Skip to main content Accessibility help
×
Home
Hostname: page-component-78dcdb465f-f64jw Total loading time: 0.967 Render date: 2021-04-17T21:41:31.504Z Has data issue: true Feature Flags: { "shouldUseShareProductTool": true, "shouldUseHypothesis": true, "isUnsiloEnabled": true, "metricsAbstractViews": false, "figures": false, "newCiteModal": false, "newCitedByModal": true }

A survey of zoonotic pathogens carried by house mouse and black rat populations in Yucatan, Mexico

Published online by Cambridge University Press:  10 July 2017

J. A. PANTI-MAY
Affiliation:
Doctorado en Ciencias Agropecuarias, Campus de Ciencias Biológicas y Agropecuarias, Universidad Autónoma de Yucatán, Merida, Mexico
R. R. C. DE ANDRADE
Affiliation:
Centro de Pesquisas Gonçalo Moniz, Fundação Oswaldo Cruz, Ministério da Saúde, Salvador, Brasil
Y. GURUBEL-GONZÁLEZ
Affiliation:
Departamento de Zoología, Campus de Ciencias Biológicas y Agropecuarias, Universidad Autónoma de Yucatán, Merida, Mexico
E. PALOMO-ARJONA
Affiliation:
Departamento de Zoología, Campus de Ciencias Biológicas y Agropecuarias, Universidad Autónoma de Yucatán, Merida, Mexico
L. SODÁ-TAMAYO
Affiliation:
Departamento de Zoología, Campus de Ciencias Biológicas y Agropecuarias, Universidad Autónoma de Yucatán, Merida, Mexico
J. MEZA-SULÚ
Affiliation:
Departamento de Zoología, Campus de Ciencias Biológicas y Agropecuarias, Universidad Autónoma de Yucatán, Merida, Mexico
M. RAMÍREZ-SIERRA
Affiliation:
Laboratorio de Parasitología, Centro de Investigaciones Regionales ‘Dr. Hideyo Noguchi’, Universidad Autónoma de Yucatán, Merida, Mexico
E. DUMONTEIL
Affiliation:
Laboratorio de Parasitología, Centro de Investigaciones Regionales ‘Dr. Hideyo Noguchi’, Universidad Autónoma de Yucatán, Merida, Mexico Department of Tropical Medicine, School of Public Health and Tropical Medicine, Tulane University, New Orleans, LA, USA
V. M. VIDAL-MARTÍNEZ
Affiliation:
Laboratorio de Patología Acuática, Departamento de Recursos del Mar, Centro de Investigación y de Estudios Avanzados del Instituto Politécnico Nacional, Unidad Merida, Merida, Mexico
C. MACHAÍN-WILLIAMS
Affiliation:
Laboratorio de Arbovirología, Centro de Investigaciones Regionales ‘Dr. Hideyo Noguchi’, Universidad Autónoma de Yucatán, Merida, Mexico
D. DE OLIVEIRA
Affiliation:
Centro de Pesquisas Gonçalo Moniz, Fundação Oswaldo Cruz, Ministério da Saúde, Salvador, Brasil
M. G. REIS
Affiliation:
Centro de Pesquisas Gonçalo Moniz, Fundação Oswaldo Cruz, Ministério da Saúde, Salvador, Brasil
M. A. TORRES-CASTRO
Affiliation:
Laboratorio de Enfermedades Emergentes y Re-emergentes, Centro de Investigaciones Regionales ‘Dr. Hideyo Noguchi’, Universidad Autónoma de Yucatán, Merida, Mexico
M. R. ROBLES
Affiliation:
Centro de Estudios Parasitológicos y de Vectores, CONICET-Universidad Nacional de La Plata, La Plata, Argentina
S. F. HERNÁNDEZ-BETANCOURT
Affiliation:
Departamento de Zoología, Campus de Ciencias Biológicas y Agropecuarias, Universidad Autónoma de Yucatán, Merida, Mexico
F. COSTA
Affiliation:
Centro de Pesquisas Gonçalo Moniz, Fundação Oswaldo Cruz, Ministério da Saúde, Salvador, Brasil Instituto de Saúde Coletiva, Universidade Federal da Bahia, Salvador, Brasil
Corresponding
E-mail address:
Rights & Permissions[Opens in a new window]

Summary

The house mouse (Mus musculus) and the black rat (Rattus rattus) are reservoir hosts for zoonotic pathogens, several of which cause neglected tropical diseases (NTDs). Studies of the prevalence of these NTD-causing zoonotic pathogens, in house mice and black rats from tropical residential areas are scarce. Three hundred and two house mice and 161 black rats were trapped in 2013 from two urban neighbourhoods and a rural village in Yucatan, Mexico, and subsequently tested for Trypanosoma cruzi, Hymenolepis diminuta and Leptospira interrogans. Using the polymerase chain reaction we detected T. cruzi DNA in the hearts of 4·9% (8/165) and 6·2% (7/113) of house mice and black rats, respectively. We applied the sedimentation technique to detect eggs of H. diminuta in 0·5% (1/182) and 14·2% (15/106) of house mice and black rats, respectively. Through the immunofluorescent imprint method, L. interrogans was identified in 0·9% (1/106) of rat kidney impressions. Our results suggest that the black rat could be an important reservoir for T. cruzi and H. diminuta in the studied sites. Further studies examining seasonal and geographical patterns could increase our knowledge on the epidemiology of these pathogens in Mexico and the risk to public health posed by rodents.

Type
Original Papers
Copyright
Copyright © Cambridge University Press 2017 

INTRODUCTION

The house mouse (Mus musculus) and the black rat (Rattus rattus) are two of the most widespread mammals in the world [Reference Battersby, Hirschhorn, Amman, Bonnefoy, Kampen and Sweeney1]. These species are serious pests in urban and rural environments. They are the cause of extensive economic damage to crops, stored food, farms, industries and households [Reference Pimentel, Zuniga and Morrison2]. House mouse and black rat populations also harbour and spread zoonotic pathogens, such as viruses (e.g., Seoul hantavirus), bacteria (e.g., Leptospira interrogans), protozoa (e.g., Toxoplasma gondii) and helminths (e.g., Hymenolepis spp.) [Reference Himsworth3].

