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All issues Volume 25 (2018) Parasite, 25 (2018) 44 Full HTML
Open Access
Issue
Parasite
Volume 25, 2018
Article Number 44
Number of page(s) 8
DOI https://doi.org/10.1051/parasite/2018044
Published online 17 August 2018

© S. Kanté Tagueu et al., published by EDP Sciences, 2018

Licence Creative CommonsThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Introduction

Tsetse flies are the cyclical vector of most trypanosome species that cause human and animal African trypanosomiasis. Two species of Trypanosoma brucei s.l. are responsible for human African trypanosomiasis (HAT): Trypanosoma brucei gambiense causes the chronic form of HAT in Western and Central Africa, while Trypanosoma brucei rhodesiense is responsible for the acute form of HAT in East Africa. The third subspecies, Trypanosoma brucei brucei, is not implicated in human infection but causes African animal trypanosomiasis (AAT), also called nagana. In addition to Trypanosoma brucei brucei, Trypanosoma congolense, Trypanosoma vivax, Trypanosoma evansi and Trypanosoma simiae also cause AAT. In Africa, the economic losses resulting from the negative impact of AAT on African agriculture are estimated to be higher than US$ 4.5 billion/year [1, 36]. Moreover, African farmers spend about 35 million dollars per year on trypanocidal drugs to protect and cure their cattle [5]. If African trypanosomiases were controlled, about 7 million km2 of tsetse infested area could be suitable for livestock and agriculture in Africa [30].

For HAT and AAT, very few drugs are available and resistance phenomena have been observed for some of them [3, 18]. To prevent trypanosome infections, no vaccine is available and several approaches have been investigated in order to improve vector control. It is in this light that in depth investigations targeting bacterial flora of tsetse flies have been undertaken in the last few decades. Indeed, tsetse flies harbor three symbionts including the obligate primary symbiont (essential) Wigglesworthia glossinidia [43], the secondary (non-essential) symbiont Sodalis glossinidius [7], and the third symbiont (non-essential) known as Wolbachia [33]. The secondary and facultative symbionts S. glossinidius [7] are enterobacteria, which are widely spread in numerous tissues of the fly [2]. They are suited as paratransgenic organisms due to their ability to survive in the same organs with trypanosomes [6]. Sodalis glossinidius affect host longevity and may influence the host’s ability to establish trypanosome infections [8]. Given their close association with their host’s biology and their large tissue tropism, S. glossinidius could be used to produce and deliver some specific molecules (antibodies) by expressing foreign genes designed to block pathogen development [10].

In some HAT foci of southern Cameroon, the presence of S. glossinidius has been reported to favor trypanosome infections [12]. However, the effect of S. glossinidius on trypanosome infections could depend on the trypanosome genotype [12, 15]. Moreover, the association between S. glossinidius and trypanosome infections could vary according to sampling areas, since the environmental conditions could affect the life history traits of tsetse flies as well as the association with their symbiotic microorganisms.

In this study, S. glossinidius and different trypanosome species were identified in Glossina palpalis palpalis caught in the Fontem sleeping sickness focus of the southern Cameroon, with the overarching goal of improving our understanding of the association between S. glossinidius and trypanosome infections.

Methods

Ethical statement

This study was carried out following the strict recommendations contained in the Guide for the Care and Use of Animals of the Department of Biochemistry of the University of Dschang.

Study area

The Fontem HAT focus (5°40′12 N, 9°55′33E) is located in the Lebialem division of the Southwest region of Cameroon. In this forest region, the climate is of the tropical moist type with a relief made up of hills and valleys. The region is crossed by many fast-moving streams. The main activity in the Fontem HAT focus is agriculture and breeding of small livestock and poultry. In addition to wild animals, several domestic animal species including dogs, pigs, sheep, and goats are found in this focus. For this study, the entomological surveys were performed in four villages of the Fontem HAT focus including Bechati, Folepi, Besali and Menji (Figure 1).

thumbnail Figure 1.

Map showing villages where entomological surveys where undertaken in the Fontem sleeping sickness focus. Stars: Villages where tsetse flies were trapped; circles: other villages. The road from Mamfé to Dschang is indicated in black.

