Summary

Fleas (Siphonaptera) are important vectors of several animal and human disease pathogens, while rodents are considered as reservoirs of most pathogens, including Yersinia pestis Factors that influence the parasitism rate of fleas, ecological aspects that modulate their distribution, and host-flea relationship in Eastern Zambia remain unknown.  Furthermore, there is little information on the biodiversity and abundance of rodents and fleas in the study area. A total of 1212 mammals were sampled and examined. These included rodents (n=329), Crocidura (n=113), domestic pigs (n=254), small ruminants (n=346) and carnivores (n=168), and 1578 fleas, where five species were identified. There were nine genera and species of rodents with one genus of Crocidura captured. The results showed that 27(8.2%) and 19(5.8%) rodents and 8(7.0%) and 2(1.8%) Crocidura were positive for antibodies and pla gene for Y. pestis, respectively. Echidnophaga larina were the most mean abundant (MA=8.58), while Xenopsylla cheopis had the least mean abundant (MA=0.14), nevertheless it was the most infected with Y. pestis. Mastomys. natalensis was highest in plague positivity 31/56, followed by Crocidura spp 10/56 and Rattus rattus 6/56. The results indicated that three flea species were infected with Yersinia pestis. Shannon-Weiner (H) and dominance (D) indices of rodents were 1.5 and 0.2789, while the flea indices were 0.5310 and 0.8389, respectively. There was a strong association between richness of fleas and plague disease (p=0.01; x²=65.3). It’s established that rodents were more biodiversity than fleas while both were unevenly distributed. It’s recommended that control measures of fleas be intensified and sustained to lessen the spread of their associated diseases.

Keywords: Biodiversity Crocidura, Fleas, Plague, Rodents

Introduction

Fleas (Insecta: Siphonaptera) encompasses at least 2500 species with 15 subspecies globally (Whiting et al., 2008). They are small, laterally flattened and wingless, and highly specialized arthropods. They are of great importance as vectors of many viral, bacterial, and parasitic pathogens. Being hematophagous arthropods, fleas may transmit pathogens through soiled mouthparts (e.g., Y. pestis, and viral pathogens), regurgitation of gut contents (e.g., Y. pestis), and infectious saliva (e.g., R. felis in salivary glands) (Eisen et al., 2012). Also, transmission can occur through contaminated feces (e.g., Rickettsia typhi, Bartonella,hanselae) (Amatre et al., 2009) (Bitam et al., 2010). Fleas have complex life cycles, which include the production of non-parasitic omnivorous larvae and obligate-hematophagous imagoes (Sanchez and Lareschi, 2019). Once the flea emerges from the cocoon, it immediately seeks a potential host to get a blood meal. Most fleas prefer the host habitat (nest fleas), while others stay more or less permanently on the body of the host itself (body fleas) (Bitam et al., 2010).

Fleas have tremendous medical and economic importance because they are vectors of several causative agents of diseases in animals and humans.  Such diseases as  the cat scratch fever  caused by B. henselae, Q fever (Coxiella burnetii), murine typhus (R. typhii), flea-borne spotted fever (R. felis),and bubonic plague (Y. pestis) (Bitam et al., 2010). Due to the potential vector capacity of fleas, their distribution and biodiversity may have essential consequences for host survival and disease dynamics (Foottit and Galloway, 2018). Arthropod-transmitted diseases represented approximately 17% of all infectious diseases globally and are influenced by a complex and dynamic ecosystem that involves vectors, hosts, and infectious agents, as well as environmental factors(Young et al., 2015). Fleas are able  to bite animals, suck blood and mechanically or biologically transmit disease pathogens such as Yersinia pestis to susceptible hosts, including humans and rodents (Ago et al., 2015). The abundance of fleas in an area may suggest an increased number or frequency of flea-borne diseases. This happens when fleas are numerous and consequently look for animals and humans to suck blood and subsequently transmit disease-causing pathogens (Nziza et al., 2019; Miarinjara and Boyer, 2016).

Rodents are small mammals of the order Rodentia, which comprises more than 2000 species and approximately 30 families. There are diverse biological and ecological differences among rodents in shape, size, weight, and habitat. The smallest rodents (Mus minutoide) may weigh 5 grams, while the largest (American cabybard) weighs more than 70 kg. Rodent's ecological requirements vary widely, which include domestic and semi-domestic field and forest habitats. These animals are potential reservoirs of infectious diseases of humans and other mammals and are also suitable hosts for flea and other ectoparasites (Esfandiari et al., 2017).

Shrews are insectivores and able to harbor disease pathogens, such as bacteria, virus, and protozoa. These disease pathogens could be transmitted to a suitable host and cause disease through the vector such as fleas (Moore et al., 2015).

