Skip to main content

Genetic characterization of Toxoplasma gondii in meat-producing animals in Iran

Abstract

Background

The consumption of uncooked or undercooked food from infected intermediate hosts can result in Toxoplasma gondii infection in humans. However, few studies have investigated the genetic diversity of this protozoan parasite in Iran. The aim of the present study was to genetically characterize isolates of T. gondii from intermediate host animals in Mazandaran Province, Iran.

Methods

Blood and heart tissue samples were collected from 204 ruminants, and brain tissue was collected from 335 birds. The prevalence of T. gondii infection in these samples was determined serologically using the modified agglutination test and by conventional PCR assays. Those PCR samples positive for T. gondii DNA and 13 DNA samples extracted from aborted fetuses in a previous study were genotyped with 12 genetic markers using the multilocus-nested PCR-restriction fragment length polymorphism (Mn-PCR–RFLP) technique.

Results

Antibodies for parasites were found in 35.7% of the ruminant (39.1% of sheep and 26.4% of goats) samples and in 51.3% of the bird (100% of geese, 52.3% of free-range chickens and 46% of ducks) samples. Molecular detection by PCR of the repetitive 529-bp DNA fragment revealed contamination of 13.2% of ruminant (14.6% of sheep and 9.4% of goats) samples and of 9.6% of bird (11.1% of chickens, 5.7% of ducks and 0% of geese samples). The results from 30 DNA samples revealed five ToxoDB (genome database for the genus Toxoplasma) PCR–RFLP genotypes, including #1 (Type II), #2 (Type III), #10 (Type I), #27 and #48, with genotype #1 the most predominant.

Conclusions

As evidenced by the results of this study, ruminants and birds are infected with T. gondii in the region, suggesting that they are likely to be involved in the transmission of T. gondii to humans through meat consumption. The identification of different genotypes may suggest a higher genetic diversity of this parasite in Mazandaran, reflecting local environmental contamination. These results have important public health implications for the prevention and control strategies of infection.

Graphical Abstract

Background

Toxoplasma gondii is a zoonotic parasite with a worldwide distribution that infects warm-blooded vertebrate animals, including ruminants and birds [1]. The majority of human T. gondii infections occur via accidental ingestion of food and water contaminated with sporulated oocysts of T. gondii and the ingestion of raw or undercooked meat from T. gondii-infected intermediate hosts. Transplacental transmission of tachyzoites occurs from non-immune mothers during primary maternal infection [2]. Infection with this parasite leads not only to clinical signs in healthy animals, but can also result in abortion and neonatal mortality in several farm animals and even death in adult animals [3]. Since Toxoplasma is considered to be one of the most important food-borne pathogens, meat-producing animals serve as one of the main sources of human infections [4]. Therefore, it has widespread public health significance since it causes opportunistic zoonosis infections in people with compromised immune systems, such as those with acquired immunodeficiency syndrome (AIDS) [5].

The unique population structure of this parasite is unpredictably oligoclonal, and was initially grouped into three clonal lineage types (I, II, and III) with limited genetic diversity. Among these three main designations, Type II is predominant in North America and Europe [6, 7]. On the contrary, it has been reported that parasite isolates from some African and Central and South American countries have a high genetic diversity with no dominance of lineages [8,9,10].

Information on parasite population structure is important from a public health perspective; nonetheless, such data on Asian parasite populations is currently insufficient. Recent efforts have focused on genetically characterizing parasite isolates from different regions and hosts, providing new information on the genetic diversity and structure of T. gondii [11,12,13]. Nonetheless, little research has been conducted on the distribution of parasite genotypes in Iran and even fewer studies have focused on the genotypes circulating in animals. Food-producing animals are a major source of infection since they reflect specific geographical areas of parasitic zoonosis. In this study, the multilocus-nested PCR-restriction fragment length polymorphism (Mn-PCR–RFLP) technique was used to genotype isolates in intermediate hosts (i.e. ruminants and birds) collected from different areas in Mazandaran, Iran. To the best of our knowledge, we report here the first research to investigate the genotypes of T. gondii in meat-producing animals. Therefore, the present study aimed to examine this domain, elucidating possible sources and routes of infection in northern Iran.

Methods

Study area

All of the tissue samples used in the study were collected from Mazandaran districts since this region is one of the most important foci of the farming and poultry industry of Iran. The geographical locations and natural climatic conditions of the areas the samples were collected have been described in previous studies [14, 15]. Inhabitants in this area have a custom of consuming undercooked meat as a meal called ‘Iranian kebab’.

Sampling protocol

This retrospective study was conducted on experimental samples (n = 204) obtained from the blood and hearts of ruminants (151 sheep and 53 goats) between October 2017 and April 2019. The investigations were conducted with the approval of the Animal Ethics Committee of Mazandaran University of Medical Sciences, Mazandaran, Iran (IR.MAZUMS.REC.94.1714). Permission to collect the biological samples at abattoirs was granted by the Chief Veterinary Officer of Mazandaran. Data related to each animal were documented for four independent variables, namely age, gender, animal species and region.

Sheep and goats of the study regions were slaughtered for human consumption at one of three abattoirs located in the western (62 animals), central (135 animals) and eastern (7 animals) regions of the province, respectively. The sampling was conducted using the data supplied by the Veterinary Officer and the abattoir manager. All of the animals were brought to the slaughter house by traders, who in turn had obtained them from farmers in the same area. In our study, sheep and goats, aged > 6 months, were selected randomly from among the slaughtered animals. Almost all slaughtered animals were male since female animals are normally kept for breeding purposes. All animals had been maintained predominantly in a semi-extensive type of management system (i.e. they were fed by grazing in communal natural grasslands during the daytime and kept in sheds at night). Data on sampling methods, the results of serological surveys of birds, and conventional PCR assays on aborted fetuses have been published in previous reports [14, 15].

Approximately 3–5 ml of blood was drawn directly from the jugular vein just before slaughter, and the heart of each animal was removed immediately after slaughter, placed in an individual zip-lock bag on ice packs, labeled appropriately and transported to the parasitology laboratory of Mazandaran University of Medical Sciences. Once in the laboratory, the sera were extracted from the blood samples, transferred to Eppendorf tubes and kept in a freezer at − 20 °C, and the collected hearts were kept refrigerated at + 4 °C until used. Different parts of the brain tissue were collected from 335 free-range birds, including 243 chickens, 87 ducks and five geese. The samples were labeled and stored in 70% ethanol until used for the molecular investigation.

