Anopheles gambiae lipophorin: Characterization and role in lipid transport to developing oocyte

ARTICLE IN PRESS Insect Biochemistry and Molecular Biology Insect Biochemistry and Molecular Biology 36 (2006) 375–386 www.elsevier.com/locate/ibmb

Anopheles gambiae lipophorin: Characterization and role in lipid transport to developing oocyte Georgia C. Atellaa,b, Ma´rio Alberto C. Silva-Netoa,b, Daniel M. Golodneb, Shamsul Arefinc, Mohammed Shahabuddina,c, a

Laboratory of Malaria and Vector Research, National Institute of Allergy and Infectious Diseases, National Institutes of Health, 4 Center Drive MSC 0425, Bethesda, MD 20892, USA b Departamento de Bioquı´mica Me´dica , Instituto de Cieˆncias Biome´dicas, Universidade Federal do Rio de Janeiro 21941-590, Brasil c Department of Biology, Boston College, 140 Commonwealth Avenue, Chestnut Hill, MA 02467, USA Received 20 June 2005; received in revised form 26 January 2006; accepted 27 January 2006

Abstract Lipid transport in arthropods is achieved by highly specialized lipoproteins, which resemble those described in vertebrate blood. Here, we describe purification and characterization of the lipid–apolipoprotein complex, lipophorin (Lp), in the malaria vector mosquito Anopheles gambiae. We also describe the Lp-mediated lipid transfer to developing eggs and the distribution of the imported lipid in developing embryos. The density of the Lp complex was 1.135 g/ml with an apparent molecular weight of 630 kDa. It is composed of two major polypeptides, apoLp I (260 kDa) and apoLp II (74 kDa) and composed of 50% protein, 48% lipid and 2% carbohydrate (w/w). Hydrocarbon, cholesterol, phosphatidyl choline, phosphatidyl ethanolamine, cholesteryl ester and diacylglyceride were the major Lpassociated lipids. Using fluorescently tagged lipids, we observed patterns that suggest that in live developing oocytes, the Lp was taken up by a receptor-mediated endocytic process. Such process was blocked at low temperature and in the presence of excess unlabeled Lp, but not by bovine serum albumin. Imported Lp was segregated in the spherical yolk bodies (mean size 1.8 mm) and distributed evenly in the cortex of the oocyte. In embryonic larvae, before hatching, a portion of the fatty acid in vesicles was found evenly distributed along the body, whereas portion of phospholipids was accumulated in the intestine. r 2006 Elsevier Ltd. All rights reserved. Keywords: Lipoprotein; Mosquito; Oogenesis; Lipid transport; Insect; Disease vector; Malaria

1. Introduction Mosquitoes are vectors of many human diseases including malaria, yellow fever, filariasis, dengue and West Nile Virus. Attempts to eradicate mosquitoes by insecticide have failed, because of the emergence of insecticideresistant mosquitoes. An effective strategy to control mosquito population is to interfere with egg laying and reducing larval density. Rapid increase of mosquito population in optimal seasons is due to the laying of several hundreds of eggs in a few days. In most vector Corresponding author. Department of Biology, Boston College, 140 Commonwealth Avenue, Chestnut Hill, MA 02467, USA. Tel.: +1 617 552 1921; fax: +1 617 552 2011. E-mail address: [email protected] (M. Shahabuddin).

0965-1748/$ - see front matter r 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.ibmb.2006.01.019

mosquitoes, egg development requires vertebrate blood components. Therefore, understanding mosquito egg development and the role of host blood components in the process may identify novel ways to control the rapid proliferation of these potentially deadly disease-carriers. It is known that lipids are carried as lipoprotein complexes and delivered to target tissues. However, it is not clear how different egg-laying animals transport nutrients to developing eggs and store nutrients for neonates to use until it ingests its first meal. Lipid compositions in eggs and neonates of different egg-laying and live-bearing lizards was found to be remarkably different (Speake and Thompson, 2000; Thompson et al., 2000). In the liver of the neonate chick, lipid composition showed an increase in cholesteryl ester and a decrease in triglycerides after birth (Noble and Ogunyemi, 1989; Noble and Cocchi, 1990). Lipid transport in vertebrates

