Vitellogenesis in the hematophagous Dipetalogaster maxima (Hemiptera: Reduviidae), a vector of Chagas’ disease

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Journal of Insect Physiology 54 (2008) 393–402 www.elsevier.com/locate/jinsphys

Vitellogenesis in the hematophagous Dipetalogaster maxima (Hemiptera: Reduviidae), a vector of Chagas’ disease Silvina A. Aguirre, Silvia Frede, Edilberto R. Rubiolo, Lilia´n E. Canavoso Departamento de Bioquı´mica Clı´nica, Centro de Investigaciones en Bioquı´mica Clı´nica e Inmunologı´a (CIBICI-CONICET), Facultad de Ciencias Quı´micas,Universidad Nacional de Co´rdoba, Co´rdoba, CP 5000, Argentina Received 26 June 2007; received in revised form 2 October 2007; accepted 22 October 2007

Abstract Oocyte extracts of anautogenous Dipetalogaster maxima were chromatographed on an ion-exchange column in order to purify vitellin (Vt), the main insect yolk protein precursor. Purified Vt (Mr 443 kDa) was composed of four subunits with approximate molecular weights of 174, 170, 50, and 44 kDa. Polyclonal anti-Vt antibody, which cross-reacted equally with fat body extracts and hemolymph vitellogenin (Vg), was used to measure the kinetics of Vg expression in the fat body and the levels in hemolymph. In addition, morphological and immunohistochemical changes that took place in the ovary during vitellogenesis were analyzed. The study was performed between 2 and 8 days post-ecdysis and between 2 and 25 days post-blood feeding. During the post-ecdysis period, D. maxima showed decreased synthesis of Vg and concomitantly, low levels of Vg in hemolymph (4.5  103 mg/ml at day 4). After a blood meal, Vg synthesis in the fat body and its levels in hemolymph increased significantly, reaching an average of 19.5 mg/ml at day 20. The biochemical changes observed in the fat body and hemolymph were consistent with the histological and immunohistochemical finds. These studies showed noticeable remodeling of tissue after blood feeding. r 2007 Elsevier Ltd. All rights reserved. Keywords: Vitellogenesis; Vitellin; Oocyte; Blood meal; Hematophagous vector

1. Introduction During oogenesis insects accumulate large amounts of proteins, lipids, carbohydrates as well as other minor components in an organized manner inside the eggs. The formation of these resources or ‘‘yolk’’ is critical since the egg can only survive if it contains enough nutrients to support embryonic growth, which occurs in isolation from the maternal body (Sappington and Raikhel, 1998). Vitellogenesis is a period during which oocyte growth is rapid due to the deposition of yolk. It is a heterosynthetic process and a central event of egg maturation. Thus, during vitellogenesis yolk protein precursors (YPPs) are produced in large amount by extraovarian tissues, mostly the fat body, secreted into the hemolymph and accumulated by developing oocytes (Raikhel and Dhadialla, 1992; Raikhel, 2005). Internalization of YPPs occurs via receptorCorresponding author. Tel.: +54 351 434 4974; fax: +54 351 433 3048.

E-mail address: [email protected] (L.E. Canavoso). 0022-1910/$ - see front matter r 2007 Elsevier Ltd. All rights reserved. doi:10.1016/j.jinsphys.2007.10.012

mediated endocytosis, an active process for the uptake and processing of large molecules that is ubiquitous among cells (reviewed by Snigirevskaya and Raikhel, 2005). Although several types of YPPs are accumulated by insect oocytes, vitellogenins (Vgs) are the most abundant in the majority of the insects. Vgs are large glycolipophosphoproteins that once selectively taken up by the oocytes are stored as vitellin (Vt) inside mature yolk bodies (also called yolk granules or platelets) until initiation of embryonic development. Vt therefore constitutes the main protein component of insect eggs that serves for embryonic and, in some cases, early larval development (Postlethwait and Giorgi, 1985; Oliveira et al., 1989; Giorgi and Nordin, 2005). Vg synthesis as well as its storage as Vt, have been extensively studied in Aedes aegypti. They both are under hormonal control and involve extensive cytological remodeling of the fat body and ovary (Belle´s, 2005; Giorgi et al., 2005; Raikhel, 2005). Hematophagous insects must surpass a threshold level regarding both the amount and quality of a blood meal in

