P. betel, A. indica, and A. catechu exhibited the richest phytochemical profiles. These findings substantiate the ethnopharmacological relevance of these plants as a potential source of plant-based anthelmintics. Future studies should include quantitative phytochemical assays and controlled in vitro and in vivo studies to isolate active compounds, determine dose-dependent efficacy, and assess safety. Limitations of this study include its reliance on qualitative screening methods, potential seasonal variability in phytochemical expression, and the absence of biological efficacy testing.

ACKNOWLEDGEMENTS

The authors sincerely acknowledge the Caraga State University – College of Agriculture and Agri-Industries for logistical assistance and laboratory access. Appreciation is also extended to the Provincial Veterinary Office of Agusan del Norte and the respective Municipal Agriculture Offices of the study sites for their assistance in field coordination. We also express our gratitude to the native chicken farmers who participated in the ethnoveterinary survey and shared valuable knowledge on traditional plant-based remedies.

NOVELTY STATEMENTS

This study provides a region-specific phytochemical profile of eight ethnoveterinary plants used against gastrointestinal helminths in native chickens in Agusan del Norte, Philippines. Although their ethnoveterinary use has been previously reported in various contexts, limited data exist on their comparative phytochemical composition using water-based extractions that mimic conventional decoction techniques. By coupling farmer-derived knowledge with laboratory evidence, the study provides a data-driven shortlist of botanical leads for dose-response, mechanism-of-action, and safety trials aimed at sustainable parasite control in low-input poultry systems.

AUTHOR’S CONTRIBUTIONS

All authors contributed substantially to the study’s conception and design, data collection, and analysis or interpretation of results. They were involved in drafting the manuscript or revising it critically for important intellectual content. All authors approved the final version for publication and agreed to be accountable for all aspects of the work.

Conflict of Interest

The authors have no conflict of interest to declare.

REFERENCES

Abdullah SM, Barman A, Ali MY, Islam MS, Mohanta UK (2021). Morphological identification and prevalence of gastrointestinal helminths in backyard chicken from selected areas of Bangladesh. Res. Agric. Livestock. Fish., 8: 145–155. https://doi.org/10.3329/ralf.v8i1.53276

Ahmad S, Humak F, Ahmad M, Altaf H, Qamar W, Hussain A, Ashraf U, Abbas RZ, Siddique A, Ashraf T, Mughal MAS (2023). Phytochemicals as alternative anthelmintics against poultry parasites: A review. Agrobiol. Rec., 12: 34–45. https://doi.org/10.47278/journal.abr/2023.015

Ayoola GA, Coker HAB, Adesegun SA, Adepoju-Bello AA, Obaweya K, Ezennia EC, Atangbayila TO (2008). Phytochemical screening and antioxidant activities of some selected medicinal plants used for malaria therapy in Southwestern Nigeria. Trop. J. Pharm. Res., 7(3): 1019–1024. https://doi.org/10.4314/tjpr.v7i3.14686

Bekalo TH, Woodmatas SD, Woldemariam ZA (2009). An ethnobotanical study of medicinal plants used by local people in the lowlands of Konta Special Woreda, southern nations Ethiopia. J. Ethnobiol. Ethnomed., 5: 26. https://doi.org/10.1186/1746-4269-5-26

Cochran WG (1963). Sampling Techniques. 2nd Ed. John Wiley and Sons, New York.

Dela Vega JP, Gonzales MA (2019). Response of native chicken to selected saponin-containing herbs extracts under partially ranged system. Mindanao J. Sci. Technol., 17: 183–194.

Doughari JH (2012). Phytochemicals: Extraction methods, basic structures and mode of action as potential chemotherapeutic agents. In: Rao V (ed), Phytochemicals – A Global Perspective of Their Role in Nutrition and Health. InTech. https://doi.org/10.5772/26052

Dube S, Zindi P, Mbanga J, Dube C (2010). A study of scavenging poultry gastrointestinal and ecto-parasites in rural areas of Matebeleland Province, Zimbabwe. Int. J. Poult. Sci., 9: 911–915.

Escosteguy AP (2014). Potential use of medicinal plants in animal production: results in Brazil. In: Proc. 4th ISOFAR Sci. Conf., Organic World Congress, Istanbul, Turkey, pp. 81–84.

Gasaliyu KA, Ajanusi OJ, Suleiman MM, Dahiru S, Yusuf KH, Kyari S, Ogwiji M, Orakpoghenor O (2022). Effects of Vernonia amygdalina methanol leaf extract and fractions on Ascaridia galli in experimentally infected birds with regard to its pathological effect. Bull. Natl. Res. Cent., 46: 131. https://doi.org/10.1186/s42269-022-00819-8

Giday M, Ameni G (2003). An ethnobotanical survey on plants of veterinary importance in two Woredas of Southern Tigray, Northern Ethiopia. SINET: Ethiop. J. Sci., 26: 123–136.