Serological analysis

The in-house modified agglutination test (MAT) was performed to detect T. gondii-specific IgG antibodies using an antigen prepared from formalin-fixed whole RH strain tachyzoites as described previously [16]. The sheep and goat sera were diluted twofold, starting with 1:10 and ending with 1:640 or higher. The cut-off titer of 1:20 was considered to indicate seropositive cases [16, 17].

DNA extraction and conventional PCR assay

For the PCR assay, 50 g of heart tissue from each sheep and goat (free of fat mass and connective tissue) was ground into 1- to 2-cm-sized pieces and homogenized via a hand-held blender. A 100-ml aliquot of 0.9% saline was then added to the homogenized tissue, the mixture was homogenized again for 30 s and then the homogenate mixed with 250 ml of acid pepsin solution. After incubation at 37 °C for 1 h, the product was filtered immediately through two layers of cotton gauze and then centrifuged at 1200 g for 10 min. The digest was neutralized with 15 ml of 1.2% sodium carbonate solution before being centrifuged at 2000 g for 10 min [18]. A 2-ml volume of this homogenate was used for DNA extraction. For the assay on brain tissue, first, the entire sample of brain tissue collected from the birds was mixed and homogenized for 5 min, following which 100-mg samples were macerated using a mortar and pestle chilled with liquid nitrogen. The DNA extraction was performed using the DynaBio Tissue Kit (Takapouzist Co., Tehran, Iran) in accordance with the volumes used in the manufacturer's protocol. Finally, the purified nucleic acid sample was easily dissolved in 50 μl of sterile TE buffer (10 mM Tris–HCl, 1 mM EDTA), its concentration assessed by ultraviolet (UV) spectrophotometric absorbance at 260/280 nm and then the sample was stored at − 20 °C before further investigation.

The quality of the extracted DNA was verified by PCR using the host gene [i.e. receptor tyrosine kinase 2 (erbB-2)] as previously described [15].

Toxoplasma gondii molecular detection was performed by analyzing the DNA isolated from each sample using the PCR method with the amplification of the RE gene (200- to 300-fold repetitive sequence). The PCR assay was run in a 25-μl reaction volume consisting of 12.5 μl 2× Master Mix (Ampliqon, Odense, Denmark), 1 μl of purified DNA template, 0.6 μl of each PCR primer (10 pmol/μl; Bioneer, Daejeon, South Korea) and 10.3 μl of double-deionized water; the two primers used were Tox4 (forward: 5′-CGCTGCAGGGAGGAAGACGAAAGTTG-3′) and Tox5 (reverse 5′-CGCTGCAGACACAGTGCATCTGGATT-3′); fragment size: 529 bp. PCR amplification cycling consisted of pre-denaturation at 93 °C for 5 min, followed by 30 cycles at 93 °C for 30 s (denaturation), 55 °C for 30 s (annealing) and 72 °C for 30 s, and an extension step for 5 min (BioRad Laboratories, Hercules, CA, USA) [19]. The PCR fragments were separated by 1.5% agarose gel electrophoresis, and the PCR products were resolved by staining with a safe stain (CinaGen Co., Tehran, Iran) and visualized under a UV lamp on a transilluminator. In this study, RH strain (HBRC for Toxoplasma, Limoges University, France) was included as the positive control, and sterile distilled water was used as the negative control in all experiments.

Genotyping analysis

Genotyping was performed by the Mn-PCR–RFLP method using 12 genetic markers (SAG1, 5′-3′ SAG2, alt-SAG2, SAG3, GRA6, BTUB, c22-8, c29-2, PK1, L358, Apico and CS3) as previously described in detail [10, 20]. The process of genotyping was carried out on 59 DNA samples isolated from sheep, goats and birds in this study, as well as on 13 DNA samples previously collected from aborted fetuses [15]. Parasite reference strains were also used in the genotyping process, including Type I (RH), Type II (PRU) and Type III (VEG), which were kindly provided by Dr. Marie-Laure Dardé at the University of Limoges, France.

The multiplex PCR reaction was performed in a final reaction volume of 25 μl consisting of 12.5 μl 2× Master Mix, 1.5 μl genomic DNA, 0.3 μM MgCl2 and 0.3 μM external primers of each gene marker in a single reaction. The PCR cycling regimen for this analysis consisted of an initial hot start step for 4 min at 95 °C, followed by 30 cycles of 30 s at 94 °C, 1 min at 55 °C and 2 min at 72 °C, with a final extension step for 5 min at 72 °C. The products of the first round were used as templates for the second round of PCR amplification by adding 25 μl of nuclease-free water (1:1). The nested PCR reaction was performed separately for each molecular marker in 25 μl of PCR mixture containing 12.5 μl 2× Master Mix, 1 μl diluted multiplex PCR products, 0.3 μM MgCl2 and 0.75 μl (10 pmol/μl) of each of the internal primers. This PCR cycling regimen consisted of an initial denaturation at 95 °C for 4 min, followed by 35 cycles at 95 °C for 30 s, at 60 °C for 1 min (BTUB and Apico markers: 58 °C), at 72 °C for 90 s, with a final extension at 72 °C for 3 min.

For the RFLP typing procedure, 5 μl of PCR amplified products was digested with the appropriate restriction enzyme(s) in a total reaction volume of 20 μl (New England Biolabs, Ipswich, MA USA). The restriction fragments were electrophoresed in a 3% agarose gel in 1× TBE buffer at 100 V for 1 h and the products then photographed. The typing data were analyzed, compared, and categorized according to the reference strain profiles in ToxoDB, a genome database for the genus Toxoplasma (http://toxodb.org/toxo/) [20]. Moreover, 20-μl samples of the purified PCR products from 19 different samples were sequenced for 12 genetic markers in both directions (forward and reverse) by the Pishgam Company (Tehran, Iran). In this study, the sequence of the obtained samples was edited, justified and aligned using Sequencher Tmv.4.1.4 software.

