Journal of Invertebrate Pathology xxx (2010) xxx–xxx

Contents lists available at ScienceDirect

Journal of Invertebrate Pathology

journal homepage: www.elsevier .com/ locate/ j ip

A standardizable protocol for infection of Rhodnius prolixus with Trypanosomarangeli, which mimics natural infections and reveals physiological effectsof infection upon the insect

Luciana L. Ferreira a, Marcelo G. Lorenzo a, Simon L. Elliot b, Alessandra A. Guarneri a,*

a Centro de Pesquisa René Rachou, Avenida Augusto de Lima, 1715, CEP 30190-002, Belo Horizonte, MG, Brazilb Department of Animal Biology, Federal University of Viçosa, Campus Universitário, CEP 36570-000, Viçosa, MG, Brazil

a r t i c l e i n f o

Article history:Received 9 March 2010Accepted 18 May 2010Available online xxxx

Keywords:RhodniusTrypanosoma rangeliHost–parasite interactionHost manipulationBiological cycleLipidFat bodyHaemolymph volume

0022-2011/$ - see front matter � 2010 Elsevier Inc. Adoi:10.1016/j.jip.2010.05.013

* Corresponding author. Fax: +55 31 3295 3115.E-mail address: guarneri@cpqrr.fiocruz.br (A.A. Gu

Please cite this article in press as: Ferreira, L.L.,natural infections and reveals physiological effe

a b s t r a c t

Trypanosoma rangeli is a protozoan parasite that shares hosts – mammals and triatomines – with Trypan-osoma cruzi, the etiological agent of Chagas disease. Although T. rangeli is customarily considered to benon-pathogenic to human hosts, it is able to produce pathologies in its invertebrate hosts. However,advances are hindered by a lack of standardization of infection procedures and these pathologies needdocumentation. To establish a suitable, and standardizable, infection protocol, the duration of the fourthinstar was evaluated in nymphs infected by injection into the thorax with different concentrations of par-asites, and compared with nymphs infected naturally (i.e. orally). We demonstrate that delays in moultwere attributable to the presence of the parasite in the haemolymph (vs. the gut) and propose that theprotocol presented here simulates closely natural infections. This methodology was then used for theevaluation of physiological parameters and several hitherto unreported effects of T. rangeli infection onRhodnius prolixus were revealed. Haemolymph volume was greater in infected than uninfected nymphsbut this alteration could not be attributed to water retention, since infected insects lost the same amountof water as controls. However, we found that lipid content and fat body weight were both increased ininsects infected by T. rangeli. We propose that this is due to the parasite’s sequestration of host blood lip-ids and carrier proteins. With these findings, we have taken a few first steps to unravelling physiologicaldetails of the host–parasite interaction. We suggest future directions towards a fuller understanding ofmechanistic and adaptive aspects of triatomine–trypanosomatid interactions.

� 2010 Elsevier Inc. All rights reserved.

1. Introduction

Rhodnius prolixus is the main vector of Chagas disease in Vene-zuela, Colombia, Honduras, Nicaragua and El Salvador (Dujardinet al., 1998; Guhl, 2007; Schofield, 1994). This is due to its high sus-ceptibility to the infection by Trypanosoma cruzi, the etiologicalagent of this disease. In fact, R. prolixus can present T. cruzi infectionrates above 30% (Dorn et al., 2001; Monroy et al., 2003; Peñalver,1959). This triatomine species also has elevated capacities for dis-persion, reproduction and domiciliation, reaching as many as threegenerations per year (Lent and Wygodzinsky, 1979). Trypanosomarangeli, meanwhile, is a heteroxenic protozoan which shares inver-tebrate and vertebrate hosts, including man and domestic animals,with T. cruzi. Human infection by T. rangeli can induce misdiagnos-es due to serological cross-reaction with T. cruzi (Guhl and Marink-elle, 1982; Moraes et al., 2008; O’Daly et al., 1994; Zuñiga et al.,1997).

ll rights reserved.

arneri).

et al. A standardizable protocolcts of infection upon the insec

The vertebrate host phase of the life cycle of T. rangeli is stillpoorly understood although there is evidence that parasite divisionoccurs inside monocytes (Eger-Mangrich et al., 2001; Osório et al.,1995). In invertebrate hosts, the cycle begins when trypomastigoteforms are ingested during a blood meal on a mammal host. Afterreaching the gut, parasites transform to the epimastigote form,and become able to multiply. Afterwards, these forms can crossthe midgut epithelium to reach the haemocoel. Once in the haemo-lymph, parasites continue dividing and some migrate and invadesalivary glands, where they become metacyclic trypomastigotesthat can be transmitted to mammal hosts during the next bloodmeal (D’Alessandro, 1976; D’Alessandro-Bacigalupo and Saraiva,1992).

While considered non-pathogenic to man, T. rangeli is under-stood to cause different levels of pathologies to their invertebratehosts, mainly triatomine bugs belonging to the genus Rhodnius(Brecher and Wigglesworth, 1944; D’Alessandro, 1976; Eichlerand Schaub, 1998; Lake and Friend, 1967). At present, investigationof this area is hindered by non-standardization of inoculation pro-cedures, and by limited information available in studies concerning

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013

2 L.L. Ferreira et al. / Journal of Invertebrate Pathology xxx (2010) xxx–xxx

the sizes of parasite inocula used, in addition to the environmentalconditions under which published assays were developed. Theseconsiderations, and the relevance of laboratory studies to under-stand natural infections, are of prime importance as T. rangeli doesnot always invade the haemocoel in the natural course of infectionand natural infection rates can be low as 2% (Añez et al., 1987; Hec-ker et al., 1990; Tobie, 1965, 1970). Therefore, the present work ini-tially compared different infection methods in order to establish aprocedure that could mimic natural infections. Then, as a first stepin defining how pathogenic T. rangeli can be to its invertebratehost, we examined changes to the insect haemolymph as a conse-quence of infection by T. rangeli.