Neglected tropical diseases (NTDs) are communicable infections that affect mainly people living in poverty and without adequate sanitation in tropical and subtropical regions [4]. Among these, American trypanosomiasis and leptospirosis are two NTDs that affect millions of people in Latin America [Reference Sánchez-Montes5, Reference Carabarin-Lima6]. Hymenolepiasis is the most common cestodiasis in humans, particularly children living in areas of low socioeconomic status and low levels of hygiene practices [Reference Mason and Patterson7, Reference Mirdha and Samantray8]. Although hymenolepiasis is not a NTD, some authors suggest to re-evaluate its status in view of emerging issues relating to the epidemiology and impact on public health of the infection it causes [Reference Thompson9].

American trypanosomiasis (Chagas disease), is a zoonotic disease in the Americas caused by the protozoan parasite Trypanosoma cruzi [Reference Hotez10]. It is endemic in Latin America and continues to be a social and economic problem in many countries, affecting an estimated 6 million people [11]. This disease has two phases, acute and chronic. The acute phase is usually asymptomatic, but when symptoms occur the infection is characterized by an elevated parasitaemia associated with fever, headache, nausea, that is rarely lethal [Reference Carabarin-Lima6]. This phase is followed by a chronic phase, which remains asymptomatic in the majority of patients for life. Approximately 20–40% of patients in this phase present a progressive and debilitating chronic chagasic cardiomyopathy that leads to congestive cardiac failure and death [Reference Carabarin-Lima6]. The transmission to humans is mainly by hematophagous bugs of the genera Triatoma, Panstrongylus and Rhodnius (Hemiptera: Reduvidae). Trypanosoma cruzi has been documented in more than 150 domestic animals (e.g. dogs and cats) and wild mammals (e.g. marsupials and rodents). In urban settings, domiciliated and intrusive vectors and synanthropic mammals are involved in the domestic cycle, whereas in rural settings, the cycle is more complex due to the presence of vectors and synanthropic and wild mammals that invade households from (tropical) forests [Reference Waleckx, Gourbière and Dumonteil12]. The black rat and the house mouse have been reported in several countries as important carriers of T. cruzi in both domestic and peridomestic cycles [Reference Lima13, Reference Pinto14].

Leptospirosis is a widespread zoonotic disease caused by Gram-negative spirochete bacteria of the genus Leptospira [Reference Ko, Goarant and Picardeau15]. It has been estimated that 1·03 million human cases of leptospirosis and 58 900 deaths due to pulmonary haemorrhage syndrome and acute kidney injury occur annually due to leptospirosis worldwide [Reference Costa16]. Leptospira strains (serovars) are, although not totally limited, adapted to different mammalian hosts [Reference Himsworth3]. For instance, Norway and black rats are reservoirs for the Icterohaemorragiae serogroup, whereas the house mouse is the main reservoir for the Ballum serogroup [Reference Ko, Goarant and Picardeau15]. In rodents, leptospires cause a systemic infection within 7–9 days after infection but they are rapidly cleared from all tissues except the renal tubules, where bacteria persist and are shed to the environment for several months [Reference Athanazio17]. Exposure with water or soil contaminated with urine of infected rodents is the common source for human infection. Leptospirosis occurs in diverse epidemiological settings, but in low socioeconomic level/status areas with high abundance of rodents, the risk of Leptospira transmission is higher [Reference Costa18]. A 2012 study reported that the median number of leptospirosis cases notified annually in the Americas by national ministries of health was 4713·5 [Reference Costa19].

Human hymenolepiasis is a zoonosis caused by the cestodes Hymenolepis nana and H. diminuta [Reference Nkouawa20]. Infections with adult hymenolepids occur worldwide, particularly in children [Reference Thompson9, Reference Edelman21]. Synanthropic rodents are the main reservoirs for these cestodes [Reference Himsworth3]. In general, cestodes of the genus Hymenolepis require arthropod intermediate hosts in their life cycle, except for H. nana, which is the only cestode known to be transmitted directly to another definitive host [Reference Baker and Baker22]. In rodents, light infections with Hymenolepis are usually non-pathogenic, but heavy infections can cause acute catarrhal enteritis or chronic enterocolitis [Reference Baker and Baker22]. Humans can be infected with hymenolepidids by accidental ingestion of intermediate hosts (e.g. beetles or fleas) or by directly ingesting the parasite eggs as a result of contamination of food or water [Reference Nkouawa20]. Human hymenolepiasis is often asymptomatic, but can cause chronic diarrhoea, abdominal pain, irritability and itching [Reference Chero23, Reference Martínez-Barbabosa24]. In the Americas, human hymenolepiasis has been reported in several countries, such as Canada, the United States, Mexico, Peru and Argentina [Reference Edelman21, Reference Martínez-Barbabosa24Reference Luney26].

In the State of Yucatan, Mexico, it has been estimated that more than 61 000 people are infected with T. cruzi [Reference Carabarin-Lima6]. In addition, field studies have reported high abundances of vectors in urban and rural areas [Reference Guzman-Tapia, Ramírez-Sierra and Dumonteil27, Reference Dumonteil28], and rats being a common blood source for vectors [Reference Guzman-Tapia, Ramírez-Sierra and Dumonteil27]. Epidemiologic studies of human leptospirosis have reported seroprevalences of ~14%, with the icterohaemorrhagiae serovar predominant in the icteric cases [Reference Vado-solís29, Reference Vado-Solís30]. In rodents, L. interrogans serovar icterohaemorrhagiae has been reported as the predominant serovar [Reference Vado-Solís30, Reference Torres-Castro31]. In Yucatecan children, H. nana is a common cestode [Reference Duarte-Zapata, Escalante-Triay and López-Novelo de Ceballos32, Reference Rodriguez-Pérez33], whereas H. diminuta has not been reported. The only study that investigated the helminth fauna of synanthropic rodents, did not reported hymenolepids in black rats nor house mice [Reference Panti-May34]. The role of synanthropic rodents and polyparasitism in these hosts are vital issues in understanding the epidemiology of these diseases. However, in Mexico, few studies have investigated the role of these animals, especially in the tropical region. The aim of this study was to determine whether house mouse and black rat populations carry Trypanosoma cruzi, Hymenolepis spp. and Leptospira spp. in two urban neighbourhoods and a rural village of Yucatan, Mexico.