Multilocus analysis by neighbor-joining clustering

The phylogenetic network was inferred using SplitsTree software (version 4.4) using the neighbor-joining algorithm between the genotype obtained in the present study and others isolated in previous research [21]. Multilocus PCR–RFLP typing data (with or without DNA restriction fragments) were coded for an allele of the locus with a combination of 0 and 1; thereafter, the genetic distances were estimated using the Tajima-Nei method.

Statistical analysis

Statistical analysis was carried out in StatsDirect software version 2.7.2 (StatsDirect Ltd., Altrincham, UK) using a standard Chi-squared method with 95% confidence intervals (95% CI). The degree of agreement between two tests (serological tests and PCR assays) was explored using Cohen’s kappa coefficient (k). A P-value of < 0.05 was considered statistically significant.

Results

Sampling and prevalence analysis

The statistical population of this research (n = 204) consisted of 151 (74%) and 53 (26%) sheep and goats, respectively. Overall, the antibodies against T. gondii were found in 73 (35.8%) cases by MAT. In addition, the seroprevalence of the parasite was 39.1% and 26.4% in sheep and goats, respectively. The titers of positive sera were: 1:20 in five sheep, 1:40 in 11 sheep, 1:80 in 21 sheep, 1:160 in nine sheep, 1:320 in six sheep and ≥ 640 in seven sheep, and 1:20 and 1:40 in 0 goats, 1:80 in two goats, 1:160 in three goats, 1:320 in eight goats and ≥ 640 in one goat. Among this population, 34 (77%) samples had MAT titers of 1:160 or higher. No significant differences were found among the different regions and species (Additional file 1: Table S1). Seroprevalence had previously been verified in 51.3% of 335 birds (MAT titer ≥ 1:20 was seen in 100%, 52.3% and 46% of geese, free-range chickens and ducks, respectively) [14]. Subsequently, the DNA samples that were PCR-positive and 13 DNA samples (aborted fetuses) from a previous study were genotyped with 12 genetic markers using the Mn-PCR-RFLP technique.

Detection in tissues

Table S2 summarizes the findings of the present study. The required tissue samples were obtained from all 204 hearts and 335 brains of birds screened for parasite DNA by conventional PCR. The PCR assays detected parasite DNA in 22 (14.6%) sheep, five (9.4%) goats, 27 (11.1%) free-range chickens and 5 (5.7%) ducks (Additional file 1: Table S2).

PCR-positive animals were almost all seropositive in this study. Positive PCR and negative MAT results were found in 14% of sheep (3/22), 0% of goats (0/5), 15% of free-range chickens (4/27) and 20% of ducks (1/5). However, Cohen’s kappa coefficient confirmed a slight degree of concordance between positive serology and positive PCR results (k = 0.18; Table 1).

Table 1 Correlation between serum MAT results and tissue PCR results

Genotyping

Of the 72 DNA samples that tested positive for T. gondii by PCR, 30 (41.7%) were successfully amplified based on Mn-PCR primers (12 gene markers); the remaining 42 samples tested PCR-negative, which could explain the observed failure in obtaining sufficient amounts of parasite DNA. The DNA of animals was designated as TgShIr (sheep), TgGoIr (goat), TgBiIr (bird) and TgAbIr (aborted fetus). The 30 amplified T. gondii genotypes originated from sheep (n = 9), goats (n = 1), free-range chickens (n = 9), ducks (n = 1) and aborted fetuses (n = 10). Moreover, five genotypes were detected in the 30 DNA samples successfully genotyped, including ToxoDB PCR–RFLP genotype #1 (also known as clonal Type II lineage), #2 (also known as clonal Type III lineage), #10 (also known as clonal Type I lineage), #27 (also known as Type I variant) and #48 (also known as Type III variant), emphasizing the genetic variability of T. gondii in northern Iran. Genotypes #1 (13 animals, 43.3%) and #2 (11 animals, 36.7%) of T. gondii were identified in all species. Parasites with clonal Type II and III alleles were also predominantly detected in sheep and birds, respectively. Genotypes #10 (TgAbIr16, 18 and 19) and #27 (TgAbIr14 and 22) were identified in aborted fetuses; nonetheless, the genotype isolated from ducks belonged to genotype #48 (TgbiIr7). The results of genotyping the 30 parasite DNA samples at all multi-molecular markers are shown in Table 2. Among the PCR products, 19 different samples were subjected to sequencing using the forward and reverse primers mentioned in the Methods section. The genomic DNA sequences reported in this study were verified by aligning them with the relevant sequences associated with the parasite and subsequently submitted to the GenBank [accessions no. MH687540.1 (Apico), MH687541.1 (BTUB), MH687542.1 (GRA6), MH687543.1 (PK1), MH687544.1 (SAG3), MH704624.1 (GRA6), MH704645.1 (alt-SAG2), MH704646.1 (BTUB), MH704647.1 (C29-2), MH704648.1 (GRA6), MH704649.1 (CS3), MH704650.1 (3-SAG2), MH704651.1 (5-SAG2), MH704652.1 (C22-8), MH704653.1 (L358), MH704654.1 (SAG1), MH704655.1 (alt-SAG2), MH704656.1 (SAG3), MH704657.1 (GRA6)].

Table 2 Multilocus genotyping of Toxoplasma gondii isolates in animal samples from northern Iran

Multilocus PCR–RFLP analysis of parasite genotypes obtained by phylogeny network

A total of five RFLP genotypes were detected among the DNA extracted from each of the 30 samples and analyzed. The neighbor-net analysis was performed using the coded genotyping data from the 47 reference strains in the ToxoDB site. The results of the phylogenetic network, which was carried out using 11 multilocus RFLP markers plus one apicoplast genome, demonstrated that the majority of the representative strains were grouped in four genetic clusters (i.e. distinct clades and haplogroups). The neighbor-joining trees had three clonal types as references (I, II, and III), which were designated as groups 1, 2, and 3, respectively. The population structures of the fourth group explained the more complex Types I and III alleles. In the current study, phylogenetic analysis of animal isolates indicated that the samples could be classified into three phylogenies groups of 1, 2 and 3, with the majority of determined isolates in groups 2 (13/30) and 3 (12/30); the remaining five isolates were placed in group 1 (5/30), as presented in Fig. 1.