2. Materials and methods

2.1. Organisms

R. prolixus were reared in the laboratory under semi-controlledconditions (26 ± 2 �C and 65 ± 10% RH) and fed weekly on mice orchicken according to institutional ethical principles for animalcare.

T. rangeli of the CHOACHI strain, isolated from naturally infectedR. prolixus from Colombia (Schottelius, 1987) was used in the pres-ent study. The epimastigote forms used were cultured at 27 �C inliver-infusion tryptose (LIT) medium supplemented with 15% of fe-tal bovine serum, 100 mg/ml of streptomycin and 100 units/ml ofpenicillin.

2.2. Experiment 1: effect of infection route on the period betweenfeeding and moulting to fifth instar

As experiments 2–6 (below) were conducted with intracoelo-matic infection (i.e. inoculation), it was necessary first to comparethis with oral infection. For intracoelomatic infection, epimastig-otes were obtained from 10 day old culture medium, washed insterile PBS (0.15 M NaCl in 0.01 M sodium phosphate, pH 7.4)and resuspended in sterile culture medium. Different concentra-tions were made up by dilution in PBS. Fourth instar nymphs(7 days post-moult) were inoculated in the side of the thorax with1 ll of the parasite suspension, using a 50 ll Hamilton syringe con-nected to a fine needle. Groups of nymphs (n = 75 for each treat-ment) were inoculated with 10, 100 or 1000 parasites, or thesame volume of blank PBS for the control group. One day afterinfection, insects were fed on anaesthetized mice. Before assayswere conducted with these insects, haemolymph samples were ta-ken by cutting off the terminal segment of a hind leg and collectinga drop of haemolymph on a microscope slide to check the status ofinfection: only positive insects were used. For uniformity of thetreatments, control insects also had a sample of haemolymph ex-tracted by the same procedure. Insects were monitored daily tocheck whether they moulted or died.

For oral infection, epimastigotes were obtained as above butwere added to heat-inactivated blood at a concentration of1 � 107 flagellates/ml. Second instar nymphs were allowed to feedon this blood through a membrane feeder (control insects were fedon blood to which only the sterile culture medium was added) andallowed to develop to fourth instar prior to further evaluation.Since in the natural course of infection, T. rangeli does not alwaysinvade the haemocoel, insects were inspected for the presence ofparasites in their faeces and haemolymph when they moulted tofourth instar. One group of insects which presented parasites onlyin their gut was then further inoculated in their haemolymph (seeabove) and another group, that had not been infected orally, wasalso infected in this manner. Thus, four groups were generated:(a) insects that presented parasites in their gut, but not in the hae-

Please cite this article in press as: Ferreira, L.L., et al. A standardizable protocolnatural infections and reveals physiological effects of infection upon the insec

molymph (gut only, n = 56); (b) insects that had parasites in boththeir gut and haemolymph (natural, n = 10); (c) insects that onlyshowed parasites in their gut and were further inoculated with ahundred parasites into the thorax (inoculation, n = 65; see above)and (d) insects that were only infected by inoculation (100 para-sites, n = 75); this was the same group as used in the section above,as was the control group). These inoculations (c and d) were con-ducted for comparison with natural infection (i.e. group b). For uni-formity of the treatments, insects from groups a and b were alsoinoculated with PBS. Nymphs were fed 7 days after their moultto the fourth instar and were then monitored daily to checkwhether they moulted or died.

2.3. Experiment 2: parasite development in haemolymph

Fifth instar nymphs were starved for 3 days after moulting andthen inoculated with 10 or 100 parasites (n = 120 for each treat-ment). From the second day after infection, parasite and haemo-cyte counts were conducted every 2 days. For this, haemolymphwas extracted as described above but was collected with 5 ll pip-ettes. Four pools of haemolymph from 5 insects were used for eachassay and these animals were discarded. Samples were immedi-ately diluted in an anticoagulant solution (0.01 M ethylenediaminetetra-acetic acid, 0.1 M glucose, 0.062 M sodium chloride, 0.03 Mtrisodium citrate, 0.026 M citric acid, pH 4.6) in a proportion of1:10 haemolymph:anticoagulant (Mello et al., 1995). The total try-panosome and haemocyte numbers were counted directly in ahaemocytometer under a light microscope.

2.4. Experiment 3: effect of infection on haemolymph volume

Fourth instar nymphs were infected intracoelomatically with100 parasites and a control group was inoculated with blank PBS(n = 30 for both treatments). At fifth instar (10 days post-moult)they were used to estimate whether infection by T. rangeli altershaemolymph volume. They were individually weighed and thedorsal cuticle of their thorax and abdomen was removed. Theirhaemolymph was removed using a piece of filter paper and theywere re-weighed. The volume of haemolymph was estimated byweight subtraction (Naidu, 2001).

2.5. Experiment 4: effect of infection on urine/faecal loss after bloodmeal

Fourth instar nymphs were inoculated with 10, 100 or 1000parasites, and a control group was inoculated with blank PBS(n = 10 for each group). At fifth instar (10 days post-moult), theywere individually weighed, fed on anesthetized mice and weighedagain for estimation of the blood meal volume. They were immedi-ately transferred to plastic micro-tubes and maintained under con-trolled conditions (25 ± 1 �C and 65 ± 10% RH). The volume ofexcreta was measured using micro-capillaries at hourly intervalsover the next 4 h and again 24 h after the blood meal (NB). The firstdrop of urine released by triatomines after feeding is mixed withthe faeces that remained in the rectum after the last meal. Thedroplets that are subsequently eliminated consist exclusively of ur-ine, hence the use the term ‘excretion’ was adopted in the presentwork.