Goetting V, Lee KA, Tell LA (2011). Pharmacokinetics of veterinary drugs in laying hens and residues in eggs: A review of the literature. J. Vet. Pharmacol. Ther., 34: 521–556. https://doi.org/10.1111/j.1365-2885.2011.01287.x

Harborne JB (1998). Phytochemical Methods: A Guide to Modern Techniques of Plant Analysis, 3rd edn. Chapman and Hall, London.

Hazarika A, Debnath S, Sarma J, Deka D (2023). Evaluation of in vivo anthelmintic efficacy of certain indigenous plants against experimentally-induced Ascaridia galli infection in local birds (Gallus domesticus). Exp. Parasitol., 247: 108476. https://doi.org/10.1016/j.exppara.2023.108476

Jamil M, Aleem MT, Shaukat A, Khan A, Mohsin M, Rehman TU, Abbas RZ, Saleemi MK, Khatoon A, Babar WI, Yan R, Li K (2022). Medicinal plants as an alternative to control poultry parasitic diseases. Life, 12: 449. https://doi.org/10.3390/life12030449

Kabir J, Umoh VJ, Audu-Okoh E, Umoh JU, Kwaga JKP (2004). Veterinary drug use in poultry farms and determination of antimicrobial drug residues in commercial eggs and slaughtered chicken in Kaduna State, Nigeria. Food Control, 15: 99–105. https://doi.org/10.1016/S0956-7135(03)00020-3

Kaplan RM (2004). Drug resistance in nematodes of veterinary importance: A status report. Trends Parasitol., 20: 477–481. https://doi.org/10.1016/j.pt.2004.08.001

Kuralkar P, Kuralkar SV (2021). Role of herbal products in animal production – An updated review. J. Ethnopharmacol., 278: 114246. https://doi.org/10.1016/j.jep.2021.114246

Lans C, Turner N, Khan T, Brauer G, Boepple W (2007). Ethnoveterinary medicines used for ruminants in British Columbia, Canada. J. Ethnobiol. Ethnomed., 3: 11. https://doi.org/10.1186/1746-4269-3-11

Mali RG, Mehta AA (2008). A review on anthelmintic plants. Nat. Prod. Radiance, 7: 466–475.

Mesfin YM, Mitiku BA, Admasu TH (2024). Veterinary drug residues in food products of animal origin and their public health consequences: A review. Vet. Med. Sci., 10: e70049. https://doi.org/10.1002/vms3.70049

Mubarokah WW, Nurcahyo W, Prastowo J, Kurniasih K (2019). In vitro and in vivo Areca catechu crude aqueous extract as an anthelmintic against Ascaridia galli infection in chickens. Vet. World, 12: 877–882. https://doi.org/10.14202/vetworld.2019.877-882

Mund MD, Khan UH, Tahir U, Mustafa BE, Fayyaz A (2016). Antimicrobial drug residues in poultry products and implications on public health: A review. Int. J. Food Prop., 20: 1433–1446. https://doi.org/10.1080/10942912.2016.1212874

Nadela HD (2020). Survey on ethno-veterinary practices of native chicken raisers in Davao Region, Philippines. Southeast. Philipp. J. Res. Dev., 25: 167–182.

Owusu-Doubreh B, Appaw WO, Abe-Inge V (2023). Antibiotic residues in poultry eggs and its implications on public health: A review. Sci. Afr., 19: e01456. https://doi.org/10.1016/j.sciaf.2022.e01456

Permin A, Hansen JW (1998). Epidemiology, diagnosis and control of poultry parasites. FAO Anim. Health Man. No. 4. Food and Agriculture Organization of the United Nations, Rome, Italy, 160.

Raza A, Muhammad F, Bashir S, Aslam B, Anwar I, Naseer M (2015). In-vitro and in-vivo anthelmintic potential of different medicinal plants against Ascaridia galli infection in poultry birds. World’s Poult. Sci. J., 72: 115–124. https://doi.org/10.1017/S0043933915002615

Ritu S, Labony SS, Hossain MS, Ali MH, Hasan MM, Nadia N, Shirin A, Islam A, Shohana NN, Alam MM, Dey AR, Alim MA, Anisuzzaman (2024). Ascaridia galli, a common nematode in semi-scavenging indigenous chickens in Bangladesh: epidemiology, genetic diversity, pathobiology, ex vivo culture, and anthelmintic efficacy. Poult. Sci., 103: 103405. https://doi.org/10.1016/j.psj.2023.103405

Sattar S, Hassan MM, Islam SKMA, Alam M, Faruk MSA, Chowdhury S, Saifuddin AKM (2014). Antibiotic residues in broiler and layer meat in Chittagong district of Bangladesh. Vet. World, 7: 738–743. https://doi.org/10.14202/vetworld.2014.738-743

Shifaw A, Feyera T, Walkden-Brown SW, Sharpe B, Elliott T, Ruhnke I (2021). Global and regional prevalence of helminth infection in chickens over time: A systematic review and meta-analysis. Poult. Sci., 100: 101082. https://doi.org/10.1016/j.psj.2021.101082

Singh AS, Masuku MB (2014). Sampling techniques and determination of sample size in applied statistics research: An overview. Int. J. Econ. Commer. Manag., 2: 1–22.