Fig. 1
figure 1

Phylogenetic network analysis of identified genotypes of viable Toxoplasma gondii isolates in animal samples from northern Iran (SplitsTree4 software). The genotypes closely related to Type I, II and III lineages are shown in green, blue, and purple circles, respectively. The genotypes of the fourth group are in yellow circles

Discussion

Farm animals are an economically significant commodity in many countries, making major contributions to milk, meat and dairy products and playing an important role in breeding [22,23,24]. In Iran, meat production from sheep, goats and birds has shown an increasing trend in recent years (See http://faostat.fao.org). Therefore, the presence of T. gondii in different tissues of livestock species highlights the potential importance of these animals as possible sources of T. gondii transmission to humans. Based on a systematic review and meta-analysis (1977–2012), the pooled seroprevalence of toxoplasmosis was reported to be 31% (range: 20–95%) and 27% (range 14.2–30%), respectively, among sheep and goats in Mazandaran Province [25]. In the current study, the results of the MAT on tissues from slaughtered sheep (39.1%) and goats (26.4%) were in close agreement with those of the earlier study (Additional file 1: Table S1). The worldwide prevalence of T. gondii among small ruminants varies widely across countries, in both sheep (minimum of 3% in China to a maximum of 94.8% in the USA) and goats (minimum of 7.8% in Spain to a maximum of 90% in Egypt) [26,27,28,29]. We hypothesized that many factors could be associated with this variation, such as geographical and climatic factors, sample size, age of animals, density of infected cats shedding oocysts, type of production or management system, access to contaminated feed and water, techniques used to diagnose infection, as well as the sensitivity–specificity of the testing kits used and their cut-off points. In the present study, semi-extensive rearing systems were predominant for herds, and seropositivity for toxoplasmosis antibodies was higher in sheep than in goats. Other related studies generally reported a higher prevalence in sheep, which may be explained by differences in feeding behavior, susceptibility to diseases and breeds. For example, sheep are mostly grazers and tend to consume the bottoms of grasses (greater exposure to sporulated oocysts); in comparison, goats are browsers and feed off the tops of plants and small trees [30,31,32,33,34].

The results of this study demonstrate that Toxoplasma DNA (Toxo-DNA) could be detected by PCR assay in the tissues of naturally infected animals since we detected Toxo-DNA in 27 heart samples from 204 small ruminants and in 32 brain samples from 335 birds. The prevalence rates of Toxo-DNA in the sheep, goats and birds analyzed in the present study were 14.6%, 9.4% and 9.6%, respectively (Additional file 2: Table S2), which are higher than those reported in studies performed in other countries, such as Poland (6.9%) [35], China (5.2%) [36] and India (2.3%) [37] in sheep, goats and birds, respectively, but lower than those reported previously in Iran [38] and in Tunisia (32.5%) [39] and Kenya (79%) [40] in sheep, goats and birds, respectively. Therefore, the findings of these studies illustrate the prevalence of viable parasites, whereas serological tests primarily detect chronic toxoplasmosis in animals [41]. It is worth noting that our positive PCR findings are not definitive in terms of estimating a true prevalence of infection since only one organ of each animal was selected for analysis. This study also found a slight correlation (k = 0.18) between serological and molecular approaches for the detection of infection; however, it used a larger size of the fragments (50 g) to extract and then evaluate DNA (Table 1). The primers TOX4 and TOX5 (a multi-copy repetitive 529-bp fragment) were used to detect the parasite due to their high sensitivity and specificity [19]. The repetitive 529-bp fragment is capable of detecting the limit of ≥ 1/50 of a parasite genome equivalent [42].

Since it is essential to understand the genotypes of parasites involved in infection for epidemiological investigations, and there was limited parasite DNA available in the biological samples, we used the Mn-PCR–RFLP assay with more genetic markers [43]. Nevertheless, two systematic review studies indicated that the genotypic diversity of animal isolates differ according to geographical and host origin [12, 13]. In the present study, DNA sequencing analysis of 19 isolates with 12 markers was completed successfully and although there were limitations, all of the sequencing data were in agreement with the RFLP results. The genotyping results in this study were based on Mn-PCR–RFLP (12 gene markers) and indicated that out of 30 parasite isolates from farm animals for human consumption and aborted fetuses in Mazandaran, 24 cases were PCR–RFLP genotypes #1 (known as Type II) and #2 (known as Type III), and the remaining six isolates were genotypes #10 (known as Type I), #27 and #48 (Table 2). These findings suggest that these genotypes may be common lineages circulating in this part of Iran. However, in earlier studies these genotypes were also detected in residents of northern Iran, indicating the genetic diversity and possible circulation of T. gondii genotypes in this area. The genotyping process in these studies identified four genotypes of parasite, including four genotypes (#1, #2, #10 and #27), in donors’ blood and six different genotypes, namely genotypes #1, #2, #10, #27, #35 and #48, in HIV-positive patients [44, 45]. In our study, the analysis of the isolates from sheep suggested that clonal Type II was overwhelmingly the predominant lineage in this region. The results of studies performed in Europe and the USA indicated that Type II strains are the most frequently identified genotype in humans and animals [12, 46, 47]. These findings are in line with those obtained in the present study. Nonetheless, the existence of these classic clonal lineages, especially Type II, is rare in South America [10]. Although a small number of isolates were studied in this research, T. gondii isolated from a goat presented genotype #2 and was clustered with Type III lineage (Table 2). The results of studies reported by Dubey et al. in the USA and by Mancianti et al. in Italy, with both groups investigating T. gondii among goats, found T. gondii of clonal Type III [48, 49].

The birds analyzed in this research were free-range animals that were also infected through ground feeding. Such free-range birds are considered one of the most important animal intermediate hosts in parasite epidemiology and they play a special role in Toxoplasma transmission to different species. The findings of isolates collected from these birds demonstrated that out of the 10 cases analyzed, all but three were genotype #2; the three exceptions were genotypes #1 (2 cases) and #48 (1 case) (Table 2). The predominance of lineage Type III over Type II isolates has been reported in studies conducted on diverse bird species in Egypt and Iran [50, 51]. Genotype #48 was identified and positioned close to clonal Type III by Bernstein et al. [52] in chickens, rabbit and rats in Argentina.