2.6. Experiment 5: effect of infection on water loss between moults(indirect estimation via weight loss)

Since the production of urine is inhibited at the end of diuresis,the material eliminated by bugs afterwards becomes viscous and itis not possible to use capillaries to measure subsequent water loss.

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013

L.L. Ferreira et al. / Journal of Invertebrate Pathology xxx (2010) xxx–xxx 3

Therefore, the weight loss measured during the inter-moult inter-val was used to obtain an indirect estimation of water loss.

Fourth instar nymphs were inoculated with 100 parasites orwith blank PBS (n = 10 for both groups). At fifth instar (10 dayspost-moult), they were individually weighed, fed on anesthetizedmice, re-weighed immediately after feeding and then weighedagain every 3 days until ecdysis. During this period, the occurrenceof ecdysis or death was recorded.

2.7. Experiment 6: effect of infection on lipid content and fat bodyweight

To measure lipid content, fourth instar nymphs were inoculatedwith 100 parasites (n = 53) or blank PBS (n = 51). Seven days aftermoulting to fifth instar, they were individually weighed and main-tained at 75 �C for 6 days to obtain their dry weight. The lipid con-tents of these insects were measured using the ether extractionmethod of David et al. (1975) with modifications. Briefly, insectswere transferred to flasks containing 20 ml of ethyl ether for7 days. Afterwards, they were dried and weighed again and theestimation of lipid contents was done by weight subtraction.

To measure fat body weights, fourth instar nymphs were inoc-ulated with 100 parasites (n = 23) or blank PBS (n = 25). At fifth in-star, they were individually dissected under a stereo microscope;their fat bodies were removed with tweezers and transferred topieces of filter paper of known weights. These were dried at50 �C and re-weighed. Estimations of fat body weights were doneby weight subtraction.

2.8. Statistical analyses

The variables that followed a normal distribution were com-pared by performing a two-tailed t-test or by ANOVA for indepen-dent samples. Proportion data were arc-sine transformed prior toanalysis. Mann–Whitney or Kruskal–Wallis non-parametric testswere used for variables with a non-normal distribution. In the caseof ANOVA tests, post hoc comparisons were made using Scheffétest. Times taken for insects to moult were analysed with Kap-lan–Meier survival analyses in SPSS 8.0, and comparisons weremade with log-rank tests. The Chi-square test was used to comparefrequencies. Throughout, the accepted significance level wasp < 0.05.

3. Results

3.1. Experiment 1: effect of infection route on the period betweenfeeding and moulting to fifth instar

The effect of intracoelomic infection by T. rangeli on the dura-tion of fourth instar in R. prolixus was evaluated first (Fig. 1A). In-fected insects had a prolonged moult when compared with thosefrom the control group. For the lowest parasite dose, the differencewas slight: median times to moult were 19 ± 0.52 days for the con-trol and 20 ± 0.43 days for the 10 parasite dose (log-rank,p = 0.028). At the higher doses, times to moult were substantiallyprolonged, at 25 ± 0.39 days for 100 parasites (log-rank vs. controlp < 0.0001 and vs. 10 parasites p = 0.0076) and at 23 ± 0.36 daysfor 1000 parasites (log-rank, vs. control p < 0.0001). There was nostatistically significant difference between the two higher doses(log-rank, p = 0.69).

To evaluate whether gut infection by T. rangeli also promotes adelay in the moulting process, second instar nymphs were orallyinfected with culture epimastigotes. Approximately 85% of the in-sects showed flagellates in faeces. In 13% of them, the cycle of T.rangeli was completed and trypomastigote forms were seen devel-

Please cite this article in press as: Ferreira, L.L., et al. A standardizable protocolnatural infections and reveals physiological effects of infection upon the insec

oping in salivary glands. The duration of fourth instar was evalu-ated in insects that only presented parasites in the intestinaltract (gut only) and insects with parasites in both gut and haemo-lymph (natural). Additionally, a group of insects that only pre-sented parasites in the intestinal tract was inoculated with 100parasites and was also evaluated (inoculation). Here, we also con-sider the group infected intracoelomatically with 100 parasitesfrom the first experiment for comparison (both experiments wererun concurrently in the same locale). The presence of the parasitein the gut had a very marginal effect on time to moult, visible onlyin Fig. 1B (19 ± 0.19 days vs. control 19 ± 0.52 days; log-rank,p = 0.046). Meanwhile, confirming the outcome of the previous as-say, the presence of the parasite in the haemolymph induced a pro-longed moult. For these insects, times to moult were:22 ± 1.58 days for natural infection (log-rank, p = 0.013 vs. control),22 ± 0.93 days for inoculation (p < 0.0001 vs. control) and25 ± 0.39 days for 100 parasites (p < 0.0001 vs. control). Mostimportantly for the present study, these three groups presentedno significant differences in time to moult, independent of themode of infection.

No statistical differences were found when mortality andmoulting rates were compared among the groups. The observedvalues for these parameters were approximately 10% and 97%,respectively (v2, n.s.).

3.2. Experiment 2: parasite development in haemolymph

Since insects inoculated with 10 parasites did not present a pro-longed moult, while those that received 100 or 1000 flagellates did,parasite growth and haemocyte counts were evaluated in the hae-molymph of fourth instar nymphs that received 10 or 100 parasites(Fig. 2). Four days after inoculation, the first flagellates could be ob-served in the haemolymph of insects injected with 100 parasites(in numbers too low to be visible on the figure). From this time,the amount of parasites in insects of this group was consistentlyhigher than that recorded in those that received an inoculation of10 parasites (Fig. 2A; Mann–Whitney U test, p < 0.05, n = 4; 4 poolswith samples of 5 insects each) except for days 8 and 12, whenboth groups showed a similar number of flagellates in the haemo-lymph. Infected insects presented a gradual increase in their totalhaemocyte counts that was maintained until day 12 (Fig. 2B).These counts, irrespective of dose, were higher than those for con-trol insects for days 10 and 12 (Mann–Whitney U tests, p = 0.02and p = 0.03, respectively; n = 4; 4 pools with samples of 5 insectseach).

