Association between naturally acquired gastrointestinal nematode infection and skeletal muscle biochemical composition in chickens (Gallus gallus)

Introduction

Poultry production is one of the most accessible sources of animal protein worldwide, particularly in small-scale and traditional production systems, where it contributes to household nutrition and income1,2. Despite improvements in management and disease control, parasitic infections remain common under field conditions3,4. Gastrointestinal helminths are frequently reported in domestic and free-range flocks, where continuous exposure to contaminated environments increases the risk of infection3–6. Studies from Ethiopia and South Africa, for example, have documented a high prevalence of gastrointestinal helminths in village and outdoor production systems4–6. Among these parasites, Ascaridia galli is commonly reported in chickens kept under non-intensive systems5,6. Previous studies have shown that helminth infection can affect performance, health status, behaviour, and social ranking in affected birds7,8. These effects suggest that gastrointestinal nematodes may influence not only productivity but also the physiological condition of infected chickens. Parasitic infections can also alter host metabolism. During infection, parasites compete for host-derived nutrients, while the host redirects energy toward immune responses9. Such changes may influence nutrient utilization and tissue composition. Because skeletal muscle composition reflects nutrient allocation and metabolic regulation, infection-related changes in lipid, carbohydrate, or protein metabolism may affect the nutritional characteristics of poultry meat10. Although the prevalence and performance-related effects of gastrointestinal nematodes have been widely studied, direct field-based evidence linking naturally acquired infection to skeletal muscle biochemical composition remains limited. Therefore, this study aimed to investigate the association between naturally acquired gastrointestinal nematode infection and muscle biochemical composition in chickens (Gallus gallus) under field conditions. This issue is relevant beyond poultry production, as changes in muscle composition may affect the nutritional quality of meat intended for human consumption.

Materials and Methods

Study design and sample collection

This study was conducted as a cross-sectional field investigation in the Zulfi region of Saudi Arabia between March and May 2025. A total of 25 indigenous chickens intended for human consumption were obtained from local small-scale farms after routine slaughter. All birds were at a similar commercial stage, with body weights ranging from 1.2 to 1.5 kg. Birds with comparable body condition and general health were included, while those showing signs of severe illness unrelated to parasitic infection were excluded. Post-mortem examination of the gastrointestinal tract was used to determine infection status. Six birds were naturally infected with gastrointestinal nematodes. For comparison, six uninfected birds were selected from the same source population. Although formal matching was not performed, the selected birds were broadly similar in size and condition. Information on age, sex, and diet was not available because of the field-based sampling design.

Necropsy and parasite recovery

Post-mortem examination was performed following standard parasitological procedures as described by Al-Hussaini and Demian11. Briefly, a longitudinal incision was made in the abdominal region, and the gastrointestinal tract was carefully removed and examined. Intestinal contents were transferred into Petri dishes containing 0.9% saline and maintained at approximately 37 °C. Recovered nematodes were repeatedly washed with saline to remove debris and prepare them for examination. Parasites were identified based on morphological characteristics using established taxonomic criteria. Molecular confirmation was not performed; therefore, recovered specimens were classified at the genus level as Ascaridia spp. (Supplementary Figures S1/S2).

Muscle sampling and biochemical analysis

Skeletal muscle samples were collected from the breast region of infected chickens (n = 6) and uninfected controls (n = 6). Approximately 60 g of muscle tissue was obtained from each bird and homogenized. From each sample, representative aliquots (~250 mg) were prepared for biochemical analysis. Separate aliquots were used to determine lipid, carbohydrate, and protein content to avoid cross-assay interference. All measurements were performed on tissue extracts, and concentrations are expressed as mg/mL of extract prepared from standardized muscle tissue weights.

Determination of lipid content

Total lipid content in muscle samples was determined using the sulfo-phospho-vanillin (SPV) colorimetric method following lipid extraction according to Folch et al.12. Briefly, 250 mg of muscle tissue was extracted using an alcohol–ether mixture (3:1) and incubated in a water bath at 65 °C for 2 h. After cooling, samples were centrifuged at 10,000 g for 15 min at 4 °C, and the supernatant was collected. The remaining residue was re-extracted using alcohol–ether (3:1) and chloroform–methanol (1:1), and all supernatants were pooled. The final volume was adjusted to 25 mL using chloroform–methanol. For the SPV reaction, 20 µL of lipid extract was mixed with 180 µL of concentrated sulfuric acid and heated at 100 °C for 10 min. After cooling to room temperature, phospho-vanillin reagent was added, and the mixture was incubated at 37 °C for 15 min to allow color development. Absorbance was measured at 530 nm using a multilabel plate reader (PerkinElmer, USA), and lipid concentrations were calculated using an oleic acid standard curve.