METHODS

Study sites

This study was carried out in the residential neighbourhoods of San Jose Tecoh (SJT; 20°53′16·0″N, 89°37′19·9″W) and Plan Ayala Sur (PAS; 20°54′54·0″N, 89°37′22·8″W), in the south of the city of Merida, Yucatan, Mexico. A 2007 study found that T. dimidiata, the main vector of T. cruzi, infested 38% of houses in the south of Merida and its infection rate by T. cruzi was 48% [Reference Guzman-Tapia, Ramírez-Sierra and Dumonteil27]. SJT is an urban area of 1·11 km2 and ~6001 inhabitants, whereas PAS is a suburban area of 1·32 km2 and has ~3037 inhabitants [35]. The neighbourhoods are situated in a low socioeconomic level/status area of the south of the city of Merida and are characterized by having paved streets, many small businesses, households in poor conditions (with cracks or holes in doors or windows) and vacant lots. In these neighbourhoods it is common to find pets (i.e. dogs and cats), chickens, weeds, shrubs, fruit trees and unserviceable domestic appliances in the yards. Additionally the rural village of Opichen (OPI, 20°33′05·26″N, 89°51′21·76″W) was surveyed as a part of a collaboration between researchers of the Universidad Autonoma de Yucatan. OPI is a rural area of 1·46 km2 and has ~4761 inhabitants. This village is located in the western part of the Yucatan. The majority of inhabitants live in houses constructed with stones, wooden poles and thatched with palm leaves that are adjacent to small bedrooms constructed with blocks of concrete. It is common to find chickens, pigs, cattle, weeds, shrubs, trees and vegetable patch plots in the yards.

Trapping methodology

In the two urban neighbourhoods (SJT and PAS), rodents were trapped intensively during a 6-month period from May to October 2013. Thirty households in each neighbourhood were selected at random from spatial maps and sampled monthly. At each household, six Sherman traps (two sizes were used, 8 × 9 × 23 and 8 × 9·5 × 30·5 cm3; HB Sherman Traps Inc., Tallahassee, Florida, USA) were set for three consecutive nights [Reference Panti-May36]. Traps were baited with a mixture of oatmeal and vanilla essence and were distributed in the house and yard close to signs of rodent activity or potential sources of food and/or harbourage. In the rural village (OPI), rodents were non-intensively (one night of trapping) trapped in 50 households in August and September 2013. The rodent trapping was conducted under license from the Mexican Ministry of Environment (SGPA/DGVS/02528/13). Trapped rodents were transported to the laboratory, anaesthetized with an intraperitoneal injection of sodium pentobarbital, and euthanized by cervical dislocation (mice) or with an overdose of anaesthesia (rats) [Reference Leary37].

Data collection

After anaesthesia, a blood sample was obtained by cardiac puncture. Subsequently, animals were euthanized, and heart, kidneys and intestinal tract were removed for pathogen determinations as described below. The blood, heart and kidneys were stored at –80 °C and the intestinal tract at –20 °C until final use. For financial reasons, not all animals were tested. So, animals were selected at random. Additionally, it was not possible to obtain enough blood and feces samples from all small mammals, especially individuals of the house mouse.

Pathogen survey

Trypanosoma cruzi

The presence of T. cruzi DNA in blood and heart samples was detected by polymerase chain reaction (PCR) at the Centro de Investigaciones Regionales ‘Dr. Hideyo Noguchi’, Mexico. For DNA extraction, we used standardized homemade protocols. Briefly, a half of each heart sample was macerated and homogenized in 400 µl of extraction buffer (1 M Tris–HCl, 5 M NaCl, 0·5 M EDTA, 10% SDS and distilled water). This mixture was allowed to stand at room temperature for 2 h and centrifuged for 10 min at 14 000 rpm. After that, it was transferred to a 1·5 ml microcentrifuge tube with 300 µl of isopropanol and centrifuged at 14 000 rpm. The sediment was dried and re-suspended in 60 µl of TE buffer (0·5 M EDTA 1 M Tris–HCl pH 7·0). To extract DNA from blood, 100 µl of each sample were denaturalized at 95 °C for 10 min in a boiling water bath and centrifuged at 14 000 rpm for 10 min. The supernatant was processed following the methodology described for the heart samples.

In the PCR reaction, we used the primers proposed by Moser et al. [Reference Moser, Kirchhoff and Donelson38]: TCZ-F and TCZ-R, which amplified a fragment of 188 pb belonging to a region of T. cruzi satellite DNA. The reaction (40 µl) included: 1× PCR Buffer (10 mM Tris–HCL pH 8·4 and 50 mM KCl, Promega, USA), 3 mM MgCl2, 0·1 mM dNTP, 250 µM both primers and molecular grade water. The template DNA was used in two different amounts: for heart samples 10 µl were used, whereas for blood samples 1 µl was used. Cycling parameters were one step of 5 min at 94 °C, 35 cycles of 10 s at 94 °C, 30 s at 55 °C and 30 s at 72 °C, and one final extension step of 5 min at 72 °C. All reactions included positive (DNA extracted from a culture of T. cruzi lineage I) and negative (sterile water) controls. PCR products were analysed in 1% agarose gels stained with ethidium bromide. Rodents from Opichen were no tested for T. cruzi.

