Detection of Toxocara species larvae in four Iranian free-range broiler farms

doi:10.21203/rs.3.rs-2004282/v1

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Detection of Toxocara species larvae in four Iranian free-range broiler farms

https://doi.org/10.21203/rs.3.rs-2004282/v1

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Background

The epidemiology of Toxocara canis and Toxocara cati in food animals, associated products, and their zoonotic potential are poorly understood. A cross sectional study was designed to ascertain the prevalence of Toxocara spp. larvae from free-range broiler chickens of traditional farms using conventional techniques and molecular method. Eight-hundred tissue samples including liver, gizzard, lungs and heart were collected from 200 chickens belongs to various zones of Zanjan Province, Iran and were processed by conventional and molecular techniques.

Results

Out of 800 chicken tissues, 49 samples (6.1%) were found positive for the nematode larvae. Polymerase chain reaction for the species-specific identification of the Toxocara larvae was performed. The findings showed 10.5% of chickens were infected with Toxocara spp. so that 12 samples were positive for Toxocara canis and 9 samples for Toxocara cati.

Conclusion

Considering the significant contamination of free-range broiler chickens with Toxocara larvae, consumption of the chicken's meat and viscera, especially liver and gizzards, can play an important role in transmission of the infection to humans. Prevention and control measures focused on regular deworming of dogs and cats, enhancing public awareness of Toxocara infection are recommended.

Toxocara

PCR

Tissue

Free-range broiler chickens

Poultry farms

The dog and cat roundworms, T. canis and T. cati respectively, are a public health concern in Asia, America, and parts of Europe [1]. There parasites are responsible for VT, OT, NLM and/or HT in more than hundred species of birds and mammals [2]. Humans usually acquire the infection when they accidentally ingest infective eggs present in natural environments. Moreover, humans become infected by the consumption of contaminated raw or undercooked meat and giblets of paratenic hosts containing Toxocara third-stage larvae [3]. Paratenic hosts of Toxocara are birds, chickens, pigs and other mammals. Larvae can move to these tissues and survive for a long period of time [4, 5]. The larvae in the tissues (chicken gizzard, lungs, kidneys, liver, and heart) are thought to be a source of infection to humans [6]. The consumption of raw or undercooked meat of economic animals increases the prevalence of toxocariasis [7]. It has been studied that in chicken, most of the T. canis larvae migrate to and remain in the liver [8–10], whereas most of the T. cati larvae migrate to and remain in the muscles [4, 11].

Owing to difficulties with conventional protocols for diagnosis of diseases, nowadays researchers utilize DNA based methods because of its rapidity and reliability. Polymerase chain reaction for identification and confirmation of the helminthic parasites were used by various researchers throughout the world targeting specific genes [12–14]. Molecular tools can then provide the level of discrimination that is often not achieved by microscopy alone and which is needed to differentiate between species or intraspecific variants of the relevant parasites. Molecular characterization may provide important information about the risk of zoonotic potential and transmission dynamics of parasites within a human community.

Molecular epidemiological studies involving animal reservoirs are crucial for determining the sources of helminthic infection in an ecological niche, deducing the pathogenic characteristics of the species, and developing effective control strategies. It is the most promising method for the epidemiological studies of parasitic infection such as toxocariasis. Molecular epidemiological data on Toxocara contaminating raw meat used for human consumption is limited in Iran, and to our knowledge, this is the first study to suggest that prevalence of toxocariasis in free-range broiler chickens. This study was therefore undertaken to investigate the prevalence and identification of Toxocara spp. larvae in four Iranian free-range broiler traditional farms in Zanjan Province, North-western Iran.

Ethical consideration

The tissue samples in this study were collected from chickens slaughtered in the traditional markets with the approval of the local center of animal disease control, and animal experiments were not conducted, therefore an animal rights statement is not applicable. Permission to conduct this study was also given by the Research Ethics Committee of the Zanjan University of Medical Sciences (IR.ZUMS.REC.1400.272).