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order to successfully produce eggs (Friend et al., 1965; Stoka et al., 1987). In the case of the vectors of Chagas’ disease, each gonotrophic cycle and egg development is strongly coupled with the intake of a blood meal (Stoka et al., 1987). However, autogeny, the capacity of a female to lay eggs without intake of any blood in the adult stage, has been observed in some species of Triatominae (Stoka et al., 1987; Noriega, 1992). Recently, great progress has been made in studying the biochemical, molecular, and biological aspects related to vitellogenesis in many species, but rather specifically in A. aegypti (Raikhel et al., 2002). However, with the exceptions of Rhodnius prolixus (reviewed by Valle, 1993; Atella et al., 2005) and to a lesser extent, Triatoma infestans (Salomo´n and Stoka, 1986; Stoka et al., 1987), vitellogenesis has been scarcely studied in Chagas’ disease vectors. In South and Central America, the vectors of Chagas’ disease are insects of importance in public health as they transmit the parasite Trypanosoma cruzi, the etiological agent of the disease (WHO, 2002). At present, the disease affects about 10 million individuals while more than 50 million are at risk of contracting the infection (Schofield et al., 2006). In order to understand how vitellogenesis takes place in Chagas’ disease vectors, and as a first step to understand its regulation, we have isolated and characterized Vt from oocyte extracts of Dipetalogaster maxima (Hemiptera: Reduviidae). This female, which is the largest among triatomines, is anautogenous under our parameters of rearing and exhibits a reproductive cycle that allows studies to be performed under standardized laboratory conditions. In addition to Vt characterization we have analyzed the relationships between Vg expression in the fat body and the levels in the hemolymph post-ecdysis and after females received a blood meal. Finally, these biochemical findings were correlated with ovarian development through histological and immunohistochemical studies. 2. Materials and methods 2.1. Reagents or chemicals Horseradish peroxidase (HPR) conjugated goat anti-rabbit IgG, o-phenylenediamine, bovine serum albumin, protease inhibitors, DEAE-Trisacryl-M, and DAB (3,30 -diaminebenzidine) were obtained from Sigma-Aldrich (St. Louis, MO). Electrophoresis protein standards and microtitre plates were from Biorad (Hercules, CA) and Delta Lab, respectively. Centriprep Centrifugal Filter Devices were purchased from Millipore-Amicon (Bedford, MA). Enhanced chemiluminescence (ECL) detection kit was from Perkin-Elmer. All other chemicals were analytical grade. 2.2. Insects Insects were taken from a colony of D. maxima, which is maintained under standardized conditions (28 1C, 70%

humidity, 8:16 h light: dark photoperiod) and fed every two weeks on hens blood (Canavoso and Rubiolo, 1995). For the experiments, males and females were separated before feeding at the last larval instar. Experimental females were segregated individually at emergence and placed together with two recently fed males for a period of 48 h. Thereafter, mated females (checked by observation of the spermatophore) were maintained in individual containers until they were able to feed a blood meal (day 10–12 post-ecdysis). Afterwards, hemolymph, fat bodies and ovaries were sampled from females in two physiological conditions: (a) at different times post-ecdysis (unfed period), (b) at different times after females received a blood meal, which represented between 3.0 and 5.5 times its body weight. In all cases, the size of the meal was estimated by determining the difference among the weight of insects before and after feeding. 2.3. Sampling collection At different times after molting or after blood feeding, insects were immobilized and the hemolymph was collected with a Hamilton syringe, from cut legs while gently pressing the abdomen. Hemolymph was received in cold microtubes, in the presence of 10 mM Na2EDTA, 5 mM dithiotreitol and protease inhibitors. Samples were centrifuged at 10,000  g for 5 min at 4 1C to remove hemocytes and then stored for one week at 70 1C for Vg quantification. After hemolymph collection, fat bodies and ovaries were carefully dissected out under cold phosphate buffered saline (PBS, 66 mM Na2HPO4/KH2PO4, 150 mM NaCl, pH 7.4) using a standard stereoscope with an optic fiber. Fat bodies were homogenized using disposable hand homogenizers in buffer Tris–NaCl (20 mM Tris, 150 mM NaCl, pH 7.4) containing a mixture of protease inhibitors (0.3 mM pepstatin, 1 mM aprotinin, 0.5 mM Na-tosyl-Llysine chloromethyl ketone (TLCK) and 1 mM benzamidine). The homogenates were first centrifuged at 2500  g for 10 min at 4 1C and the floating fat cake and pellet were discarded. The resulting material was centrifuged at 15,000  g for 30 min at 4 1C and the supernatants collected and used for Western blotting. Protein concentration was determined by Bradford (1976), using albumin serum bovine as standard. Ovaries were washed in cold PBS, fixed in 4% paraformaldehyde-PBS and processed for histological and immunohistochemical studies as described below. 2.4. Vt purification All steps were carried out in an ice bath or at 4 1C. Vt was obtained according to Salerno et al. (2002) with few modifications. Briefly, chorionated oocytes were removed from vitellogenic ovaries and washed with NaCl 150 mM. The oocytes were homogenized in 2 ml of buffer Tris– Na2EDTA (50 mM Tris, 2.5 mM Na2EDTA, 150 mM