Determination of carbohydrate content

Total carbohydrate content was estimated as glucose using the colorimetric method described by Kemp and Van Heijningen13. Approximately 250 mg of muscle tissue was homogenized in 5% trichloroacetic acid using an automated homogenizer. The homogenate was heated in a boiling water bath for 15 min and then centrifuged at 10,000 g for 15 min at 4 °C. The supernatant was filtered through Whatman filter paper. An aliquot of 1 mL of the filtrate was mixed with 3 mL of concentrated sulfuric acid, shaken immediately, and heated for 6.5 min in a boiling water bath. After cooling, absorbance was measured at 560 nm, and glucose concentrations were calculated using a standard curve prepared from pure glucose.

Determination of protein content

Protein concentration was determined using the Bradford method14. Muscle tissue samples were homogenized in 50 mM phosphate buffer (pH 7.4) containing 1 mM EDTA and centrifuged at 10,000 g for 15 min at 4 °C. The supernatant was collected, and protein levels were measured spectrophotometrically at 625 nm using bovine serum albumin as the standard. Lipid, carbohydrate, and protein values were calculated from their respective standard curves (Fig. 1A–C).

(Fig. 1 here)

Fig. 1. Calibration curves for skeletal muscle biochemical assays. Standard curves were generated using oleic acid (A), glucose (B), and bovine serum albumin (C). The x-axis represents concentration, and the y-axis represents absorbance.

Statistical analysis

Biochemical data are presented as mean ± SD for six birds per group. Each biochemical assay was performed in three replicate measurements. The Shapiro–Wilk test was used to assess normality. Differences between infected and uninfected chickens (n = 6 per group) were evaluated using an independent samples t-test, with statistical significance set at P ≤ 0.05. Effect sizes (Cohen’s d) and exact P-values were also reported to support interpretation of the results, given the exploratory nature of the study and the small sample size. Statistical analyses were performed using SPSS (version 27.0; IBM Corp., Armonk, NY, USA).

Ethics statement

According to institutional guidelines, formal ethical approval was not required, as the study was conducted exclusively on post-mortem samples obtained from chickens slaughtered for human consumption.

Results

Prevalence of gastrointestinal nematode infection

Out of the 25 chickens examined, six (24%) were found to be infected with gastrointestinal nematodes. Infected birds harbored between 15 and 26 adult worms per bird. No significant correlations were observed between worm burden and muscle biochemical parameters. The recovered nematodes were identified as Ascaridia spp. based on morphological characteristics. Molecular confirmation was not performed; therefore, identification was limited to the genus level.

Muscle biochemical composition

Differences were observed between infected and uninfected birds (Table 1). Muscle lipid concentration was significantly lower in infected chickens (146.3 ± 39.9 mg/mL) compared with uninfected chickens (240.1 ± 85.5 mg/mL) (t(10) = 2.40, P = 0.037; d = 1.39). A similar pattern was observed for carbohydrates, with lower values in infected birds (0.41 ± 0.28 mg/mL) compared with controls (1.00 ± 0.38 mg/mL) (t(10) = 3.01, P = 0.013; d = 1.74). Protein concentration was also lower in infected chickens (0.12 ± 0.06 mg/mL) than in uninfected chickens (0.21 ± 0.12 mg/mL); however, this difference was not statistically significant (t(10) = 1.58, P = 0.146; d = 0.91).

Table 1. Skeletal muscle biochemical composition of infected and uninfected chickens
Parameter	Infected (n=6)	Uninfected (n=6)	t (df=10)	P-value	Cohen’s d
Lipids (mg/mL)	146.3 ± 39.9	240.1 ± 85.5	2.40	0.037	1.39
Carbohydrates (mg/mL)	0.41 ± 0.28	1.00 ± 0.38	3.01	0.013	1.74
Protein (mg/mL)	0.12 ± 0.06	0.21 ± 0.12	1.58	0.146	0.91

*Values are presented as mean ± SD (n = 6 per group). Concentrations are expressed as mg/mL of tissue extract prepared from standardized muscle samples.

