- Open Access
Does Trypanosoma cruzi (Chagas, 1909) (Kinetoplastida: Trypanosomatidae) modify the antennal phenotype of Triatoma dimidiata (Latreille, 1811) (Hemiptera: Triatominae)?
Parasites & Vectors volume 15, Article number: 466 (2022)
Triatoma dimidiata is a vector of the protozoan parasite Trypanosoma cruzi, the etiologic agent of Chagas disease. Phenotypic plasticity allows an organism to adjust its phenotype in response to stimuli or environmental conditions. Understanding the effect of T. cruzi on the phenotypic plasticity of its vectors, known as triatomines, has attracted great interest because of the implications of the parasite–triatomine interactions in the eco-epidemiology and transmission of the etiologic agent of Chagas disease. We investigated if the infection of the vector with T. cruzi may be associated with a change in the antennal phenotype of sylvatic, domestic, and laboratory-reared populations of T. dimidiata.
The abundance of each type of sensillum (bristles, basiconic, thick- and thin-walled trichoid) on the antennae of T. cruzi-infected and non-infected T. dimidiata reared in the laboratory or collected in sylvatic and domestic ecotopes were measured under light microscopy and compared using Kruskal–Wallis non-parametric tests and permutational multivariate analysis of variance.
We found significant differences between sensilla patterns of infected and non-infected insects within sylvatic and domestic populations. Conversely, we found no significant differences between sensilla patterns of infected and non-infected insects within the laboratory-reared population. Besides, for sylvatic and domestic populations, sexual dimorphism tended to be increased in infected insects.
The differences observed in infected insects could be linked to higher efficiency in the perception of odor molecules related to the search for distant mates and hosts and the flight dispersal in search of new habitats. In addition, these insects could have a positive effect on population dynamics and the transmission of T. cruzi.
Phenotypic plasticity is of great interest in ecology and evolution because it allows an organism to actively adjust its phenotype in response to stimuli or environmental conditions [1,2,3,4,5,6,7,8]. The response may or may not be adaptive, and it may involve changes in morphology, physiological state, behavior, or some combination of these . Besides, phenotypic plasticity is also widely recognized as an important factor for the evolution, population biology, and ecological interactions of many species [10,11,12,13]; thus, it is a major mechanism of ecological adaptation . Most information on phenotypic plasticity comes mainly from social insects [14,15,16], triatomines [17,18,19,20], grasshoppers [21, 22], and butterflies [13, 23].
Triatomine nymphs and adults of both sexes are strict blood-sucking insects that feed on vertebrate species available in their habitat . Their olfactory system plays an important role in many behavioral contexts, such as host-seeking, refuges and mate finding, alarm and aggregation behaviors, as well as avoidance of natural enemies . In triatomines, the antennal phenotype (AP) comprises the type and number of sensilla (classified as mechanoreceptors and chemoreceptors) distributed on the antennae. Sensilla act as an interface between the external and internal environments of insects (inter and intraspecific communication), capturing different stimuli from the external environment and directing them to the central nervous system [24,25,26]. This then triggers specific behavioral responses, such as the selection of a host for feeding, oviposition behavior, mate finding, and alarm and aggregation behaviors [27,28,29,30,31,32,33].
AP has been widely used as a sensitive marker to distinguish populations of triatomines [33,34,35]. In certain species or complexes, AP analysis complements other phenotypic and genetic characteristics [34, 36,37,38,39] or provides evidence for species differentiation [40, 41]. On the other hand, previous studies have established that the antennal sensilla of triatomines may show a degree of morphological variability between populations that seem to be associated with adaptations based on the sensorial requirements of different habitats [17, 37]. The number of sensilla may also vary because of selection pressure, sex, infection by a microorganism, and feeding habits [37, 39, 42,43,44,45,46]. Such changes show the degree of phenotypic plasticity exhibited by the species . Importantly, different studies have shown an absence of a correlation between the number of chemoreceptors and the total antenna length, length of antenna segments, and the number of each type of sensillum arranged over them [47, 48].
As vectors of the parasite Trypanosoma cruzi (Chagas, 1909) (Kinetoplastida: Trypanosomatidae), the causal agent of Chagas disease, the insects of the subfamily Triatominae (Hemiptera: Reduviidae), have special relevance in Latin America . The parasite is transmitted to humans and other animals when feces or urine of infected insects come into contact with mucous membranes or damaged areas of mammal skin . The co-evolution between triatomines and T. cruzi has promoted the development of powerful and sophisticated strategies, which can modify a wide range of physiological processes of the insects, including those related to the input, development, and discharge of the parasite . The existence of these modifications as a characteristic of an association between T. cruzi and triatomines could be the consequence of different adaptive or nonadaptive scenarios (e.g., adaptive host manipulation) [52, 53]. While several works have analyzed the mechanisms associated with T. cruzi–vector dynamics (e.g., biotic and abiotic factors) to understand the T. cruzi–triatomine interactions, under a co-evolutionary scenario , literature about how the parasites may influence the insects is more limited, and the studies have mainly been focused on the parasite’s effects on four patterns of the vector behavior: life-history traits, feeding, defecation, and dispersion/locomotion . Different studies have found negative effects of T. cruzi infection on vector survival [56,57,58,59], fecundity [59, 60], post-embryonic development [59, 61, 62], behavior [55, 63,64,65,66,67,68], and physiological processes [55, 60, 69,70,71], while other studies have not identified these effects on patterns of alimentation/defecation [56, 72, 73], development, and reproduction [74,75,76]. Overall, most of these studies determined that the effects of T. cruzi are species-dependent, age-dependent, sex-dependent, and even environment/physiology-dependent.
Although the AP, effects of T. cruzi, and phenotypic plasticity of the triatomines have been extensively studied [17, 34, 54], the phenotypic plasticity linked to the infection with T. cruzi in triatomines has not been investigated so far. In this study, we evaluated the changes in the AP of Triatoma dimidiata (Latreille, 1811) according to T. cruzi infection. More specifically, we investigated whether T. cruzi infection was associated with AP and sexual dimorphism modifications.