The abundance of rodents and fleas depends on climatic and environmental conditions. Climate changes caused by global warming and human intervention have contributed to vicissitudes in the biological parameters and distribution ranges of vector/fleas and their hosts (Kausrud et al., 2007). These conditions are likely to affect the biodiversity and abundance or dominance of both rodents and their flea ectoparasites. At times of heavy rainfall and availability of food crops, there are corresponding large numbers of rodents and their fleas, contrary to times of adverse climate when the rodents and their fleas diminish in number. Rodents have the potential to transmit sylvatic plague and other zoonotic diseases directly to humans and other animals. Such diseases can be transmitted by contamination of foods with rodent fecal materials or urine, direct contacts with infected animals, droplets, or bites by appropriate insect vectors such as fleas in case of plague disease (Backhans and Fellström, 2012). These vectors then transmit the pathogens from one reservoir host to another susceptible animal, including humans, small carnivores, and other animals.  In view of the aforegoing description of the relationship between rodents and fleas regarding plague epidemiology, it was felt desirable to investigate and establish the current status of biodiversity and abundance of these creatures in relation to plague epidemiology; therefore the aim of the study was to accomplish this desirability in the eastern region of Zambia.

Materials and methods

Area and time of study

This study took place in two districts, Nyimba and Sinda, both in the eastern province of Zambia from 2013 to 2017. The choice of these areas was based on their recent history of plague outbreaks, whereby the last human plague was detected in Nyimba and Sinda districts in 2015 and 2008, respectively. The eastern Zambia districts experience high rainfall between January and March of each year. The communities grow crops such as maize, groundnuts, cassava, which are all suitable for rodents and other animals.

Study design

A cross-sectional study design was conducted, where the abundance and biodiversity of the rodents, shrews, and flea ectoparasites were examined.

Sample collection

Domestic animals

Villages in the study area were randomly selected in which 20 or more domestic animals of the selected species, comprising pigs, dogs, cats, goats, and sheep were sampled. From each selected client, a verbal consent was acquired (as per approved Plague Research policy provided in Assurance No. FWA0000338).  Each selected animal was assigned a unique identification number. The first animal was randomly selected, followed by a systematic technique (Systematic Random Sampling). The selected animal was immobilized and rested on the white plastic sheet for blood collection and inspection of fleas and other ectoparasites. The animal was groomed with cotton wool containing 90% diethyl ether to sedate the ectoparasites. The appropriate brush was used to scrub the animal to detach fleas and other ectoparasites from its fur and skin. A pair of forcepswas used to gently removed fleas that fell onto the white sheet and those which remained attached on the animal’s fur/skin into vials with 70% ethanol. The study did not include animals that came from other villages in the past six months (Nyirenda et al., 2017).

Rodents and shrews trapping

The study villages were selected randomly and were alienated into six arbitrary sectors from where three zones were randomly nominated for trapping rodents and shrews. Sherman’s live traps (50×65×157 mm) were baited with peanut butter mixed with soya beans flour and were placed at a distance of 10m apart in nearby bushes overnight. Wire cage traps (145×100×230 mm) (Hoga-lab, Kyoto, Japan), baited with fish, Stolothrissa tanganicae (Kapenta), and ripe tomatoes, were set in chosen houses in the zones. Inspection of traps was conducted the following morning. The captured animals were taken for flea ectoparasite inspection and collection in a mobile laboratory installed at a distant place away from the community to collect sera, organs, and fleas. Other processes included identification and enumeration of rodents and fleas were carried out in such a laboratory. The ensnaring process was continuous for three consecutive days in the same area as previously described (Kilonzo, 1976). Sera and organs were kept at -20^oC until needed for ELISA and PCR, respectively (Nyirenda et al., 2018).

Collection of flea ectoparasites from rodents and shrews

Fleas and other ectoparasites from the rodents and shrews were collected by introducing the latter into a plastic bag with a soaked cotton wool, in 90% diethyl ether to sedate both the animal and fleas. Once the animals were sedated, they were placed into a silver basin and brushed thoroughly with a hard toothbrush to disengage the ectoparasites. Fleas that dropped into the basin were gently picked using either a pair of forceps or fine camel brush into vials with 70% ethanol as a preservative. Other ectoparasites were also collected into separate vials with the same concentration of ethanol (Nyirenda et al., 2018).

Rodent identification

All the captured rodents were identified to genus and/or specie level using the key features described by Kingdon (Kingdon, 1974). In addition, the body and tail lengths were measured and recorded. The carcasses were then counted, and numbers of each species were established.