Discussion

The present findings suggest that naturally acquired gastrointestinal nematode infection may be associated with changes in the biochemical composition of skeletal muscle in chickens. Although only 24% of the examined birds were infected, clear differences in lipid and carbohydrate concentrations were observed between infected and uninfected birds. Muscle lipid content was significantly lower in infected chickens, which is consistent with previous reports of reduced lipid reserves in hens naturally infected with Ascaridia galli15. Although that study examined liver tissue rather than skeletal muscle, it supports the broader view that parasitic infection can affect host energy balance. These changes may be partly explained by the increased energetic demands of immune responses during infection16. The reduction in carbohydrate concentration may also indicate altered energy metabolism in infected birds. Previous metabolomic work has shown that gastrointestinal nematode infection can disturb energy-related pathways in chickens17. Field-based observations have also reported changes in meat composition and reductions in muscle mass in parasitized birds18. Taken together, these findings suggest that gastrointestinal nematodes may influence nutrient allocation and tissue composition, possibly through a combination of nutrient competition, reduced absorption, and immune-related energy costs. However, because the present study was conducted under field conditions, causality cannot be established. Protein concentration was lower in infected birds, but the difference was not statistically significant. Therefore, this result should be interpreted cautiously. The relatively large effect size suggests that changes in protein metabolism cannot be excluded, but confirmation requires a larger sample size. One possible explanation is that amino acids may be redirected toward immune-related processes rather than muscle deposition during infection. Experimental studies have shown that A. galli infection can affect growth performance and nutrient utilization in chickens19. Other studies have reported intestinal changes, including altered morphology and reduced nutrient absorption capacity, during A. galli infection20,21. These mechanisms may help explain the tissue-level differences observed in the present study. This study has several limitations. The sample size was small, which may have reduced the statistical power, particularly for protein content. In addition, the field-based design did not allow control over age, sex, diet, or management conditions, all of which may influence muscle composition. Parasite identification was based on morphology only, and molecular confirmation was not performed. Despite these limitations, the study provides preliminary field-based evidence that naturally acquired gastrointestinal nematode infection may be associated with changes in muscle lipid and carbohydrate composition. Since poultry is an important source of dietary protein, especially in small-scale production systems, such changes may have implications for the nutritional quality of meat. Further controlled studies with larger sample sizes are needed to confirm these findings and clarify the underlying mechanisms.

Conclusion

This field-based study suggests that naturally acquired gastrointestinal nematode infection is associated with lower muscle lipid and carbohydrate concentrations in chickens. Although the study was limited by its small sample size and lack of control over age, sex, and diet, the observed differences indicate that infection may influence muscle biochemical composition under field conditions. These changes may have implications for the nutritional quality of poultry meat intended for human consumption. Further controlled studies with larger sample sizes are needed to confirm these findings and clarify the mechanisms involved.

Competing Interests

The author declares no competing interests.

Authors’ Contributions

The author conceived and designed the study, performed data collection and laboratory analyses, interpreted the data, and drafted and approved the final manuscript.

References

1. Farrell D. The role of poultry in human nutrition. In: Poultry development review. Rome: Food and Agriculture Organization of the United Nations (FAO); 2013. Available from: https://www.fao.org/4/al709e/al709e00.pdf

1. Mnisi CM. Nutraceuticals as components of sustainable poultry production. Agriculture & Food Security. 2024;13:77. https://doi.org/10.1186/s40066-024-00477-1

1. Tomza-Marciniak A, Pilarczyk B, Tobiańska B, Tarasewicz N. Gastrointestinal parasites of free-range chickens. Annals of Parasitology. 2014;60(4):303–306.

