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Article

Natural Trypanosoma cruzi Infection and Climatic Season Influence the Developmental Capacity in Field-Caught Mepraia spinolai Nymphs

by
Juan Botzotz
1,
Gabriel Méndez-Valdés
1,
Sylvia Ortiz
1,
Angélica López
1,
Carezza Botto-Mahan
2,* and
Aldo Solari
1,*
1
Programa de Biología Celular y Molecular, Instituto de Ciencias Biomédicas, Facultad de Medicina, Universidad de Chile, Santiago 8380000, Chile
2
Departamento de Ciencias Ecológicas, Facultad de Ciencias, Universidad de Chile, Santiago 7800003, Chile
*
Authors to whom correspondence should be addressed.
Insects 2023, 14(3), 272; https://doi.org/10.3390/insects14030272
Submission received: 28 January 2023 / Revised: 28 February 2023 / Accepted: 1 March 2023 / Published: 9 March 2023
(This article belongs to the Section Medical and Livestock Entomology)
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Abstract

:

Simple Summary

Triatomines are hemimetabolous and hematophagous insect vectors of Trypanosoma cruzi, the parasite that causes Chagas disease. The life cycle of triatomines is composed of five nymphal stages and an adult stage. The wild kissing bug Mepraia spinolai is distributed in a vast area of the semiarid Mediterranean ecosystem of South America, characterized by strongly marked seasons. The population structure of this vector changes during the different seasons, suggesting its development is controlled under natural conditions. In this study, we performed a biological assay to study this vector development after two laboratory feedings. We collected nymphs, first to fourth instars, in eight M. spinolai populations during the transition of late fall to winter (cooling period) and spring (warming period) of three consecutive years. The gathered information with vectors molting twice suggests that development is differentially controlled in nymphs from first and fourth instars depending on the climatic period of collection. Additionally, T. cruzi infection changes development forward only in nymphs of second and fourth instars of the warming period. Our findings suggest that the effect of the climatic period and T. cruzi infection on the development of this vector species is a stage-dependent phenomenon.

Abstract

In this study, we evaluated the effect of the climatic season and infection by Trypanosoma cruzi, etiological agent of Chagas disease, on the molting capacity of the triatomine vector Mepraia spinolai endemic to Chile. We used wild-caught first-to-fourth instar nymphs during cooling (fall and winter) and warming (spring) periods. After capturing, nymphs were fed at the laboratory, and maintained under optimal rearing conditions. Feeding was repeated 40 days later. We followed-up the molting events on 709 nymphs, recording one, two or the absence of molts after two feeding opportunities. Within the same climatic period, only infected second- and fourth-instar nymphs from the warming period showed a larger proportion of double molting compared to uninfected nymphs. Regarding the climatic period, infected and uninfected first- and fourth-instar nymphs exhibited a larger proportion of double molting in the warming and cooling periods, respectively. The pattern of non-molting nymph occurrence suggests they probably reach diapause by environmental stochasticity. The effect of the climatic period and T. cruzi infection on the development of M. spinolai is an instar-dependent phenomenon, highlighting the occurrence of finely synchronized processes at different moments of the life cycle of such an hemimetabolous insect as triatomines.

Graphical Abstract

1. Introduction

Triatomine bugs (Hemiptera: Reduviidae) are hemimetabolous and hematophagous insects distributed in the Americas, capable of transmitting Trypanosoma cruzi, the etiological agent of Chagas disease [1]. Triatomine development includes five nymphal stages and an adult stage. Several demographic factors can modify triatomine population dynamics such as age structure, life history trait variation caused by environmental fluctuations, dispersal in spatially heterogeneous environments, and local extinction and colonization, as well as T. cruzi infection [2,3,4,5,6,7,8,9].
In vector-borne infections, vectors can also be considered as an alternative host for the parasite; therefore, some degree of vector exploitation is to be expected [10,11]. In the case of T. cruzi, the pathogenicity to the insect vector can be gauged via mortality, reproduction, and developmental arrests (diapause). In nature, developmental delays are observed in triatomines submitted to different conditions such as low environmental temperature [3], density-dependent access to blood sources [12], or diapause triggered by environmental factors such as changes in weather [7]. Almost a century ago, a study on the molting process of the triatomine Rhodnius prolixus was carried out, detecting that a full engorgement was necessary to trigger ecdysis [13]. Currently, it is known that the initial steps of triatomine development involve the release of the neuropeptide prothoracicotropic hormone and ecdysteroids in the hemolymph within the neuroendocrine system [14].
Experimental studies evaluating the effect of T. cruzi in triatomine development have shown variable results under optimal climatic conditions [15,16,17,18,19], mostly depending on the experimental design. However, experimental studies assessing the effect of T. cruzi on triatomine development submitted to variable abiotic conditions are less represented. For example, in R. prolixus, a development retardment was observed in T. cruzi-infected insects and the molting time was shorter at 27–30 °C than at lower temperatures [11]. Regarding the effect of environmental temperature on the development of uninfected Triatoma brasiliensis, it was shown that no ecdysis occurred between 12 °C and 21 °C [20]. Few studies on the development of triatomines submitted to natural climatic conditions are available. Laboratory-reared Triatoma infestans studied in the field under the variable environmental conditions of the austral winter of Argentina detected developmental arrests in nymphs of all instars [3]. Similarly, studies on laboratory-reared T. infestans submitted to temperatures and photoperiods found that under natural conditions, feeding ceases below 16–18 °C; the same is observed for reproduction [21,22]. Insects might regulate development forward, causing molting arrests or changing vital rates that translate into different ratios of developmental stages during the life cycle [7,23,24]. The developmental response of holometabolous insects to the photoperiod length has been studied, detecting regulations of seasonal morphs, growth rates, and diapause, among others [24,25,26]. Notwithstanding, no information has been reported on how and when development regulations are controlled in triatomine bugs (hemimetabolous insects), depending on climatic conditions.
Mepraia spinolai is considered the most epidemiologically important triatomine species in Chile due to its wide distribution, 26°–34° S, from the coast up to 3000 m.a.s.l. [27], an area with four marked climatic seasons [28], and the high frequency of T. cruzi infection in its populations [29,30,31]. This triatomine species mainly use stockpiles and bromeliads as refuge, where it coexists with small mammals and reptiles; both groups of vertebrates are included in its diet [32,33,34] and are part of the sylvatic transmission of T. cruzi in the semiarid Mediterranean ecosystem [35,36,37,38,39]. In addition, wild-caught M. spinolai individuals exhibit blood stealing between conspecifics, increasing the complexity of T. cruzi transmission under natural conditions [40].
In nature, M. spinolai populations exhibit a spatially patchy distribution with variable densities and a population structure of predominantly early-instar nymphs in late austral summer–fall, and one represented by all developmental stages in austral spring–early summer [29,41]. Adult females of this species are macropterous and adult males present three alary morphs: macropterous, brachypterous, and macropterous; mostly macropterous males invade human dwellings [42]. Reproduction occurs from late austral spring to early fall, when adults are present. During the warm seasons, the home range of M. spinolai colonies is larger and their blood sources are more diverse than in the cold seasons [32,43]. Under optimal laboratory rearing conditions, the generation time of M. spinolai varies from 9 to 10 months, while in nature—characterized by high thermal oscillation and variable photoperiods—it takes up to 12 months to molt to the fifth instar [19,44,45].
In this study, we examined the developmental process (molting events)—under optimal laboratory-controlled conditions (27 °C; 70% relative humidity (RH))—of wild-caught M. spinolai nymphs (first to fourth instars) either naturally infected with T. cruzi or uninfected. Nymphs were collected in late fall and winter (i.e., cooling period) and early and late spring (i.e., warming period) over three consecutive years (2016–2018). To this end, nymphs were fed ad libitum twice after capture to evaluate molting capacity. The goals of our study were: (i) to determine how natural infection with T. cruzi affects the molting capacity of M. spinolai nymphs of various instars collected in cooling and warming periods; and (ii) to determine if M. spinolai nymphs collected in cooling and warming periods regulate development at a particular nymphal stage.