There is meager evidence on the genotypes of parasites circulating in aborted fetuses worldwide. The findings of the present study suggested that clonal Type II was the dominant lineage in aborted cases and was capable of causing spontaneous abortion [53]. The results of studies carried out from 1999 to 2002 demonstrated that all isolates were Type II based on one locus [54, 55]. This result was in accordance with the findings of a previous animal study conducted in Brazil [56]. In contrast to these results, there are reports of parasite strains named atypical genotypes from goat abortion cases in Argentina [57]. In the current research, one sample was infected with genotype #2, which was previously recognized from cases of ovine abortion in Ireland [53]. In the fetal analysis, isolated genotype #10 was a high pathogenicity strain, which has been mainly observed in Asia [20]. However, Type I was recognized in ovine aborted fetuses in Qazvin and Fars provinces, Iran [58, 59]. It is noteworthy that genotype #27, which is closely related to the clonal Type I of the TgAbIr14 and TgAbIr22, was recovered previously from a bird and cat in South America (Table 2) [10].

These data were supported by network analyses, which showed genetic diversity in several studied populations and identified three groups of ancestral types and related genotypes by SplitsTree analysis (Fig. 1). The comparison of identified genotypes among animal isolates revealed overlaps, except for genotypes #10, #27 and #48, the latter being identified in aborted cases and birds. The results of genotyping in this study suggested that Mazandaran province had an epidemic population structure of the parasite. Host-parasite interactions between resistant hosts and virulent T. gondii strains could likely render current population structures to the parasite. Regarding this, further research needs to be performed to gain more comprehensive knowledge in this domain.

Conclusion

In general, the findings of this study indicate that the estimated prevalence of parasite infection among livestock is widespread in the study area. The DNA of T. gondii was detected in tissue samples from all tested animals (sheep, goats, birds and aborted fetuses), indicating that these animals might pose a risk to human health by transmitting human toxoplasmosis if their infected meat is eaten or raw meats are handled without proper hygienic procedures. To the best of our knowledge, this is the first report on the genotypes of T. gondii circulating in animals. The results of this study can be used for further epidemiological surveys since they reflect a specific geographical origin, elucidate possible sources and routes of parasite infection for humans and may have important implications for public health in Iran. Nevertheless, more research is needed to assess the pathological aspects of these genotypes in an animal model.

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Abbreviations

DNA:

Deoxyribonucleic acid

PCR:

Polymerase chain reaction

MAT:

Modified agglutination test

Mn-PCR–RFLP:

Multilocus-nested PCR-restriction fragment length polymorphism

References

  1. Dubey JP. Toxoplasmosis of animals and humans. 2nd ed. Boca Raton: CRC Press; 2016.

  2. Tenter AM, Heckeroth AR, Weiss LM. Toxoplasma gondii: from animals to humans. Int J Parasitol. 2000;30:1217–58. https://doi.org/10.1016/S0020-7519(00)00124-7.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  3. Dubey JP, Beattie C. Toxoplasmosis of animals and man. Boca Raton: CRC Press; 1988.

  4. Tenter AM. Toxoplasma gondii in animals used for human consumption. Mem Inst Oswaldo Cruz. 2009;104:364–9. https://doi.org/10.1590/S0074-02762009000200033.

    PubMed  Article  Google Scholar 

  5. Jones JL, Lopez A, Wilson M, Schulkin J, Gibbs R. Congenital toxoplasmosis: a review. Obstet Gynecol Surv. 2001;56:296–305.

    CAS  PubMed  Article  Google Scholar 

  6. Ajzenberg D, Banuls A-L, Tibayrenc M, Dardé ML. Microsatellite analysis of Toxoplasma gondii shows considerable polymorphism structured into two main clonal groups. Int J Parasitol. 2002;32:27–38. https://doi.org/10.1016/S0020-7519(01)00301-0.

    CAS  PubMed  Article  Google Scholar 

  7. Sibley LD, Mordue DG, Su C, Robben PM, Howe DK. Genetic approaches to studying virulence and pathogenesis in Toxoplasma gondii. Philos Trans R Soc Lond B Biol Sci. 2002;357:81–8. https://doi.org/10.1098/rstb.2001.1017.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  8. Ajzenberg D, Banuls AL, Su C, Dumetre A, Demar M, Carme B, et al. Genetic diversity, clonality and sexuality in Toxoplasma gondii. Int J Parasitol. 2004;34:1185–96. https://doi.org/10.1016/j.ijpara.2004.06.007.

    CAS  PubMed  Article  Google Scholar 

  9. Al-Kappany YM, Rajendran C, Abu-Elwafa SA, Hilali M, Su C, Dubey JP. Genetic diversity of Toxoplasma gondii isolates in Egyptian feral cats reveals new genotypes. J Parasitol. 2010;96:1112–4. https://doi.org/10.1645/GE-2608.1.

    CAS  PubMed  Article  Google Scholar 

  10. Pena HF, Gennari SM, Dubey JP, Su C. Population structure and mouse-virulence of Toxoplasma gondii in Brazil. Int J Parasitol. 2008;38:561–9. https://doi.org/10.1016/j.ijpara.2007.09.004.

    CAS  PubMed  Article  Google Scholar 

  11. Khan A, Dubey JP, Su C, Ajioka JW, Rosenthal BM, Sibley LD. Genetic analyses of atypical Toxoplasma gondii strains reveal a fourth clonal lineage in North America. Int J Parasitol. 2011;41:645–55. https://doi.org/10.1016/j.ijpara.2011.01.005.

    PubMed  PubMed Central  Article  Google Scholar 

  12. Sharif M, Amouei A, Sarvi S, Mizani A, Aarabi M, Hosseini SA, et al. Genetic diversity of Toxoplasma gondii isolates from ruminants: a systematic review. Int J Food Microbiol. 2017;258:38–49. https://doi.org/10.1016/j.ijfoodmicro.2017.07.007.