Laboratory-reared T. dimidiata came from a colony maintained for the past 10 years at the Parasitology Laboratory of the Regional Research Center Dr. Hideyo Noguchi, Autonomous University of Yucatan. New insects have been periodically added to this colony to avoid inbreeding depression. The insects were reared and maintained for 11 generations under controlled conditions (27 ± 1 °C, 70 ± 5% RH, a photoperiod of 12:12 [L:D] h), and were fed on immobilized pigeons [Columba livia Gmelin, 1789 (Aves, Columbidae)]. The domestic and sylvatic populations were composed of insects collected during entomological surveillance in 2018 inside and outside human dwellings of the rural village of Teya (25° 02′ 55″ N, 89° 04′ 25″ W), Yucatan, Mexico. The closest human dwellings were 3.5 km from the sylvatic site. The study was approved by the Institutional Bioethics Committee of the Autonomous University of Yucatan.
For infection of triatomines, the “V strain,” a TcI strain of T. cruzi originally isolated from a T. dimidiata specimen and maintained in the laboratory by cyclical passages in BALB/c adult mice, was used.
Infection of the laboratory-reared triatomines with T. cruzi
After a 2-week starvation period, the initial infection of the laboratory-reared triatomines was carried out with nymphs that had just molted to their fifth instar. Nymphs were fed ad libitum on BALB/c mice 15 days after they were infected with 1 × 106 parasites ml−1 of blood (i.e., during the parasite's exponential stage of growth ). Approximately 30 days after infection, we corroborated the infection status through examination of a fecal drop observed under a light microscope at ×40 magnification. Control group insects were fed under the same conditions on non-infected mice. The nymphs of both groups were maintained under rearing conditions and were fed fortnightly on infected/non-infected mice until they molted to the adult stage.
Assessment of the infection of domestic and sylvatic populations with T. cruzi
For T. dimidiata collected in natural conditions (i.e., domestic and sylvatic populations), we evaluated the infection with T. cruzi by amplifying parasite DNA from each bug midgut by polymerase chain reaction (PCR), using TCZ primers as described previously . Based on the results obtained by PCR, each population had a group of infected and non-infected insects.
We examined a total of 130 antennae of T. cruzi-infected and non-infected females and males from the sylvatic, domestic, and laboratory-reared populations of T. dimidiata (Table 1). One right antenna per specimen was removed using fine forceps and scissors. Antennae were processed with sodium hydroxide 4% for 6 h at 60 °C and then neutralized with glacial acetic acid 5% for 2 min. This procedure allowed cuticle diaphanization and enabled the identification and counting of the sensilla using a Zeiss Primostar® stereo microscope at ×400 magnification. The number and type of sensilla on antennal segments was counted manually using a procedure reported in previous works . The ventral side of the three distal segments of the antennae (P: pedicel, F1: flagellum 1, and F2: flagellum 2) was evaluated by identifying and counting sensilla including bristles (BR), thin-walled trichoid (TH), thick-walled trichoid (TK), and basiconic (BA) (nomenclature according to Catalá and Schofield ), thus giving a total of 12 morphological variables. The person who performed the measurements was unaware of the bug’s infection status to avoid any bias (MJET).
Differences in the AP between T. cruzi-infected (I) and non-infected (NI) insects were explored in the overall population, within each sex, within each population (i.e., sylvatic, domestic, and laboratory-reared), and within each sex within each population using univariate and multivariate analyses. Means and standard deviations of abundance were calculated for each type of sensilla (chemoreceptors: BR, TH, TK, and mechanoreceptors: BA) and antennal segment (pedicel, flagellum 1, and flagellum 2). As original data and their transformations were not normally distributed using Shapiro–Wilk tests , Kruskal–Wallis non-parametric tests were used for univariate analyses, followed by pairwise comparisons . Data were analyzed with the Minitab Statistical Software, version 17 (Minitab, Inc., PA, USA). In all cases, P < 0.05 was considered statistically significant. Moreover, the sources of variation of the AP were assessed using two-way permutational multivariate analysis of variance (PERMANOVA) on Bray–Curtis similarity matrices of square root with 9999 permutations. These analyses were conducted in PAST version 3.05.
Abundances of the sensilla found for all the T. dimidiata specimens included in this study are shown in Additional file 1: Table S1. All the insects’ antennae presented three types of chemoreceptors (TH, TK, and BA) and one mechanoreceptor (BR) on the three segments. The average number of sensilla per insect was 669.52 ± 176.45. Overall, the TH sensillum of the pedicel (P-TH) was the most abundant (183.42 ± 92.70), while the BR sensillum of the flagellum 2 (F2-BR) was the least abundant (17.45 ± 12.51). The pedicel was the segment with the highest number of sensilla (322.42 ± 115.54) while the flagellum 2 was the segment with the lowest number of sensilla (149.63 ± 54.43).
AP of T. cruzi-infected and non-infected T. dimidiata
Differences in each sensillum on the three antennal segments between infected and non-infected insects in the overall population, within each sex, within each population (i.e., sylvatic, domestic, and laboratory-reared), and within each sex within each population are summarized in Table 2.
When infected and non-infected insects were compared, significantly more BA sensilla on pedicel (P-BA) and TK sensilla on flagellum 1 (F1-TK) were observed in infected insects (Kruskal–Wallis test, P = 0.007 and P = 0.01, respectively).
Within each sex
When infected and non-infected insects were compared for each sex (I females vs, NI females; I males vs, NI males), significantly fewer TH sensilla on pedicel (P-TH) were observed in infected females (Kruskal–Wallis test, P = 0.04). Conversely, significantly more TK sensilla on flagellum 1 (F1-TK) were observed in infected males (Kruskal–Wallis test, P = 0.008).
Within each population
In the domestic population, when infected and non-infected insects were compared, significantly more BR sensilla on pedicel (P-BR) were observed in infected insects (Kruskal–Wallis test, P = 0.01). On the other hand, significantly fewer TH and TK sensilla on pedicel, BR, BA, TK sensilla on flagellum 1, and BR, BA, TH sensilla on flagellum 2 were observed in infected insects (Kruskal–Wallis test, P < 0.05 in all cases). Additionally, the two-way PERMANOVA test associated the infection with T. cruzi with the AP of the domestic population (F = 7.15; P = 0.0001), while the sex and the interaction infection*sex did not have a significant association with the AP (F = 1.51; P = 0.177 and F = 1.188; P = 0.299, respectively; Table 3).