Study area

This study was conducted in Zanjan Province in Northwest of Iran. The province is geographically located between latitude 36° 40′ 30.36″ North and longitude 48° 29′ 4.2″ East. Agriculture and animal husbandry are the principal occupation. The current population of Zanjan Province is a more than a million people. Poultry and sheep are the main sources of animal protein for the inhabitants [15].

Sample collection and necropsy

This cross-sectional study was conducted from April to October, 2021. In each agricultural zone, a traditional market for the supply of local and free range chickens was selected. In these traditional markets, free-range broiler chickens are offered and delivered to customers after butchery. The selected centers from each agricultural zone were visited once per week and 50 chickens were selected using a 1 in 4 systematic random sampling method [5]. The chicken carcasses were transferred to a parasitology laboratory for examination. Each carcass was dipped in the container of water for washing. Then, the visceral organs of the animals including the liver, lungs, gizzard, and heart were isolated individually. The tissues were fragmented into small pieces of less than 5 mm³ with pointed forceps for pepsin digestion.

Digestion method and larval recovery

The digestion method used was modified after that reported by Azizi et al. [4] and Taira et al. [9]. Briefly, the fragmented tissues were put into digestive solution [1 gr pepsin (Sigma-Aldrich, USA) and 10 mL HCl 37% (Merck, Germany) in 1000 mL distilled water] under constant stirring with a magnetic stirrer and kept 2 h at 39 ℃ to recover the remaining. The sediments were filtered through system of sieves with 246-µm apertures (60-mesh) and allowed to settle for 40 min. After the supernatant was removed, sedimented liquids were poured into tubes and centrifuged for 2 min at 1,500 rpm. Each of the sediments was transferred to rectangular plastic petri dishes (Polystyrene, BIOTEST, Zist-Azmoon, Iran), and the presence of any nematode larvae was monitored under a light microscope (Olympus CX23, Japan).

DNA extraction

The DNA from the tissue samples was extracted using the Tissue Genomic DNA Extraction Mini Kit (Favorgen Biotech, Ping-Tung, Taiwan) according to the manufacturer’s instruction. In Brief, 0.25 mL of digested sample were transferred to 1.5 mL Eppendorf tube, suspended in 200 µL of lysis buffer (FATG1 Buffer) and 20 µL of Proteinase K. The solution was mixed in a vortex and incubated for up to 3 hr at 60 ºC to lyse the sample. Then, 200 µL FATG2 buffer was added to the sample, mixed thoroughly by pulse-vortexing, and incubated at 70 ºC for 10 minutes. The solution was combined with 200 µL of absolute ethanol (Merck, Germany) and it was mixed thoroughly. The mixture transferred to the FATG Mini Column (supported by the kit) and centrifuged at full speed (~ 18,000 x g) for 1 min, then discarded flow-through. After an additional washing with 750 µL of wash buffer, the samples were treated with 100 µL of elution buffer for 3 min at room temperature and centrifuged at full speed for 2 min to elute DNA. Finally, purified DNA concentration measured using a NanoDrop microvolume spectrophotometer (BECO, Germany).

Molecular characterization

Identification of Toxocara spp. isolates was performed by PCR of the ITS region and a portion of the ribosomal large subunit. The specific forward primer JW4 (5′-ACTGTCGAGGATGAGCGTGA-3′) was used, which is specific for T. cati and reverse primer NC2 (5′-TTAGTTTCTTTTCCTCCGCT-3′) to amplify partial internal transcribed spacer-1 (ITS-1), complete 5.8S and ITS-2 of rDNA as described by previous studies [16, 17]. The forward primer YY1 (5′-CGGTGAGCTATGCTGGTGTG − 3′) which is specific for T. canis as previously designed and combined with reverse primer NC2 (5′-TTAGTTTCTTTTCCTCCGCT-3′) was used to amplify partial ITS-2 as well [18]. PCR reaction was performed in Super master mix-2X 12.5 µL, primers (forward and reverse) (10 pmol/ µL) 1 µL, DNA template 5 µL, DW 5.5 µL. The PCR conditions were 95°C for 5 min (primary denaturation), followed by 94°C for 45 s (denaturation), 58°C for 35 s (annealing), and 72°C for 35 s for 35 cycles (extension) with a final elongation step of 72°C for 10 min.