Hymenolepis spp.

The faecal and caecum contents were examined for Hymenolepis eggs using the formalin–ethyl acetate sedimentation technique [39] at the Centro de Investigaciones Regionales ‘Dr. Hideyo Noguchi’. One gram of the content was homogenized in a centrifuge tube containing 10 ml of 10% formalin. After homogenization, 3 ml of ethyl acetate were added to the suspension in the tube and the resulting suspension was centrifuged at 1200 rpm for 3 min. Subsequently, the fatty pug was removed and the supernatant discarded. Finally, ~1 ml of saline solution was added to the sediment and three drops were transferred to a slide for examination. Hymenolepis eggs were measured and identified as H. diminuta by light microscopy [39].

Leptospira spp.

Leptospires in kidneys were detected at the Institute Gonçalo Moniz, Brazil, using the imprint method previously described [Reference Chagas-Junior40]. Briefly, we obtained kidney imprints by pressure of the cut surface of the tissue onto poly-L-lysine-coated glass slides. Slides were dried at room temperature and fixed in acetone for 3 min prior blocking with 1% bovine serum albumin (BSA) for 40 min. Then they were incubated for 1 h with a primary rabbit polyclonal anti-leptospiral antibody to Leptospira interrogans serovar Icterohaemorrhagiae strain RGA diluted 1:1000. Following three phosphate-buffered saline (PBS) washes, the slides were incubated for 1 h with goat anti-rabbit IgG Alexa 488conjugate (Invitrogen, USA) at a 1:500 dilution. After final washings, the slides were mounted with anti-fading medium (ProLong Molecular Probes, Thermo Fisher Scientific, USA) and examined for leptospires using fluorescent microscopy (Olympus BX51 microscope, Olympus America, USA) at a magnification ×400 and ×1000. Samples from non-infected laboratory rats and kidney-positive wild rats were similarly treated as negative and positive controls, respectively. Positive samples were determined by microscopic observation of intact leptospires.

Data analysis

Trap success (TS) was used to estimate the relative rodent abundance as follows: number of rats trapped × 100/(number of traps × number of nights) [Reference Gómez Villafañe41]. The non-parametric Mann–Whitney U-test was used to compare the TS between rodent species.

The proportion of positive animals was compared between species and sites, using a Fisher's exact test due to their low frequencies [Reference McDonald42]. In all statistical analyses, the level of significance was P < 0·05.

RESULTS

A total of 302 house mice and 161 black rats were trapped from the three sites (house mice: 159 in SJT, 80 in PAS and 63 in OPI; black rats: 38 in SJT, 109 in PAS and 14 in OPI). The house mouse was significantly more abundant, as suggested by the median trap success, in SJT (TS = 5·2%) than the black rat (TS: 1·1% , P = 0·005), whereas in PAS and OPI, the black rat (TS: 3·5% in PAS, 1·2% in OPI) and the house mouse (TS: 2·4% in PAS, 5·3% in OPI) had similar abundances (PAS, P = 0·093; OPI, P = 0·221). Table 1 shows the number and the percentage of trapped rodents tested for zoonotic pathogens.

Table 1. Number and percentage (in parenthesis) of house mice and black rats examined for zoonotic pathogens

Trypanosoma cruzi DNA was detected in 15 of 278 (5·4%) rodent hearts. The overall prevalence in black rats was 6·2% (7/113), whereas in-house mice were 4·9% (8/165) (Table 2). All blood samples tested by PCR were negative. Hymenolepis diminuta was the most prevalent pathogen among rodents (5·6%, 16/288). Black rats were more frequently infected with H. diminuta (14·2%, 15/106) than house mice (0·5%, 1/182) (Fisher's exact test, P < 0·001). Leptospires were detected only in 1 of 118 black rats (0·9%). A co-infection was detected in one individual, a black rat, carrying both T. cruzi and H. diminuta.

Table 2. Prevalence of zoonotic pathogens in house mice and black rats from Yucatan, Mexico

Data are presented as % positive (n positive/N analysed).

In SJT, 26·9% (7/26) of black rats were positive for T. cruzi, whereas in PAS only house mice were found positive (15·7%; 8/51) (Table 2). No animals from OPI were tested for this infection. There was a significant difference in the prevalence of infection with H. diminuta in rats and the site of trapping. The prevalence of SJT, 31·1% was higher than the 4·7% of PAS (Fisher's exact test, P = 0·001). There were no statistical differences between the prevalence of SJT and OPI (P = 0·723), and between OPI and PAS (P = 0·057). Hymenolepis diminuta eggs were found in a house mouse in OPI (1·2%, 1/52). The sole rat infected with Leptospira was trapped in PAS.

DISCUSSION

The house mouse and the black rat are a threat to public health; however, few studies have evaluated their role as carriers of zoonotic pathogens in urban and rural settlements of Mexico [Reference Torres-Castro31, Reference Panti-May34, Reference Panti-May43, Reference Torres-Castro44]. In this study, we report the presence of T. cruzi, H. diminuta and L. interrogans among house mouse and black rat populations from two urban neighbourhoods and a rural village from Yucatan, Mexico.

In this study, we detected the presence of T. cruzi in hearts of house mice and black rats, but not in blood samples. This suggests that rodents were in the chronic phase of the infection, which is characterized by a low parasitaemia and a high invasion of cardiac cells [Reference Andrade and Andrews45, Reference Zhang and Tarleton46]. Several studies have reported that synanthropic rodents are the main reservoir for T. cruzi in domestic and peridomestic cycles [Reference Herrera47]. Particularly, black rats had a high prevalence (27%), which has been noted in Brazil (24%), Chile (28%), Ecuador (12%) and Yucatan (47%) [Reference Lima13, Reference Pinto14, Reference Galuppo48, Reference Zavala-Velázquez49]. Some studies have suggested that the black rat could be a possible link between the domestic and sylvatic cycles of T. cruzi due to its synanthropic behaviour, its high reproductive rates and its preference to areas with trees [Reference Battersby, Hirschhorn, Amman, Bonnefoy, Kampen and Sweeney1]. On the other hand, the house mouse could be an important reservoir in the domestic cycle due to its preference to establish its colonies inside or close to the dwelling and its small home range (3–10 m) [Reference Battersby, Hirschhorn, Amman, Bonnefoy, Kampen and Sweeney1].