    CAS  PubMed  Article  Google Scholar 

  13. Amouei A, Sarvi S, Sharif M, Aghayan SA, Javidnia J, Mizani A, et al. A systematic review of Toxoplasma gondii genotypes and feline: geographical distribution trends. Transbound Emerg Dis. 2020;67:46–64. https://doi.org/10.1111/tbed.13340.

    PubMed  Article  Google Scholar 

  14. Amouei A, Sharif M, Hosseini SA, Sarvi S, Mizani A, Salehi S, et al. Prevalence of Toxoplasma gondii infection in domestic and migrating birds from Mazandaran province. Northern Iran Avian Biol Res. 2018;11:12–5. https://doi.org/10.3184/175815617X15105842200290www.

    Article  Google Scholar 

  15. Amouei A, Sharif M, Sarvi S, Bagheri Nejad R, Aghayan SA, Hashemi-Soteh MB, et al. Aetiology of livestock fetal mortality in Mazandaran province. Iran PeerJ. 2019;6:e5920. https://doi.org/10.7717/peerj.5920.

    CAS  PubMed  Article  Google Scholar 

  16. Dubey JP, Desmonts G. Serological responses of equids fed Toxoplasma gondii oocysts. Equine Vet J. 1987;19:337–9. https://doi.org/10.1111/j.2042-3306.1987.tb01426.x.

    CAS  PubMed  Article  Google Scholar 

  17. Lopes AP, Cardoso L, Rodrigues M. Serological survey of Toxoplasma gondii infection in domestic cats from northeastern Portugal. Vet Parasitol. 2008;155:184–9. https://doi.org/10.1016/j.vetpar.2008.05.007.

    PubMed  Article  Google Scholar 

  18. Hamilton CM, Kelly PJ, Bartley PM, Burrells A, Porco A, Metzler D, et al. Toxoplasma gondii in livestock in St. Kitts and Nevis, West Indies. Parasit Vectors. 2015;8:166. https://doi.org/10.1186/s13071-015-0776-7.

    PubMed  PubMed Central  Article  Google Scholar 

  19. Homan W, Vercammen M, De Braekeleer J, Verschueren H. Identification of a 200-to 300-fold repetitive 529 bp DNA fragment in Toxoplasma gondii, and its use for diagnostic and quantitative PCR. Int J Parasitol. 2000;30:69–75. https://doi.org/10.1016/S0020-7519(99)00170-8.

    CAS  PubMed  Article  Google Scholar 

  20. Su C, Shwab EK, Zhou P, Zhu XQ, Dubey JP. Moving towards an integrated approach to molecular detection and identification of Toxoplasma gondii. Parasitology. 2010;137:1–11. https://doi.org/10.1017/S0031182009991065.

    CAS  PubMed  Article  Google Scholar 

  21. Huson DH, Bryant D. Application of phylogenetic networks in evolutionary studies. Mol Biol Evol. 2006;23:254–67. https://doi.org/10.1093/molbev/msj030.

    CAS  PubMed  Article  Google Scholar 

  22. Guo M, Mishra A, Buchanan RL, Dubey JP, Hill DE, Gamble HR, et al. A systematic meta-analysis of Toxoplasma gondii prevalence in food animals in the United States. Foodborne Pathog Dis. 2016;13:109–18. https://doi.org/10.1089/fpd.2015.2070.

    PubMed  Article  Google Scholar 

  23. Belluco S, Mancin M, Conficoni D, Simonato G, Pietrobelli M, Ricci A. Investigating the determinants of Toxoplasma gondii prevalence in meat: a systematic review and meta-regression. PLoS ONE. 2016;11:e0153856. https://doi.org/10.1371/journal.pone.0153856.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  24. Deng H, Devleesschauwer B, Liu M, Li J, Wu Y, van der Giessen JWB, et al. Seroprevalence of Toxoplasma gondii in pregnant women and livestock in the mainland of China: a systematic review and hierarchical meta-analysis. Sci Rep. 2018;8:6218. https://doi.org/10.1038/s41598-018-24361-8.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  25. Sharif M, Sarvi S, Shokri A, Teshnizi SH, Rahimi M, Mizani A, et al. Toxoplasma gondii infection among sheep and goats in Iran: a systematic review and meta-analysis. Parasitol Res. 2015;114:1–16. https://doi.org/10.1007/s00436-014-4176-2.

    CAS  PubMed  Article  Google Scholar 

  26. Wang C, Qiu J, Gao J, Liu L, Wang C, Liu Q, et al. Seroprevalence of Toxoplasma gondii infection in sheep and goats in northeastern China. Small Rumin Res. 2011;97:130–3. https://doi.org/10.1016/j.smallrumres.2011.02.009.

    Article  Google Scholar 

  27. Edwards JF, Dubey JP. Toxoplasma gondii abortion storm in sheep on a Texas farm and isolation of mouse virulent atypical genotype T. gondii from an aborted lamb from a chronically infected ewe. Vet Parasitol. 2013;192:129–36. https://doi.org/10.1016/j.vetpar.2012.09.037.

    PubMed  Article  Google Scholar 

  28. Rodriguez-Ponce E, Conde M, Corbera J, Jaber JR, Ventura M, Gutiérrez C. Serological survey of antibodies to Toxoplasma gondii and Neospora caninium in goat population in Canary Islands (Macaronesia Archipelago, Spain). Small Rumin Res. 2017;147:73–6. https://doi.org/10.1016/j.smallrumres.2016.11.020.

    Article  Google Scholar 

  29. Saad NM, Hussein AAA, Ewida RM. Occurrence of Toxoplasma gondii in raw goat, sheep, and camel milk in Upper Egypt. Vet World. 2018;11:1262–5. https://doi.org/10.14202/vetworld.2018.1262-1265.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  30. Lopes AP, Dubey JP, Neto F, Rodrigues A, Martins T, Rodrigues M, et al. Seroprevalence of Toxoplasma gondii infection in cattle, sheep, goats and pigs from the North of Portugal for human consumption. Vet Parasitol. 2013;193:266–9. https://doi.org/10.1016/j.vetpar.2012.12.001.