2. Materials and Methods

2.1. Study Area and Triatomine Collection

Triatomine nymphs were collected from three localities (Las Chinchillas National Reserve, Los Pozos, and El Cuyano; Coquimbo Region, north-central Chile), covering a total area of 61.5 km2 (Figure 1). M. spinolai populations, defined as a group of bugs captured in an area of approximately 50 m2 within a specific period of time, were collected from different north-facing slopes in the area. The area is a hyper-endemic zone of Chagas disease, with M. spinolai the most abundant triatomine species, and characterized by a semiarid Mediterranean climate with rocky outcrops and scarce vegetation cover, mostly represented by bromeliads (Puya sp.) and cacti. Domestic (goats, cats, dogs, sheep), native (rodents, marsupials, foxes, bats), free-ranging introduced (European rabbits) mammals and lizard species can be found in this area, with several of these vertebrate species acting as blood meal of M. spinolai and T. cruzi reservoirs [30,34,35,38,39].
Triatomine collection was manually performed by one trained researcher over the course of one week in the cooling period and over the course of three days in the warming period, from 11:00 to 16:00 h in rocky outcrops, the period of maximum activity [46]. Triatomines were collected from rocks as they came closer to the researcher (i.e., without removing the rocks). Captured bugs were transported to the laboratory in polystyrene foam boxes, with folded papers as refuge, within one to two days after capture, and kept individually in plastic boxes in a climatic chamber at 27 °C, 70% RH, and a 14:10 h light:dark photoperiod until feeding.

2.2. Triatomine Fecal Sample Collection

Within the next two weeks, nymphs were fed with uninfected mice (Mus musculus) anesthetized with 2% sodium pentothal (Laboratorios RICHMOND, Buenos Aires, Argentina). Mice of 2 months of age were obtained from the vivarium of the Faculty of Medicine, University of Chile. Procedures of animal handling were performed according to the rules and with the permission of the Animal Ethics Committee of the University of Chile (CBA#0987-FMUCH-2016, CBA#1027-FMUCH-2019). After feeding, each nymph was classified by nymphal stage and kept separately in a plastic container with small compartments of 3.2 × 3.6 cm to allow for individual follow-up. Fecal samples were collected 30 min after feeding without pressing the abdomen to avoid internal damage, i.e., as a result of spontaneous defecation, and mixed with 100 μL of distilled water. This procedure was repeated for each nymph after a second feeding 40–45 days later. Nymphs were maintained in a growth chamber under the same conditions previously described. Death and ecdysis were recorded after each feeding. Only fed nymphs visually engorged and surviving the two feeding events were used for further analysis.

2.3. DNA Extraction and T. cruzi Infection Detection

Fecal samples were individually extracted in a final volume of 200 μL, using conditions already described by manufacturers of the EZNA Blood DNA kit (OMEGA BIO-TEK, Norcross, GA, USA). T. cruzi presence was tested three times by PCR analysis directed to minicircle kDNA with oligos 121 and 122, using different volumes of extracted DNA as templates in a final volume of 50 μL to improve detection [47,48]. A volume of 10 μL of each reaction mixture was run in agarose gel electrophoresis (Lafken, Santiago, Chile) and stained with ethidium bromide (Invitrogen, Waltham, MA, USA). Each PCR assay was repeated when negative results were obtained. In those samples, the extracted DNA was concentrated three times by evaporation, and then three new PCR assays were repeated using variable volumes of the concentrated DNA as a template. A negative sample resulted when all PCR assays failed to detect a 330 bp amplicon. This kind of amplicon (i.e., pattern of bands) is characteristic of T. cruzi minicircles, ensuring the amplification is not Trypanosoma rangeli, which also amplify with this couple of primers [49].

2.4. Statistical Analysis

Abiotic variables (temperatures and photoperiod) from cooling and warming periods were compared with t-tests. The proportion of T. cruzi-infected nymphs (all instars combined) from the cooling and warming periods were compared by χ2 tests, as well as the proportion of nymphs (all instars combined) molting twice (after two feeding events), depending on the infection status within each climatic period and vice versa (i.e., depending on the climatic period, keeping infection status constant).
For each developmental stage (first to fourth), the association between double molting and T. cruzi infection was tested by χ2 tests within the same climatic period (cooling or warming). The same test was used to evaluate the association between double molting and climatic period—for each developmental stage—within the same infection status (uninfected or T. cruzi-infected). The same test was used to examine the association between complete absence of molting (after two feeding events) and infection status (or climatic period), but considering all developmental stages combined due to the low number of non-molting nymphs. The significance level (or alpha level) considered statistically significant in this study was 0.05. Analyses were performed with R (version 3.6.0, R Development Core Team 2019) or JMP-Pro (version 14) (SAS Institute Inc., Cary, NC, USA)

3. Results

3.1. M. spinolai Populations and Climatic Conditions

Between 2016 and 2018, a total of eight M. spinolai populations were sampled, four in each climatic period (cooling: late fall and winter; warming: spring). A total of 343 and 366 nymphs were collected in the cooling and warming periods, respectively. Details on each M. spinolai population used in this study (period, date of capturing, total number of captures) and abiotic conditions (photoperiod, minimum and maximum temperatures one week prior to collection events) are summarized in Table 1. Only the mean maximum temperatures showed statistically significant difference between warming and cooling periods (mean min T° ± SE: cooling = 5.00 ± 0.42, warming = 5.30 ± 0.36, t(54) = −0.59, p = 0.280; mean max T° ± SE: cooling = 20.86 ± 0.62, warming: 23.23 ± 0.53, t(54) = −2.90, p = 0.003). As expected, the warming period presented, on average, more daylight hours than the cooling period (mean daylight hours ± SE: cooling = 10.30 ± 0.12, warming = 12.75 ± 0.46, t(6) = −5.18, p = 0.001). The complete dataset of minimum and maximum temperatures is available in Table S1.