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is orchestrated by chylomcron, very-low-density (VLDL), low-density (LDL), intermediate-density (IDL) and highdensity (HDL) lipoproteins. Whereas in insects, lipid transport occurs via lipophorin (Lp), a class of HDL (Chino et al., 1981; Kawooya and Law, 1988; Arrese et al., 2001). Lps are complexes of lipids and apoproteins that carry and distribute different classes of lipids through insect hemolymph to various tissues (Chino et al., 1981; Shapiro et al., 1988; Atella et al., 1992). The role of Lp as the major lipid vehicle for lipid transport to the oocytes was shown in Philosamia cynthia (Chino et al., 1977) and Manduca sexta (Kawooya et al., 1988). Most insect Lps contain two apolipoproteins—apolipoprotein I (
250 kDa) and apolipoprotein II (
80 kDa), which comprise about 60% of the mass of the lipid/protein complex (Chino, 1985). A third apoprotein (apolipoprotein III) of molecular mass
17 kDa is present in adults of some species (Chino et al., 1986). Also, it is well demonstrated that the molar ratio of the apoproteins is a function of the insect physiological state (Shapiro and Law, 1983; Chino, 1985; Kawooya et al., 1986). Like the apoproteins, lipid composition also varies depending on the insect and its physiological state. Generally, the major lipids associated with lipophorin are diacylglycerides, cholesterol, hydrocarbons and phospholipids (Thomas and Gilbert, 1968; Chino et al., 1969; Peled and Tietz, 1973; Gondim et al., 1989). Insect Lps may function as reusable shuttles for lipid transport (Blacklock and Ryan, 1994; Soulages et al., 1994; Arrese et al., 2001; van der Horst et al., 2002). For example, Lp complexes transfer diacylglycerols to flight muscles and the protein can be recycled back to the hemolymph. Also in M. sexta, low-density Lps (LDLp) first unloads its lipids to the fat body. Then the apolipoprotein is recycled back to the hemolymph (Kawooya et al., 1988). During oogenesis, a rapid accumulation of proteins and lipids occur in the oocytes, hence facilitating the eggs to mature in a relatively short time. This process is complex and the involvement of several tissues coordinated by hormones has been demonstrated (Engelmann, 1979). High-density Lp (HDLp) also transfers lipids without recycling of apoproteins (Kawooya et al., 1988). The presence of a Lp receptor Aedes aegypti suggested that in this mosquito, the internalization of Lp occurs via endocytosis (Cheon et al., 2001). The internalized Lp acts as yolk protein precursor (Kulakosky and Telfer, 1990; Sun et al., 2000). Despite the evidence suggesting that Lp is internalized in a developing oocyte by endocytosis (Van Antwerpen et al., 1993), few clear observations of endocytic vesicles in oocytes have been demonstrated. Recently, Van Hoof et al. (2005) examined lipoprotein and transferrin trafficking in insect cells. Using a lipoprotein-specific antibody with Lp receptor gene transfected cultured cells, endocytic vesicles were observed by these authors. However, studies of endocytosis have proven to be difficult with fat body due to the fragility and irregular composition of the tissue (Locke, 1998).