Amplified products were separated by electrophoresis on a 1.5% agarose gel and stained with ethidium bromide with 0.5 µg/mL ethidium bromide solutions, and a 100 kb ladder was used as DNA sizemarker for estimating the size of the amplicons and photographed using a gel documentation system (UV Transilluminator, QUANTUM SD4-1000, VILBER, France).

Statistical methods

The data and frequencies of Toxocara species were entered into SPSS Statistics 20.0 and subjected to descriptive statistics to determine their percentages. Mann-Whitney, Kruskal-Wallis, and Chi-square (Fisher’s exact) tests were used to evaluate associations of the variables. Differences in all statistical tests were considered significant at P < 0.05.

Microscopic examination of tissue samples. Of the 200 chickens were examined in the present study, 44 (22.0%) were male and 156 (78.0%) were female. Their age range was between 2 and 48 (16.1 ± 8.4) months. The weight of the study subjects were between 150 and 4000 (1740.5 ± 799.7) grams.

In the present study, out of 800 tissue samples screened, 49 samples (6.1%) were positive for the nematode larvae by conventional microscopic examination (Fig. 1). The presences of the larvae were demonstrated in the gizzard (11.5%), liver (9.5%), lungs (2.0%) and heart (1.5%), respectively (Table 1).

Table 1

Investigated parameters and recovery of larvae from tissues of free-range broiler chickens by digestion method and microscopic observation
Subject (No.)	Parameters
Chickens (200)	Age, month	Weight, gram	Female	Male	Infection rate
	Min-max (Mean ± SD)	Min-max (Mean ± SD)	No. (%)	No. (%)	No. (%)
	2–48 (16.1 ± 8.4)	150–4000 (1740.5 ± 799.7)	156 (78.0)	44 (22.0)	33 (16.5)
Organs (800)	Infection rate, No. (%)
	Gizzard	Liver	Lungs	Heart	Total
	23 (11.5)	19 (9.5)	4 (2.0)	3 (1.5)	49 (6.1)

PCR for specific detection. The 49 positive tissue samples for the nematode larvae were subjected to PCR for detection by targeting ITS-1 and ITS-2 regions which are specific for T. canis and T. cati. The amplification of target fragments revealed that 21 chickens are positive for Toxocara species. Under the optimized cycling conditions, primer set YY1/NC2 amplified a product of 330 bp uniquely from T. canis (57.14%, 12 out of 21) and primer set JW4/NC2 amplified a product of 600 bp solely from T. cati (42.86%, 9 out of 21) (Fig. 2).

The age-wise distribution of T. canis larvae in the subjects showed higher prevalence in younger chickens than older one (P ˂ 0.05). On the other hand, the weight-wise distribution of the subjects revealed that chickens with lesser weight were highly infected with Toxocara species compared to that of larger body weight chickens. This was statistically significant (P ˂ 0.05) (Table 2).

Table 2

Species wise prevalence of toxocariasis in free-range broiler chickens by molecular method (PCR)
Parameter	Species	Infected	None-infected	P-value
		No. (Mean ± SD)	No. (Mean ± SD)
Age	T. canis	12 (9.25 ± 5.1)	188 (16.5 ± 8.9)	0.002
	T. cati	9 (11.88 ± 7.9)	191 (16.3 ± 8.4)	0.074
Wight	T. canis	12 (1254.2 ± 759.3)	188 (1771.5 ± 794.0)	0.027
	T. cati	9 (1311.1 ± 760.7)	191 (1760.7 ± 797.6)	0.123

Consumption of raw or undercooked meat, especially chicken meat or giblets containing the larvae is a significant route for transmission of Toxocara species [7, 12]. In Iran, the percentage of Toxocara seropositivity has been found to range from 0 to 64% among humans and from 3 to 31% in dogs and cats as the definitive hosts [19]. In some areas of northwestern Iran, the prevalence of anti-Toxocara IgG has been reported 6.9% in dog and cat owners [20], and 4.5% in children with allergic manifestations [21]. However, no data exists on the prevalence of Toxocara in free-range broiler chickens in Iran with regard to housing systems on farms. In addition, no research has been conducted on the molecular prevalence of Toxocara in chicken tissues supplied at traditional markets in Iran.