In the sylvatic population, when infected and non-infected insects were compared, significantly more BA sensilla on pedicel, BR, BA, and TK sensilla on flagellum 1, and BR, BA, TH and TK sensilla on flagellum 2 were observed in infected insects (Kruskal–Wallis test, P < 0.05 in all cases). The two-way PERMANOVA test associated the infection with T. cruzi and the sex with the AP of the sylvatic population (F = 7.41; P = 0.0001 and F = 4.28; P = 0.002, respectively), while the interaction infection*sex did not have a significant association with the AP (F = 0.368; P = 0.125; Table 3).
Finally, in the laboratory-reared population, when infected and non-infected insects were compared, no difference in the number of sensilla was observed (Kruskal–Wallis test, P > 0.05 in all cases). In the same way, the two-way PERMANOVA test did not reveal significant association of the infection with T. cruzi, of the sex and of the interaction infection*sex with the AP of laboratory-reared insects (P > 0.05; Table 3).
Within each sex within each population
Differences in the abundance of each sensillum on the three antennal segments between infected and non-infected insects within each sex within each population are shown in Additional file 1: Table S1 and are summarized in Table 2.
When infected and non-infected females of the domestic population were compared, significantly fewer TH sensilla on pedicel and flagellum 2, and BA sensilla on flagellum 2 (Kruskal–Wallis, P < 0.05 in all cases) were observed. On the other hand, when infected and non-infected males of the domestic population were compared, significantly more BR sensilla on pedicel (P-BR) (Kruskal–Wallis test, P = 0.01) were observed. Moreover, when infected and non-infected males of the domestic population were compared, significantly fewer BR sensilla on flagellum 1 and flagellum 2, and BA sensilla on flagellum 2 (Kruskal–Wallis, P < 0.05 in all cases) were observed.
When infected and non-infected females of the sylvatic population were compared, significantly more BA sensilla on the three segments of the antennae, BR sensilla on flagellum 1 and flagellum 2, and TK sensilla on flagellum 2 (Kruskal–Wallis test, P < 0.05 in all cases) were observed. On the other hand, when infected and non-infected males of the sylvatic population were compared, significantly more BA sensilla on pedicel, BR and TK sensilla on flagellum 1, and BR and TH sensilla on flagellum 2 (Kruskal–Wallis test, P < 0.05 in all cases) were observed.
In the laboratory-reared population, there were no differences in the abundance of each sensillum between infected and non-infected females and between infected and non-infected males (Kruskal–Wallis test, P > 0.05).
Sexual dimorphism of T. cruzi-infected and non-infected insects
Differences in the abundances of each sensillum between non-infected females and males, and between infected females and males in the overall population, and within each population, are summarized in Table 4.
When non-infected females and males were compared, no significant difference in the abundance of each sensillum was observed (Kruskal–Wallis test, P > 0.05). However, when infected females and males were compared, significantly more TH sensilla on pedicel (P-TH) were observed in males (Kruskal–Wallis test, P = 0.002).
When non-infected females and males were compared, no significant difference in the abundance of each sensillum was observed (Kruskal–Wallis test, P > 0.05). However, when infected females and males were compared, significantly more TH sensilla on flagellum 1 (F1-TH) were observed in males (Kruskal–Wallis test, P = 0.01).
When non-infected females and males were compared, significantly more TH sensilla on pedicel (P-TH) and flagellum 1 (F1-TH) were observed in males (Kruskal–Wallis test, P = 0.02 and P = 0.003, respectively).
When infected females and males were compared, a significant difference in the abundance of TH sensilla on pedicel (P-TH) was still observed (Kruskal–Wallis test, P = 0.04), while the difference in the abundance of TH sensilla on flagellum 1 (F1-TH) was not observed anymore. However, significantly more BA sensilla on pedicel (P-BA) and flagellum 1 (F1-BA) were observed in females (Kruskal–Wallis test, P = 0.003 and P = 0.04, respectively).
In the laboratory-reared population, there was no sexual dimorphism in infected and non-infected insects (Kruskal–Wallis test, P > 0.05).
Phenotypic plasticity has been analyzed in different triatomine species in response to ecological factors [17, 35, 80], or to assess the effect of ecotope , food source , environment [35, 43, 81, 82], and sex . The present study is the first to analyze phenotypic plasticity related to the infection with T. cruzi in domestic, sylvatic, and laboratory-reared populations of T. dimidiata.
Our results show that there is an association between the infection status and the AP of T. dimidiata, at least in natural conditions. Indeed, we observed that infected and non-infected insects from the domestic and sylvatic populations showed significant differences in the abundance of some sensilla types. Besides, our results show that the sexual dimorphism tends to increase in T. cruzi-infected natural populations. Nevertheless, these differences in the abundance of some sensilla types between infected and non-infected insects, and the increased sexual dimorphism in infected insects was not observed in the laboratory-reared population. Consequently, while we were unable to evidence in this study a causal relationship between the infection with T. cruzi and the observed AP differences between infected and non-infected insects in natural populations of T. dimidiata, we could not exclude it either because the laboratory-reared insects were infected during their fifth development stage. If a causal relationship between T. cruzi infection and AP exists, this suggests that we should have established the infection in the earliest development stages to observe this effect, since insects infected in early development stages are more likely to be manipulated, as Poulin et al.  have suggested. However, more laboratory research is needed to understand how long it would take for T. cruzi to modify the AP of T. dimidiata and if there is a relation between AP changes and the parasite load in these vectors .
Therefore, the question remains whether the observed morphological differences are explained by the direct effect of T. cruzi infection and host manipulation, are evolutionary responses to selection at the population level, or are the consequences of different causal factors such as microecological influences .
Determining why and how host manipulation by parasites evolves is a fascinating but challenging question for evolutionary biologists. Pioneer authors addressing this question [86, 87] proposed that host changes probably occurred after the establishment of complex life cycles involving more than one host species. Ramirez-Gonzalez et al.  determined the effect of T. cruzi on the motor activity of fifth-stage nymphs of Triatoma longipennis Usinger, 1939 and Triatoma phyllosoma (Burmeister, 1835) infected during the second-stage nymph. On the other hand, Depickere et al.  determined the effect of T. cruzi on the aggregation behavior of Triatoma infestans (Klug, 1834) captured in the field and naturally infected. Recent studies with T. infestans have shown that infected insects after 45 days present changes in their circadian locomotor activity and feeding and defecation patterns [68, 73].