3.3. Experiment 3: effect of infection on haemolymph volume

The haemolymph of infected insects represented 18.7 ± 1.3% oftheir body weight, being higher than that of control insects(12.3 ± 0.9%; t = �4.13, p = 0.00011; df = 57).

3.4. Experiment 4: effect of infection on urine/faecal loss after bloodmeal

To evaluate whether the observed increase in haemolymphweight of infected insects was due to an alteration of diuresis,water loss was measured in recently fed nymphs. All insects,whether infected or not, lost higher amounts of urine/faeces duringthe first 2 h after feeding, excretion being gradually reduced overthe 24 h of observation. No statistical differences were foundamong the groups in relation to excretion rate or parasite counts(ANOVA, F = 0.55, p = 0.64, df = 3); as the data are not qualitativelydifferent from previously published work, they are not shown here(Maddrell, 1963; Wigglesworth, 1931).

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013

Fig. 1. Experiment 1: Effect of infection route of the parasite Trypanosoma rangeli on time to moult of the triatomine bug Rhodnius prolixus. (A) Infection was intracoelomic inthe fourth instar, with doses of 0 (control, i.e. blank PBS), 10, 100 or 1000 parasites per inoculation (n = 75 for each dosage). Insects receiving the higher doses (100 and 1000)took significantly longer to moult than the control insects and those receiving 10 parasites (survival analyses; see text for details). (B) Infection was oral (except wherestated), in the second instar, and insects were reared until fourth instar, given a bloodmeal, and observed as in (A). Treatments: control – fed on uncontaminated blood; gutonly – insects that presented parasites in their gut, but not in the haemolymph; natural – insects that had parasites in both their gut and haemolymph; inoculation – insectsthat only showed parasites in their gut and were further inoculated with 100 parasites into the thorax; and haemolymph only – insects that were only inoculated with 100parasites into the thorax.

Fig. 2. Experiment 2: Development of Trypanosoma rangeli in the haemolymph of Rhodnius prolixus, following intracoelomic inoculation at 5th instar. (A) Number of T. rangeliepimastigotes. (B) Total haemocyte counts.

4 L.L. Ferreira et al. / Journal of Invertebrate Pathology xxx (2010) xxx–xxx

3.5. Experiment 5: effect of infection on water loss between moults(indirect estimation via weight loss)

The duration of the fifth instar was prolonged for infected in-sects, as seen when fourth instar insects were studied (Fig. 3A; con-trol 19 ± 0.47 days vs. infected 28 ± 2.57 days; log-rank,p < 0.0001). During this period, both groups showed a similarweight reduction of approximately 60% of the blood ingested(62.5 ± 1.1% and 63.5 ± 1.07%, for insects of control and infectedgroups, respectively; t = 0.61, p = 0.54, df = 18). The daily rate ofweight loss was calculated by dividing total weight loss by thenumber of days taken to reach the fifth instar. As shown inFig. 3B, infected insects reduced their daily rates of weight loss incomparison with controls (t = �4.41, p = 0.0003; df = 18).

3.6. Experiment 6: effect of infection on lipid contents and fat bodyweight

Lipid contents of insects were determined to evaluate whetherthis could account for the observed increase in haemolymph vol-

Please cite this article in press as: Ferreira, L.L., et al. A standardizable protocolnatural infections and reveals physiological effects of infection upon the insec

ume in bugs with T. rangeli present in the haemolymph. Since in-fected insects of this group were lighter than control ones(t = �2.65, p = 0.009, df = 104), data were calculated as percentageof lipids in relation to body weight. As shown in Fig. 4A, infectedinsects presented an increase in lipid content of approximately48% when compared to those from the control group (t = �40.82,p = 0.00001; df = 104). Similarly, fat bodies in infected nymphswere 60% heavier than those of the insects from control group(Fig. 4B; t = 5.05, p < 0.000007; df = 46).

4. Discussion

The first aim of this study was to compare oral and inoculationinfection procedures with T. rangeli in R. prolixus. Only 13% of theinsects infected orally that presented parasites in the intestinaltract had parasites infecting their salivary glands. This is in accor-dance with previously reported infection rates ranging between 2%and 50% (Añez et al., 1987; Groot, 1954; Hecker et al., 1990; Mar-inkelle, 1968; Tobie, 1965, 1970). The mechanisms that prevent thepassage of the parasite from the lumen of the midgut to the

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013

Fig. 3. Experiment 5: Determination of weight loss during the inter-moult period. (A) Interval between feeding and moult to the fifth instar of R. prolixus infected by T. rangeli.(B) Daily weight loss of R. prolixus infected by T. rangeli. Data were calculated as a percentage of weight loss in relation to blood ingested.

Fig. 4. Experiment 6: Effects of infection of Rhodnius prolixus by Trypanosoma rangeli on: (A) lipids as a proportion of body weight and (B) fat body weight.

L.L. Ferreira et al. / Journal of Invertebrate Pathology xxx (2010) xxx–xxx 5

haemocoel are not understood yet. Gomes et al. (2002) showedthat gamma irradiation induces changes in the ultrastructuralorganisation of the insect perimicrovillar membranes; in this case,flagellates were present in the haemolymph of irradiated insectsearlier than non-irradiated ones. In this sense it is possible thatperimicrovillar membrane represents an essential structure inthe control of T. rangeli invasion and that alterations in these mem-branes may facilitate the penetration of the parasite into the hae-mocoel of the vector. Another factor that may interfere ininfection rates is the identity of the T. rangeli strain and Rhodniusspecies used. The ability of T. rangeli to reach the insect’s haemo-lymph is dependent on both the parasite strain and the triatominespecies, such that insects are more susceptible to strains isolatedfrom the same geographic region (Cuba Cuba et al., 1972; D’Ales-sandro, 1972; Grisard et al., 1999; Machado et al., 2001). This isthe case in the present work where we used the CHOACHI strainthat was isolated from R. prolixus in Colombia.