3.2. Molting Events: Climatic Period and T. cruzi Infection

Overall, more than half of nymphs (all instars and infection status combined) from the cooling and warming periods molted only once, 63.9% and 60.9% after two feeding events, respectively. Only 27.4% and 31.4% of the nymphs from cooling and warming periods were capable of molting twice under the same conditions, respectively. Less than 10% of the nymphs were unable to molt (8.8% in cooling and 7.7% in warming). Details (of numbers) of nymphs by developmental stage (first to fourth) and type of molting (one, twice, none) in each M. spinolai population are shown in Table 2.
Regarding T. cruzi infection, nymphs (all instars combined) from the warming period were twice more infected with T. cruzi than those from the cooling period (56.8% vs. 21.9%, respectively; χ2 = 139.81, p < 0.0001). Within the warming period, a larger proportion of infected nymphs molted twice than those uninfected (36.1% vs. 25.3%, respectively; χ2 = 4.81, p = 0.028), but this molting behavior was not detected in nymphs from the cooling period (33.3% vs. 25.7%, respectively; χ2 = 1.70, p = 0.193). Altogether (combining all developmental stages from the same climatic period by infection status), regardless of the infection status, we did not detect statistically significant differences between cooling and warming periods in the proportion of nymphs exhibiting double molting (infected: 33.3% vs. 36.1%, respectively, χ2 = 0.1791, p = 0.672; uninfected: 25.7% vs. 25.3%, respectively, χ2 = 0.01, p = 0.922). Details on total number of nymphs by developmental stage (first to fourth instar), type of molting (one, twice, none), and T. cruzi infection combining populations from the same climatic period (cooling and warming) are summarized in Table 3. The complete dataset at the individual basis is available in Table S1.

3.3. Nymphs Molting Twice: Developmental Stage, T. cruzi Infection, and Climatic Period

Overall, 209 nymphs (first to fourth instars) molted twice after the two feeding events (94 from cooling and 115 from warming). Within the same climatic period, only infected second- and fourth-instar nymphs from the warming period showed a larger proportion of double molting compared to uninfected nymphs of the same instar (second instar: 50.0% vs. 26.8%, respectively, χ2 = 5.61, p = 0.018; fourth instar: 16.4% vs. 2.2%, respectively; χ2 = 5.59, p = 0.018; Figure 2). No nymphs of the remaining instars from the warming (first and third) and cooling (first to fourth) periods showed statistically significant differences between the probability of double molting depending on the infection status (p > 0.05).
Separated analyses by developmental stages showed that first-instar nymphs, regardless of their infection status, exhibited a larger proportion of double molting in the warming period compared to those from the cooling period (infected: 54.6% vs. 12.5%, respectively, χ2 = 8.45, p = 0.004; uninfected: 59.1% vs. 25.8%, respectively, χ2 = 11.92, p < 0.001; Figure 2). On the other hand, fourth-instar nymphs—regardless of their infection status—exhibited a larger proportion of double molting in the cooling period compared to those from the warming period (infected: 42.9% vs. 16.4%, respectively, χ2 = 4.75, p = 0.029; uninfected: 26.8% vs. 2.2%, respectively, χ2 = 10.82, p = 0.001; Figure 2). No other infected or uninfected nymphs of remaining instars (second and third) showed statistically significant differences between the probability of double molting depending on climatic period of collection (p > 0.05).

3.4. Non-Molting Nymphs: Infection and Climatic Period

Overall, 58 nymphs across all developmental stages did not molt after the two feeding events, 30 and 28 from the cooling and warming periods, respectively (Table 2). Within the same climatic period, we did not detect differences between infected and uninfected nymphs in their probability of not molting (cooling: 6.7% vs. 9.6%, respectively, χ2 = 0.63, p = 0.427; warming: 6.7% vs. 8.9%, respectively, χ2 = 0.58, p = 0.448; Table 3). Within the same infection status, we did not detect differences between nymphs from cooling and warming periods in their probability of not molting (infected: 6.7% for both climatic periods, χ2 = 0.0004, p = 0.95; uninfected: 9.6% vs. 8.9%, respectively, χ2 = 0.07, p = 0.792; Table 3). Because we did not detect differences by infection and collecting period in the frequency of non-molting nymphs, we tested for differences among nymphal stages (combining both infection status and periods). We detected overall differences among the frequency of non-molting nymphs depending on the nymphal stage (first: 2.4%; second: 6.2%; third: 12.6%; fourth: 12.4%; χ2 = 16.30, p = 0.001; Table 2).