In our study, infected insects of the domestic and sylvatic populations showed, in general, significantly more BR sensilla compared with non-infected insects. These mechanoreceptors are associated with habitat selection rather than host selection , with the perception of mechanical stimuli related to the microhabitat . This suggests that infected insects could have greater capacities for adaptation or colonization of new habitats as compared with non-infected insects.
On the other hand, these mechanoreceptors also allow insects to perceive vibratory signals (through stridulation) during mating, and variations in the air current [88,89,90,91]. Moreover, they play an important role in the orientation toward odor-laden currents . Various studies have determined that the infection by T. cruzi, can impair the fecundity, fertility, and mating performance of triatomines (e.g., Fellet et al. ). An increase in these mechanoreceptors suggests that infected insects may benefit from copulation frequency and searching mating pairs, although reproductive success could be affected because of the infection. However, several functional aspects of these mechanoreceptors are unknown, and for this reason, further studies aimed at analyzing these contexts are needed to gain a better understanding of the functionality associated with habitat selection and the search for mating pairs.
Concerning the chemoreceptors (BA, TH and TK), it has previously been reported that BA sensilla have an olfactory and/or gustative function for the detection of habitats, shelters, hosts, mating pairs and related to the perception of sex-pheromone [93,94,95]. Besides, these sensilla seem involved in the detection of presumed pheromones in conspecific feces [96, 97]. The multiparous TH sensilla were first described in T. infestans by Bernard . The function of these sensilla may be associated with reproductive activities . They respond to a range of fatty acids—particularly pyruvic and lactic—and amyl acetate, and to breathing [93, 98]. On the other hand, although TK sensilla have been shown to predominate in triatomines , their chemosensory function has not been confirmed [93, 99]. However, Bernard  suggested that they may respond only to special compounds such as pheromones, thus acting as olfactory sensilla , as has been shown in the insect Cimex lectularius (Linnaeus, 1758) (Hemiptera: Cimicidae) . In our study, variation in the olfactory sensitivity associated with T. cruzi infection in the domestic and sylvatic populations is suggested. Indeed, in these populations, infected and non-infected insects showed significant differences in the abundance of some specific chemoreceptors. In the sylvatic population, the infection with T. cruzi was associated with more abundance of chemoreceptors. In a natural context, triatomines rely heavily on their sense of smell to locate, detect, and orientate toward a host from which they feed . The evidence indicates infected T. longipennis and Triatoma pallidipennis react more quickly to human odor than non-infected . Infected Mepraia spinolai (Porter, 1934) orient toward their host twice as fast, and their number of bites is duplicated . Some of these behavioral changes could promote vector competence, which encompasses the ability to acquire, maintain, and transmit a pathogen . Assuming the hypothesis of sensory modulation by the parasite [71, 92], it is possible that the infected insects of the sylvatic population generate an increase in the antennal receptors because of the wide range of hosts in the sylvatic ecotope and to enhance parasite transmission probability [77, 102, 103]. Moreover, this increase may enhance the capacity for dispersal and invasion of different habitats [47, 81], and efficiency in the perception of odor molecules in the search of distant mates and hosts and for flight dispersal in search of new habitats, as it has been suggested by other authors [82, 102, 104,105,106], thus conferring an advantage to T. cruzi.
Nevertheless, infected insects of the domestic populations showed a decrease in some chemoreceptors. The occurrence of these modifications as characteristic of a parasite–host system may be the consequence of an adaptive process (i.e., adaptive host manipulation by the parasite or compensatory response by the host). Also, it may be an after-effect of the presence of the parasite that allows insects to reduce their investment in costly or reduced-use structures [52, 107].
Several studies have provided information about the sexual dimorphism in non-infected triatomines from different species, populations, rearing, and ecotopes [38, 80, 88]. However, our study reports for the first time the sexual dimorphism in the AP of infected insects of T. dimidiata. In general, the sexual dimorphism observed in infected insects of T. dimidiata was based on an increase in the abundance of TH sensilla in infected males and/or an increase of BA sensilla in infected females. These chemoreceptors have an olfactory function for the detection of sexual pairs, habitats, and hosts as mentioned above. Evidence of this study and previous works [48, 81, 108] suggest that the sexual dimorphism in the AP may be linked to the perception of molecules related to sexual behavior and to differences in sensing sexual pheromones, as has been suggested by other authors (e.g., May-Concha et al. ; Souza et al. ). May-Concha  provided information on a chemical signal produced during T. dimidiata mating, since fewer mating attempts were observed when the opening of female glands was occluded. Besides, that study describes a chemical signal which promotes the attraction of males to volatiles emitted by females and to mating couples . On the other hand, based on previous works on olfactory receptors [24,25,26], we can propose that the increased abundance of TH chemo-sensilla in infected males contributes to a greater efficiency in the perception of odor molecules involved in sexual communication compared with infected females. In contrast, we can hypothesize that the increased abundance of BA chemo-sensilla in infected females contributes to a greater efficiency in the perception of host odors compared with infected males. Therefore, the increase in the odor perception in infected insects may elicit a positive effect on vector population dynamics and could enhance the vectorial transmission of T. cruzi. Future studies should examine in-depth the effect of the parasite on other aspects of the behavior of triatomine insects, such as aggregation, alarm, feeding, excretion/defecation, and host foraging, which could constitute epidemiologically relevant behavioral changes, and evaluate sexual behavioral changes in adults, which could impact the growth of triatomine populations.
To our knowledge, this is the first work that relates an association of the AP with the T. cruzi infection status of T. dimidiata. Although we could not demonstrate any causal relationship, we revealed a clear association between the natural infection status of T. dimidiata and its antennal phenotypic variation. The differences observed in infected insects could be linked to higher efficiency in the olfactory perception related to the search for distant mates and hosts and the flight dispersal in search of new habitats. In addition, these insects could have a positive effect on population dynamics and the transmission of T. cruzi.
Availability of data and materials
The datasets used and/or analyzed during the present study are available from the corresponding author upon reasonable request.
Via S, Gomulkiewicz R, De Jong G, Scheiner SM, Schlichting CD, Van Tienderen TH. Adaptive phenotypic plasticity: consensus and controversy. Trends Ecol Evol. 1995;10:212–7.
West-Eberhard MJ. Developmental plasticity and evolution. New York, NY: Oxford University Press; 2003.