Here, we describe the purification and characterization of malaria vector mosquito Anopheles gambiae Lp and the use of fluorescently labeled Lp to examine the process of lipid transfer in live developing oocytes. The results showed that lipophorin is taken up via endocytosis by mosquito eggs. We also examined the distribution of imported lipids in maturing eggs and in developing embryos. This showed that imported endocytic vesicles fuse to form the yolk bodies until they reach to a certain size. Imported fatty acids and phospholipids remained stored in yolk bodies during egg development. However, some of these lipids apparently remain unused during egg and embryonic development, and are distributed in different tissues in newly hatched larvae. 2. Materials and methods 2.1. Mosquito rearing The An. gambiae (G3 strain) colony was maintained under standard insectary conditions (27 1C and 80% relative humidity, photoperiodism: 12L–12D) and fed on a diluted Karo syrup (CPC International Inc., Englewood Cliffs, NJ) saturated cotton ball. For blood feeding, 6–8 days after emergence from pupae, adult female mosquitoes were starved overnight and fed ad libitum on 4- to 5-weekold White Leghorn chickens. Fed mosquitoes were separated and kept in the insectary until used. 2.2. Lp purification and composition analysis An. gambiae Lp was purified as described earlier with minor modifications (Atella and Shahabuddin, 2002). Five to 10 days after emergence from pupae, sugar-fed mosquitoes (4 g) were snap frozen under liquid nitrogen and ground to a powdered form with a ceramic grinder (Fisher Scientific, Pittsburgh, PA). The powder was immediately transferred to the homogenization solution (10 mM phosphate, 0.15 M NaCl, pH 7.4) containing glutathione (20 mM), EDTA (5 mM), PMSF (2 mM), antipain (0.5 mg/ml), pepstatin (5 mM) and leupeptin (0.5 mg/ml) on an ice bath. The homogenate was centrifuged at 100,000  g, for 30 min at 4 1C. Solid KBr was added to the supernatant to a final concentration of 0.4 g/ml and the mixture was again centrifuged at 125,000  g in a Beckman ultracentrifuge (Optima L-90 ultracentrifuge, Beckman Coulter, Palo Alto, CA) with a fixed angle Beckman rotor Ti50.2 at 4 1C for 20 h. Supernatant was collected from the top as fractions. Presence of Lp in each fraction was checked using SDS polyacrylamide gel. Fractions with typical banding pattern of Lp were pooled and extensively dialyzed against PBS. Samples were then concentrated using a vacuum dryer (speed vac, Savant) and stored under liquid nitrogen. Protein concentration was estimated using micro BCA Kit (Pierce, Rockford, IL) in the presence of 0.5% SDS, using bovine serum albumin (BSA) as standard. Each

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fraction was analyzed by PAGE both under denaturing (Laemmli, 1970) and nondenaturing conditions (Davies, 1964). Molecular weights of the apoproteins were determined by comparing with prestained full range (250–10 kDa) protein molecular weight markers (Biorad Laboratories, Richmondm, CA). For radioactive samples, the gels were stained, dried and autoradiographed. Total carbohydrate was determined colorimetrically by the sulfuric acid/phenol method using glucose as standard (Dubois et al., 1956). Molecular mass of the Lp was estimated in a Superose 6 HR 10/30 HPLC column equilibrated with 20 mM Tris–HCl, 1.0 M NaCl, pH 8.0, at a flow rate of 0.5 ml/ min, and calibrated using the following protein standards: thyroglobulin (669 kDa); apoferritin (440 kDa); b-amylase (200 kDa); BSA (66 kDa); soybean trypsin inhibitor (20 kDa). Lp density was determined according to the method described earlier (Gondim et al., 1989) with minor modifications. Briefly, purified 32P-Lp was added to a PBS containing 44% (w/v) KBr, in a final volume of 5 ml. This solution was put into a centrifuge tube, overlayed with 5 ml of PBS, and centrifuged for 20 h in a Beckman 40 Ti at 125,000  g at 4 1C. After centrifugation, the KBr gradient was fractionated. The density of each fraction containing radioactivity was determined from the refractive index of KBr at 25 1C. 2.3. Preparation of

32

P-phospholipid-labeled Lp

32

Pi, 0.1 ml (1 mCi, Amersham, Piscataway, NJ), was injected into the hemolymph of adult females using a PLI100 microinjector (Harvard Apparatus, Holliston, MA). One day later, 32P-phospholipid-labeled Lp (32P-Lp) was purified from the injected mosquitoes, as described above, from the total mosquitoes homogenate on a KBr ultracentrifugation gradient. The incorporation of radioactivity was measured by scintillation counting. 2.4. Preparation and analysis of 3H-lipid-labeled Lp 3

H-lipid-labeled Lp (3H-Lp) was prepared as follows. H-palmitic acid (0.1 ml, 1 mCi, Perkin Elmer, USA) was injected using a PLI-100 microinjector (Harvard Apparatus, Holliston, MA) on the first day after a blood meal. After 24 h, of 3H-Lp was purified from the injected mosquito, as described above, from the total mosquitoes homogenate. The incorporation of radioactivity was measured by scintillation counting. Routinely, 3H-Lp was subjected to a lipid extraction and high performance thinlayer chromatography (HPTLC). The spot of each lipid was scrapped and the radioactivity was determined.