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NaCl, pH 7.2) containing a mixture of protease inhibitors (0.3 mM aprotinin, 1 mM benzamidine, 10 mM PMSF, and 1 mM pepstatin). The homogenate was centrifuged for 10 min at 3000  g to remove large debris and then, centrifuged again for 30 min at 15,000  g. The resulting floating fat cake was discarded and the infranatant was subjected to a double passage through a DEAE-TrisacrylM column (1.5  14.5 cm), equilibrated with buffer Tris–HCl–Na2EDTA (20 mM Tris, 2.5 mM Na2EDTA, pH 8.3). The column was exhaustively washed with the same buffer and proteins were eluted using a linear gradient 0–300 mM NaCl (flow rate 20 ml/h). Fractions of 1.5 ml were collected and their protein content was estimated by the absorbance at 280 nm. The degree of purification was monitored by SDS-PAGE and fractions showing a banding compatible with Vt were pooled, concentrated by ultrafiltration (Centriprep-50) and stored at 70 1C until use. Molecular weights of vitellin apoprotein subunits as well as the mass of native Vt were estimated by PAGE under both denaturing (Laemmli, 1970) and non-denaturing conditions (Bollag and Edelstein, 1992), using protein molecular weight markers (Biorad) for the comparison. 2.5. Preparation of anti-Vt antibody Purified Vt was subjected to SDS-PAGE, loading about 15 mg of proteins per lane in the slab gels. Gels were stained with Coomassie brilliant blue and the two bands with Mr 170 and 174 kDa (quantitatively the most important bands), were sliced out of the gel and pooled. The bands were neutralized (Tris–HCl 100 mM, ethanol 10%, pH 8.0), homogenized and mixed with 1 ml of Freund’s complete adjuvant. The emulsion was inoculated on the back of two New Zealand rabbits. A second injection was applied 20 days later, but using Freund’s incomplete adjuvant. A final booster was made at day 35, inoculating approximately 150 mg of protein. Rabbits were bleeding 10 days after the third injection. The serum was collected after centrifugation at 2500  g for 20 min and the gglobulin fraction was obtained by precipitation with ammonium sulfate, according to Rubiolo (1981). Finally, the antibody was concentrated and stored at 20 1C in aliquots. 2.6. ELISA Vg concentrations in the hemolymph of females at different times after molting or after a blood meal were determined by an indirect enzyme-linked immunosorbent assay (Canavoso and Rubiolo, 1998). Microtiter plates were loaded with 200 ml/well of standard Vt or with appropriate hemolymph dilutions in buffer carbonate (15 mM Na2CO3, 35 mM NaHCO3, pH 9.6) and incubated for 90 min at 37 1C. Plates were washed four times with PBS-Tween (PBST: 8.2 mM Na2HPO4, 1.5 mM KH2PO4,