Pigliucci M. Evolution of phenotypic plasticity: where are we going now? Trends Ecol Evol. 2005;20:481–6.
Laland K, Uller T, Feldman M, Sterelny K, Müller GB, Moczek A, et al. Does evolutionary theory need a rethink? Nature. 2014;514:161–4.
Lande R. Evolution of phenotypic plasticity in colonizing species. Mol Ecol. 2015;24:2038–45.
Beaman JE, White CR, Seebacher F. Evolution of plasticity: mechanistic link between development and reversible acclimation. Trends Ecol Evol. 2016;31:237–49.
Gadenne C, Barrozo RB, Anton S. Plasticity in insect olfaction: to smell or not to smell? Annu Rev Entomol. 2016;61:317–33.
Colautti RI, Alexander JM, Dlugosch KM, Keller SR, Sultan SE. Invasions and extinctions through the looking glass of evolutionary ecology. Philos Trans R Soc B Biol Sci. 2017;372:20160031.
West-Eberhard MJ. Phenotypic plasticity. In: Jørgensen E, Fath B, editors. Encyclopedia of ecology. Amsterdam: Elsevier Sci; 2008. p. 2701–7.
Baldwin JM. A new factor in evolution. Am Nat. 1986;30:441–51.
West-Eberhard M. Phenotypic plasticity and the origins of diversity. Annu Rev Ecol Syst. 1989;20:249–78.
Weiss LC. Sensory ecology of predator-induced phenotypic plasticity. Front Behav Neurosci. 2019;12:330.
Bhardwaj S, Jolander LS, Wenk MR, Oliver JC, Nijhout HF, Monteiro A. Origin of the mechanism of phenotypic plasticity in satyrid butterfly eyespots. Elife. 2020;2019:e49544.
Manfredini F, Arbetman M, Toth AL. A potential role for phenotypic plasticity in invasions and declines of social insects. Front Ecol Evol. 2019;7:375.
Stern DL, Foster WA. The evolution of soldiers in aphids. Biol Rev. 1996;71:27–76.
Moczek AP. Phenotypic plasticity and diversity in insects. Philos Trans R Soc B Biol Sci. 2010;365:593–603.
Dujardin JP, Panzera P, Schofield CJ. Triatominae as a model of morphological plasticity under ecological pressure. Mem Inst Oswaldo Cruz. 1999;94:223–8.
Batista VSP, Fernandes FA, Cordeiro-Estrela P, Sarquis O, Lima MM. Ecotope effect in Triatoma brasiliensis (Hemiptera: Reduviidae) suggests phenotypic plasticity rather than adaptation. Med Vet Entomol. 2012;27:247–54.
Nattero J, Malerba R, Rodríguez C, Crocco L. Phenotypic plasticity in response to food source in Triatoma infestans (Klug, 1834) (Hemiptera, Reduviidae: Triatominae). Inf Gen Evol. 2013;19:39–44.
Nattero J, Dujardin JP, Fernández MP, Gürtler RE. Host-feeding sources and habitats jointly affect wing developmental stability depending on sex in the major Chagas disease vector Triatoma infestans. Inf Gen Evol. 2015;36:539–46.
Bernays EA, Chapman RF. Phenotypic plasticity in numbers of antennal chemoreceptors in a grasshopper: effects of food. J Comp Physiol A. 1998;183:69–76.
Hochkirch A, Depperman J, Gröning J. Phenotypic plasticity in insects: the effects of substrate colour on the colouration of two ground-hopper species. Evol Dev. 2008;10:350–9.
Jorge LR, Cordeiro-Estrela P, Klaczko LB, Moreira GRP, Freitas AVL. Host-plant dependent wing phenotypic variation in the neotropical butterfly Heliconius erato. Biol J Linn Soc. 2011;102:765–74.
May-Concha IJ, Guerenstein PG, Malo EA, Catalá S, Rojas JC. Electroantennogram responses of the Triatoma dimidiata complex to volatiles produced by its exocrine glands. Acta Trop. 2018;185:336–43.
Guidobaldi F, May-Concha IJ, Guerenstein PG. Morphology and physiology of the olfactory system of blood-feeding insects. J Insect Physiol. 2014;106:96–111.
Pontes G, Minoli S, Ortega Insaurralde I, de Brito Sanchez MG, Barrozo RB. Bitter stimuli modulate the feeding decision of a blood-sucking insect via two sensory inputs. J Exp Biol. 2014;217:3708–17.
Guerenstein PG, Guerin PM. Olfactory and behavioural responses of the bloodsucking bug Triatoma infestans to odours of vertebrate hosts. J Exp Biol. 2001;204:585–97.
Guidobaldi F, Guerenstein PG. Oviposition in the blood-sucking insect Rhodnius prolixus is modulated by host odors. Parasit Vectors. 2015;8:265.
Guidobaldi F, Guerenstein PG. A CO2-free synthetic host odor mixture that attracts and captures triatomines: effect of emitted odor ratios. J Med Entomol. 2016;53:770–5.
May-Concha IJ, Rojas JC, Cruz-López L, Millar JG, Ramsey JM. Volatile compounds emitted by Triatoma dimidiata, a vector of Chagas disease: chemical identification and behavioural analysis. Med Vet Entomol. 2013;27:165–74.
May-Concha IJ, Rojas JC, Cruz-López L, Ibarra-Cerdeña CN, Ramsey JM. Volatile compound diversity and conserved alarm behaviour in Triatoma dimidiata. Parasit Vectors. 2015;8:84–98.
May-Concha IJ, Loobia PA, Mougabure-Cueto G. Interaction between two aggregation chemical signals in Triatoma infestans (Hemiptera: reduviidae). J Insect Physiol. 2018;109:79–84.
May-Concha IJ, Cruz-López LC, Rojas JC, Ramsey JW. “Sweeter than a rose”, at least to Triatoma phyllosoma complex males (Triatominae: Reduviidae). Parasit Vectors. 2018;11:95.
May-Concha IJ, Guerenstein PG, Ramsey JM, Rojas JC, Catalá S. Antennal phenotype of Mexican haplogroups of the Triatoma dimidiata complex, vector of Chagas disease. Infect Genet Evol. 2016;40:73–9.