3

2.5. Lipid analysis Lipid extraction was performed according to Bligh and Dyer (1959) with modifications. Samples were mixed for

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2 h in a stoppered tube using 5 ml of chloroform–methanol–water solution (2:1:0.8, v/v/v), with intermittent agitation. The mixture was centrifuged and the supernatant was collected. The pellet was subjected to a second lipid extraction (1 h). To the pooled supernatant, 2.5 ml of water and 2.5 ml of chloroform were added. The mixture was vigorously shaken. After centrifugation, the organic phase was removed and dried under a stream of nitrogen. Total lipid content was determined gravimetrically. Lipids were analyzed by HPTLC on Silica gel 60 plates (Merck, Darmstadt, Germany). Plates were first developed in hexane–ethyl ether–acetic acid (60:40:1, by volume) until the solvent front reached the middle of the plate and then in hexane–chloroform–acetic acid (80:20:1, by volume). HPTLC plates were stained by spraying with sulfuric acid (30%) and heating at 120 1C for 10–15 min. Quantitation of different lipids was performed by analyzing digital image of the HPTCL plates using Quantiscan software (Biosoft, Cambridge, UK). The plates were also stained with iodine and autoradiographed. The spots were scraped, the lipids eluted, and the radioactivity associated with each spot was determined by scintillation counting. 2.6. Lp labeling and in vivo observation Purified Lp was labeled with fluorescent fatty acid (BODIPYs FL C16) or phospholipids (TRITC-DHPE) (Molecular Probes, Eugene, OR, USA) using the method described by Martin-Nizard (Martin-Nizard et al., 1987). The apoprotein of the Lp was also labeled with fluorescein 50 -isothiocyanate (FITC) using a Fluorescein-EX Protein Labeling Kit (FluoReporters, Molecular Probes, Eugene, OR, USA). Distribution of the fluorescent lipids and Lps was observed using the methods described (Atella and Shahabuddin, 2002). Following the manufacturer’s instructions for radiolabeling (Sigma, St Louis, MO), purified Lp was iodinated with 125I-iodine (100 mCi/ml, Amersham, Piscataway, NJ), using 200 mCi/mg of protein. The iodogen was used at a concentration of 100 mg/mg of protein. To remove the free iodine, the reaction mixture was passed through Sephadex G-50 spin columns. As a control experiment, Lp was also labeled with nonradioactive iodine, following the same procedure used for the radioactive labeling. 2.7. Lp binding to the ovaries Nonradioactive ovaries were dissected and incubated for different periods on ice under three different experimental conditions: (i) with purified 125I-Lp (0.1 mg/ml); (ii) with 125 I-Lp in the presence of different concentrations of nonradioactive Lp or (iii) in the presence of 10 mg/ml BSA. After incubation, the ovaries were washed in 1 ml of culture medium at 28 1C and homogenized in PBS. The radioactivity was determined using a scintillation counter.

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2.8. Lp and lipid uptake by ovaries At different days after blood feeding, fluorescent or radioactive labeled Lp was injected into the hemolymph of vitellogenic females using a PLI-100 microinjector (Harvard Apparatus, Holliston, MA). At the desired time, the ovaries were dissected in Medium 199 (Sigma Chemical Co., St. Louis, MO) and follicles were separated according to their stages of development (Christophers, 1911), washed in the medium and let stand for 30 min at 28 1C in 1 ml of the same culture medium. The fluorescence distribution in the ovary was observed using confocal microscopy (TCS-NT/SP, Leica Microsystems GmbH, Heidelberg, Germany). The radioactive ovaries were homogenized in 0.15 M sodium chloride and the radioactivity was estimated.