Although in the present work just one instar was studied, theresults obtained with insects infected by intracoelomic inoculationor by the oral route suggest that the presence of the parasite in thehaemolymph was sufficient to produce a delay in insect moult(Fig. 1). In these insects, whichever method was used to establishinfection of the haemolymph, and whether or not the gut pre-sented parasites, the time to moult was similar. Meanwhile, insectsinfected in the gut alone had a similar moulting time to uninfectedcontrols. Eichler and Schaub (2002) correlated the delay in devel-

Please cite this article in press as: Ferreira, L.L., et al. A standardizable protocolnatural infections and reveals physiological effects of infection upon the insec

opment of T. rangeli-infected nymphs to a decrease in the numberof intestinal symbionts. Indeed, the lack of these symbionts pro-motes a delay in the development of these insects in addition toan increase in mortality rates (Brecher and Wigglesworth, 1944;Dasch et al., 1984; Lake and Friend, 1967). Based on our resultswe suggest that triatomines inoculated via coelomic cavity canbe used to mimic the effects of the T. rangeli in the natural infec-tion. After characterising the different infection routes, inoculationwas chosen for use in subsequent experiments. The first studyshowed that the size of a parasite dose affects the course of infec-tion. Moult was delayed in bugs inoculated with 100 or 1000 par-asites, but not in those that received 10 parasites. The fact that thetwo higher dose treatments were similar to each other suggeststhat the parasite population needs to reach a threshold size beforethese pathogenic effects are seen in that instar. Interestingly, thebugs inoculated with 10 parasites in fact had a delayed moultingprocess in the following instar, after their infection became chronic(data not shown). Meanwhile, parasite growth was measured atthe beginning of infection for insects inoculated with 10 or 100parasites (Fig. 2A), showing that these parasite populations wereequal after 12 days. Together, these observations suggest that theamount of flagellates may be related with the pathological effects.In this way, insects at the beginning of coelomatic infection or withfewer parasites could have a similar development to uninfectedones. For those insects with a high parasitic load, however, thepathological effects became apparent quickly.

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013

6 L.L. Ferreira et al. / Journal of Invertebrate Pathology xxx (2010) xxx–xxx

Some classical studies that analysed the interaction between R.prolixus and T. rangeli described an infection-induced increase inthe volume of haemolymph but did not quantify it (Grewal,1957; Ormerod, 1967; Tobie, 1965; Watkins, 1971). Therefore,we evaluated whether this effect could be confirmed under ourexperimental conditions, i.e. if insects infected by intracoelomicinoculation with 100 parasites would show increased haemolymphvolumes. In fact, the haemolymph volume of infected insects pre-sented an increase of 34% in relation to that of insects from thecontrol group. Watkins (1971) demonstrated that R. prolixus in-fected by intracoelomic injection of T. rangeli flagellates presentedan extreme reduction in excretion during the first 3 h after feeding.Therefore, the first hypothesis that could explain the haemolymphincrease was that infection promoted the retention of water in in-fected insects. To test this hypothesis, we first evaluated the effectof the parasite on excretion in infected insects, through a period oftime after feeding. However, no effect of infection was found, evenusing three distinct parasite levels. Weight loss was used as anindirect method to estimate inter-moult water loss. During thefifth instar, both infected and control insects lost approximately60% of the weight of the blood ingested – this was in spite of theinfected insects’ moult being delayed, i.e. daily water loss was low-er in infected insects but the moult was longer. This provided fur-ther evidence that water retention was not responsible for theincrease on haemolymph volume. The increase in haemolymphvolume cannot be attributed to the parasites’ own volume as thiswould respond for approximately 0.1% of the volume of haemo-lymph (calculations not shown).

During the examination of insects to confirm infection, anapparently higher amount of lipids was normally observed in thehaemolymph of infected insects and our quantitative measure-ments showed this increase to be almost twofold. This was despiteinfected insects being lighter than controls. Moreover, and eventaking into account this latter fact, infected insects had heavierfat bodies than controls.

Therefore, we propose some possible physiological explanationsfor these findings. The blood ingested by triatomines is stored inthe anterior midgut and digested in the thin and coiled posteriormidgut, the site of protein and lipid digestion (Billingsley, 1988,1990). After a blood meal, diacylglycerol and phospholipids aretransferred from the posterior midgut to lipophorin, the lipid car-rier lipoprotein circulating in the haemolymph (Atella et al.,1995; Coelho et al., 1997). Lipophorin then distributes lipids to dif-ferent organs, including fat bodies, by interacting with specificbinding sites at cell surfaces (Machado et al., 1996; Pontes et al.,2002, 2008). Lipids stored in the fat body may then be used whenno more nutrients are available in the midgut, and delivery is alsoachieved by circulating lipophorins (Atella et al., 1992; Coelhoet al., 1997; Grillo et al., 2007). Studies of lipid flow inside verte-brate hosts have shown that members of the family Trypanosomat-idae lack the complete synthesis pathways for some sterols andfatty acids, implying that these molecules must be taken from hostfluids (Coppens and Coutoy, 1995; Coppens et al., 1995; Paul et al.,2001; Vial et al., 2003). Folly et al. (2003) showed that T. rangeli up-takes either neutral lipids or phospholipids from R. prolixus haemo-lymph and internalises the lipophorin protein moiety through aspecific receptor on the parasite surface. In this sense, the increaseor at least part of the increase in the amount of lipids observed ininfected R. prolixus in the present study could be in fact, an accu-mulation of lipids as a result of an imbalance of available lipopho-rin molecules that have been taken by parasites. In addition,trypanosomatids are also able to incorporate some lipids directlyfrom culture media (Morita and Englund, 2001; Paul et al., 2001).Therefore, T. rangeli might be also able to redirect insect nutritiontowards an increased production of fat. Since infected insects werelighter but not smaller (data not shown) than controls, it is possible

Please cite this article in press as: Ferreira, L.L., et al. A standardizable protocolnatural infections and reveals physiological effects of infection upon the insec

that the fat reserves for muscles or even organs could be accumu-lating in the fat body and haemolymph for use by the parasites.Additional studies will be performed to test these hypotheses.