    PubMed  Article  Google Scholar 

  31. Fereig RM, Mahmoud H, Mohamed SGA, AbouLaila MR, Abdel-Wahab A, Osman SA, et al. Seroprevalence and epidemiology of Toxoplasma gondii in farm animals in different regions of Egypt. Vet Parasitol Reg Stud Rep. 2016;3–4:1–6. https://doi.org/10.1016/j.vprsr.2016.05.002.

    Article  Google Scholar 

  32. Rasti S, Marandi N, Abdoli A, Delavari M, Mousavi SGA. Serological and molecular detection of Toxoplasma gondii in sheep and goats in Kashan, Central Iran. J Food Saf. 2018;38:e12425.

    Article  Google Scholar 

  33. Tzanidakis N, Maksimov P, Conraths FJ, Kiossis E, Brozos C, Sotiraki S, et al. Toxoplasma gondii in sheep and goats: seroprevalence and potential risk factors under dairy husbandry practices. Vet Parasitol. 2012;190:340–8. https://doi.org/10.1016/j.vetpar.2012.07.020.

    PubMed  Article  Google Scholar 

  34. Ramzan M, Akhtar M, Muhammad F, Hussain I, Hiszczynska-Sawicka E, Haq AU, et al. Seroprevalence of Toxoplasma gondii in sheep and goats in Rahim Yar Khan (Punjab). Pakistan Trop Anim Health Prod. 2009;41:1225–9. https://doi.org/10.1007/s11250-009-9304-0.

    CAS  PubMed  Article  Google Scholar 

  35. Sroka J, Bilska-Zajac E, Wojcik-Fatla A, Zajac V, Dutkiewicz J, Karamon J, et al. Detection and Molecular Characteristics of Toxoplasma gondii DNA in Retail Raw Meat Products in Poland. Foodborne Pathog Dis. 2019;16:195–204. https://doi.org/10.1089/fpd.2018.2537.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  36. Qian W, Yan W, Lv C, Bai R, Wang T, Wei Z, et al. Molecular Detection and Genotyping of Toxoplasma gondii and Neospora caninum in Slaughtered Goats in Central China. Foodborne Pathog Dis. 2020;17:348–56. https://doi.org/10.1089/fpd.2019.2726.

    CAS  PubMed  Article  Google Scholar 

  37. Thakur R, Sharma R, Aulakh RS, Singh BB. Toxoplasma gondii in Chickens (Gallus domesticus) from North India. Acta Parasitol. 2021;66:185–92. https://doi.org/10.1007/s11686-020-00266-3.

    CAS  PubMed  Article  Google Scholar 

  38. Azizi H, Shiran B, Boroujeni AB, Jafari M. Molecular survey of Toxoplasma gondii in sheep, cattle and meat products in Chaharmahal and Bakhtiari Province. Southwest of Iran Iran J Parasitol. 2014;9:429–34.

    PubMed  Google Scholar 

  39. Amdouni Y, Rjeibi MR, Rouatbi M, Amairia S, Awadi S, Gharbi M. Molecular detection of Toxoplasma gondii infection in slaughtered ruminants (sheep, goats and cattle) in Northwest Tunisia. Meat Sci. 2017;133:180–4. https://doi.org/10.1016/j.meatsci.2017.07.004.

    CAS  PubMed  Article  Google Scholar 

  40. Mose JM, Kagira JM, Karanja SM, Ngotho M, Kamau DM, Njuguna AN, et al. Detection of natural Toxoplasma gondii infection in chicken in Thika region of Kenya using nested polymerase chain reaction. BioMed Res Int. 2016;2016:7589278. https://doi.org/10.1155/2016/7589278.

  41. Robert-Gangneux F, Darde ML. Epidemiology of and diagnostic strategies for toxoplasmosis. Clin Microbiol Rev. 2012;25:264–96. https://doi.org/10.1128/CMR.05013-11.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  42. Kasper DC, Sadeghi K, Prusa AR, Reischer GH, Kratochwill K, Forster-Waldl E, et al. Quantitative real-time polymerase chain reaction for the accurate detection of Toxoplasma gondii in amniotic fluid. Diagn Microbiol Infect Dis. 2009;63:10–5. https://doi.org/10.1016/j.diagmicrobio.2008.09.009.

    CAS  PubMed  Article  Google Scholar 

  43. Liu Q, Wang ZD, Huang SY, Zhu XQ. Diagnosis of toxoplasmosis and typing of Toxoplasma gondii. Parasit Vectors. 2015;8:292. https://doi.org/10.1186/s13071-015-0902-6.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  44. Hosseini SA, Golchin E, Sharif M, Sarvi S, Ahmadpour E, Rostamian A, et al. A serological investigation and genotyping of Toxoplasma gondii among Iranian blood donors indicates threat to health of blood recipients. Transfus Apher Sci. 2020;59:102723. https://doi.org/10.1016/j.transci.2020.102723.

  45. Hosseini SA, Sharif M, Sarvi S, Abediankenari S, Hashemi-Soteh MB, Amouei A, et al. Genetic characterization of Toxoplasma gondii in Iranian HIV positive patients using multilocus nested-PCR-RFLP method. Parasitology. 2020;147:322–8. https://doi.org/10.1017/S0031182019001598.

    CAS  PubMed  Article  Google Scholar 

  46. Dardé M. Toxoplasma gondii, “new” genotypes and virulence. Parasite. 2008;15:366–71. https://doi.org/10.1051/parasite/2008153366.

  47. Dubey JP, Lago EG, Gennari SM, Su C, Jones JL. Toxoplasmosis in humans and animals in Brazil: high prevalence, high burden of disease, and epidemiology. Parasitology. 2012;139:1375–424. https://doi.org/10.1017/S0031182012000765.

    CAS  PubMed  Article  Google Scholar 

  48. Dubey JP, Rajendran C, Ferreira LR, Martins J, Kwok OC, Hill DE, et al. High prevalence and genotypes of Toxoplasma gondii isolated from goats, from a retail meat store, destined for human consumption in the USA. Int J Parasitol. 2011;41:827–33. https://doi.org/10.1016/j.ijpara.2011.03.006.