3. Results 3.1. An. gambiae Lp purification and characterization The clear supernatant from homogenates of sugar-fed adult An. gambiae was subjected to a KBr ultracentrifugation gradient and the fractions were analyzed by SDSPAGE (Fig. 1A). As expected, the fractions collected from the top of the gradient showed primarily two bands with molecular weights of 260 and 74 kDa. The molecular weights were consistent with the apoLp from other insect species of order Diptera (the fruit fly Drosophila melanogaster, the mosquito Ae. aegypti, the midges Chaoborus maximus, the black fly Simulium vittatum and the crane fly Nephrotoma abbreviata) (Pennington and Wells, 2002). Lps from these insects also consisted of two proteins of approximately 240 and 75 kDa each. The banding pattern on the SDS-PAGE suggested that the preparation of the An. gambiae Lp was essentially pure. The two apolipoproteins were then called AgapoLp-I (260 kDa) and AgapoLpII (74 kDa) (Fig. 1A). The purified Lp was subjected to analytical gel filtration to determine the native molecular mass of the Lp complex. Results suggested an estimated size of
636 kDa (Fig. 1B). The estimated density of the purified Lp from adult mosquitoes was 1.13570.005 g/ml (N ¼ 4), which suggests that the Anopheles Lp is predominantly of high density (HDL) similar to those found in other dipterans (Pennington and Wells, 2002). The composition of the purified lipophorin was estimated gravimetrically. It contained 50% protein, 48% lipid and 2% neutral sugar. Hydrocarbon (HC), free cholesterol (CH), cholesteryl ester (CE), diacylglycerol (DG), triacylglycerol (TG) were the neutral lipids found in the Anopheles Lp (Fig. 1C). Although relatively low, significant amounts of free fatty acid (FA) were also present. Among the phospholipids, phosphatityl choline (PC) and phosphatidyl ethanolamine (PE) were the major phospholipids found (data not shown).

Fig. 1. Purification and characterization of An. gambiae lipophorin. (A) Mosquito homogenate was subjected to KBr gradient ultracentrifugation. After centrifugation, fractions were collected from the top of the gradient, analyzed by SDS-PAGE (5–15% polyacrylamide gel), and stained with Coomassie brilliant blue. (B) Molecular mass determination of the Lp. Lp and the protein standards: (a) thyroglobulin, (b) apoferritin, (c) b-amylase, (d) bovine serum albumin and (e) soybean trypsin inhibitor were separately applied on a Superose 6 HR 10/30 HPLC column. Absorbance at 280 nm was monitored and the peak elution volume was determined for each standard. The arrow indicates the elution position of Lp; Vo, void volume; and Ve, elution volume. (C). Lipid composition of the lipophorin. Total extracted lipids were analyzed by high performance thinlayer chromatography (HPTLC). Result presented as a percent of total lipid. HC, hydrocarbon; CE, cholesteryl ester; TG, tryglyceride; FA, fatty acid; CH, cholesterol; DG, diglyceride; ND, undetected; and PL, phospholipids.

3.2. Role of the Lp in mobilization of lipid Anautogenous mosquitoes require vertebrate blood for reproduction. Ingested blood provides nutrients for survival, flight and egg development. Hemolymph Lps play key roles in mobilizing lipids from the fed vertebrate blood in the midgut to storage space in fat body and to the sites of usage. Therefore, the extent of lipid distribution in various organs is likely to vary depending on the demand and activity of the tissue that ensures optimal utilization of the limited resource. This is especially interesting for a mosquito that has lived only on sugar water and never had a blood meal. We examined distribution of phospholipids in several active tissues after the first blood meal. We fed 5 to 9-day-old mosquitoes on chicken ad libitum. Processing of the ingested blood begins immediately in the

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Fig. 2. Uptake and distribution of lipids from the lipophorin. (A) Uptake of radiolabeled lipids by different organs in vivo. Six hours after injection of 32PLp into vitellogenic females, different organs were dissected, washed and homogenized. The radioactivity present in each organ was estimated by scintillation counting. Results are average of three independent experiments: MG, midgut; OV, ovary; FB, fat body; MS, muscle. (B) Transfer of lipid into oocytes at different stages of egg development. Stages described according to Christophers (1911). (C) Kinetics of lipid uptake by stage 2 oocytes at 28 1C. A parallel experiment was conducted at 4 1C to determine the basal levels of lipophorin binding to the tissue. Result presented is the value obtained at 28 1C minus the value at 4 1C. (D) Uptake of radiolabeled lipophorin at 28 1C without any competitor (’), in presence of unlabeled lipophorin (m) or BSA (K).