Müller JN, Gonçalves TCM, Ricardo-Silva AH, Souza CA, Santos FM, Santos R, et al. Does antennal sensilla pattern of different populations of Triatoma maculata (Hemiptera: Reduviidae) reveal phenotypic variability? Parasit Vectors. 2019;12:602.
Catalá S, Schofield CJ. The antennal sensilla of Rhodnius. J Morphol. 1994;219:193–203.
Catalá S. Antennal sensilla of Triatominae. A comparative study of five genera. Int J Insect Morphol Embryol. 1997;26:67–73.
Carbajal de la Fuente AL, Noireau F, Catalá SS. Inferences about antennal phenotype: the “Triatoma maculata complex” (Hemiptera: Triatominae) is valid? Acta Trop. 2008;106:16–21.
Hernández ML, Abrahan L, Moreno M, Gorla D, Catalá S. Phenotypic variability associated to genomic changes in the main vector of Chagas disease in the southern cone of South America. Acta Trop. 2008;106:60–7.
Catalá S, Torres M. Similitude of the patterns of sensilla on the antennae of Triatoma melanosoma and Triatoma infestans. Ann Trop Med Parasitol. 2001;95:287–95.
Martínez-Hernández F, Martínez-Ibarra JA, Villalobos G, De la Torre P, Laclette JP, Alejandré-Aguilar R, et al. Natural crossbreeding between sympatric species of the Phyllosoma complex (Insecta: Hemiptera: Reduviidae) indicate the existence of one species with morphologic and genetic variations. Am J Trop Med Hyg. 2010;82:74–82.
Chapman R. Chemoreception: the significance of receptors numbers. Adv Insect Physiol. 1982;16:247–356.
Catalá SS, Maida DM, Caro-Riaño H, Jaramillo N, Moreno J. Changes associated with laboratory rearing in antennal sensilla patterns of Triatoma infestans, Rhodnius prolixus, and Rhodnius pallescens (Hemiptera, Reduviidae, Triatominae). Mem Inst Oswaldo Cruz. 2004;99:25–30.
Abrahan L, Hernández L, Gorla D, Catalá S. Phenotypic diversity of Triatoma infestans at the microgeographic level in the Gran Chaco of Argentina and the Andean valleys of Bolivia. J Med Entomol. 2008;45:660–6.
Dujardin JP, Costa J, Bustamante D, Jaramillo N, Catalá S. Deciphering morphology in Triatominae: the evolutionary signals. Acta Trop. 2009;110:101–11.
Cantillo-Barraza O, Garcés E, Gómez-Palacio A, Cortés LA, Pereira A, Marcet PL, et al. Eco-epidemiological study of an endemic Chagas disease region in northern Colombia reveals the importance of Triatoma maculata (Hemiptera: Reduviidae), dogs and Didelphis marsupialis in Trypanosoma cruzi maintenance. Parasit Vectors. 2015;8:482.
Arroyo CM, Esteban L, Catalá S, Angulo VM. Variación del fenotipo antenal de poblaciones del domicilio, peridomicilio y silvestres de Triatoma dimidiata (Hemiptera: Reduviidae) en Santander. Colombia Biomed. 2007;27:92–100.
Carbajal de la Fuente AL, Catalá S. Relationship among the habitat and the antenal sensilla pattern of six species of Triatominae (Hemiptera: Reduviidae). Mem Inst Oswaldo Cruz. 2002;97:1073–7.
Guhl F. Enfermedad de Chagas: Realidad y perspectivas. Rev Biomed. 2009;20:228–34.
Silva-Neto MAC, Fampa P, Caiaffa CD, Carneiro AB, Atella GC. Cell signaling during Trypanosoma cruzi development in triatominae. Open Parasitol J. 2010;4:188–94.
Córdoba-Aguilar A. Chagas bugs and Trypanosoma cruzi: puppets and puppeteer? Acta Trop. 2020;211:105600.
Poulin R. Parasite manipulation of host behavior: an update and frequently asked questions. Adv Study Behav. 2010;41:151–86.
Heil M. Host manipulation by parasites: cases, patterns, and remaining doubts. Front Ecol Evol. 2016;4:80.
De Fuentes-Vicente JA, Gutiérrez-Cabrera AE, Flores-Villegas AL, Lowenberger C, Benelli G, Salazar-Schettino PM, et al. What makes an effective Chagas disease vector? Factors underlying Trypanosoma cruzi triatomine interactions. Acta Trop. 2018;183:23–31.
Depickère S, Ramírez-Ávila GM, Deneubourg J. Alteration of the aggregation and spatial organization of the vector of Chagas disease, Triatoma infestans, by the parasite Trypanosoma cruzi. Sci Rep. 2019;9:17432.
Elliot SL, Rodrigues JdO, Lorenzo MG, Martins-Filho OA, Guarneri AA. Trypanosoma cruzi, etiological agent of Chagas disease, is virulent to its triatomine vector Rhodnius prolixus in a temperature-dependent manner. PLoS Negl Trop Dis. 2015;9:e0003646.
Hinestroza G, Ortiz MI, Molina J. Behavioral fever response in Rhodnius prolixus (Reduviidae: Triatominae) to intracoelomic inoculation of Trypanosoma cruzi. Rev Soc Bras Med Trop. 2016;49:425–32.
Peterson JK, Graham AL, Elliott RJ, Dobson AP, Chávez OT. Trypanosoma cruzi–Trypanosoma rangeli co-infection ameliorates negative effects of single trypanosome infections in experimentally infected Rhodnius prolixus. Parasitology. 2016;143:1157–67.
Cordero-Montoya G, Flores-Villegas AL, Salazar-Schettino PM, Vences-Blanco MO, Rocha-Ortega M, Gutiérrez-Cabrera AE, et al. The cost of being a killer’s accomplice: Trypanosoma cruzi impairs the fitness of kissing bugs. Parasitol Res. 2019;118:2523–9.
Fellet MR, Lorenzo MG, Elliot SL, Carrasco D, Guarneri AA. Effects of infection by Trypanosoma cruzi and Trypanosoma rangeli on the reproductive performance of the vector Rhodnius prolixus. PLoS ONE. 2014;9:e105255.
Eichler S, Schaub GA. Development of symbionts in triatomine bugs and the effects of infections with trypanosomatids. Exp Parasitol. 2002;100:17–27.