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150 mM NaCl, 2.7 mM KCl, 0.05% Tween 20, pH 7.4) and incubated with anti-Vt antibody (1:10,000, 60 min at 37 1C) in PBST containing 0.1% of bovine serum albumin. Plates were then washed as described above and loaded with antirabbit immunoglobulin conjugated to horseradish peroxides in PBST (1:5000; 30 min at 37 1C). After washing, plates were incubated with o-phenylenediamine in substrate buffer (81.5 mM Na2HPO4, 33 mM citric acid, 0.08% H2O2, pH 5.0). The enzyme reaction was allowed to proceed for 30 min and then stopped with 4 N H2SO4. Plates were read at 492 nm using an ELISA microplate reader. Vg concentration in the hemolymph was calculated on the basis of a standard curve of Vt. The linear response for the assay ranged between 2.0  103 and 40.0  103 mg/ml. 2.7. Western blotting Protein extracts from fat bodies were subjected to SDSPAGE (7.5%) and then transferred onto a nitrocellulose membrane at a constant current of 400 mA for 1 h. After rinsing the membrane with 10 mM Tris–HCl buffer containing 150 mM NaCl (TBS, pH 7.5), Western-blot analysis was performed according to Towbin et al. (1979). Blocking steps were performed with TBS-5% non-fat milk under gentle agitation at room temperature. Incubations with primary and secondary antibodies were performed for 1 h each at room temperature as follows: (a) polyclonal anti-Vt antiserum (1:5000) diluted in TBSTween 20 (0.1%) and (b) HPR conjugated goat anti-rabbit IgG (secondary antibody, 1:5000) in the same medium. After three washes with TBS, visualization of immunoreaction was performed by chemiluminescence, using the ECL detection kit (Perkin-Elmer) according to manufacturer instructions. 2.8. Histological and immunohistochemical studies Fixed ovaries in PBS-4% paraformaldehyde were embedded in paraffin, processed routinely for hematoxylin–eosin (H&E) and examined under a light microscope (Zeiss). In addition, tissue sections were immunostained with polyclonal anti-Vt antibody obtained as described previously (1:200). After incubation with HPR-conjugated goat anti-rabbit IgG the color was developed with the substrate chromogen DAB (3,30 -diaminobenzidine). Slides were examined under a Nikon Eclipse microscope. 2.9. Statistical analyses The experiments were performed in triplicate, using 6–8 insects per point. Except for Western blotting, all the samples from each insect were individually analyzed. Unless otherwise stated, the results are expressed as mean7SEM. Po 0.05 was considered the level of significant difference between means by Student’s t-test.

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When the extract of D. maxima oocytes was subjected to the first chromatography on DEAE-Trisacryl M two peaks, FI and FII were eluted with a linear salt gradient (Fig. 1A). FI eluted at an NaCl concentration 50–100 mM and showed one predominant protein band (490%), compatible with Vt by native-PAGE. Therefore, this peak was tentatively named D. maxima Vt. FI was subjected to a second chromatography using the same anion exchanger, leading to the elution of a single peak at a concentration of NaCl corresponding to 50–80 mM (Fig. 1B).

Purified Vt was analyzed by native-PAGE on a 6% acrylamide gel to determine its native molecular mass. Results showed an estimated size of 443 kDa on the basis of comparison with standards (Fig. 2A). When purified Vt was subjected to SDS-PAGE on a 7.5% acrylamide gel, it showed two major subunits that appeared to be arranged in doublets of 174 and 170 kDa, and two minor components of approximately 50 and 44 kDa (Fig. 2B). Analysis of the hemolymph and fat body protein extracts of D. maxima females and males showed that the antibody against the two major bands of Vt (174 and 170 kDa) recognized Vg in both the fat body and the hemolymph of females (Fig. 2D, lanes 2 and 3). However,

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Fig. 1. Vitellin purification by ion-exchange chromatography. (A) The homogenate of eggs was applied on a DEAE-Trisacryl M column, equilibrated with Tris–Na2EDTA buffer, pH 8.3. Proteins were eluted by a linear gradient of 0–300 mM NaCl (- - -) and fractions of 1.5 ml were collected. These were monitored at 280 nm (-J-) and pooled (FI and FII). (B) FI peak eluted from (A) was subjected to a second chromatography step using the same anion exchanger. Vitellin (Vt) was eluted at a gradient of 50–80 mM NaCl.