To conclude, we have presented an inoculation procedurewhich is simple and is standardizable as it excludes insect behav-iour (i.e. feeding) as a variable. Based on our observations of com-paratively subtle effects of infection on the insect host’sphysiology, we propose that this method can be used to investigatemore fully the pathology and physiology induced by this interac-tion. For studies of within-host dynamics of a parasite population,and especially for studies of multiple infections (e.g. with T. cruziand T. rangeli, or with different strains of one species), it is essentialto have such experimental control. It increasingly being shownthat infection of insects (or other organisms) can have very subtleeffects (e.g. Elliot et al., 2005). This is true for systems where theinsect is primarily considered as a vector (e.g. Elliot et al., 2003;Lambrechts and Scott, 2009); in the case of triatomines and trypan-osomatids, there is much to be discovered about the pathologicaleffects of the parasites on the insects. We have shown severalphysiological effects of T. rangeli on R. prolixus moulting and waterloss. Meanwhile, we have previously shown that ambient humidityand water loss can affect development and survival of other triato-mines (e.g. Guarneri et al., 2002; Xavier et al., 2005). We have ta-ken some steps towards a mechanistic explanation of thesephenomena, but it will be important, for a fuller understanding,to integrate these different aspects of the insects’ physiology. It willalso be interesting to explore possible adaptive explanations forthe observations by examining how physiological effects of para-sitism impact upon the insects’ survival and reproduction, on theone hand, and on measures of the parasite’s fitness on the other.

Acknowledgments

We thank Dr. Mário Steindel, UFSC, Brazil, for supplying the T.rangeli strain used here. This work was supported by grants fromFundação de Amparo à Pesquisa do Estado de Minas Gerais (FAP-EMIG, APQ-3549-5.03/07), Conselho Nacional de DesenvolvimentoCientífico e Tecnológico (CNPq, 478786/2007-7) and Fundação Os-waldo Cruz (FIOCRUZ), Brazil.

References

Añez, N., Nieves, E., Cazorla, D., 1987. Studies on Trypanosoma rangeli Tejera, 1920.IX. Course of infection in different stages of Rhodnius prolixus. Mem. Inst.Oswaldo Cruz 82, 1–6.

Atella, G.C., Gondim, K.C., Masuda, H., 1992. Transfer of phospholipids from the fatbody to lipophorin in Rhodnius prolixus. Arch. Insect Biochem. Physiol. 19, 133–144.

Atella, G.C., Gondim, K.C., Masuda, H., 1995. Loading of lipophorin particles withphospholipids at the midgut of Rhodnius prolixus. Arch. Insect Biochem. Physiol.30, 337–350.

Billingsley, P.F., 1988. Morphometric analysis of Rhodnius prolixus Stal (Hemiptera:Reduviidae) midgut cells during blood digestion. Tissue Cell 20, 291–301.

Billingsley, P.F., 1990. The midgut ultrastructure of hematophagous insects. Annu.Rev. Entomol. 35, 219–248.

Brecher, G., Wigglesworth, V.B., 1944. The transmission of Actinomyces rhodniiErikson in Rhodnius prolixus Stal (Hemiptera) and its influence on the growth ofthe host. Parasitology 35, 220–224.

Coelho, H.S.L., Atella, G.C., Moreira, M.F., Gondim, K.C., Masuda, H., 1997. Lipophorindensity variation during oogenesis on Rhodnius prolixus. Arch. Insect Biochem.Physiol. 35, 301–313.

Coppens, I., Coutoy, P.J., 1995. Exogenous and endogenous sources of sterols in theculture-adapted procyclic trypomastigotes of Trypanosoma brucei. Mol.Biochem. Parasitol. 73, 179–188.

Coppens, I., Levade, T., Courtoy, P.J., 1995. Host plasma low density lipoproteinparticles as an essential source of lipids for the bloodstream forms ofTrypanosoma brucei. J. Biol. Chem. 270, 5736–5741.

Cuba Cuba, C.A., Morales, N., Fernández, E., Fernández, W., 1972. Hallazgo deRhodnius ecuadoriensis Lent & León, 1958 infectado naturalmente porTrypanosomas semejantes a Trypanosoma rangeli Tejera, 1920 en caseríos de laProvíncia de Cascas, Contumazá, Perú. Rev. Inst. Med. Trop. São Paulo 14, 191–202.

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013

L.L. Ferreira et al. / Journal of Invertebrate Pathology xxx (2010) xxx–xxx 7

D’Alessandro, A., 1972. New experimental vectors of Colombian Trypanosomarangeli. J. Med. Entomol. 9, 187–195.

D’Alessandro, A., 1976. Biology of Trypanosoma (Herpetosoma) rangeli Tejera, 1920.In: Lumsden, W.H.R., Evans, D.A. (Eds.), Biology of Kinetoplastida. AcademicPress, London, pp. 328–403.