1. Yadeta W, Sori T, Debelo M, Michael BG, Jilo K. Prevalence of major gastrointestinal parasites in small-scale commercial poultry farms from Jimma town, southwest Ethiopia. International Journal of Research Studies in Biosciences. 2019;7(10):13–21. https://doi.org/10.20431/2349-0365.0710002

1. Mwale M, Masika PJ. Point prevalence study of gastrointestinal parasites in village chickens of Centane district, South Africa. African Journal of Agricultural Research. 2011;6(9):2033–2038. https://doi.org/10.5897/AJAR09.495

1. Sarba EJ, Bayu MD, Gebremedhin EZ, et al. Gastrointestinal helminths of backyard chickens in selected areas of West Shoa Zone, Central Ethiopia. Veterinary Parasitology: Regional Studies and Reports. 2019;15:100265. https://doi.org/10.1016/j.vprsr.2019.100265

1. Gauly M, Duss C, Erhardt G. Influence of Ascaridia galli infections and anthelmintic treatments on the behaviour and social ranks of laying hens (Gallus gallus domesticus). Veterinary Parasitology. 2007;146(3–4):271–280. https://doi.org/10.1016/j.vetpar.2007.03.005

1. Kaufmann F, Daş G, Sohnrey B, Gauly M. Helminth infections in laying hens kept in organic free-range systems in Germany. Livestock Science. 2011;141(2–3):182–187. https://doi.org/10.1016/j.livsci.2011.05.015

1. Samuel FU, Mensah CA, Mohammed IA. Principles of parasitism in parasitic diseases. In: Handbook of Poultry Parasites. Bentham Science Publishers; 2025. p.46–76. https://doi.org/10.2174/9798898811235125010007

1. Choi J, Kong B, Bowker BC, Zhuang H, Kim WK. Nutritional strategies to improve meat quality and composition in the challenging conditions of broiler production: A review. Animals. 2023;13(8):1386. https://doi.org/10.3390/ani13081386

1. Al-Hussaini AH, Demian ES. Practical manual of parasitology. Cairo: Faculty of Science, Cairo University; 1982.

1. Folch J, Lees M, Stanley GHS. A simple method for the isolation and purification of total lipids from animal tissues. Journal of Biological Chemistry. 1957;226:497–509. https://doi.org/10.1016/S0021-9258(18)64849-5

1. Kemp A, Van Heijningen AJMK. A colorimetric micro-method for the determination of carbohydrates. Biochemical Journal. 1954;56:646–648. https://doi.org/10.1042/bj0560646

1. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Analytical Biochemistry. 1976;72:248–254. https://doi.org/10.1016/0003-2697(76)90527-3

1. Sharma N, Hunt PW, Hine BC, Ruhnke I. Performance, egg quality, and liver lipid reserves of free-range laying hens naturally infected with Ascaridia galli. Poultry Science. 2018;97(6):1914–1921. https://doi.org/10.3382/ps/pey068

1. Sharma N, Hunt PW, Hine BC, Ruhnke I. The impacts of Ascaridia galli on performance, health, and immune responses of laying hens: New insights into an old problem. Poultry Science. 2019;98(12):6517–6526. https://doi.org/10.3382/ps/pez422

1. Oladosu OJ, Barbosa Correia BS, Grafl B, Liebhart D, Metges CC, Bertram HC, Daş G. Metabolomic alterations in chickens infected with gastrointestinal nematodes. Gut Pathogens. 2023;15:56. https://doi.org/10.1186/s13099-023-00584-7

1. Grigoryan V, Grigoryan L, Abrahamyan V, Melkonyan Z, Yeribekyan S, Ghazaryan A, et al. Impact of intestinal parasites on chicken meat quality and restoration by yogurt supplementation. Functional Food Science. 2025;5(7):275–285. https://doi.org/10.31989/ffs.v5i7.1695

1. Daş G, Kaufmann F, Abel H, Gauly M. Effect of extra dietary lysine in Ascaridia galli-infected grower layers. Veterinary Parasitology. 2010;170(3–4):238–243. https://doi.org/10.1016/j.vetpar.2010.02.026

1. Attia MM, Mahmoud MA, Abdelsalam M, Almutairi LA, Alqahtani MA, Areshi SM, El-Saadony MT, El-Tarabily KA, Salem HM. Molecular and histopathological characterization of Ascaridia galli infection and its impact on intestinal integrity and nutrient absorption. Frontiers in Veterinary Science. 2025;12:1622170. https://doi.org/10.3389/fvets.2025.1622170

1. Napolean A, Wong ZY, Zamani NA, Abdul Rahman N, Silip JJ, Abdul Rasid R, Along FN, Farisand AS, Bhuiyan MSA. Ascaridia galli in chickens: Co-infection dynamics, effect of production, and mitigation strategies: A review. BIO Web of Conferences. 2026;218:02003. https://doi.org/10.1051/bioconf/202621802003