posterior midgut and continues till about 60 h. Peak blood digestion takes place during 24–48 h after feeding. Around 24 h after the blood meal, we supplemented the hemolymph Lp pool with exogenous 32P-labeled Lp or 3H-lipid labeled Lp by injection into the hemocoel. About 24 h after the injection, the midgut, ovary, fat body and muscle were isolated. The radioactivity associated with each tissue was then measured (Fig. 2A). Highest amount of the lipid was found in the developing ovary, which was almost half of the total lipid measured. A remarkable accumulation of lipid was also noted in the fat bodies. Radioactivity associated with midgut was about a third of the ovary and two-thirds of the fat-body-associated radioactivity. Thoracic muscle is involved in flight. A significant amount of lipid from the Lps also absorbed by the muscle, although the amount was less than the other tissues. The results are in agreement with the fact that the majority of the nutrients from the ingested vertebrate blood are invested in reproduction. 3.3. Kinetics of lipid transfer to the ovaries Since the ovary is the major target tissue of the Lpbound lipids, we studied the mode of transfer of lipids from the Lp to the ovary. First, we examined the kinetics of transfer at different stages of the egg development to

determine when major lipid transfer takes place after a blood meal. Following injection of radiolabeled Lp into the hemolymph, the mosquitoes were kept in the insectary for 5 h. The ovaries were then dissected out and the amount of incorporated radioactivity was determined. Lipid incorporation by different stages of ovaries from injected Lp is shown in Fig. 2B. The incorporation began immediately after activation of the oocytes. At stage 2, which roughly corresponds to the second day after the blood feeding, the rate of lipid incorporation increased about two-fold. The incorporation gradually declined during the 3rd and 4th stages of development, but still was significant. When the first egg is already laid or ready to be laid on day five, the lipid incorporation to the ovary was minimal. To further examine the kinetics of lipid incorporation at stage 2 when the oocyte is most active, we dissected the developing ovaries on the second day after the blood meal. Ovaries were incubated in a medium supplemented with 32Plabeled purified Lp. After incubation for different periods, the associated radioactivity was measured (Fig. 2C). The incorporation was essentially linear during the 4 h period. 3.4. Mechanism of lipid uptake in developing eggs Fig. 2C shows a linear kinetics of lipid uptake at the most active stage of egg development indicating an active

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internalization of the lipid. To examine if the Lp is internalized by the eggs and if Lp-mediated lipid delivery is a receptor-mediated process, we performed a competition assay with unlabeled Lp and 125I-labeled Lp. Unlabeled BSA as a nonspecific protein competitor was used in control experiments. Fig. 2D shows that in the presence of unlabeled Lp, labeled Lp uptake was blocked in a dose-dependent manner. This showed that the unlabeled Lps competed effectively with the labeled molecules for entry into the oocyte. Similar competition was not observed with BSA. Together the competition studies suggested that a receptor-mediated process is involved in Lp-mediated lipid transfer to the developing eggs. Receptor-mediated internalization of molecules is an energy-dependent active process and significantly reduced at lower temperatures (Goldstein et al., 1976). We then examined the uptake of the Lp at 28 and 4 1C. At 28 1C, the uptake of Lp was linear for at least 6 h. At 4 1C, no increase of lipid uptake was observed. Together these results demonstrated that the Lp internalization by developing eggs in An. gambiae is an energy-dependent active process. Use of radiolabeled Lp provided information on the kinetics of uptake of the lipid and Lp in developing eggs. However, using this technique, it was difficult to comprehend spatial distribution of the Lps in the developing eggs. For example, it is difficult to know if individual eggs in the ovary are more active than the other or if any specific area of the developing oocyte surface is particularly active in the uptake process. To gain more insight into the spatial distribution, we supplemented the hemolymph Lp after a blood meal with fluorescent phospholipids- and fatty acidlabeled Lp. At different stages of development, the oocytes were dissected out and examined with confocal microscopy. When oocytes were dissected 1 day after the blood feeding and a few hours after the labeled Lp injection, accumulation of Lp was observed around the vitellin membrane (Fig. 3B and C). Labeled Lp was found attached to the follicular membrane around the oocytes. At this stage, a fluorescent punctate pattern was visible around the oocyte. The accumulation around the vitellin membrane was uneven and appeared patchy. This pattern is consistent with the endocytic vesicles observed in cells with Lp receptor importing labeled Lp (Van Hoof et al., 2005). As the oocytes grow, more lipids were found accumulated as yolk bodies in the cortex (Fig. 3D–F). The lipid-filled yolk bodies were also observed in freshly laid eggs (Fig. 3G–I). Examining the entire ovary showed similar fluorescence in all oocytes (Fig. 3J–L), suggesting that most eggs in the mosquito mature concurrently. This is consistent with the laying of a batch of eggs within a short period. If lipid deposition in mosquito eggs occurs by an endocytic process, it is likely that fusion of the internalized endocytic vesicles form the spherical yolk bodies. We examined the mosquito yolk bodies with confocal microscopy. Fig. 4A shows a stage 2 oocyte. Measurement of 308 yolk bodies