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Fig. 2. Characterization of vitellin from D. maxima. (A) Analysis of vitellin (Vt) by Native PAGE (6%). (B) Analysis of Vt by SDS-PAGE (7.5%) after purification by ion-exchange chromatography. Lane M, molecular weight markers. The molecular mass of native Vt or its subunits are indicated by arrows. (C) SDS- PAGE (7.5%) of Vt (lane 1), extract of fat body and hemolymph from vitellogenic females (lanes 2 and 3), extract of fat body and hemolypmph from mature males of D. maxima (lanes 4 and 5). (D) Western-blot analysis of the samples showed in (C) using a polyclonal anti-vitellin antiserum. Visualization of immunoreactive bands (indicated by an arrow head) was performed by chemiluminescence.

the doublet form was less evident by SDS-PAGE fractionation (Fig. 2C, lanes 2 and 3), most likely due to the heterogeneity of the protein material. Double immunodiffusion tests also showed that polyclonal anti-Vt antibody cross-reacted equally with purified Vt as well as with ovarian extracts of vitellogenic females (results not shown). Taken together, the results indicated that Vt and Vg shared similar immunological properties. By contrast, Vg was not present in fat body or hemolymph of D. maxima mature males (Fig. 2C–D, lanes 4 and 5). 3.2. Vg expression in the fat body and the levels in hemolymph In our laboratory colony, the period between eclosion to the adult stage and the first adult feed is normally 10–12 days for anautogenous females of D. maxima. During this period, b and g-oocytes reached a critical size, entering into the previtellogenic phase. After the insect takes a blood meal, the terminal oocytes (a position) begin active

vitellogenesis and the oviposition takes place between days 8 and 12 post-blood meal. Although some heterogeneity can be observed, usually the females lay an average of 30–35 eggs during about 25–28 days post-feeding, surviving to produce another batch of eggs if a second blood meal is ingested. Western-blot analyses were performed to demonstrate the effect of feeding on the expression of Vg protein in the fat body. During the unfed period, a faint immunoreactive band was detected for the protein at days 2, 4, and 6 postecdysis. Vg was not detected in fat body extracts from day 8 onwards (Fig. 3A). In agreement with these results, levels of Vg in hemolymph measured by ELISA were very low at different times post-molting, ranging between 2.8  103 and 5.7  103 mg/ml with the maximum value seen on day 4. However, under our experimental conditions, a small amount of Vg (2.3  103 mg/ml) was still detectable in the hemolymph at day 8 (Fig. 3B). After the insects were fed, Vg became detectable in the fat body and its level progressively increased. The pattern of

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Fig. 3. Vitellogenin in the fat body and hemolymph of D. maxima. The expression of vitellogenin (Vg) in the fat body was analyzed by Western blot (A and C), whereas the levels in the hemolymph were measured by ELISA (B and D). Studies were performed after ecdysis (unfed period) and at different times after females fed a blood meal. Values are means7SEM (n ¼ 8), performed in triplicate. *Po0.01 vs. day 2 post-ecdysis; #Po 0.001 vs. days 2 and 8 post-ecdysis; **Po 0.001 vs. days 4, 6, 8, 15 and 20 post-blood feeding.

protein expression obtained at different times after feeding showed a clearly visible immunoactive band for Vg at day 2, and sustained high levels of the protein between days 4 and 20 post-feeding. Vg protein expression then decreased, being comparable on day 25 to day 2 post-feeding (Fig. 3C). As shown in Fig. 3D, titers of Vg in the hemolymph also increased significantly after feeding. They increased from an average level of 6.5 mg/ml (day 2)–17 mg/ml (day 4 post-feeding) and then remained steady until day 20. Higher Vg levels were found at days 15 and 20 post-feeding (19.5 mg/ml), although they were not significantly different from those found at days 4, 6, and 8 post-feeding. However, at day 25, the Vg titer decreased reaching values comparable to day 2 post-feeding (Fig. 3D). 3.3. Morphological and immunohistochemical changes in the ovary during vitellogenesis In an attempt to correlate the above biochemical findings with ovarian development, histological and immunohistochemical studies were performed on D. maxima during the unfed period and during active vitellogenesis (post-feeding period). The gross anatomy of the female reproductive system of D. maxima is typical of hemipterans in that it consists of two ovaries each of which is made up of seven ovarioles of the telotrophic type (Huebner and Anderson, 1972). Fig. 4A displays the poor development of the ovary at day 2 post-molting. A histological examination of one ovariole section at this time showed the tropharium which contained large nurse cells, and a vitellarium, which was further away from the nurse cells. In this region, there was a transition zone where small oocytes were embedded, surrounded at the base by prefollicular cells (Fig. 4B).