In the present study, 16.5% (33/200) of the free-range broiler chickens were positive for Toxocara spp. by conventional microscopic examination for the nematode larvae. Zibaei et al. [12] has recorded somewhat lower (15.2%) prevalence than the present study which might be attributed to the sample size, organs and target population. In fact, the high proportion of the infection were found in the present study may be related to exposure of the free-range chickens to the natural environment, which is known to increase the probability of Toxocara infection [12]. This risk is associated with a higher probability of ingestion of infective eggs from contaminated soil [22-24]. Dogs and cats may roam freely in hay and farms. The high percentage of infected poultry identified in farms may be a consequence of inadequate housing protection with low hygiene standards for chickens, which may lead to ingestion of larval eggs excreted by dogs and cats or ingestion of invertebrate paratenic hosts of Toxocara [25].

The tissue samples which were positive for larvae by microscopy were re-subjected to molecular method to identification of Toxocara species by targeting ITS1 and ITS2 regions. As a result, 21 chickens were confirmed positive for Toxocara species (10.5%, 21/200) with relative frequency of 57.1% for T. canis and 42.7% for T. cati. Similarly, Zibaei et al. [12] isolated Toxocara larvae from tissue samples of 33 broiler chickens from Lorestan Province of Iran and identified the Toxocara species by PCR. Amongst the larvae originating from chickens, 5 (83.3%) were determined as T. canis and 1 (16.7%) as T. cati. In a recent study by Okada et al. [13], the liver, breast meat and thigh meats of culled chickens from a commercial layer farm were investigated by microscopy and PCR. T. cati and T. tanuki larvae were detected from 2 out of 50 chickens.

Many researchers have exploited PCR assay for differentiation of Toxocara species which helps in molecular epidemiological investigations and to better understand the species wise distribution of the parasite. According to the results of various studies including the present study for detection of Toxocara species by PCR, it can be conclude that there is a possibility of infection with both T. canis and T. cati in paratenic hosts, which is necessary to be differentiated using species specific primers.

In the current survey, the most infected organs were gizzard and liver. This is in accordance with the study conducted in Japan by Taira et al. [7] in which the liver were infected with T. cati larvae in the initial migration stage. Some studies reported that the gastro-intestinal tract of chickens including the crop, gizzard, duodenum, large intestine and cloaca has a higher chance or risk of infection for Toxocara larvae than other organs [4, 9].

One of the main findings of the study was a significant difference between recovered larvae in infected and non-infected chickens in terms of age and body weight. The results indicated that the viscera of young and less weighed chickens are more affected by larvae. Although not much data has been published about the weight and age of naturally infected chickens with Toxocara larvae, in a number of experimental studies, infection of younger chickens with infective eggs indicated the early accumulation of larvae in the internal organs based on the migration pattern of larvae [4, 7, 9]. Taira et al. showed that although there was no decrease in total larval recovery over time in chickens experimentally infected with T. cati eggs, predilection sites changed from liver at 1 day post-infection to muscles at 29 days post-infection, and finally 99.6% of larvae were recovered from muscles on 176 days post-infection [7]. Further studies are needed, if the more infection of internal organs in naturally infected younger chickens is related to the larval migration pattern or is due to their high sensitivity.

Limitations

The current study faced limitations such as resource constraints and lack of access to sequencing tests. Molecular testing with high sensitivity/specificity has been done to overcome some limitations. Furthermore, the migration pattern of Toxocara species larvae on carcasses and brain will be studied in the future studies.

Detection of Toxocara spp. larvae as zoonotic helminths in free-range broiler chickens indicates that the chickens can serves as one of the important paratenic hosts of infections to human beings. Consumption of farm-raised chicken viscera, especially gizzards, liver and heart can be associated with a high risk of human infection to toxocariasis. Finally, we recommend prevention and control measures including disinfection of chicken farm facilities with hot water to kill the parasite eggs and also preventing the contamination of poultry feed by dog and cat faeces, enhancing public awareness about toxocariasis to minimize the risk of the parasite transmission from consuming undercooked chicken's viscera.