Sampling of tsetse

Two entomological surveys were conducted in four villages (Figure 1) of the Fontem HAT focus using 30 traps in February 2015 and 12 traps in November 2016. In each of these villages, pyramidal traps [19] were set up for four consecutive days in various tsetse fly-favorable biotopes. The geographical coordinates of each trap were recorded with a global positioning system. In each village, tsetse flies were collected twice a day (from 9 to 10 am and from 3 to 4 pm). The collected flies were identified and numbered according to traps. Thereafter, the flies were sorted into teneral (young flies that had never taken a blood meal) and non-teneral flies. Each identified tsetse fly was subsequently put into a microtube containing ethanol at 95%. The microtubes were maintained at room temperature in the field. In the laboratory, they were stored at −20 °C until use.

DNA extraction

DNA was extracted from whole tsetse fly using the cetyl trimethyl ammonium bromide (CTAB) method as described by Navajas et al. [29]. Briefly, the alcohol used to preserve each fly was evaporated by incubating the opened microtubes containing whole fly at 80 °C in an oven for about 1 h. Thereafter, each fly was disrupted with a pestle in CTAB buffer (CTAB 2%; 1 M Tris, pH 8; 0.5 M EDTA pH 8; 5 M NaCl). The disrupted tissues were incubated at 60 °C for 30 min before the addition of chloroform/isoamylic alcohol mixture (24/1; V/V). DNA was precipitated by addition of isopropanol (V/V) and a centrifugation at 13,000 rpm for 15 min. The DNA pellets were washed twice with 70% cool ethanol and then dried at room temperature. DNA pellets were finally re-suspended in 50 μL of sterile water before their storage at −20 °C until use.

Detection of S. glossinidius

The presence of S. glossinidius was revealed by PCR with pSG2 direct (5′-TGAAGTTGGGAATGTCG-3′) and reverse (5′-AGTTGTAGCACAGCGTGTA-3’) primers as described by Darby et al. [9]. The PCR reactions were carried out in a DNA thermal cycler (Prime). Each amplification reaction was performed in a total volume of 25 μL containing 20 pmol of each primer, 2.5 μL of 10× reaction buffer, 2 mM of MgCl2, 200 mM of each dNTPs, 4 μL of DNA template, and 0.5 units of Taq DNA polymerase (New England Biolab 5 U/μL). The amplification reactions involved a denaturation step at 94 °C for 5 min followed by 40 amplification cycles made up of a denaturation step at 94 °C for 30 s, an annealing step at 56 °C for 30 s, and an extension step at 72 °C for 45 s. These amplifications were followed by a final extension step at 72 °C for 5 min. The amplified products were resolved by electrophoresis at 100 volts for 30 min on 2% agarose gel containing ethidium bromide. DNA bands were visualized under ultraviolet light.

Detection of trypanosomes

Four sets of specific primers (Table 1) were used to identify Trypanosoma brucei s.l., Trypanosoma vivax, and Trypanosoma congolense “forest” and “savannah” types. This identification was done by PCR as described by Herder et al. [16]. For this identification, each PCR reaction was carried out in a final volume of 15 μL containing 1.5 μL of 10× PCR reaction buffer, 1.5 mM of MgCl2, 200 mM of each dNTP, 10 picomoles of each primer (Table 1), 0.3 units of Taq DNA polymerase (New England Biolab 5 U/μL) and 3 μL of DNA extract. For each amplification reaction, a denaturation step at 94 °C for 5 min was followed by 40 amplification cycles. Each of these cycles included a denaturation step at 94 °C for 30 s, an annealing step for 30 s at 60 °C for T. brucei s.l., T. vivax, and T. congolense “forest” and “savannah” types, and an extension step at 72 °C for 1 min. A final extension step was performed at 72 °C for 10 min. The amplified products were separated on 2% agarose gel containing ethidium bromide and visualized under UV illumination.

Table 1.

Primers used for the identification of different trypanosome species.

Statistical analysis

The statistical analyses were performed using StataCorp 2015 statistical software, release 14 (StataCorp LP; College Station, TX, USA). Chi-squared tests were used to compare the infection rates of S. glossinidius and different trypanosome species between villages. The differences were considered significant when the p-values were lower than 0.05. To see whether the presence of S. glossinidius could favor trypanosome infections, a generalized linear model using StataCorp 2015 software with 95% confidence intervals (CIs) was used. For these analyses, T. vivax was excluded because its lifecycle is exclusively completed within the mouthparts of the tsetse fly.