Hymenolepis diminuta was the most prevalent pathogen among rodents, particularly among black rats (14·2%). This parasite, which has a worldwide distribution, parasitizes mainly synanthropic rats of the genus Rattus [Reference Hancke and Suárez50]. This cestode has been reported in black rats from different habitats such as households [Reference Hancke and Suárez50], markets [Reference Mohd Zain, Behnke and Lewis51] and farms [Reference Milazzo52], with a prevalence varying from 14·3% to 33·3%. In this study, the prevalence among black rat populations varied from 4·7% (95% confidence interval (CI) 1–13·1%) in PAS to 23·1% (95% CI 5·0–53·8%) in OPI and 31·1% (95% CI 15·3–50·8%) in SJT. Hymenolepis diminuta requires an arthropod intermediate host to complete its life cycle. The main intermediate hosts are the mealworm beetle (Tenebrio molitor), the four beetle (Tribolium confusum) and the northern rat flea (Nosopsyllus fasciatus) [Reference Baker and Baker22]. The variation found in the prevalence could be related to the abundance of intermediated hosts in each area. Further studies investigating the species and abundance of intermediate hosts present in the studied sites could help us to understanding the epidemiology of H. diminuta in Yucatan.

The leptospiral carriage among R. rattus trapped in this work was low (0·9%). In a rural community close to the sampled neighbourhoods (San Jose Tecoh and Plan de Ayala Sur), L. interrogans was reported with a prevalence of 12·8% in R. rattus by PCR [Reference Torres-Castro31]. Although R. rattus has been reported as carrier of pathogenic Leptospira, in the Americas several studies have reported that R. norvegicus is the main reservoir in urban slums of Brazil, Colombia and Peru [Reference Johnson53Reference Costa55]. The low prevalence of Leptospira in R. rattus could be explained by the fact that R. rattus is an arboreal animal in contrast to R. norvegicus that is typically more terrestrial [Reference Battersby, Hirschhorn, Amman, Bonnefoy, Kampen and Sweeney1]. In Yucatan, there are no records of R. norvegicus, which suggests that R. rattus could be the main reservoir of Leptospira in absence of R. norvegicus as has been noted in some islands [Reference Foronda56, Reference Desvars57].

In this study we used different methodologies to detect different pathogens. PCR amplification of the 188 pb T. cruzi repetitive element is a highly sensitive technique for detecting small numbers of parasites, only a 1/200 of the DNA of the parasite is necessary for a positive identification [Reference Moser, Kirchhoff and Donelson38]. However, it is more applicable for acute infections than in chronically infected mammals; in chronically mammals, parasitaemias are intermittent and contain few or no parasites [Reference Moser, Kirchhoff and Donelson38]. On the other hand, T. cruzi lineage I, the predominant lineage in Mexico, has a tropism for the cardiac cells during the chronic phase of the infection [Reference Bosseno58], which indicates the utility of PCR for detection of tissue parasite in chronically infected hosts [Reference Herrera47]. The formalin–ether/ethyl acetate is a widely used sedimentation technique for the diagnosis of intestinal parasite eggs [39, Reference Navone59]. Its sensitivity ranging from 72% to 85%, depending on several factors such as the parasite species, the number of eggs/cysts per gram of faeces, and the time of infection [Reference Navone59]. For H. nana, this technique has shown a sensitivity ranging from 61% to 72% [Reference Navone59, Reference Steinmann60]. The immunofluorescent imprint method is a rapid technique for the direct observation of Leptospira spp. by microscopy. This method has been used to study experimental and natural infections [Reference Chagas-Junior40, Reference Costa61]. A comparative study with the real-time PCR (qPCR) showed that for the detection, the imprint method is equivalent to qPCR in both acute and chronic rodent models [Reference Chagas-Junior62]. Nevertheless, this method was restricted to the serovar Icterohaemorrhagiae, and consequently the prevalence of Leptospira could be underestimated. As the capacity to detect parasites is different between techniques, prevalence data of the three parasites are not comparable and may be compared only with studies using similar techniques.

Several studies have reported that changes in rodent demography, intermediate host populations and environmental factors could alter the risk of zoonotic pathogen transmission [Reference Costa61, Reference Himsworth63]. In this study, we found an overall low prevalence of zoonotic pathogens in rodent populations; however, previous ecological studies in Merida, Yucatan have shown that the reproductive rates of synanthropic rodents are high in low socioeconomic areas, which could increase the public health risks. Of the pathogens examined, T. cruzi and H. diminuta could represent a risk to inhabitants. Trypanosoma cruzi is a serious threat in Latin America due to the irreversible damage caused by the parasite, the low efficacy of the antiparasitic treatment during the chronic phase of the disease, and the presence of intrusive vectors, which lead to considerable morbidity and mortality rates [Reference Carabarin-Lima6]. Case reports of H. diminuta infection in humans are uncommon and are limited to rural and urban areas with high levels of poverty; however, in these areas, the environmental characteristics favour the abundance of rodents and intermediate hosts, facilitating the reinfection [Reference Martínez-Barbabosa24, Reference Marangi64]. Conversely, L. interrogans was the less prevalent pathogen among rodents. Further studies are required to assess whether humans are becoming infected within the studied sites. Our results suggest that the black rat could be an important reservoir for T. cruzi and H. diminuta in the studied sites. Nevertheless, both mice and rats live in close contact with inhabitants invading kitchens, bedrooms and consuming human foodstuff, which could increase the risk for a pathogen to be transmitted to inhabitants. It would be advisable to conduct further studies examining seasonal and geographical patterns. This could increase our knowledge on the epidemiology of these pathogens in Mexico.