    CAS  PubMed  Article  Google Scholar 

  49. Mancianti F, Nardoni S, D’Ascenzi C, Pedonese F, Mugnaini L, Franco F, et al. Seroprevalence, detection of DNA in blood and milk, and genotyping of Toxoplasma gondii in a goat population in Italy. Biomed Res Int. 2013;2013:905326. https://doi.org/10.1155/2013/905326.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  50. Dubey JP, Graham DH, Dahl E, Hilali M, El-Ghaysh A, Sreekumar C, et al. Isolation and molecular characterization of Toxoplasma gondii from chickens and ducks from Egypt. Vet Parasitol. 2003;114:89–95. https://doi.org/10.1016/s0304-4017(03)00133-x.

    CAS  PubMed  Article  Google Scholar 

  51. Khademvatan S, Saki J, Yousefi E, Abdizadeh R. Detection and genotyping of Toxoplasma gondii strains isolated from birds in the southwest of Iran. Br Poult Sci. 2013;54:76–80. https://doi.org/10.1080/00071668.2013.763899.

    CAS  PubMed  Article  Google Scholar 

  52. Bernstein M, Pardini L, More G, Unzaga JM, Su C, Venturini MC. Population structure of Toxoplasma gondii in Argentina. Infect Genet Evol. 2018;65:72–9. https://doi.org/10.1016/j.meegid.2018.07.018.

    PubMed  Article  Google Scholar 

  53. Gutierrez J, O’Donovan J, Proctor A, Brady C, Marques PX, Worrall S, et al. Application of quantitative real-time polymerase chain reaction for the diagnosis of toxoplasmosis and enzootic abortion of ewes. J Vet Diagn Invest. 2012;24:846–54. https://doi.org/10.1177/1040638712452730.

    PubMed  Article  Google Scholar 

  54. Owen MR, Trees AJ. Genotyping of Toxoplasma gondii associated with abortion in sheep. J Parasitol. 1999;85:382–4. https://doi.org/10.2307/3285654.

    CAS  PubMed  Article  Google Scholar 

  55. Jungersen G, Jensen L, Rask MR, Lind P. Non-lethal infection parameters in mice separate sheep Type II Toxoplasma gondii isolates by virulence. Comp Immunol Microbiol Infect Dis. 2002;25:187–95. https://doi.org/10.1016/s0147-9571(01)00039-x.

    CAS  PubMed  Article  Google Scholar 

  56. de Oliveira JMB, de Almeida JC, de Melo RPB, de Barros LD, Garcia JL, Andrade MR, et al. First description of clonal lineage type II (genotype #1) of Toxoplasma gondii in abortion outbreak in goats. Exp Parasitol. 2018;188:21–5. https://doi.org/10.1016/j.exppara.2018.03.008.

    PubMed  Article  Google Scholar 

  57. Unzaga JM, More G, Bacigalupe D, Rambeaud M, Pardini L, Dellarupe A, et al. Toxoplasma gondii and Neospora caninum infections in goat abortions from Argentina. Parasitol Int. 2014;63:865–7. https://doi.org/10.1016/j.parint.2014.07.009.

    CAS  PubMed  Article  Google Scholar 

  58. Habibi G, Imani A, Gholami M, Hablolvarid M, Behroozikhah A, Lotfi M, et al. Detection and identification of Toxoplasma gondii type one infection in sheep aborted fetuses in qazvin province of Iran. Iran J Parasitol. 2012;7:64–72.

    PubMed  PubMed Central  Google Scholar 

  59. Arefkhah N, Pourabbas B, Asgari Q, Moshfe A, Mikaeili F, Nikbakht G, et al. Molecular genotyping and serological evaluation of Toxoplasma gondii in mothers and their spontaneous aborted fetuses in Southwest of Iran. Comp Immunol Microbiol Infect Dis. 2019;66:101342. https://doi.org/10.1016/j.cimid.2019.101342.

    PubMed  Article  Google Scholar 

Download references

Acknowledgements

The authors are grateful to Professor Marie-Laure Dardé (Head of Biological Resource Center for Toxoplasma, Limoges University, France) for kindly supplying the reference strains for Types I, II and III (RH, PRU, and VEG strains). We would like to thank Professor Chunlei Su (Department of Microbiology, the University of Tennessee, Knoxville, USA) for assistance with the T. gondii genotyping. The authors also would like to extend their gratitude of the Central Laboratory of the Department of Veterinary Medicine in the Mazandaran province for their collaboration during sampling and the Institute of Experimental Animal Research, Mazandaran University of Medical Sciences (MUZUMS), Sari, Iran for providing the T. gondii MAT antigen. Project support was provided in part by grants from (No. 1714) from the deputy of research, MUZUMS, Sari, Iran.

Funding

Elite Researcher Grant Committee under award number (No. 963443) from the National Institutes for Medical Research Development (NIMAD), Tehran, Iran.

Author information

Authors and Affiliations

Authors

Contributions

AA, AD, and ShS conceived and designed the study. AA, AM, ShSh and SS collected the samples and performed the experiments. AA and SAH analyzed the data. MBH-S, FA and SGh contributed reagents/materials/analysis tools. AA, ShS and MBH-S collected and recorded the gene data and performed phylogenetic network. JJ participated in technological guidance and coordination. AA and AD wrote the manuscript. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Ahmad Daryani.

Ethics declarations

Ethics approval and consent to participate

This study was conducted under the Animal Ethics Committee of Mazandaran University of Medical Sciences, Mazandaran, Iran (IR.MAZUMS.REC.94.1714).

Consent for publication

Not applicable.

Competing interests

All authors declare no competing interests.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1

: Table S1. Seroprevalence of T. gondii IgG antibodies by MAT in sheep and goats from northern Iran.

Additional file 2

: Table S2. Detection of T. gondii DNA in livestock from northern Iran.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Amouei, A., Sarvi, S., Mizani, A. et al. Genetic characterization of Toxoplasma gondii in meat-producing animals in Iran. Parasites Vectors 15, 255 (2022). https://doi.org/10.1186/s13071-022-05360-1

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s13071-022-05360-1

Keywords

  • Toxoplasma gondii
  • Genotype
  • Diversity
  • Meat-producing animals
  • Mazandaran
  • Iran