Results

Entomological surveys

During the two entomological surveys, 274 tsetse flies were collected. Details regarding results of entomological surveys are reported in Table 2. Of the 274 tsetse flies collected, 9 (3.28%) teneral flies were identified (Table 2). No teneral flies were identified at Besali. The mean apparent fly density per trap per day (ADT) varied from 0.2 to 2.79, with an average of 1.63. The highest ADT was recorded at Folepi.

Table 2.

Results of entomological surveys and infection rates of S. glossinidius according to villages

Molecular identification of S. glossinidius

Of the 274 flies analyzed, 96 were positive for S. glossinidius, yielding an overall infection rate of 35.04%. The highest infection rate of 39.44% [95% CI = 27.96% – 50.4%] was observed at Menji, and the lowest rate of 25% [95% CI = 3.35% – 76.22%] at Besali. Despite the variations observed in the infection rates, no significant difference (p-value: 0.5801) was observed between villages (Table 2).

Molecular detection of different trypanosome species

Of the 265 non-teneral tsetse flies analyzed, 100 (37.73%) were infected by at least one trypanosome species. Of these 100 infected flies, 69 (26.04%) were infected due to T. congolense “forest type”, 48 (18.11%) to T. vivax, and 17 (6.41%) were mixed infections (Table 3). No infections due to T. brucei s.l. were observed. The highest infection rates of 40% [95% CI = 29.24% – 51.82%] for T. congolense and 25% [95% CI = 3.35% – 76.22%] for T. vivax were observed at Menji and Besali, respectively. The lowest infection rates of 16.98% [95% CI = 9.08% – 29.53%] for T. congolense and 15.94% [95% CI = 10.73% – 23.03%] for T. vivax were observed at Bechati and Folepi, respectively. For T. congolense “forest type”, a significant difference was found between villages. For T. vivax, no significant difference was found between villages. Of the 100 tsetse flies with trypanosome infections, 17 (17%) were co-infected by T. vivax and T. congolense “forest type”, yielding an overall co-infection rate of 6.41% (17/265). Between villages, significant differences (p-value: 0.0454) were observed in the co-infection rates (Table 3).

Table 3.

Trypanosome infections according to villages.

Co-infection of trypanosomes and S. glossinidius

Of the 265 tsetse flies that were simultaneously analyzed for the presence of trypanosomes and S. glossinidius, 92 (34.72%) harbored S. glossinidius and 100 (37.73%) were infected by at least one trypanosome species. The number of tsetse flies with trypanosome infections is higher than the number of flies with S. glossinidius. Considering the fact that T. vivax is found exclusively in the mouthparts, the 21 tsetse flies with only T. vivax infections were excluded from the analyses performed here. As a consequence, only 69 flies with T. congolense “forest type” were considered during investigations on the association between S. glossinidius and trypanosome infections. Of the 69 flies infected by T. congolense “forest type”, 23 (33.33%) also harbored S. glossinidius (S + Tcf +), while the remaining 46 (66.67%) were without S. glossinidius (S-Tcf +) (Table 4). About 71.86% (69/96) of flies harboring S. glossinidius were not infected by trypanosomes. The analyses performed to see if the presence of S. glossinidius could have an impact on the trypanosome infections (Tcf +) revealed no significant association (r = −0.0831; p = 0.7785; [95% CI = −0.66 – 0.5]) between these two micro-organisms (Table 4).

Table 4.

Sodalis glossinidius and T. congolense co-infections according to villages.

Discussion

We carried out several studies on tsetse flies to understand their biology and their bacterial flora, and also to identify the parasites infecting these flies. S. glossinidius and different trypanosome species were investigated in tsetse flies caught in the Fontem HAT focus of Cameroon, with the overarching goal of improving our knowledge on the vector competence of tsetse flies. Results of entomological surveys confirm G. p. palpalis as the only tsetse species in this focus. They are in agreement with previous observations [26, 32], highlighting the role of G. p. palpalis in the transmission of African trypanosomiases in the Fontem HAT focus. The apparent density of the tsetse per trap per day (ADT) of 1.63 is very low when compared to 7.9 and 4.85 obtained 20 (1998) and 10 (2007) years ago in the same area by Morlais et al. [26] and Njitchouang et al. [32]. This decrease of ADT could be linked to local disturbance resulting more likely from bush clearing and population growth which has induced climatic and environmental modifications that affected tsetse biotopes. These modifications occurred with time and subsequently, have induced some changes in the composition and host availability, the nutritional behavior of tsetse, and the transmission dynamics of trypanosomes [28, 38, 41].