ACKNOWLEDGEMENTS

We would like to thank the families from San Jose Tecoh, Plan de Ayala Sur and Opichen for their participation and cooperation in this research.

This work was funded by Consejo Nacional de Ciencia y Tecnología (grant numbers 2014-247005 and 2008-108929), the National Institutes of Health (R01 TW009504) and the Wellcome Trust (102330/Z/13/Z). J.A. Panti-May was supported by a doctoral grant from Consejo Nacional de Ciencia y Tecnología (grant no. 259164).

DECLARATION OF INTEREST

None.

ETHICAL STANDARDS

The authors assert that all procedures contributing to this work comply with the ethical standards of the relevant national and institutional guides on the care and use of laboratory animals.

References

1. Battersby, S, Hirschhorn, RB, Amman, BR. Commensal rodents. In: Bonnefoy, X, Kampen, H, Sweeney, K, eds. Public Health Significance of Urban Pests. Copenhagen: World Health Organization, 2008, pp. 387419.Google Scholar
2. Pimentel, D, Zuniga, R, Morrison, D. Update on the environmental and economic costs associated with alien-invasive species in the United States. Ecological Economics 2005; 52: 273288.CrossRefGoogle Scholar
3. Himsworth, CG, et al. Rats, cities, people, and pathogens: a systematic review and narrative synthesis of literature regarding the ecology of rat-associated zoonoses in urban centers. Vector Borne and Zoonotic Diseases 2013; 13: 349359.CrossRefGoogle ScholarPubMed
4. World Health Organization. Neglected tropical diseases (http://www.who.int/neglected_diseases/diseases/en/). Accessed 25 November 2016.Google Scholar
5. Sánchez-Montes, S, et al. Leptospirosis in Mexico: epidemiology and potential distribution of human cases. PLoS ONE 2015; 10: e0133720.CrossRefGoogle ScholarPubMed
6. Carabarin-Lima, A, et al. Chagas disease (American trypanosomiasis) in Mexico: an update. Acta Tropica 2013; 127: 126135.CrossRefGoogle ScholarPubMed
7. Mason, PR, Patterson, BA. Epidemiology of Hymenolepis nana infections in primary school children in urban and rural communities in Zimbabwe. Journal of Parasitology 1994; 80: 245250.CrossRefGoogle Scholar
8. Mirdha, BR, Samantray, JC. Hymenolepis nana: a common cause of paediatric diarrhoea in urban slum dwellers in India. Journal of Tropical Pediatrics 2002; 48: 331334.CrossRefGoogle ScholarPubMed
9. Thompson, RCA. Neglected zoonotic helminths: Hymenolepis nana, Echinococcus canadensis and Ancylostoma ceylanicum . Clinical Microbiology and Infection 2015; 21: 426432.CrossRefGoogle ScholarPubMed
10. Hotez, PJ, et al. An unfolding tragedy of Chagas disease in North America. PLoS Neglected Tropical Diseases 2013; 7: e2300.CrossRefGoogle ScholarPubMed
11. World Health Organization. Chagas disease in Latin America: an epidemiological update based on 2010 estimates. Weekly Epidemiological Record 2015; 90: 3344.Google ScholarPubMed
12. Waleckx, E, Gourbière, S, Dumonteil, E. Intrusive versus domiciliated triatomines and the challenge of adapting vector control practices against Chagas disease. Memorias do Instituto Oswaldo Cruz 2015; 110: 324338.CrossRefGoogle ScholarPubMed
13. Lima, MM, et al. Investigation of Chagas disease in four periurban areas in northeastern Brazil: epidemiologic survey in man, vectors, non-human hosts and reservoirs. Transactions of the Royal Society of Tropical Medicine and Hygiene 2012; 106: 143149.CrossRefGoogle ScholarPubMed
14. Pinto, CM, et al. Infection by trypanosomes in marsupials and rodents associated with human dwellings in Ecuador. Journal of Parasitology 2006; 92: 12511255.CrossRefGoogle ScholarPubMed
15. Ko, AI, Goarant, C, Picardeau, M. Leptospira: the dawn of the molecular genetics era for an emerging zoonotic pathogen. Nature Reviews Microbiology 2009; 7: 736747.CrossRefGoogle ScholarPubMed
16. Costa, F, et al. Global morbidity and mortality of leptospirosis: a systematic review. PLoS Neglected Tropical Diseases 2015; 9: e0003898.CrossRefGoogle ScholarPubMed
17. Athanazio, DA, et al. Rattus norvegicus as a model for persistent renal colonization by pathogenic Leptospira interrogans . Acta Tropica 2008; 105: 176180.CrossRefGoogle ScholarPubMed
18. Costa, F, et al. Influence of household rat infestation on Leptospira transmission in the urban slum environment. PLoS Neglected Tropical Diseases 2014; 8: e3338.CrossRefGoogle ScholarPubMed
19. Costa, F, et al. Surveillance for leptospirosis in the Americas, 1996–2005: a review of data from ministries of health. Revista Panamericana de Salud Publica 2012; 32: 169177.CrossRefGoogle ScholarPubMed
20. Nkouawa, A, et al. Cryptic diversity in hymenolepidid tapeworms infecting humans. Parasitology International 2016; 65: 8386.CrossRefGoogle ScholarPubMed
21. Edelman, MH, et al. Hymenolepis diminuta (rat tapeworm) infection in man. American Journal of Medicine 1965; 38: 951953.CrossRefGoogle ScholarPubMed
22. Baker, DG. Parasites of rats and mice. In: Baker, DG, ed. Flynn's Parasites of Laboratory Animals, 2nd edn. Iowa: Blackwell Publishing, 2007, pp. 303397.CrossRefGoogle Scholar
23. Chero, JC, et al. Hymenolepis nana infection: symptoms and response to nitazoxanide in field conditions. Transactions of the Royal Society of Tropical Medicine and Hygiene 2007; 101: 203205.CrossRefGoogle ScholarPubMed
24. Martínez-Barbabosa, I, et al. Infección por Hymenolepis diminuta en una estudiante universitaria. Revista Biomédica 2012; 23: 6164.Google Scholar
25. Bacigalupo, J. Infestation by Hymenolepis nana . Archivos Argentinos de Enfermedades del Apararato Disgestivo y Nutricion 1932; 7: 359364.Google Scholar
26. Luney, FW. Hymenolepis diminuta (rat tapeworm) in man. Canadian Medical Association Journal 1934; 30: 385386.Google ScholarPubMed
27. Guzman-Tapia, Y, Ramírez-Sierra, MJ, Dumonteil, E. Urban infestation by Triatoma dimidiata in the city of Mérida, Yucatán, México. Vector Borne and Zoonotic Diseases 2007; 7: 597606.CrossRefGoogle Scholar
28. Dumonteil, E, et al. Geographic distribution of Triatoma dimidiata and transmission dynamics of Trypanosoma cruzi in the Yucatan peninsula of Mexico. American Journal of Tropical Medicine and Hygiene 2002; 67: 176183.CrossRefGoogle Scholar
29. Vado-solís, IA, et al. Estudio de casos clínicos e incidencia de leptospirosis humana en el estado de Yucatán, México durante el período 1998 a 2000. Revista Biomédica 2002; 13: 157164.Google Scholar
30. Vado-Solís, I, et al. Clinical-epidemiological study of leptospirosis in humans and reservoirs in Yucatán, México. Revista do Instituto de Medicina Tropical Sao Paulo 2002; 44: 335340.CrossRefGoogle ScholarPubMed
31. Torres-Castro, MA, et al. First molecular evidence of Leptospira spp. in synanthropic rodents captured in Yucatan, Mexico. Revue de Medecine Veterinaire 2014; 7–8: 213218.Google Scholar