D’Alessandro-Bacigalupo, A., Saraiva, N.G., 1992. Trypanosoma rangeli. In: Kreir, J.P.,Baker, J. (Eds.), Parasitic Protozoa. Academic Press, London, pp. 1–54.

Dasch, G.A., Weiss, E., Chang, K., 1984. Endosymbionts of insects. In: Krieg, N.R.(Ed.), Bergey’s Manual of Systematic Bacteriology. Williams e Wilkins,Baltimore, MD, pp. 811–833.

David, J., Cohet, Y., Fouillet, P., 1975. Physiologie de l’inanition et utilisation dereserves chez les adultes de Drosophila melanogaster. Arch. Zool. Exp. Gen. 116,579–590.

Dorn, P.L., Flores, J., Brahney, B., Gutierrez, A., Rosales, R., Rodas, A., Monroy, C., 2001.Comparison of polymerase chain reaction on fresh tissue samples and fecaldrops on filter paper for detection of Trypanosoma cruzi in Rhodnius prolixus.Mem. Inst. Oswaldo Cruz 96, 503–505.

Dujardin, J.P., Muñoz, M., Chávez, T., Ponce, C., Moreno, J., 1998. The origin ofRhodnius prolixus in Central America. Med. Vet. Entomol. 12, 113–115.

Eger-Mangrich, I., Oliveira, M.A., Grisard, E.C., De Souza, W., Steindel, M., 2001.Interaction of Trypanosoma rangeli Tejera, 1920 with different cells line in vitro.Parasitol. Res. 87, 505–509.

Eichler, S., Schaub, G.A., 1998. The effects of aposymbiosis and of an infection withBlastocrithidia triatomae (Trypanosomatidae) on the tracheal system of thereduviid bugs Rhodnius prolixus and Triatoma infestans. J. Insect Physiol. 44,131–140.

Eichler, S., Schaub, G.A., 2002. Development of symbionts in Triatomine bugs andthe effects of infections with trypanosomatids. Exp. Parasitol. 100, 17–27.

Elliot, S.L., Adler, F.R., Sabelis, M.W., 2003. How virulent should a parasite be to itsvector? Ecology 84, 2568–2574.

Elliot, S.L., Horton, C.M., Blanford, S., Thomas, M.B., 2005. Impacts of fever on locustlife-history traits: costs or benefits? Biol. Lett. 1, 181–184.

Folly, E., Cunha e Silva, N., Lopes, A.C.S., Silva-Neto, M., Atella, G.C., 2003.Trypanosoma rangeli uptakes the main lipoprotein from the hemolymph of itsinvertebrate host. Biochem. Biophys. Res. Commun. 310, 555–561.

Gomes, S.A.O., Graciano, G.L., Nogueira, N.F.S., De Souza, W., Garcia, E.S., Azambuja,P., 2002. Effects of gamma irradiation on the development of Trypanosomarangeli in the vector Rhodnius prolixus. J. Invertebr. Pathol. 79, 86–92.

Grewal, M.S., 1957. Pathogenicity of Trypanosoma rangeli Tejera, 1920 in theinvertebrate host. Exp. Parasitol. 6, 123–130.

Grillo, L.A.M., Majerowicz, D., Gondim, K.C., 2007. Lipid metabolism in Rhodniusprolixus (Hemiptera: Reduviidae): role of a midgut triacylglycerol-lipase. InsectBiochem. Mol. Biol. 37, 579–588.

Grisard, E.C., Steindel, M., Guarneri, A.A., Eger-Mangrich, I., Campbell, D.A., Romanha,A.J., 1999. Characterization of Trypanosoma rangeli strains isolated in Central andSouth America: an overview. Mem. Inst. Oswaldo Cruz 94, 203–209.

Groot, H., 1954. Estudios sobre los trypanosomas humanos clasificados como T.Rangeli con especial referencia a su evolución in Rhodnius prolixus y a sucomparación con T. ariarii. An. Soc. Biol. Bogota 6, 109–126.

Guarneri, A.A., Lazzari, C., Diotaiuti, L., Lorenzo, M.G., 2002. The effect of relativehumidity on the behaviour and development of Triatoma brasiliensis. Physiol.Entomol. 27, 142–147.

Guhl, F., 2007. Chagas disease in Andean countries. Mem. Inst. Oswaldo Cruz 102(Suppl. 1), 29–37.

Guhl, F., Marinkelle, C.J., 1982. Antibodies against Trypanosoma cruzi in miceinfected with T. Rangeli. Ann. Trop. Med. Parasitol. 76, 361.

Hecker, H., Schwarzenbach, M., Rudin, W., 1990. Development and interactions ofTrypanosoma rangeli in and with the reduviid bug Rhodnius prolixus. Parasitol.Res. 76, 311–318.

Lake, P., Friend, W.G., 1967. A monoxenic relationship, Nocardia rhodnii Erikson inthe gut of Rhodnius prolixus Stal (Hemiptera: Reduviidae). Proc. Entomol. Soc.Ontario 98, 53–57.

Lambrechts, L., Scott, T.W., 2009. Mode of transmission and the evolution of arbovirusvirulence in mosquito vectors. Proc. Roy. Soc. Lond. B 276, 1369–1378.

Lent, H., Wygodzinsky, M., 1979. Revision of the Triatominae (Hemiptera:Reduviidae) and their significance as vector of Chagas disease. Bull. Am. Mus.Nat. Hist. 163, 123–520.

Machado, E.A., Atella, G.C., Gondim, K.C., de Souza, W., Masuda, H., 1996.Characterization and immunocytochemical localization of lipophorin binding

Please cite this article in press as: Ferreira, L.L., et al. A standardizable protocolnatural infections and reveals physiological effects of infection upon the insec

sites in the oocytes of Rhodnius prolixus. Arch. Insect Biochem. Physiol. 31, 185–196.