4. Discussion

In this study, we have designed a simple biological test to study the development of field triatomines. Previously, we have performed follow-up studies in wild-caught M. spinolai after successive feedings to test the effect of T. cruzi infection on insect development and survivorship [50,51,52]. The gathered results from the mentioned studies showed that all nymphal stages are able to molt under laboratory-standard conditions. In the present study, nymphs were fed twice (the first feeding right after capture) and maintained under optimal abiotic conditions; therefore, any detected effect in the development should mostly depend on biotic or abiotic factors the insect experienced in the field. The results of first-to-fourth instar nymph development after the two feeding events generated three kinds of molting patterns: nymphs molting once (N = 442; 62.3%), molting twice (N = 209; 29.5%), or not molting at all (N = 58; 8.2%). This result suggests that M. spinolai nymphs are heterogeneous in nature regarding their nutritional and physiological conditions. The first group is split into two subgroups of insects, those molting after the first feeding and those after the second feeding. Previous studies using field triatomines in other latitudes have revealed that blood meals are often difficult to obtain and starvation seems to be frequent [53,54]. However, the mechanisms underlying our results of one-molt events are not straightforward.
Regarding the group of nymphs molting twice, these insects were more homogeneous since they probably molted with extensive abdominal distensions, after the two feeding events [55,56]. Overall, T. cruzi-infected nymphs (all stages combined) from the warming period presented double molting (i.e., maximum molting capacity) more frequently than equivalent uninfected nymphs. A detailed analysis for each nymphal stage revealed that second- and fourth-instar nymphs infected by T. cruzi presented higher molting capacity than equivalent uninfected insects, but only in those captured in the warming period. This result is surprising since this implies T. cruzi infection is associated with increased molting capacity, contrary to what is expected for a potentially detrimental pathogen. In fact, the experimental infection with T. cruzi of laboratory-reared M. spinolai (first generation), under optimal conditions, generated development retardment in the last three nymphal stages [19]. Other alterations in life history traits of T. cruzi-infected M. spinolai have been reported, including the shortening of defecation time, increasing biting rate, and faster approaching behavior to humans [57,58]. These results suggest that the presence of T. cruzi might favor parasitic transmission towards hosts [59]. The contrasting results between experimentally infected insects and naturally infected insects suggest that the origin of M. spinolai (laboratory-reared versus wild-caught nymphs), the type of T. cruzi infection (e.g., DTU identity, amount of T. cruzi entering with the blood meal, or frequency of infected blood meals), microbiota changes (i.e., bacteria composition after triatomine feeding), and/or the blood source are important factors that might explain, at least in part, the observed differences [60,61,62,63,64].
The higher molting capacity of second- and fourth-instar T. cruzi-infected nymphs found only in insects collected in the warming period is interesting. In this study, we showed that M. spinolai nymphs (all instars combined) exhibited higher T. cruzi prevalence in this period, and since insects are ectotherms, this translates to them having higher metabolic activity and mobility in the warming period. In parallel, T. cruzi also positively responds to higher temperatures; therefore, infected M. spinolai nymphs compensate the exerted T. cruzi exploitation by means of higher feeding efficiency (i.e., more voracious nymphs searching for more feeding opportunities, probably until engorgement), triggering more molting events and development forward [43,57,58,65,66,67].
Regarding the climatic period, nymphs did not differ between cooling and warming periods in the proportion of double molting, regardless of infection status, when all developmental stages were analyzed in combination. However, a detailed analysis separated by nymphal stage revealed that first-instar nymphs—regardless of their infection status—presented maximum molting capacity, reaching the third nymphal stage, when collected in the warming period. On the other hand, fourth-instar nymphs showed a maximal molting capacity, reaching the adult stage (i.e., incomplete metamorphosis), when collected in the cooling period. To the best of our knowledge, no other studies using field triatomines have examined the molting pattern separated by nymphal stage. These results suggest that development is controlled forward in first-instar nymphs, increasing the proportion of the next nymphal stages (e.g., second- and third-instar) in M. spinolai populations during warmer periods, and in the fourth-instar nymphs, increasing double molting in the cooling period, translating into adult occurrence when the warming period initiates in early spring. Adults of M. spinolai are observed in the field from mid-spring until late summer when the appropriate abiotic conditions for reproduction occur, and vertebrate abundance is optimal for blood meals [41,43]. In other latitudes, the population dynamics of Triatoma dimidiata in the field revealed positive associations of nymphs’ abundance and the presence of adults with the occurrence of the dry and hot seasons of Mexico [6]. In austral latitudes of Argentina, T. infestans fecundity and population numbers are associated with rising temperatures in early spring [68].
Developmental events in holometabolous insects such as egg hatching, pupation, and metamorphosis occur only once in their lives (circannual), and clock control could explain how a rhythm in such an event is also produced by a circadian clock [69]. In this line of evidence, the response to photoperiod has been identified in several insect species; it is demonstrated to be involved in growth rate regulation, migration strategy, and diapause [25]. We propose that environmental temperatures, the photoperiod, or both factors could regulate the developmental process of this hemimetabolous wild triatomine species.
In this study, a third group of nymphs are those that did not molt after either of the two feeding occasions. This is a small number of insects, prevalent neither in the warming period nor the cooling period but, interestingly, first-to-fourth-instar nymphs within the same M. spinolai population presented this behavior. Notwithstanding, a higher proportion of third- and fourth-instar nymphs presented molting absence compared to first- and second-instar nymphs. This result suggests that in our study, molting failure might be due to environmental stochasticity, and not to demographic stochasticity, since the latter phenomenon arises only in individuals of a given developmental stage [23]. Interestingly, three out of the five M. spinolai populations showing absence of molting were time-related. Specifically, C3, C4, and W4—all 2018′ populations—presented maximum rates of molting failure in a year with an unusually warm fall [48]. Therefore, our results of molting failure could be interpreted as diapause by environmental stochasticity, which seems to end after an unknown period of time after failure.
In conclusion, development is differentially controlled in nymphs from first and fourth stages depending on the climatic period of collection, and T. cruzi infection changes development forward only in nymphs of second and fourth instars of the warming period. Therefore, the effect of the climatic period and T. cruzi infection on the development of the austral kissing bug M. spinolai is an instar-dependent phenomenon, highlighting the occurrence of finely synchronized processes at different moments of the life cycle of such an hemimetabolous insect as this wild triatomine.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/insects14030272/s1, Table S1: Climatic records from Las Chinchillas National Reserve, CONAF, Chile, and dataset of captured nymphs, developmental stage, and infection status at an individual basis for each population in each season.

Author Contributions

Conceptualization, A.S.; investigation, A.S., J.B., G.M.-V., S.O. and A.L.; methodology, A.S., J.B., G.M.-V., S.O. and A.L.; data curation, C.B.-M.; formal analysis, C.B.-M.; supervision, A.S.; writing—original draft, A.S. and C.B.-M.; writing—review and editing, A.S. and C.B.-M., funding acquisition, A.S.; project administration, A.S. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Chilean National Agency for Research and Development (ANID) program’s National Fund for Scientific and Technological Development (FONDECYT), grant numbers 1190392 (A.S.) and 1221045 (C.B.-M.). Partial support was obtained from Vicerrectoría de Investigación y Desarrollo (VID) from the Universidad de Chile, Enlace-FONDECYT-ENL01/21 (C-B.-M.).

Institutional Review Board Statement

Procedures of animal handling were performed according to the rules and with the permission of the Animal Ethics Committee of the University of Chile CBA#0987-FMUCH-2016, CBA#1027-FMUCH-2019.

Informed Consent Statement

Not applicable.

Data Availability Statement

The data presented in this study are available in the manuscript and in its Supplementary Material (Table S1).