32. Duarte-Zapata, L, Escalante-Triay, F, López-Novelo de Ceballos, M. Prevalencia de parasitosis intestinal en población de clase media de la ciudad de Mérida. Gaceta Medica de Mexico 1984; 120: 193197.Google Scholar
33. Rodriguez-Pérez, MA, et al. Lessons from a study in a rural community from southern Mexico: risk factors associated to transmission and reinfection of gastrointestinal parasites after albendazole treatment. Research and Reports in Tropical Medicine 2011; 2: 147153.CrossRefGoogle Scholar
34. Panti-May, JA, et al. Infection levels of intestinal helminths in two commensal rodent species from rural households in Yucatan, Mexico. Journal of Helminthology 2015; 89: 4248.CrossRefGoogle ScholarPubMed
35. Instituto Nacional de Estadística y Geografía. Inventario nacional de viviendas (http://www3.inegi.org.mx/sistemas/mapa/inv/Default.aspx?bi=1). Accessed 25 May 2013.Google Scholar
36. Panti-May, JA, et al. Population characteristics of human-commensal rodents present in households from Mérida, Yucatán, México. Manter Journal Parasite Biodiversity 2016; 5: 16.Google Scholar
37. Leary, S, et al. AVMA Guidelines for the Euthanasia of Animals: 2013 Edition. Schaumburg, Illinois: American Veterinary Medical Association, 2013, p. 102.Google Scholar
38. Moser, DR, Kirchhoff, LV, Donelson, JE. Detection of Trypanosoma cruzi by DNA amplification using the polymerase chain reaction. Journal of Clinical Microbiology 1989; 27: 14771482.Google ScholarPubMed
39. World Health Organization. Bench Aids for the Diagnosis of Intestinal Parasites, 4th edn. Geneva: World Health Organization, 2012, p. 20.Google Scholar
40. Chagas-Junior, AD, et al. An imprint method for detecting leptospires in the hamster model of vaccine-mediated immunity for leptospirosis. Journal of Medical Microbiology 2009; 58: 16321637.CrossRefGoogle ScholarPubMed
41. Gómez Villafañe, IE, et al. Differences in population parameters of Rattus norvegicus in urban and rural habitats of central Argentina. Mammalia 2013; 77: 187193.Google Scholar
42. McDonald, JH. Handbook of Biological Statistics, 3rd edn. Baltimore, Maryland: Sparky House Publishing, 2014, p. 299.Google Scholar
43. Panti-May, JA, et al. Detection of Rickettsia felis in wild mammals from three municipalities in Yucatan, Mexico. Ecohealth 2015; 12: 523527.CrossRefGoogle ScholarPubMed
44. Torres-Castro, MA, et al. First molecular evidence of Toxoplasma gondii in synanthropic rodents (Mus musculus and Rattus rattus) captured in Yucatan, Mexico. Revue de Medecine Veterinaire 2016; 169: 250255.Google Scholar
45. Andrade, LO, Andrews, NW. The Trypanosoma cruzi–host-cell interplay: location, invasion, retention. Nature Reviews Microbiology 2005; 3: 819823.CrossRefGoogle Scholar
46. Zhang, L, Tarleton, RL. Parasite persistence correlates with disease severity and localization in chronic Chagas’ disease. Journal of Infectious Diseases 1999; 180: 480486.CrossRefGoogle ScholarPubMed
47. Herrera, CP, et al. Genotype diversity of Trypanosoma cruzi in small rodents and Triatoma sanguisuga from a rural area in New Orleans, Louisiana. Parasites & Vectors 2015; 8: 19.CrossRefGoogle ScholarPubMed
48. Galuppo, S, et al. Predominance of Trypanosoma cruzi genotypes in two reservoirs infected by sylvatic Triatoma infestans of an endemic area of Chile. Acta Tropica 2009; 111: 9093.CrossRefGoogle ScholarPubMed
49. Zavala-Velázquez, J, et al. Infection by Trypanosoma cruzi in mammals in Yucatan, Mexico: a serological and parasitological study. Revista do Instituto de Medicina Tropical de Sao Paulo 1996; 38: 289292.CrossRefGoogle ScholarPubMed
50. Hancke, D, Suárez, OV. Infection levels of the cestode Hymenolepis diminuta in rat populations from Buenos Aires, Argentina. Journal of Helminthology 2016; 90: 199205.CrossRefGoogle ScholarPubMed
51. Mohd Zain, SN, Behnke, JM, Lewis, JW. Helminth communities from two urban rat populations in Kuala Lumpur, Malaysia. Parasites & Vectors 2012; 5: 47.CrossRefGoogle ScholarPubMed
52. Milazzo, C, et al. Helminths and Ectoparasites of Rattus rattus and Mus musculus from Sicily, Italy. Comparative Parasitology 2003; 70: 199204.CrossRefGoogle Scholar
53. Johnson, MAS, et al. Environmental exposure and leptospirosis, Peru. Emerging Infectious Diseases 2004; 10: 10161022.CrossRefGoogle ScholarPubMed
54. Agudelo-Flórez, P, et al. Prevalence of Leptospira spp. in urban rodents from a groceries trade center of Medellín, Colombia. American Journal of Tropical Medicine and Hygiene 2009; 81: 906910.CrossRefGoogle ScholarPubMed
55. Costa, F, et al. Patterns in Leptospira shedding in Norway rats (Rattus norvegicus) from Brazilian slum communities at high risk of disease transmission. PLoS Neglected Tropical Diseases 2015; 9: e0003819.CrossRefGoogle ScholarPubMed
56. Foronda, P, et al. Pathogenic Leptospira spp. in wild rodents, Canary Islands, Spain. Emerging Infectious Diseases 2011; 17: 17811782.CrossRefGoogle ScholarPubMed
57. Desvars, A, et al. Similarities in Leptospira serogroup and species distribution in animals and humans in the Indian Ocean island of Mayotte. American Journal of Tropical Medicine and Hygiene 2012; 87: 134140.CrossRefGoogle ScholarPubMed
58. Bosseno, M-F, et al. Predominance of Trypanosoma cruzi Linage I in Mexico. J Clin Microbiol. 2002; 40: 627632.CrossRefGoogle Scholar
59. Navone, GT, et al. Estudio comparativo de recuperación de formas parasitarias por tres diferentes métodos de enriquecimiento coproparasitológico. Parasitologia Latinoamericana 2005; 60: 178181.Google Scholar
60. Steinmann, P, et al. FLOTAC for the diagnosis of Hymenolepis spp. infection: proof-of-concept and comparing diagnostic accuracy with other methods. Parasitology Research 2012; 111: 749754.CrossRefGoogle Scholar
61. Costa, F, et al. Infections by Leptospira interrogans, Seoul virus, and Bartonella spp. among Norway rats (Rattus norvegicus) from the urban slum environment in Brazil. Vector Borne and Zoonotic Diseases 2014; 14: 3340.CrossRefGoogle Scholar
62. Chagas-Junior, AD, et al. Detection and quantification of Leptospira interrogans in hamster and rat kidney samples: immunofluorescent imprints versus real-time PCR. PLoS ONE 2012; 7: 15.CrossRefGoogle ScholarPubMed
63. Himsworth, CG, et al. An investigation of Bartonella spp., Rickettsia typhi, and Seoul Hantavirus in rats (Rattus spp.) from an inner-city neighborhood of Vancouver, Canada: is pathogen presence a reflection of global and local rat population structure? Vector Borne and Zoonotic Diseases 2015; 15: 2126.CrossRefGoogle ScholarPubMed
64. Marangi, M, et al. Hymenolepis diminuta infection in a child living in the urban area of Rome, Italy. Journal of Clinical Microbiology 2003; 41: 39943995.CrossRefGoogle Scholar