Flea processing and identification

The fleas were preliminary identified and pooled (1–5) according to their species, location of the host, and species of its host. One to two (1–2) fleas were removed and processed from each pool to confirm the identity. The fleas were processed and identified to specie level using the method described by Kilonzo (Kilonzo, 1999), which basically involve drying the insects on the No. 1 filter paper, boiling them for 10 minutes in heat resistant tubes containing 10% Sodium Hydroxide. The fleas were then put in cold water for one hour and into acidified water (water + glacial acetic acid of equal volume) for 30 minutes to neutralize Sodium Hydroxide. After this, fleas were dehydrated through increasing strengths of ethanol (from 50%, 70%, 95%, and absolute alcohol) for one hour at each stage. After this process, the fleas were transferred into the vial  with  clove oil and incubated at room temperature overnight or until they were clear and transparent, after which they were mounted with Dibutylphthalate Polystyrene Xylene (DPX) on the glass slide with a coverslip for examination under the light microscope using dry objectives (x4, x10 or x40) magnification for detection of common taxonomic features (Kilonzo, 1999).

Identification of the fleas was based on  main flea features such as pronotal combs, genal combs, and shape of the head and reproductive organs (spermathecae in females and penis plates in males (Nyirenda et al., 2018; Kilonzo, 1999; Pratt and Stojanovich, 1966; Traub, 1962).

DNA extraction from organs and fleas

The DNA from tissues of rodents was extracted using the DNA extraction kit (ZR genomic DNA-Tissue Mini-Prep Catalog No. D3051) following the manufacturer’s instructions (Zymo Research Irvine, CA, USA). In the case of fleas, these were identified before preparing them for DNA extraction using the heat treatment method as described elsewhere (Nyirenda et al., 2017).

PCR Technique

PCR was performed using Phusion flash high fidelity master mix (Finnzymes Oy, Finland) in a highly PCR specialized laboratory. The primers Yp pla1 (5'TGC TTT ATG ACG CAG AAA CAG G3') as the forward primer and Yp pla2 (5'CTG TAG CTG TCC AAC TGA AAC G3') as the reverse primer, that amplifies a 344-bp region spanning residues 425 to 769 of the plasminogen activator gene, were used (Sodeinde and Goguen, 1989). The PCR reactions were performed as described elsewhere (Nyirenda et al., 2018) (Nyirenda et al., 2017). The positive and negative controls, which came together with the kit, were also included in the test.

ELISA testing

The stored sera were removed from the freezer (−20 °C) and left at room temperature to thaw and processed using the protocol as previously described (Chu, 2000).

Data analysis

Data were entered in Microsoft Excel software and analyzed using Epi info^TM 7.0.8.0, a computer statistical package from the Centre for Disease Control and Prevention (CDC), where confidence intervals (CI) and positive percentages were generated. The p-value and the chi-square (x²) were also calculated.

Flea diversity was estimated by calculating the Specific richness (S=number of flea species) and Shannon diversity species index (H = -Σpi ln pi), where pi =proportion of each flea species in the sample). Flea evenness was estimated by calculating the Simpson dominance index (D = Σp²i), where pi is the proportion of individuals found in species ‘i' as calculated elsewhere (Sanchez and Lareschi, 2019; Ralaizafisoloarivony et al., 2014).

For each host species, the following indices and parameters were calculated: flea species richness (S=number of flea species), Simpson index (Mean Abundance (MA) = total number of individuals’ of a parasite species on a host/total number of host species, including infested and non-infested species), and prevalence ([P = number of infested animals with one or more individuals of a parasite species/total number of examined mammals for that parasite species] multiplied by 100).

Results

In this study, 1212 mammals were examined for flea vectors, which comprised rodents (n=329), shrews (n=113), domestic pigs (n=254), small domestic ruminants (n=364), and domestic carnivores (n=168) were examined for fleas. A total of 1578 fleas were collected from such animals. The rodent population composed of nine genera and species, while all the shrews belonged to the genera of Crocidura spp. (n=113) The rodent species included: Mastomys natalensis (n=189), Rattus rattus (n=60), Saccostomus spp (pouched mouse (n=43), Tatera spp (n=2), Graphiurus spp (n=5), Mus spp (n=2), Acomys spp (n=1), Gerbillurus (Gerbil) spp (n=22) and Steastomys parvus (fat mouse) (n=5). On ELISA and PCR, 27 (8.2%) and 19 (5.8%) were positive for Yersinia pestis antibodies and pla gene, respectively, while 8 (7.0%) and 2 (1.8%) shrews were also positive for Yersinia pestis antibodies and pla genes, respectively. Five species of fleas collected from the animals in the study area were; X. cheopis (n=59), C. canis (n=64), E. gallinacea (n=389), E. larina (n=1064),and C. felis (n=2).The results also showed E. gallinacea had the highest mean abundance (MA) of the (8.58), while X. cheopis had the lowest (MA=0.14)in both districts (Table 1). It was revealed that Mastomys natalensis (31/56) was the highest-ranking rodent species carrying Y. pestis. Mus spp, Acomy spp, and Steatomys parvus were negative for Y. pestis on both tests. Statistical analysis presented no significant statistical difference between the rodents and plague disease (p = 0.347; x²=10) (Table 2).