Acknowledgments

We especially thank Nora Peña for her invaluable work collecting M. spinolai in the field, and Esteban San Juan for compiling the map and figure. We are very grateful to Jorge Silva (National Forest Corporation, CONAF, Illapel, Chile) for providing climatic data from Las Chinchillas National Reserve, and José Fernández from the Faculty of Medicine, University of Chile, for providing bibliographic resources.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Telleria, J.; Tibayrenc, M. American Trypanosomiasis Chagas Disease: One Hundred Years of Research, 2nd ed.; Academic Press: Amsterdam, The Netherlands, 2017; p. 844. [Google Scholar]
  2. Rabinovich, J.E. Vital statistics of Triatominae (Hemiptera: Reduviidae) under laboratory conditions. I. Triatoma infestans Klug. J. Med. Entomol. 1972, 9, 351–370. [Google Scholar] [CrossRef] [PubMed]
  3. Gorla, D.E.; Schofield, C.J. Population dynamics of Triatoma infestans under natural climatic conditions in the Argentine Chaco. Med. Vet. Entomol. 1989, 3, 179–194. [Google Scholar] [CrossRef] [PubMed]
  4. Figueiras, A.N.L.; Lazzari, C.R. Aggregation in the haematophagous bug Triatoma infestans: A novel assembling factor. Physiol. Entomol. 1998, 23, 33–37. [Google Scholar] [CrossRef]
  5. Lorenzo, M.G.; Guarneri, A.A.; Pires, H.H.; Diotaiuti, L.; Lazzari, C.R. Microclimatic properties of the Triatoma brasiliensis habitat. Cad. Saude Publica 2000, 16, 69–74. [Google Scholar] [CrossRef] [PubMed]
  6. Dumonteil, E.; Gourbiere, S.; Barrera-Perez, M.; Rodriguez-Felix, E.; Ruiz-Pina, H.; Banos-Lopez, O.; Ramirez-Sierra, M.J.; Menu, F.; Rabinovich, J.E. Geographic distribution of Triatoma dimidiata and transmission dynamics of Trypanosoma cruzi in the Yucatan peninsula of Mexico. Am. J. Trop. Med. Hyg. 2002, 67, 176–183. [Google Scholar] [CrossRef] [PubMed]
  7. Menu, F.; Ginoux, M.; Rajon, E.; Lazzari, C.R.; Rabinovich, J.E. Adaptive developmental delay in Chagas disease vectors: An evolutionary ecology approach. PLoS Negl. Trop. Dis. 2010, 4, e691. [Google Scholar] [CrossRef] [PubMed]
  8. Lazzari, C.R.; Pereira, M.H.; Lorenzo, M.G. Behavioural biology of Chagas disease vectors. Mem. Inst. Oswaldo Cruz 2013, 108, 34–47. [Google Scholar] [CrossRef]
  9. Pelosse, P.; Kribs-Zaleta, C.M.; Ginoux, M.; Rabinovich, J.E.; Gourbière, S.; Menu, F. Influence of vectors’ risk-spreading strategies and environmental stochasticity on the epidemiology and evolution of vector-borne diseases: The example of Chagas’ disease. PLoS ONE 2013, 8, e70830. [Google Scholar] [CrossRef]
  10. Elliot, S.L.; Adler, F.R.; Sabelis, M.W. How virulent should a parasite be to its vector? Ecology 2003, 84, 2568–2574. [Google Scholar] [CrossRef]
  11. Elliot, S.L.; Rodrigues, J.d.O.; Lorenzo, M.G.; Martins-Filho, O.A.; Guarneri, A.A. 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. [Google Scholar] [CrossRef] [PubMed]
  12. Schofield, C.J. The role of blood intake in density regulation of populations of Triatoma infestans (Klug) (Hemiptera:Reduviidae). Bull. Ent. Res. 1982, 72, 617–629. [Google Scholar] [CrossRef]
  13. Wigglesworth, V.B. Factors controlling moulting and ‘metamorphosis’ in an insect. Nature 1934, 133, 725–726. [Google Scholar] [CrossRef]
  14. Steel, C.G.; Vafopoulou, X. Circadian orchestration of developmental hormones in the insect, Rhodnius prolixus. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 2006, 144, 351–364. [Google Scholar] [CrossRef] [PubMed]
  15. Reis dos Santos, J.; Lacombe, D. Estudos relativos à duraçao da ecdise e ovopisiçao de Triatoma infestans infectado pela Trypanosoma cruzi. An. Acad. Brasil. Ciênc. 1985, 57, 127. [Google Scholar]
  16. Schaub, G.A. Developmental time and mortality of larvae of Triatoma infestans infected with Trypanosoma cruzi. Trans. R. Soc. Trop. Med. Hyg. 1988, 82, 94–96. [Google Scholar] [CrossRef]
  17. Lima, M.M.; Pereira, J.B.; Santos, J.A.A.; Pinto, Z.T.; Braga, M.V. Development and reproduction of Panstrongylus megistus (Hemiptera: Reduviidae) infected with Trypanosoma cruzi, under laboratory conditions. Ann. Entomol. Soc. Am. 1992, 85, 458–461. [Google Scholar] [CrossRef]
  18. Oliveira, T.G.; Carvalho-Costa, F.A.; Gomes, T.F.; Sarquis, O.; Sposina, R.; Lima, M.M. Developmental and reproductive patterns of Triatoma brasiliensis infected with Trypanosoma cruzi under laboratory conditions. Mem. Inst. Oswaldo Cruz 2010, 105, 1057–1060. [Google Scholar] [CrossRef]
  19. 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, 505–510. [Google Scholar] [CrossRef]
  20. Guarneri, A.A.; Lazzari, C.; Xavier, A.A.P.; Diotaiuti, L.; Lorenzo, M.G. The effect of temperature on the behaviour and development of Triatoma brasiliensis. Physiol. Entomol. 2003, 28, 185–191. [Google Scholar] [CrossRef]
  21. Joerg, M.E. Influencia de temperaturas fijas en períodos anuales sobre metamorfosis y fertilidad de Triatoma infestans. Bol. Chil. Parasitol. 1962, 17, 17–19. [Google Scholar] [PubMed]
  22. Catalá, S. The biting rate of Triatoma infestans in Argentina. Med. Vet. Entomol. 1991, 5, 325–333. [Google Scholar] [CrossRef]
  23. Lande, R. Genetics and demography in biological conservation. Science 1988, 241, 1455–1460. [Google Scholar] [CrossRef] [PubMed]
  24. Numata, H.; Miyazaki, Y.; Ikeno, T. Common features in diverse insect clocks. Zool. Lett. 2015, 1, 10. [Google Scholar] [CrossRef]
  25. Saunders, D.S.; Bertossa, R.C. Deciphering time measurement: The role of circadian ‘clock’ genes and formal experimentation in insect photoperiodism. J. Insect. Physiol. 2011, 57, 557–566. [Google Scholar] [CrossRef]
  26. Goto, S.G. Roles of circadian clock genes in insect photoperiodism. Entomol. Sci. 2013, 16, 1–16. [Google Scholar] [CrossRef]
  27. Garrido, R.; Bacigalupo, A.; Peña-Gómez, F.; Bustamante, R.O.; Cattan, P.E.; Gorla, D.; Botto-Mahan, C. Potential impact of climate change on geographical distribution of two wild vectors of Chagas disease in Chile: Mepraia spinolai and Mepraia gajardoi. Parasit. Vectors 2019, 12, 478. [Google Scholar] [CrossRef] [PubMed]