During the post-feeding period, histological and immunohistochemical changes in the ovaries were in agreement with the changing pattern of expression of Vg in the fat body and its levels in the hemolymph. After a blood meal the ovarian tissue underwent noticeable development and oocytes in active vitellogenesis were observed at day 2 post-feeding (Fig. 4C). As vitellogenesis proceeded, the enlarging oocytes were gradually encompassed by a follicular epithelium (Fig. 4C, day 4 post-feeding). In addition, Vt stores in the oocytes became more abundant and were clearly observed by immunohistochemistry (Fig. 4D, day 4 post-feeding). At day 6, ovarioles contained oocytes with different degrees of maturation, showing a typical asynchronous development (Fig. 4E). Immunolocalization of Vt in a section of the ovarian tissue showed some differential deposition of the protein according to the size of the follicles. Thus, in smaller follicles, Vt became more uniformly distributed inside oocytes, although no complete packing of the core was observed (Fig. 4F). In more developed oocytes Vt was present in the outer space of the oocyte and yolk granules (Fig. 4F, inset). Occasionally, it was observed that some follicular cells reacted positively with the anti-Vt antibody raised. This immunostaining pattern was not detected in follicular epithelium of previtellogenic or early vitellogenic oocytes (before day 4 post-feeding) (results not shown). 4. Discussion Chagas’ disease remains one of the most serious public health problems in Latin-American countries (WHO, 2002). The vectors of the disease are hematophagous insects that belong to the Triatominae. Although R. prolixus is the member of this subfamily most studied,

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Fig. 4. Morphological and immunohistochemical changes on ovaries of D. maxima during vitellogenesis. (A and B) One hemi-ovary (20X) and a ovariole section at day 2 post-ecdysis (H&E, 200X). (C) Ovarian tissue 2 days post-feeding (H&E, 200X). (D) Immunohistochemistry showing vitellin accumulation in follicles at day 4 post-feeding (200X). (E) One hemi-ovary at day 6 post-feeding exhibiting ovarioles with typical asynchronism (30X). At this time, immunohistochemistry shows follicles at different stages of development (F, 150X). Magnification of an area corresponding to larger follicle shows vitellin deposition in the outer layer of oocyte and inside yolk granules (inset, 400X). NC, nurse cells; PFC, prefollicular cells; FC, follicular cells; FL, follicles; Oo, oocyte; Vo, vitellarium; Tro, tropharium; Tc, trophic cords; *Vt, vitellin; YG, yolk granules.

our understanding about the biology of reproduction in Chagas’ disease vectors is still limited. In this work, we have analyzed the process of vitellogenesis in D. maxima, a species that can be found in wild areas of Baja California, Mexico (Jime´nez et al., 2003). Among arthropods, evolution from predaceous to blood-sucking habits is associated with a series of morphological, physiological, behavioral, and demographic changes (Schofield, 1994). Thus, hematophagy entailed mechanical modifications in the mouthparts and the development of metabolic strategies in order to obtain blood from vertebrate hosts and use it as a food source (Lehane, 1991a; Ribeiro, 1995). The majority of triatomines ingest a large amount of blood in a single meal, comprising several times their own body weight (Schofield,

1994; Canavoso et al., 2004). In adult insects, the blood meal is largely used to provide resources for the reproductive effort; thus, in females, digestion and ovarian development are physiologically integrated (Lehane, 1991b). After blood feeding, a large amount of amino acids arising from protein digestion is used to produce molecules such as Vg, the main YPP. In most insects, Vg is synthesized in the fat body, secreted to the hemolymph, taken up by the growing oocytes, and stored as Vt (Raikhel, 2005; Atella et al., 2005). Regarding this process, it is important to keep in mind some of the pioneering studies that largely contributed to our understanding of how egg protein deposition takes place during vitellogenesis in many insects (Snigirevskaya and Raikhel, 2005). The deposition of extra-ovarian protein in insect oocytes was