Distilled water

HCl

Hydrochloric acid

Hidden toxocariasis

ITS

Internal transcribed spacer

NLM

Neural larva migrans

Ocular toxocariasis

PCR

Polymerase chain reaction

SPSS

Statistical package for the social sciences

T. canis

Toxocara canis

T. cati

Toxocara cati

Visceral toxocariasis.

Acknowledgements: The authors would like to express their gratitude and appreciation to Dr. Farzaneh Firoozeh, Academic member of the Department of Microbiology, as well as Lila Moshki members of the Department of Parasitology and Mycology, School of Medicine, Alborz University of Medical Sciences for technical assistant. We also would like to thank Dr. Ooi Hong-Kean, retired professor at Azbabu University in Japan for their comments and suggestions which greatly improved the manuscript from its original version.

Authors’ contributions

ES, AH, and MZ worked on the data collection. ES, AH, and MZ wrote the manuscript following discussions with KT. KM did the statistical analysis. AH and MZ revised the paper and improved the technical quality of the manuscript. ES, AH, and MZ were the project coordinator and participated in all parts of the work. All authors read and approved the final manuscript.

Funding

There was no any funding or sponsoring organization for this paper.

Availability of data and materials

All the data generated or analyzed in this study are included in this published article. The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Ethics approval and consent to participate

The study was carried out in compliance with the ARRIVE guidelines. Informed consent was obtained from the owner or legal custodian of all animals described in this study for the procedures undertaken. The ethical statement of this study was approved and written by the Ethical Committee for Zanjan University of Medical Sciences with the code of IR.ZUMS.REC.1400.272. All methods were carried out in accordance with relevant guidelines and regulations. This project was partially supported by the Vice Chancellor for Research of Zanjan University of Medical Sciences by the code A-12–95-16.

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Author details

¹ Department of Parasitology and Mycology, School of Medicine, Zanjan University of Medical Sciences, Zanjan, Iran. ² Department of Parasitology and Mycology, School of Medicine, Alborz University of Medical Sciences, P.O. Box: 3149779453, Karaj, Iran. ³ Department of Epidemiology and Biostatistics, School of Medicine, Zanjan University of Medical Sciences, Karaj, Iran. ⁴ Laboratory of Parasitology, School of Veterinary Medicine, Azabu University, Fuchinobe, Sagamihara, Kanagawa, Japan.