The presence of S. glossinidius in G. p. palpalis caught in Cameroon confirms results obtained in two other HAT foci of the forest regions of southern Cameroon [12]. The S. glossinidius infection rate of 35.04% obtained here is lower than the 64.4% and 45.3% reported in G. p. palpalis caught at the Bipindi and Campo HAT foci of Cameroon, which are located more 400 km from the Fontem HAT focus [12]. This is higher than the 9.3% reported in Liberia for the same tsetse species [24]. These differences could be linked to sampling areas since each area is characterized by specific environmental factors that affect tsetse biology as well as the symbiotic association, and subsequently the vertical transmission of S. glossinidius from mother to offspring. When environmental factors are stable, like in insectariums, the transmission rate of symbiotic micro-organisms from mother to offspring is quite high [37]. For colonies of G. p. gambiense and G. m. morsitans from Burkina Faso that were maintained in insectariums, the infection rates of S. glossinidius reached 100% [14].

Comparing the S. glossinidius infection rates between different tsetse species [20, 24, 42], the high variation observed could be explained by the intrinsic characters of each tsetse species. For the same stimuli (internal or external), tsetse species will respond differently (differential behaviors) because of their specific biological characters that induce variations in the molecular interactions between tsetse and its symbiotic microorganisms and consequently, in the infection rates of different symbionts. The high variation reported above could also result from certain differences in analytical methods. In our study for instance, whole tsetse fly was used while in previous studies, tsetse flies were dissected and investigations were performed on isolated tissues.

The identification of T. congolense and T. vivax confirms results obtained in the same area [26, 40]. In the same villages, these trypanosomes have previously been detected in tsetse and different domestic animals; indicating their active transmission. Our results corroborate those reported in western, eastern and central African where the same trypanosome species were detected in different tsetse species [11–13, 23, 26, 34, 35]. They are also in agreement with results obtained in a variety of wild and domestic animals despite the fact that none of these animals were investigated in this study [17, 31, 39, 40]. This wide distribution of T. vivax and T. congolense indicates their ubiquity and the presence of appropriate vertebrate hosts. The high infection rate of T. congolense forest “type” is linked to the geographical localization of the Fontem HAT focus because this species is mainly found in the forest regions. It can also be explained by the fact that whole tsetse fly were analyzed, and the infection rates reported here are the combination of infections occurring in different tissues such as mouthparts and midguts.

Although single infections were predominant, about 6.41% mixed infections involving T. congolense forest and T. vivax were identified. This result corroborates data reported in tsetse [25, 31] and domestic animals [39] of Cameroon and other African countries [21, 22, 35]. It is important to point out that the identification of different trypanosome species was performed on whole tsetse fly and consequently, we do not know whether these infections were immature or mature, and which organ or tissue was infected. It is also unknown whether the mixed infections identified were from the same organ or tissue. Without such information, it becomes difficult to foresee the impact of mixed infections on the transmission and the dynamics of trypanosomes from tsetse to vertebrate hosts. Remarkably, the trypanosomes (T. vivax and T. congolense) reported in mixed infections can be found in their metacyclic forms in the mouthparts of tsetse flies. In such conditions, these trypanosomes can be simultaneously transmitted to vertebrate hosts. The mixed infections identified in this study therefore highlight a high probability that tsetse flies harbor or transmit several trypanosome species. If such transmission occurs, it becomes important to know which parasite could develop rapidly and what could be the impact of such infections on the transmission dynamics and animal health. Investigations on mixed infections in vertebrate hosts have shown mutual suppression and their advantages for the infected host [4]. As for vertebrate hosts, understanding how mixed infections evolve in tsetse flies and their potential impacts on the transmission dynamics of trypanosomes are areas for future investigation. The high infection rate observed in this study indicates active transmission of different trypanosome species. It highlights that the animal African trypanosomiases remain a serious threat to animal health and to the rural economy in the villages of the Fontem HAT focus.