Stephen K, Ajanusi OJ, Suleiman MM (2022). In vitro and in vivo anthelmintic effects of Sterospermum kunthianum (Cham-Holl) leaf extract against Ascaridia galli in experimentally infected broiler chickens. J. Parasitol. Dis., 46: 152–158. https://doi.org/10.1007/s12639-021-01426-6

Tchetan E, Olounladé PA, Azando EVB, Khaliq HA, Ortiz S, Houngbeme A, Alowanou GG, Koura BI, Akouedegni GC, Houinato MRB, Hounzangbe-Adote SM, Gbaguidi FA, Quetin-Leclercq J (2022). Anthelmintic activity, cytotoxicity, and phytochemical screening of plants used to treat digestive parasitosis of small ruminants in Benin (West Africa). Animals, 12: 2718. https://doi.org/10.3390/ani12192718

Thapa S, Hinrichsen L, Brenninkmeyer C, Gunnarsson S, Heerkens JLT, Verwer C, Niebuhr K, Willett A, Grilli G, Thamsborg SM, Sørensen JT, Mejer H (2015). Prevalence and magnitude of helminth infections in organic laying hens (Gallus gallus domesticus) across Europe. Vet. Parasitol., 214: 118–124. https://doi.org/10.1016/j.vetpar.2015.10.009

Tiuria R, Sutardi LN, Nugraha AB, Subangkit M (2024). In vitro anthelmintic activity of Phyllanthus niruri Linn., Andrographis paniculata, Curcuma xanthorrhiza Roxb., and Curcuma aeruginosa Roxb. ethanol extracts on the motility and cuticle damage of Ascaridia galli. Vet. World, 17: 2488–2496. https://doi.org/10.14202/vetworld.2024.2488-2496

Trease GE, Evans WC (2002). Pharmacognosy, 15th edn. Saunders, London.

Wadekar R, Wani NS, Bagul UB, Bagul SD, Bedmutha RK (2011). Phytochemical investigation and screening of in vitro anthelmintic activity of Plectranthus amboinicus leaf extracts. Int. J. Pharmacogn. Phytochem. Res., 3: 35–38.

Wongrak K, Gauly M, Daş G (2015). Diurnal fluctuations in nematode egg excretion in naturally and in experimentally infected chickens. Vet. Parasitol., 208: 195–203. https://doi.org/10.1016/j.vetpar.2015.01.020

Wuthijaree K, Lambertz C, Vearasilp T, Anusatsananun V, Gauly M (2019). Prevalence of gastrointestinal helminths in Thai indigenous chickens raised under backyard conditions in Northern Thailand. J. Appl. Poult. Res., 28(1): 221–229. https://doi.org/10.3382/japr/pfy062

Wuthijaree K, Tatsapong P, Yung-Rahang S, Thirawong P, Pongmanee K (2024). Prevalence of natural gastrointestinal helminth infection of Thai indigenous chickens aged 12–18 weeks in small-scale chicken farms on river plains in Central Thailand. Adv. Anim. Vet. Sci., 12(4): 693–702. https://doi.org/10.17582/journal.aavs/2024/12.4.693.702

Ybañez RHD, Resuelo KJG, Kintanar APM, Ybañez AP (2018). Detection of gastrointestinal parasites in small-scale poultry layer farms in Leyte, Philippines. Vet. World, 11: 1587–1591. https://doi.org/10.14202/vetworld.2018.1587-1591

Zalizar L, Winaya A, Malik A, Widodo W, Suyatno, Anggraini A (2021). Species identification and prevalence of gastrointestinal helminths in Indonesian native chickens, and its impact on egg production. Biodiversitas, 22: 4363–4369. https://doi.org/10.13057/biodiv/d221029

Zirintunda G, Kateregga J, Nalule S, Acai-Okwee J, Ghoneim MM, Abdelhamid M, Zaghlool SS, Alshehri S, Abdelgawad MA (2025). Extracts of Carica papaya L. and Capsicum annuum L. showed comparable efficacy to piperazine citrate and levamisole hydrochloride in treatment of poultry helminths. Beni-Suef Univ. J. Basic Appl. Sci., 14: 25. https://doi.org/10.1186/s43088-025-00607-z