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first observed by Wigglesworth (1943) in R. prolixus. This work further led to the characterization of Vg uptake by oocytes of Hyalophora cecropia (Telfer, 1960). Later on, it was established that the internalization of YPPs in A. aegypti occurred by receptor-mediated endocytosis, a mechanism now recognized as a general pathway employed by eukaryotic cells for internalizing specific macromolecules (Roth and Porter, 1964). Vg and Vt have been isolated from several species; high levels of these proteins are indicative of achieve oogenesis. Among triatomines, R. prolixus is the most studied insect with respect to Vg and Vt (Atella et al., 2005). However, scarce information is available regarding the other triatomine species. In D. maxima, characterization of Vt from chorionated oocytes showed a molecular mass of the native protein (Fig. 2A) close to those estimated for T. protracta (437 kDa; Mundall and Engelmann, 1977) and R. prolixus (430 and 260 kDa; Chalaye, 1979; Valle, 1993). However, our results were quite different from those reported for T. infestans (220 kDa; Salomo´n and Stoka, 1986). It is possible that where a small molecular mass was reported for Vt, this may have been due to proteolytic degradation during sample preparation (Valle, 1993). Interestingly, three different populations of Vt have been reported in R. prolixus oocyte extract, after purification by ion-exchange chromatography (Salerno et al., 2002). Vts, named VT1, VT2, and VT3 showed the same electrophoretical profile but differed in their phosphorylation levels as well as sugar composition. Although no definitive evidence can explain the role of three Vt classes in R. prolixus oocytes, it seems likely that they are the result of different post-endocytic processing pathways for Vgs arising from other sites rather than the fat body, such as follicular cells and the tropharium (reviewed by Atella et al., 2005). Under our experimental conditions only one kind of Vt was obtained on ion-exchange purification of D. maxima oocyte extracts (Fig. 1). Further work is needed to determine whether other triatomines follow the same pattern. In most insects, native Vgs or Vts dissociate into several subunits under denaturing conditions, with a range from 150–200 kDa to 50–60 kDa, respectively (reviewed by Raikhel and Dhadialla, 1992; Valle, 1993; Atella et al., 2005). The yolk proteins of the higher Diptera are an exception, since they consist of three to five peptides that range in molecular weight from 44 to 51 kDa (Bownes, 1982; Raikhel and Dhadialla, 1992; Tufail et al., 2005). In D. maxima, the analysis of Vt by SDS-PAGE showed four subunits with molecular weights similar to those reported for R. prolixus (Masuda and Oliveira, 1985; Valle et al., 1987), being in agreement with the majority of insects (Fig. 2B). In most insects, Vgs and Vts share immunological properties and are similar in physical and chemical characterization, although differences in lipid content and phosphorylation have been observed (Raikhel and Dhadialla, 1992). As expected, antibodies raised against

the large subunit of Vt from D. maxima also recognized immunologically Vgs from the female fat body and hemolymph (Fig. 2C–D). It is interesting to note that previous studies in the moth Manduca sexta used antibodies directed specifically against different subunits of Vts or Vgs to determine their relative positioning in the native molecule (Osir et al., 1986). In D. maxima, the antibody against a large subunit of Vt cross-reacted with Vg suggesting that this subunit is more exposed to the aqueous environment rather than residing within the native molecule. The fat body, a tissue hormonally regulated and functionally analogous to vertebrate liver, is the exclusive site of Vg synthesis in the majority of insects (Tufail et al., 2005; Giorgi et al., 2005). However, in some species the synthesis can also be accomplished by the ovarian follicular epithelium, as is well documented in higher Diptera (Bownes, 2005). Additionally, hematophagous insects, like vectors of Chagas’ disease require large blood meals to activate genes involved in blood digestion, the synthesis of YPPs and ultimately, the production of eggs (Raikhel, 2005). Vg synthesis in the fat body of D. maxima during the unfed period was minimal, as indicated by a reduced expression of the protein by Western-blot analysis (Fig. 3). However, expression of Vg in the fat body and its levels in the hemolymph increased significantly in insects after a blood meal, most likely in response to the secretion of juvenile hormone (JH) from the corpora allata. It is well known that in R. prolixus, JH released after a blood meal activates Vg gene expression and synthesis in the fat body (Coles, 1965; Wang and Davey, 1993; Belle´s, 2005). It is known that in R. prolixus JH also acts on the follicular epithelium surrounding the oocyte, causing cell shrinkage and the appearance of large inter-follicular spaces. This condition, known as patency, allows Vg synthesized in the fat body and other macromolecules to reach the surface of oocyte (Davey and Huebner, 1974; Davey, 1996, 2007). However, the ovarian follicle cells of some species are capable of intensely secreting yolk polypeptides into the perioocytic space of the ovarian follicles (Giorgi et al., 2005). Recently, the follicle cells of R. prolixus have been shown to be capable of synthesizing Vg (Melo et al., 2000). Unlike many higher Diptera, in which the yolk protein synthesis is initiated simultaneously in both the fat body and the ovary (Bownes, 2005), only late follicles that had largely ceased Vg uptake from hemolymph were involved in the Vg production of R. prolixus (Melo et al., 2000). In D. maxima, as seen in the hemipterans R. prolixus (Pratt and Davey, 1972) and T. infestans (Stoka et al., 1987), the ovarioles exhibited asynchronous development so that oocytes at different stages of maturation are simultaneously present (Fig. 4E). Interestingly, we observed that a few follicular epithelial cells surrounding the largest oocytes immunoreacted positively when challenged with an anti-Vt antibody (results not shown). This observation suggests that at least some follicular cells could synthesize Vg. However, unlike