The identification of tsetse flies with co-infections of S. glossinidius and trypanosomes, and other with trypanosome infections and without S. glossinidius, or with S. glossinidius and no trypanosome infection, corroborates results obtained elsewhere [12]. These results show that different scenarios in the tripartite association between tsetse, trypanosomes and symbiotic microorganisms can occur in natural populations of tsetse flies. In addition to these scenarios, the teneral or non-teneral status of tsetse fly and the first blood meal taken on non-infected vertebrate host could also affect the ability of tsetse to become infected and therefore, could negate any positive influence that S. glossinidius might have on tsetse susceptibility.

Our results showing no significant association (r = −0.083; p = 0.778) between the presence of S. glossinidius and trypanosomes infections indicate that the presence of S. glossinidius seems not absolutely necessary for trypanosome infections in the Fontem HAT focus. These findings are in line with those reported in other tsetse species like G. austeni [42], G. brevipalpis, G. morsitans morsitans and G. pallidipes [11] where no significant association has been reported between the presence of S. glossinidius and trypanosome infections. These results contrast with previous ones where the presence of S. glossinidius was reported to favor trypanosome infections not only in G. p. palpalis of other HAT foci of Cameroon [12], but also in other tsetse species [42]. These results highlight differences in the tripartite association between tsetse fly, S. glossinidius and trypanosomes. As a result of tsetse biology and environmental factors impacting the association between tsetse fly and its symbiotic micro-organisms, the tripartite association between tsetse, S. glossinidius and trypanosomes seems to vary according to tsetse species, and also to different populations (from different tsetse infested areas) of the same tsetse species. To better understand this tripartite association, more in-depth investigations on natural populations of different tsetse species in various tsetse infested areas are becoming important.

Our study on the tripartite association was based on presence/absence of trypanosome or S. glossinidius. Instead of focusing on this presence/absence, the genetic characterization of S. glossinidius strains could add additional value. In fact, Geiger et al. [15] have demonstrated that the tripartite association could be affected by specific genotypes of S. glossinidius and some trypanosome species, such as T. b. gambiense and T. b. brucei. Some specific S. glossinidius genotypes could affect the vectorial competence of G. p. gambiensis and G. m. morsitans for some trypanosome species [14]. The genetic characterization of bacteria populations could therefore enable us to improve our understanding of the tripartite association, and to better understand the real contribution of S. glossinidius to this association.

Conclusion

This study has shown that, within the same tsetse infested area, the infection rates of S. glossinidius and different trypanosome species do not vary significantly between villages. Our results also show that in natural conditions, some tsetse flies simultaneously harbor S. glossinidius and trypanosomes, while others have no infection or can be infected by only one of these microorganisms. They do not confirm previous results reporting that the presence of S. glossinidius seems to favor trypanosome infections in G. p. palpalis. The relationship between S. glossinidius and trypanosome infections seems to vary according to the tsetse infested areas. Genetic comparisons between S. glossinidius populations found in tsetse flies co-infected with both S. glossinidius and trypanosomes, and those found in tsetse flies without trypanosome infections, could enable us to deepen our understanding of the role of S. glossinidius in the vector competence of G. p. palpalis.

Acknowledgments

This study was supported by IRD through the “Jeune Équipe de Recherche Associée; JEAI EpiReTryp”, the “UMR INTERTRYP” of IRD, and the University of Dschang.

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Cite this article as: Kanté Tagueu S, Farikou O, Njiokou F & Simo G. 2018. Prevalence of Sodalis glossinidius and different trypanosome species in Glossina palpalis palpalis caught in the Fontem sleeping sickness focus of the southern Cameroon. Parasite 25, 44.

All Tables

Table 1.

Primers used for the identification of different trypanosome species.

In the text
Table 2.

Results of entomological surveys and infection rates of S. glossinidius according to villages

In the text
Table 3.

Trypanosome infections according to villages.

In the text
Table 4.

Sodalis glossinidius and T. congolense co-infections according to villages.

In the text

All Figures

thumbnail Figure 1.

Map showing villages where entomological surveys where undertaken in the Fontem sleeping sickness focus. Stars: Villages where tsetse flies were trapped; circles: other villages. The road from Mamfé to Dschang is indicated in black.
In the text

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