Ma G, Rostami A, Wang T, Hofmann A, Hotez PJ, Gasser RB. Global and regional seroprevalence estimates for human toxocariasis: A call for action. Adv Parasitol. 2020;109:275-90.
Despommier DD. Toxocariasis: clinical aspects, epidemiology, medical ecology, and molecular aspects. Clin Microbiol Rev. 2003;16:265-72.
Schantz P.M. Toxocara larva migrans now. Am J Trop Med Hyg. 1989:41:21-34.
Azizi S, Oryan A, Sadjjadi SM, Zibaei M. Histopathologic changes and larval recovery of Toxocara cati in experimentally infected chickens. Parasitol Res. 2007;102:47-52.
Naderbandi M, Zibaei M, Haniloo A, Firoozeh F, Hatami Z, Shokri E, et al. Larva migrans in BALB/c mice experimentally infected with Toxocara cati ensured by PCR assay. BMC Vet Res. 2022;18:257.
Kusnoto K. Toxocariasis prevalence in stray cats in Surabaya via gastrointestinal surgery. Vet Med J. 2005;21:7-11.
Taira K, Saitoh Y, Kapel CM. Toxocara cati larvae persist and retain high infectivity in muscles of experimentally infected chickens. Vet Parasitol. 2011;180:287-91.
Galvin TJ. Experimental Toxocara canis infections in chickens and pigeons. J Parasitol. 1964;50:124-7.
Taira K, Permin A, Kapel CM. Establishment and migration pattern of Toxocara canis larvae in chickens. Parasitol Res. 2003;90:521-3.
Dutra GF, Pinto NS, de Avila LF, Dutra PC, Telmo Pde L, Rodrigues LH, et al. Risk of infection by the consumption of liver of chickens inoculated with low doses of Toxocara canis eggs. Vet Parasitol. 2014:,203:87-90.
Oryan A, Sadjjadi SM, Azizi S. Longevity of Toxocara cati larvae and pathology in tissues of experimentally infected chickens. Korean J Parasitol. 2010;48:79-80.
Zibaei M, Sadjjadi SM, Maraghi S. The occurrence of Toxocara species in naturally infected broiler chickens revealed by molecular approaches. J Helminthol. 2017:91:633-6.
Okada N, Ooi HK, Taira K. Detection of larvae of Toxocara cati and T. tanuki from the muscles of free-ranging layer farm chickens. Parasitol Res. 2021;120:1737-41.
Michelutti A, Sgubin S, Falcaro C, Cagnin V, Zoroaster A, Danesi P. Detection of Toxocara cati larvae from ostrich and wild boar meat intended for human consumption. Pathogens. 2021;10:1290.
Kohansal MH, Nourian A, Haniloo A, Fazaeli A. Molecular detection of Taenia spp. in dogs’ feces in Zanjan Province, Northwest of Iran. Vet World. 2017;10:445-9.
Zhu XQ, D'Amelio S, Palm HW, Paggi L, George-Nascimento M, Gasser R.B. SSCP-based identification of members within the Pseudoterranova decipiens complex (Nematoda: Ascaridoidea: Anisakidae) using genetic markers in the internal transcribed spacers of ribosomal DNA. Parasitology. 2002;124:615-23.
Li MW, Lin RQ, Chen HH, Sani RA, Song HQ, Zhu XQ. PCR tools for the verification of the specific identity of ascaridoid nematodes from dogs and cats. Mol Cell Probes. 2007;21:349-54.
18. Jacobs DE, Zhu X, Gasser RB, Chilton N. PCR-based methods for identification of potentially zoonotic ascaridoid parasites of the dog, fox and cat. Acta Trop. 1997;68:191-200.
Zibaei M, Sadjjadi SM. Trend of toxocariasis in Iran: a review on human and animal dimensions. Iran J Vet Res. 2017;18:233-42.
Nourian AA, Amiri M, Ataeian A, Haniloo A, Mosavinasab SN, Badali H. Seroepidemiological study for toxocariasis among children in Zanjan-Northwest of Iran. Pak J Biol Sci. 2008;11:1844-7.
Abbasi S, Ahmadi Afshar A, Haniloo A. Prevalence of anti-Toxocara antibodies in children with allergic manifestations referring to allergy clinics in Zanjan. SJKU. 2017;22:71-9.
Campos-da-Silva DR, da Paz JS., Fortunato VR, Beltrame MA, Valli LC, Pereira FE. Natural infection of free-range chickens with the ascarid nematode Toxocara sp. Parasitol Res. 2015;114:4289-93.
Von Sohsten AL, Vieira da Silva A, Rubinsky-Elefant G, Freitas Santana IS, Correia JE, Alves da Cruz L, et al. Chickens bred extensively as sentinels from soil contamination by Toxocara. Exp Parasitol. 2020;211:107852.
Pezeshki A, Haniloo A, Alejafar A, Mohammadi-Ghalehbin B. Detection of Toxocara spp. eggs in the soil of public places in and around of Ardabil city, Northwestern Iran. Iran J Parasitol. 2017;12:136-42.
Rasouli E, Zibaei M, Bahadory S, Hosseini H, Firoozeh F. Parasitological and molecular study of Toxocara spp. in Lumbericus terrestricus earthworms. Bulg J Vet Med. 2020;23:487-93.

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You are reading this latest preprint version

Detection of Toxocara species larvae in four Iranian free-range broiler farms

Status:

Version 1

Abstract

Background

Results

Conclusion

Figures

Background

Methods

Ethical consideration

Study area

Sample collection and necropsy

Digestion method and larval recovery

DNA extraction

Molecular characterization

Statistical methods

Results

Discussion

Conclusion

Abbreviations

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1