  28. di Castri, F.; Hajek, E.R. Bioclimatología de Chile; Editorial Universidad Católica de Chile: Santiago, Chile, 1976. [Google Scholar]
  29. Ihle-Soto, C.; Costoya, E.; Correa, J.P.; Bacigalupo, A.; Cornejo-Villar, B.; Estadella, V.; Solari, A.; Ortiz, S.; Hernández, H.J.; Botto-Mahan, C.; et al. Spatio-temporal characterization of Trypanosoma cruzi infection and discrete typing units infecting hosts and vectors from non-domestic foci of Central Chile. PLoS Negl. Trop. Dis. 2019, 13, e7170. [Google Scholar] [CrossRef]
  30. Botto-Mahan, C.; Bacigalupo, A.; Correa, J.P.; Fontúrbel, F.E.; Cattan, P.E.; Solari, A. Prevalence, infected density or individual probability of infection? Assessing vector infection risk in the wild transmission of Chagas disease. Proc. R. Soc. B 2020, 287, 20193018. [Google Scholar] [CrossRef]
  31. San Juan, E.; Araya-Donoso, R.; Sandoval-Rodríguez, A.; Yáñez-Meza, A.; Quiroga, N.; Botto-Mahan, C. Lizards and rabbits may increase Chagas infection risk in the Mediterranean-type ecosystem of South America. Sci. Rep. 2020, 10, 1853. [Google Scholar] [CrossRef] [PubMed]
  32. Canals, M.; Cruzat, L.; Molina, M.C.; Ferreira, A.; Cattan, P. Blood host source of Mepraia spinolai (Heteroptera, Reduviidae) wild vector of Chagas disease in Chile. J. Med. Entomol. 2001, 38, 303–307. [Google Scholar] [CrossRef]
  33. Chacon, F.; Bacigalupo, A.; Quiroga, M.; Ferreira, A.; Cattan, P.; Ramirez-Toloza, G. Feeding profile of Mepraia spinolai, a sylvatic vector of Chagas disease in Chile. Acta Trop. 2016, 162, 171–173. [Google Scholar] [CrossRef]
  34. De Bona, S.; Correa, J.P.; San Juan, E.; Estay-Olea, D.; Quiroga, N.; Bacigalupo, A.; Araya-Donoso, R.; Botto-Mahan, C. Opportunistic or selective? stage-dependent feeding behavior in a wild vector of Chagas disease. Int. J. Parasitol. 2023, 53, 55–64. [Google Scholar] [CrossRef]
  35. Rozas, M.; Botto-Mahan, C.; Coronado, X.; Ortiz, S.; Cattan, P.E.; Solari, A. Co-existence of Trypanosoma cruzi genotypes in wild and peridomestic mammals in Chile. Am. J. Trop. Med. Hyg. 2007, 77, 647–653. [Google Scholar] [CrossRef]
  36. Correa, J.P.; Bacigalupo, A.; Fontúrbel, F.; Oda, E.; Cattan, P.E.; Solari, A.; Botto-Mahan, C. Spatial distribution of an infectious disease in a native small mammal community. Sci. Nat. 2015, 102, 51. [Google Scholar] [CrossRef]
  37. Correa, J.P.; Bacigalupo, A.; Yefi-Quinteros, E.; Rojo, G.; Solari, A.; Cattan, P.E.; Botto-Mahan, C. Trypanosomatid infections among vertebrates of Chile: A systematic review. Pathogens 2020, 9, 661. [Google Scholar] [CrossRef]
  38. Quiroga, N.; Campos-Soto, R.; Yañez-Meza, A.; Rodríguez-San Pedro, A.; Allendes, J.L.; Bacigalupo, A.; Botto-Mahan, C.; Correa, J.P. Trypanosoma cruzi DNA in Desmodus rotundus (common vampire bat) and Histiotus montanus (small big-eared brown bat) from Chile. Acta Trop. 2022, 225, 106206. [Google Scholar] [CrossRef]
  39. Botto-Mahan, C.; Correa, J.P.; Araya-Donoso, R.; Farías, F.; Quiroga, N.; Reyes-Olivares, C.; San Juan, E.; Campos-Soto, R.; González-Acuña, D. Lizards as silent hosts of Trypanosoma cruzi. Emerg. Infect. Dis. 2022, 28, 1250–1253. [Google Scholar] [CrossRef]
  40. Garrido, R.; Campos-Soto, R.; Quiroga, N.; Botto-Mahan, C. Bloodmeal-stealing in wild-caught Mepraia spinolai (Hemiptera: Reduviidae), a sylvatic vector of Trypanosoma cruzi. Ecol. Entomol. 2021, 46, 681–683. [Google Scholar] [CrossRef]
  41. Canals, M.; Ehrenfeld, M.; Solís, R.; Cruzat, L.; Pinochet, A.; Tapia, C.; Cattan, P.E. Comparative biology of Mepraia spinolai in laboratory and field conditions: Five years study. Parasitol. Dia. 1998, 22, 72–78. [Google Scholar] [CrossRef]
  42. Frías-Lasserre, D.; González, C.R.; Valenzuela, C.R.; de Carvalho, D.B.; Oliveira, J.; Canals, M.; da Rosa, J.A. Wing polymorphism and Trypanosoma cruzi infection in wild, peridomestic, and domestic collections of Mepraia spinolai (Hemiptera: Reduviidae) from Chile. J. Med. Entomol. 2017, 54, 1061–1066. [Google Scholar] [CrossRef]
  43. Botto-Mahan, C.; Cattan, P.E.; Canals, M.; Acuña, M. Seasonal variation in the home range and host availability of the blood-sucking insect Mepraia spinolai in wild environment. Acta Trop. 2005, 95, 160–163. [Google Scholar] [CrossRef]
  44. Lent, H.; Jurberg, J. Algumas informações sôbre Triatoma spinolai Porter, 1934 com um estudo sôbre as genitálias externas (Hemiptera, Reduviidae). Rev. Brasil Biol. 1967, 28, 499–520. [Google Scholar]
  45. Ehrenfeld, M.J.; Canals, M.; Cattan, P.E. Population parameters of Triatoma spinolai (Heteroptera: Reduviidae) under different environmental conditions and densities. J. Med. Entomol. 1998, 35, 740–744. [Google Scholar] [CrossRef]
  46. Canals, M.; Solis, R.; Valderas, J.; Ehrenfeld, M.; Cattan, P.E. Preliminary studies on temperature selection and activity cycles of Triatoma infestans and T. spinolai (Heteroptera: Reduviidae), Chilean vectors of Chagas’ disease. J. Med. Entomol. 1997, 34, 11–17. [Google Scholar] [CrossRef]
  47. Wincker, P.; Britto, C.; Pereira, J.B.; Cardoso, M.A.; Oelemann, O.; Morel, C.M. Use of a simplified polymerase chain reaction procedure to detect Trypanosoma cruzi in blood samples from chronic chagasic patients in a rural endemic area. Am. J. Trop. Med. Hyg. 1994, 51, 771–777. [Google Scholar] [CrossRef] [PubMed]
  48. Cortés, V.; Cruz, A.; Onetti, S.; Kinzel, D.; Garcia, J.; Ortiz, S.; López, A.; Cattan, P.E.; Botto-Mahan, C.; Solari, A. Trypanosoma cruzi infection follow-up in a sylvatic vector of Chagas disease: Comparing early and late stage nymphs. PLoS Negl. Trop. Dis. 2021, 15, e0009729. [Google Scholar] [CrossRef]
  49. Guhl, F.; Jaramillo, C.; Carranza, J.C.; Vallejo, G.A. Molecular characterization and diagnosis of Trypanosoma cruzi and T. rangeli. Arch. Med. Res. 2002, 33, 362–370. [Google Scholar] [CrossRef]