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R. prolixus where Vg was found inside the great majority of follicular cells (Melo et al., 2000), the phenomenon in D. maxima did not appear to be generalized. Undoubtedly, the fat body serves as the major source for building Vt stores in oocytes of R. prolixus and D. maxima. Clearly the timing for Vg synthesis in both vectors changes in response to blood feeding and the size reached by the follicles. Therefore, the ability of follicular cells to accomplish Vg synthesis in D. maxima deserves to be explored using complementary biochemical and cellular approaches. The nutritional control of vitellogenesis in anautogenous mosquitoes is now understood at a molecular level. In A. aegypti, both the amount and quality of hemolymph amino acids arising from blood digestion are critical in regulating the expression of specific YPP genes (Attardo et al., 2005). In triatomines, the intake of a blood meal triggers vitellogenesis, enabling females to achieve a high productivity of eggs. However, some species like R. prolixus exhibit a typical autogenic behavior (Stoka et al., 1987; Noriega, 1992). Understanding the link between the nutritional requirements to activate vitellogenesis and biochemical and molecular factors involved in egg development in Triatominae offers an opportunity to explore the reproductive process of these vectors, including their natural feeding behavior and lifestyle. Further investigation into these areas could contribute to designing effective strategies for the control of the vector. Acknowledgments We thank Dr. Patricia Scaraffia for critical reading of the manuscript and for her many important suggestions throughout the course of the work. We also wish to thank Ana C. Donadı´ o for technical assistance in microscopy, Rau´l Stariolo for rearing of insects, and the anonymous reviewers for their helpful comments. This work was supported by Grants from Consejo Nacional de Investigaciones Cientı´ ficas y Te´cnicas (CONICET) (LEC) and from SECyT- Universidad Nacional de Cordoba (ERR). LEC is a member of the CIC-Consejo Nacional de Investigaciones Cientı´ ficas y Te´cnicas (CONICET-Argentina). S.A.A. is a research fellow from SECyT-UNC. References Atella, G.C., Gondim, K.C., Machado, E.A., Medeiros, M.N., SilvaNeto, M.A.C., Masuda, H., 2005. Oogenesis and egg development in triatomines: a biochemical approach. Anais da Academia Brasileira de Ciencias 77, 405–430. Attardo, G.M., Hansen, I.A., Raikhel, A.S., 2005. Nutritional regulation of vitellogenesis in mosquitoes: implications for anautogeny. Insect Biochemistry and Molecular Biology 35, 661–675. Belle´s, X., 2005. Vitellogenesis directed by juvenile hormone. In: Raikhel, A.S. (Ed.), Reproductive Biology of Invertebrates, vol. XII, Part B (Adiyodi, A.G., Adiyodi, R.G. (Eds.)). Science Publishers Inc., pp. 157–197. Bollag, D.M., Edelstein, S.J., 1992. Gel electrophoresis under nondenaturing conditions. In: Bollag, D.M., Edelstein, S.J. (Eds.), Protein Methods. Wiley-Liss Inc, New York, pp. 143–160.

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