Botto-Mahan C. Trypanosoma cruzi induces life-history trait changes in the wild kissing bug Mepraia spinolai: implications for parasite transmission. Vector Borne Zoonotic Dis. 2009;9:505–10.
Schaub GA. Parasitogenic alterations of vector behaviour. Int J Med Microbiol. 2006;296:37–40.
Botto-Mahan C, Cattan PE, Medel R. Chagas disease parasite induces behavioural changes in the kissing bug Mepraia spinolai. Acta Trop. 2006;98:219–23.
Pereyra N, Lobbia PA, Mougabure-Cueto G. Effects of the infection with Trypanosoma cruzi on the feeding and excretion/defecation patterns of Triatoma infestans. Bull Entomol Res. 2019;24:1–8.
Ramírez-González MG, Flores-Villegas AL, Salazar-Schettino PM, Gutiérrez Cabrera AE, Rojas-Ortega E, Córdoba-Aguilar A. Zombie bugs? Manipulation of kissing bug behavior by the parasite Trypanosoma cruzi. Acta Trop. 2019;200:105177.
Uc-Diaz SS. Efecto de Trypanosoma cruzi sobre la selección de hospederos sanguíneos por Triatoma dimidiata. BcB Thesis. Universidad Autónoma de Yucatán, Campus de Ciencias Biológicas y Agropecuarias. 2020. p. 65.
Chacón F, Muñoz-San Martín C, Bacigalupo A, Álvarez-Duhart B, Solís R, Cattan PE. Trypanosoma cruzi parasite load modulates the circadian activity pattern of Triatoma infestans. Insects. 2022;13:76.
Vallejo G, Guhl F, Schaub G. Triatominae-Trypanosoma cruzi/T. rangeli: vector–parasite interactions. Acta Trop. 2009;110:137–47.
Oliveira T, Carvalho-Costa F, Gomes T, Sarquis O, Sposina R, Lima R. Developmental and reproductive patterns of Triatoma brasiliensis infected with Trypanosoma cruzi under laboratory conditions. Mem Inst Oswaldo Cruz. 2010;105:1057–60.
Marliére NP, Latorre-Estivalis JM, Lorenzo MG, Carrasco D, Alves-Silva J, Rodrigues JO, et al. Trypanosomes modify the behavior of their insect hosts: effects on locomotion and on the expression of a related gene. PLoS Negl Trop Dis. 2015;9:e0003973.
Takano-Lee M, Edman JD. Lack of manipulation of Rhodnius prolixus (Hemiptera: Reduviidae) vector competence by Trypanosoma cruzi. J Med Entomol. 2002;39:44–51.
Chacón F, Bacigalupo A, Álvarez-Duhart B, Cattan PE, Solis R, Muñoz-San MC. The parasite load of Trypanosoma cruzi modulates feeding and defecation patterns of the Chagas disease vector Triatoma infestans. Microorganism. 2022;10:103.
Zeledón R, Guardia VM, Zúñiga A, Swartzwelde JC. Biology and ethology of Triatoma dimidiata (Latreille, 1811). II. Life span of adults and fecundity and fertility of females. J Med Entomol. 1970;7:462–9.
Schaub GA. Developmental time and mortality of larvae of Triatoma infestans infected with Trypanosoma cruzi. Trans R Soc Trop Med Hyg. 1988;82:94–7.
Lima MM, Borges-Pereira J, Albuquerque Dos Santos JA, Teixeira-Pinto Z, Vianna-Braga M. Development and reproduction of Panstrongylus megistus (Hemiptera: Reduviidae) infected with Trypanosoma cruzi, under laboratory conditions. Ann Entomol Soc Am. 1992;85:458–61.
Moo-Millan JI, Arnal A, Pérez-Carrillo S, Hernandez-Andrade A, Ramírez-Sierra MJ, Rosado-Vallado M, et al. Disentangling Trypanosoma cruzi transmission cycle dynamics through the identification of blood meal sources of natural populations of Triatoma dimidiata in Yucatán, Mexico. Parasit Vectors. 2019;12:572.
Sokal RR, Rohlf FJ. Introduction to biostatistics. 2nd ed. New York: Dover Publications Inc.; 2009. p. 384.
Conover WJ. Practical nonparametric statistics. New York: Wiley; 1999. p. 608.
Moreno ML, Gorla D, Catalá S. Association between antennal phenotype, wing polymorphism and sex in the genus Mepraia (Reduviidae: Triatominae). Infect Genet Evol. 2006;6:228–34.
Catalá S, Sachetto C, Moreno M, Rosales R, Salazar-Schettino PM, Gorla D. Antennal phenotype of Triatoma dimidiata populations and its relationship with species of Phyllosoma and Protracta Complexes. J Med Entomol. 2005;42:719–25.
Souza AC, Catalá S, Carbajal-de-la Fuente AL, Junqueira A. Phenotypic variability of the Amazonian species Rhodnius brethesi (Hemiptera: Reduviidae). J Med Entomol. 2017;54:909–16.
Poulin R, Brodeur J, Moore J. Parasite manipulation of host behaviour: should hosts always lose? Oikos. 1994;70:479–84.
Ramirez-Sierra MJ, Dumonteil E. Infection rate by Trypanosoma cruzi and biased vertebrate host selection in the Triatoma dimidiata (Hemiptera: Reduvidae) species complex. J Med Entomol. 2016;53:20–5.
Montes de Oca-Aguilar AC, González-Martínez A, Chan-González R, Ibarra-López P, Smith-Ávila S, Córdoba-Aguilar A, et al. Signs of urban evolution? Morpho-functional traits co-variation along a nature-urban gradient in a Chagas disease vector. Front Ecol Evol. 2022;10:805040.
Smith-Trail DR. Behavioural interactions between parasites and host: host suicide and evolution of complex life cycles. Am Nat. 1980;116:77–91.
Moore J. Altered behavioral responses in intermediate hosts—an acanthocephalan parasite strategy. Am Nat. 1984;123:572–7.
Catalá S, Dujardin JP. Antennal sensilla patterns indicate geographic and ecotopic variability among Triatoma infestans (Hemiptera: Reduviidae) populations. J Med Entomol. 2001;38:423–8.
Wigglesworth VB, Gillett JD. The function of the antennae in Rhodnius prolixus Hemiptera and the mechanism of orientation to the host. J Exp Biol. 1934;11:120–39.