Altmetric attention score

Full text views

Full text views reflects PDF downloads, PDFs sent to Google Drive, Dropbox and Kindle and HTML full text views.

Total number of HTML views: 285
Total number of PDF views: 524 *
View data table for this chart

* Views captured on Cambridge Core between 10th July 2017 - 17th April 2021. This data will be updated every 24 hours.

You have Access

Send article to Kindle

To send this article to your Kindle, first ensure no-reply@cambridge.org is added to your Approved Personal Document E-mail List under your Personal Document Settings on the Manage Your Content and Devices page of your Amazon account. Then enter the ‘name’ part of your Kindle email address below. Find out more about sending to your Kindle. Find out more about sending to your Kindle.

Note you can select to send to either the @free.kindle.com or @kindle.com variations. ‘@free.kindle.com’ emails are free but can only be sent to your device when it is connected to wi-fi. ‘@kindle.com’ emails can be delivered even when you are not connected to wi-fi, but note that service fees apply.

Find out more about the Kindle Personal Document Service.

A survey of zoonotic pathogens carried by house mouse and black rat populations in Yucatan, Mexico
Available formats
×

Send article to Dropbox

To send this article to your Dropbox account, please select one or more formats and confirm that you agree to abide by our usage policies. If this is the first time you use this feature, you will be asked to authorise Cambridge Core to connect with your <service> account. Find out more about sending content to Dropbox.

A survey of zoonotic pathogens carried by house mouse and black rat populations in Yucatan, Mexico
Available formats
×

Send article to Google Drive

To send this article to your Google Drive account, please select one or more formats and confirm that you agree to abide by our usage policies. If this is the first time you use this feature, you will be asked to authorise Cambridge Core to connect with your <service> account. Find out more about sending content to Google Drive.

A survey of zoonotic pathogens carried by house mouse and black rat populations in Yucatan, Mexico
Available formats
×
×

Reply to: Submit a response


Your details


Conflicting interests

Do you have any conflicting interests? *