Machado, P.E., Eger-Mangrich, I., Rosa, G., Koerich, L.B., Grisard, E., Steindel, M.,2001. Differential susceptibility of triatomines of the genus Rhodnius toTrypanosoma rangeli strains from different geographical origins. Int. J.Parasitol. 31, 632–634.

Maddrell, S.H.P., 1963. Excretion in the blood-sucking bug, Rhodnius prolixus. I. Thecontrol of diuresis. J. Exp. Biol. 40, 247–256.

Marinkelle, C.J., 1968. Pathogenicity of Trypanosoma rangeli for Rhodnius prolixusStal in nature (Hemiptera: Reduviidae) (Kinetoplastida: Trypanosomatidae). J.Med. Entomol. 5, 497–499.

Mello, C.B., Garcia, E.S., Ratcliffe, N.A., Azambuja, P., 1995. Trypanosoma cruzi andTrypanosoma rangeli: interplay with hemolymph components of Rhodniusprolixus. J. Invertebr. Pathol. 65, 261–268.

Monroy, C., Rodas, A., Mejía, M., Rosales, R., Tabaru, Y., 2003. Epidemiology ofChagas disease in Guatemala: infection rate of Triatoma dimidiata, Triatomanitida and Rhodnius prolixus (Hemiptera, Reduviidae) with Trypanosoma cruziand Trypanosoma rangeli (Kinetoplastida, Trypanosomatidae). Mem. Inst.Oswaldo Cruz 98, 305–310.

Moraes, M.H., Guarneri, A.A., Girardi, F.P., Rodrigues, J.B., Eger, I., Tyler, K.M.,Steindel, M., Grisard, E.C., 2008. Different serological cross-reactivity ofTrypanosoma rangeli forms in Trypanosoma cruzi-infected patients sera.Parasites Vectors 2008, 1–20.

Morita, Y.S., Englund, P.T., 2001. Fatty acid remodeling of glycosylphosphatidylinositol anchors in Trypanosoma brucei: incorporation of fattyacids other than myristate. Mol. Biochem. Parasitol. 115, 157–164.

Naidu, S.G., 2001. Water balance and osmoregulation in Stenocara gracilipes, awaxblooming tenebrionid beetle from the Namib Desert. J. Insect Physiol. 47,1429–1440.

O’Daly, J.A., Carrasco, H., Fernandez, V., Rodríguez, M.B., 1994. Comparison ofchagasic and non-chagasic myocardiopathies by ELISA and immunoblottingwith antigens of Trypanosoma cruzi and Trypanosoma rangeli. Acta Trop. 56,265–287.

Ormerod, W.E., 1967. Taxonomy of the sleeping sickness trypanosomes. J. Parasitol.53, 824–830.

Osório, Y., Travi, B.L., Palma, G., Saravia, N.G., 1995. Infectivity of Trypanosomarangeli in a promonocytic mammalian cell line. J. Parasitol. 81, 687–693.

Paul, K.S., Jiang, D., Morita, Y.S., Englund, P.T., 2001. Fatty acid synthesis in Africantrypanosomes: a solution to the myristate mystery. Trends Parasitol. 17, 381–387.

Peñalver, L.M., 1959. Estado actual de la enfermedad de Chagas en Guatemala. In:Aguilar, F. (Ed.), Homenaje al Cincuentenario del Descubrimiento de laEnfermedad de Chagas. Ministerio de Salud, Guatemala, pp. 32–52.

Pontes, E.G., Grillo, L.A.M., Gondim, K.C., 2002. Characterization of lipophorinbinding to the fat body of Rhodnius prolixus. Insect Biochem. Mol. Biol. 32, 1409–1417.

Pontes, E.G., Leite, P., Majerowicz, D., Atella, G.C., Gondim, K.C., 2008. Dynamics oflipid accumulation by the fat body of Rhodnius prolixus: the involvement oflipophorin binding sites. J. Insect Physiol. 54, 790–797.

Schofield, C.J., 1994. Triatominae: Biologia y control. Eurocommunica Publications,West Sussex.

Schottelius, J., 1987. Neuraminidase fluorescent test for differentiation ofTrypanosoma cruzi and Trypanosoma rangeli. Trop. Med. Parasitol. 38, 323–327.

Tobie, E.J., 1965. Biological factors influencing transmission of Trypanosoma rangeliby Rhodnius prolixus. J. Parasitol. 51, 837–841.

Tobie, E.J., 1970. Observations of the development of Trypanosoma rangeli in thehaemocel of Rhodnius prolixus. J. Invertebr. Pathol. 15, 118–125.

Vial, H.J., Eldin, P., Tielens, A.G.M., van Hellemond, J.J., 2003. Phospholipids inparasitic protozoa, review. Mol. Biochem. Parasitol. 126, 143–154.

Watkins, R., 1971. Trypanosoma rangeli: effect on excretion in Rhodnius prolixus. J.Invertebr. Pathol. 17, 67–71.

Wigglesworth, V.B., 1931. The physiology of excretion in the bloodsucking insect,Rhodnius prolixus (Hemiptera, Reduviidae). I. Composition of the urine. J. Exp.Biol. 8, 411–451.

Xavier, A.A.P., Lorenzo, M.G., Lazzari, C., Diotaiuti, L., Guarneri, A.A., 2005. Relativehumidity and water loss in Triatoma brasiliensis. Physiol. Entomol. 30, 338–342.

Zuñiga, C., Palau, T., Penin, P., Gamallo, C., Diego, J.A., 1997. Protective effect ofTrypanosoma rangeli against infections with a highly virulent strain ofTrypanosoma cruzi. Trop. Med. Int. Health 2, 482–487.

for infection of Rhodnius prolixus with Trypanosoma rangeli, which mimicst. J. Invertebr. Pathol. (2010), doi:10.1016/j.jip.2010.05.013