McIver SB, Siemicki R. Fine structure of antennal mechanosensilla of adult Rhodnius prolixus Stål Hemiptera: Reduviidae. J Morphol. 1984;180:19–28.
Lazzari CR, Nuñez JA. The response to radiant heat and the estimation of the temperature of distant sources in Triatoma infestans. J Ins Physiol. 1989;35:525–9.
Barrozo RB, Reisenmann CE, Guerenstein P, Lazzari CR, Lorenzo MG. An inside look at the sensory biology of triatomines. J Insect Physiol. 2017;97:3–19.
Bernard J. Études Electrophysiologiques des Récepteurs Impliqués dans la Orientation vers l'Hôte et dans l'Acte Hematophage chez un Hémiptère Triatoma infestans. PhD Thesis. Université de Rennes, France.1974. p. 285
Guerenstein P, Lazzari C. Host-seeking: how triatomines acquire and make use information to find blood. Acta Trop. 2009;110:148–58.
Bohman B, Weinstein AM, Unelius CR, Lorenzo MG. Attraction of Rhodnius prolixus males to a synthetic female-pheromone blend. Parasit Vectors. 2018;11:418.
Taneja J, Guerin PM. Oriented responses of triatomines bugs Rhodnius prolixus and Triatoma infestans to vertebrate odours on a servosphere. J Comp Physiol A. 1995;176:455–64.
Taneja J, Guerin PM. Ammonia attracts the haematophagous bug Triatoma infestans: behavioural and neurophysiological data on nymphs. J Comp Physiol A. 1997;181:21–34.
Mayer MS. Response of single olfactory cell of Triatoma infestans to human breath. Nature. 1968;220:924–5.
Guerenstein PG. Sensory and behavioural responses of Triatoma infestans to host and conspecific odours. Ph.D. Thesis. University of Neuchâtel, Switzerland. 1999. p. 137.
Steinbrecht RA, Stankiewicz BA. Molecular composition of the wall of insect olfactory sensilla: the chitin question. J Insec Physiol. 1999;45:785–90.
Steinbrecht R, Muller B. Fine structure of the antenna1 receptors on the bed bug, Cimex lectularius L. Tissue Cell. 1976;8:615–36.
López-Cancino SA, Tun-Ku E, De la Cruz-Felix HK, Ibarra-Cerdeña CN, Izeta-Alberdi A, Pech-May A, et al. Landscape ecology of Trypanosoma cruzi in the southern Yucatan Peninsula. Acta Trop. 2015;151:58–72.
Dumonteil E, Ramirez-Sierra MJ, Pérez-Carrillo S, Teh-Poot C, Herrera C, Gourbière S, et al. Detailed ecological associations of triatomines revealed by metabarcoding and next-generation sequencing: implications for triatomine behavior and Trypanosoma cruzi transmission cycles. Sci Rep. 2018;8:4140.
Guzmán-Tapia Y, Ramírez-Sierra MJ, Dumonteil E. Urban infestation by Triatoma dimidiata in the city of Mérida, Yucatán, México. Vector Borne Zoonotic Dis. 2007;7:597–606.
Dumonteil E, Gourbiére S, Barrera-Perez M, Rodriguez-Felix E, Ruiz-Piña H, Baños-López O, et al. Geographic distribution of Triatoma dimidiata and transmission dynamics of Trypanosoma cruzi in the Yucatan peninsula of Mexico. Am J Trop Med Hyg. 2002;67:176–83.
Ibarra-Cerdeña CN, Zaldívar-Riverón A, Peterson AT, Sánchez-Cordero V, Ramsey JM. Phylogeny and niche conservatism in North and Central American triatomine bugs (Hemiptera: Reduviidae: Triatominae), vectors of Chagas’ disease. PLoS Negl Trop Dis. 2014;8:e3266.
May-Concha I, Remón C, Mougabure-Cueto G. Behavioral response mediated by feces in Triatoma infestans (Hemiptera: Reduviidae: triatominae) susceptible and resistant to deltamethrin. Acta Trop. 2020;206:105442.
Catalá S, Carbajal A, Torres M, Moreno M, Ordoñez R, Montaña F, et al. La antena de los triatominae: caracteres ancestrales y marcadores funcionales. In: Schofield, C., Ponce, C. editors. Proceedings IV International Workshop on Population Genetics and Control of Triatominae. European Community, Cartagena de Indias. Colombia, 2000. pp. 80–81 (INDRE. Mexico).
May-Concha IJ. Compuestos volatiles emitidos por adultos en disturbio y copula de Triatoma dimidiata (Hemiptera: Reduviidae), vector de la enfermedad de Chagas. MSc Thesis. Instituto Nacional de Salud Publica. 2010. p. 27
We are grateful for the technical assistance of Bachelor of Biology Salma Uc Diaz and Victor Garrido Gonzalez, and the statistical advice of Master of Science Joel Moo-Millan.
This research was supported by Consejo Nacional de Ciencia y Tecnología de México (CONACyT) IJMC/CVU: 272733. This work received financial support from CONACYT (National Council of Science and Technology, Mexico), Basic Science (Project ID: CB2015-258752), and National Problems (Project ID: PN2015-893) programs attributed to EW.
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Additional file 1: Table S1.
Abundances of each sensillum on the three antennal segments in infected and non-infected insects of each sex within each population of T. dimidiata. The data shown are the means and standard deviation. N=130. TH: thin-walled trichoid; TK: thick-walled trichoid; BA: basiconic; BR: bristles. The number between parentheses represents the number of specimens analyzed. The number between clasps represents the standard deviation of the data. I = infected, NI = non-infected, D = domestic, L = laboratory-reared, S = sylvatic, F = female, M = male. Different letters indicate significant differences between infected and non-infected insects of the same sex within each population (Kruskal–Wallis tests; P < 0.05).
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.
About this article
Cite this article
May-Concha, I.J., Escalante-Talavera, M.J., Dujardin, JP. et al. Does Trypanosoma cruzi (Chagas, 1909) (Kinetoplastida: Trypanosomatidae) modify the antennal phenotype of Triatoma dimidiata (Latreille, 1811) (Hemiptera: Triatominae)?. Parasites Vectors 15, 466 (2022). https://doi.org/10.1186/s13071-022-05587-y
- Chagas disease
- Host manipulation
- Phenotypic plasticity