  50. Egaña, C.; Pinto, R.; Vergara, F.; Ortiz, S.; Campos-Soto, R.; Solari, A. Fluctuations in Trypanosoma cruzi discrete typing unit composition in two naturally infected triatomines: Mepraia gajardoi and M. spinolai after laboratory feeding. Acta Trop. 2016, 160, 9–14. [Google Scholar] [CrossRef]
  51. Garcia, V.; Graterol, J.; López, A.; Ortiz, S.; Solari, A. Influence of Trypanosoma cruzi (Kinetoplastida: Trypanosomatidae) infection on mortality of the sylvatic Triatomine vector, Mepraia spinolai (Heteroptera: Reuviidae), under fasting. J. Med. Entomol. 2019, 56, 1384–1388. [Google Scholar] [CrossRef]
  52. McCabe, A.; Yañez, F.; Pinto, R.; López, A.; Ortiz, S.; Muñoz-San Martin, C.; Botto-Mahan, C.; Solari, A. Survivorship of wild caught Mepraia spinolai nymphs: The effect of seasonality and Trypanosoma cruzi infection after feeding and fasting in the laboratory. Infect. Genet. Evol. 2019, 71, 197–204. [Google Scholar] [CrossRef]
  53. Noireau, F.; Dujardin, J.P. Flight and nutritional status of sylvatic Triatoma sordida and Triatoma guasayana. Mem. Inst. Oswaldo Cruz 2001, 96, 385–389. [Google Scholar] [CrossRef]
  54. Sarquis, O.; Carvalho-Costa, F.A.; Oliveira, L.S.; Duarte, R.; Andrea, P.S.D.; de Oliveira, T.G.; Lima, M.M. Ecology of Triatoma brasiliensis in northeastern Brazil: Seasonal distribution, feeding resources, and Trypanosoma cruzi infection in a sylvatic population. J. Vector Ecol. 2010, 35, 385–394. [Google Scholar] [CrossRef]
  55. Anwyl, R. The structure and properties of an abdominal stretch receptor in Rhodnius prolixus. J. Insect Physiol. 1972, 18, 2143–2147, 2149–2153. [Google Scholar] [CrossRef]
  56. Chiang, R.G.; Davey, K.G. A novel receptor capable of monitoring applied pressure in the abdomen of an insect. Science 1988, 241, 1665–1667. [Google Scholar] [CrossRef]
  57. Botto-Mahan, C.; Cattan, P.E.; Medel, R. Chagas disease parasite induces behavioural changes in the kissing bug Mepraia spinolai. Acta Trop. 2006, 98, 219–223. [Google Scholar] [CrossRef]
  58. Estay-Olea, D.; de Bona, S.; Bacigalupo, A.; Quiroga, N.; San Juan, E.; Correa, J.P.; Solari, A.; Botto-Mahan, C. Trypanosoma cruzi could affect wild triatomine approaching behaviour to humans by altering vector nutritional status: A field test. Acta Trop. 2020, 210, 105574. [Google Scholar] [CrossRef]
  59. Córdoba-Aguilar, A. Chagas bugs and Trypanosoma cruzi: Puppets and puppeteer? Acta Trop. 2020, 211, 105600. [Google Scholar] [CrossRef]
  60. Díaz, S.; Villavicencio, B.; Correia, N.; Costa, J.; Haag, K.L. Triatomine bugs, their microbiota and Trypanosoma cruzi: Asymmetric responses of bacteria to an infected blood meal. Parasites Vectors 2016, 9, 636. [Google Scholar] [CrossRef]
  61. Oliveira, J.L.; Cury, J.C.; Gurgel-Gonçalves, R.; Bahia, A.C.; Monteiro, F.A. Field-collected Triatoma sordida from Central Brazil display high microbiota diversity that varies with regard to developmental stage and intestinal segmentation. PLoS Negl. Trop. Dis. 2018, 12, e0006709. [Google Scholar] [CrossRef]
  62. Guarneri, A.A.; Schaub, G.A. Interaction of Triatomines with their bacterial microbiota and Trypanosomes. In Triatominae. The Biology of Chagas Disease Vectors; Guarneri, A., Lorenzo, M., Eds.; Springer Nature: Cham, Switzerland, 2021; pp. 345–386. [Google Scholar] [CrossRef]
  63. Vázquez, D.P.; Canale, D.; Gürtler, R.E. Effects of non-susceptible hosts on the infection with Trypanosoma cruzi of the vector Triatoma infestans: An experimental model. Mem. Inst. Oswaldo Cruz 1999, 94, 413–419. [Google Scholar] [CrossRef]
  64. de Fuentes-Vicente, J.A.; Gutiérrez-Cabrera, A.E.; Flores-Villegas, A.L.; Lowenberger, C.; Benelli, G.; Salazar-Schettino, P.M.; Córdoba-Aguilar, A. What makes an effective Chagas disease vector? Factors underlying Trypanosoma cruzi-triatomine interactions. Acta Trop. 2018, 183, 23–31. [Google Scholar] [CrossRef]
  65. Asin, S.; Catala, S. Development of Trypanosoma cruzi in Triatoma infestans: Influence of temperature and blood consumption. J. Parasitol. 1995, 81, 1–7. [Google Scholar] [CrossRef]
  66. Ferreira, R.C.; Teixeira, C.F.; de Sousa, V.F.A.; Guarneri, A.A. Effect of temperature and vector nutrition on the development and multiplication of Trypanosoma rangeli in Rhodnius prolixus. Parasitol. Res. 2018, 117, 1737–1744. [Google Scholar] [CrossRef]
  67. Tamayo, L.D.; Guhl, F.; Vallejo, G.A.; Ramírez, J.D. The effect of temperature increase on the development of Rhodnius prolixus and the course of Trypanosoma cruzi metacyclogenesis. PLoS Negl. Trop. Dis. 2018, 12, e0006735. [Google Scholar] [CrossRef]
  68. Castañera, M.B.; Aparicio, J.P.; Gürtler, R.E. A stage-structured stochastic model of the population dynamics of Triatoma infestans, the main vector of Chagas disease. Ecol. Model. 2003, 162, 33–53. [Google Scholar] [CrossRef]
  69. Pittendrigh, C.S. The circadian oscillation in Drosophila pseudosobscura pupae: A model for the photoperiodic clock. Z. Pflanzenphysiol. 1966, 54, 275–307. [Google Scholar]
Insects 14 00272 g001 550
Figure 1. Map of Chile, South America, showing the area of Mepraia spinolai collection and the location of the Weather Station at Las Chinchillas National Reserve, CONAF, Coquimbo Region. On the right side: view of a collecting site (top), M. spinolai nymph (middle), and adult female (bottom).
Figure 1. Map of Chile, South America, showing the area of Mepraia spinolai collection and the location of the Weather Station at Las Chinchillas National Reserve, CONAF, Coquimbo Region. On the right side: view of a collecting site (top), M. spinolai nymph (middle), and adult female (bottom).
Insects 14 00272 g001
Insects 14 00272 g002 550
Figure 2. Percentage of Trypanosoma cruzi-infected and uninfected Mepraia spinolai nymphs molting twice after two feeding events, combining the same developmental stage of populations from the cooling or warming periods for the three years of collection (2016–2018). Blue and red bars represent nymphs from the cooling and warming periods, respectively. Hatched bars represent T. cruzi-infected nymphs. Black contrasts represent comparisons between nymphs of different periods but with the same infection status. Red contrasts represent comparisons between nymphs of different infection status within the same climatic period. Only statistically significant contrasts are shown. Significance levels: * < 0.05, ** < 0.01, *** < 0.001.
Figure 2. Percentage of Trypanosoma cruzi-infected and uninfected Mepraia spinolai nymphs molting twice after two feeding events, combining the same developmental stage of populations from the cooling or warming periods for the three years of collection (2016–2018). Blue and red bars represent nymphs from the cooling and warming periods, respectively. Hatched bars represent T. cruzi-infected nymphs. Black contrasts represent comparisons between nymphs of different periods but with the same infection status. Red contrasts represent comparisons between nymphs of different infection status within the same climatic period. Only statistically significant contrasts are shown. Significance levels: * < 0.05, ** < 0.01, *** < 0.001.
Insects 14 00272 g002
Table
Table 1. Total number of Mepraia spinolai nymphs (first to fourth stage) captured in each collection event that survived the two feeding procedures. Photoperiod (L:D) at the time of capturing and mean (±SE) minimum/maximum temperatures (°C) one week before collection are indicated.
Table 1. Total number of Mepraia spinolai nymphs (first to fourth stage) captured in each collection event that survived the two feeding procedures. Photoperiod (L:D) at the time of capturing and mean (±SE) minimum/maximum temperatures (°C) one week before collection are indicated.
Population PeriodMonth/YearNº of CapturesL:D Min T°Max T°
C1CoolingMay 201679 10.5:13.58.37 ± 0.5321.51 ± 0.99
C2CoolingJuly 201760 10.1:13.94.34 ± 0.1517.71 ± 0.79
C3CoolingJune 201810010.1:13.94.29 ± 0.2224.00 ± 1.07
C4CoolingAugust 201810410.5:13.52.91 ± 0.2920.20 ± 0.50
W1WarmingOctober 201610012.8:11.25.09 ± 0.2724.57 ± 0.91
W2WarmingSeptember 20174711.5:12.53.73 ± 0.6320.34 ± 0.71
W3WarmingNovember 20174013.7:10.37.91 ± 0.2824.91 ± 0.76
W4WarmingOctober 201817913.0:11.04.49 ± 0.2023.09 ± 1.02
Nº: number; T°: temperature.
Table
Table 2. Mepraia spinolai nymphs molting once, twice, or not at all after the two feeding events for each population captured during the cooling and warming periods. Total numbers of nymphs are ordered by developmental stage (first to fourth stage; I to IV) and type of molting (one, two, or none) is shown in the same order. Total number represents the developmental stage at capturing.
Table 2. Mepraia spinolai nymphs molting once, twice, or not at all after the two feeding events for each population captured during the cooling and warming periods. Total numbers of nymphs are ordered by developmental stage (first to fourth stage; I to IV) and type of molting (one, two, or none) is shown in the same order. Total number represents the developmental stage at capturing.
Nº Molting Events
PopulationNymphal StageTotal NumberOneTwoNone
C1I-II-III-IV16-28-13-2214-24-12-142-4-1-80-0-0-0
C2I-II-III-IV24-16-10-1018-13-7-46-2-2-60-1-1-0
C3I-II-III-IV21-31-38-1010-13-29-48-13-1-13-5-8-5
C4I-II-III-IV15-43-33-1315-17-15-100-22-16-20-4-2-1
W1I-II-III-IV28-46-20-611-29-12-516-14-2-11-3-6-0
W2I-II-III-IV15-16-5-112-2-4-313-14-1-80-0-0-0
W3I-II-III-IV18-12-5-55-7-3-313-5-2-20-0-0-0
W4I-II-III-IV27-33-35-8419-21-27-708-11-5-00-1-3-14
Nº: number; C: cooling; W: warming.
Table
Table 3. Total number of Mepraia spinolai nymphs (first to fourth stage; I to IV) molting once, twice, or not molting after two feeding events, combining the same developmental stage of populations from the cooling or warming periods for the three years of collection (2016–2018). Parentheses indicate the number of infected nymphs within each total.
Table 3. Total number of Mepraia spinolai nymphs (first to fourth stage; I to IV) molting once, twice, or not molting after two feeding events, combining the same developmental stage of populations from the cooling or warming periods for the three years of collection (2016–2018). Parentheses indicate the number of infected nymphs within each total.
Molting Events
Cooling PeriodWarming Period
Nymphal StageOneTwoNoneOneTwoNone
I57 (14)16 (2)3 (0)37 (19)50 (24)1 (1)
II67 (9)41 (9)10 (3)59 (31)44 (33)4 (2)
III63 (15)20 (8)11 (1)46 (26)10 (8)9 (3)
IV32 (7)17 (6)6 (1)81 (43)11 (10)14 (8)
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Botzotz, J.; Méndez-Valdés, G.; Ortiz, S.; López, A.; Botto-Mahan, C.; Solari, A. Natural Trypanosoma cruzi Infection and Climatic Season Influence the Developmental Capacity in Field-Caught Mepraia spinolai Nymphs. Insects 2023, 14, 272. https://doi.org/10.3390/insects14030272

AMA Style

Botzotz J, Méndez-Valdés G, Ortiz S, López A, Botto-Mahan C, Solari A. Natural Trypanosoma cruzi Infection and Climatic Season Influence the Developmental Capacity in Field-Caught Mepraia spinolai Nymphs. Insects. 2023; 14(3):272. https://doi.org/10.3390/insects14030272

Chicago/Turabian Style

Botzotz, Juan, Gabriel Méndez-Valdés, Sylvia Ortiz, Angélica López, Carezza Botto-Mahan, and Aldo Solari. 2023. "Natural Trypanosoma cruzi Infection and Climatic Season Influence the Developmental Capacity in Field-Caught Mepraia spinolai Nymphs" Insects 14, no. 3: 272. https://doi.org/10.3390/insects14030272

APA Style

Botzotz, J., Méndez-Valdés, G., Ortiz, S., López, A., Botto-Mahan, C., & Solari, A. (2023). Natural Trypanosoma cruzi Infection and Climatic Season Influence the Developmental Capacity in Field-Caught Mepraia spinolai Nymphs. Insects, 14(3), 272. https://doi.org/10.3390/insects14030272

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