showed an estimated median diameter of 1.84 mm (Fig. 4B). The mean diameter is 1.79 mm with a standard deviation of 0.453 and a standard error of mean of 0.026. More variation in size was observed in the smaller size range than larger size range. This may suggest that the yolk bodies become larger as sequential deposition of Lp occurs in stages, until reaching a certain size. What determines the size of the yolk body is not clear. A noticeable difference in fluorescent intensity for fatty acid and phospholipids in the yolk bodies was not observed. 3.5. Distribution, conversion and storage of lipid in larvae Unlike live-bearing animals, where developing embryos are nourished by the continuous supply of nutrients, egglaying (oviparous) animals must transfer all nutrients needed for embryonic development and survival of the neonate until the first food is found and ingested. In oviparous animals, the maternal nutrients need to be mobilized in correct tissues during embryonic development. Previously, we reported that some maternal fatty acids accumulate along the body of the neonate larvae and a portion of the phospholipids are concentrated in both the gastric cecae and intestine of the newly hatched larvae (Atella and Shahabuddin, 2002). To gain insight about the differential accumulation of different lipids before hatching, we examined the developing embryos from eggs deposited by mosquitoes injected with labeled Lp. It was only possible to examine embryos of about 36 h old. Approximately 40 h after laying, Anopheles eggs begin to hatch. The 36-h-old embryo was already fully developed (Fig. 5A). Green fluorescence was almost evenly distributed along the entire body (Fig. 5C). The majority of the fluorescence seemed to be in punctate and patchy pattern. The embryonic intestine, however, had more fatty acid than the rest of the body. Confocal imaging showed more phospholipids also accumulated in the embryonic intestine (Fig. 5B and D). The distribution of the maternal phospholipids and fatty acids was more dramatic in newly hatched larvae (Fig. 6A) as described earlier (Atella and Shahabuddin, 2002). To examine the fatty-acid-related fluorescence, we looked at the larvae more closely with higher magnification. Spherical bodies appeared as droplets of fat accumulated in each of the segments of the neonate larvae that fluoresced intensely green (Fig. 6B and C). Unlike the yolk bodies in oocytes, no phospholipid fluorescence (red) was detected in the spherical bodies. This suggested that the lipid types were truly separated and accumulated in different locations. Lp of adult An. gambiae contains little free fatty acid (Fig. 6D). To examine if the injected fatty acid is converted to neutral lipids, we extracted total lipid from mosquito larvae hatched from eggs injected with exogenous fatty acid and phospholipids. Total lipids were subjected to separation in HPTLC with an acidic solvent (Fig. 6D). Most of the fluorescent fatty acid did not move from the origin in an acidic solvent, which that indicated the fatty acids were

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Fig. 3. Real-time observation of lipid uptake at different stages of egg development (Panels A, D, G and J). Vitellogenic mosquitoes were injected with lipophorins labeled with BODIPY-labeled fatty acid (green) or TRITC-labeled phospholipids (red) 1 day after the blood meal. Developing oocytes were dissected at different times and examined with confocal microscopy. About 5–6 h after injection, vesicular structures surrounded the vitellogenic membrane (Panels B and C; arrow in C), indicating endocytic uptake of lipids. Endocytic vesicles fuse to form yolk bodies, which are distributed evenly in the cortex of the more mature oocytes (Panels E and F). Yolk bodies were visible in newly laid eggs (Panels H and I). All oocytes of the same batch apparently mature and uptake lipids simultaneously (Panels K and L).

converted to neutral lipids. Further characterization of the modified fluorescent neutral lipids was not performed. In contrast to the fatty acid fluorescence, a detectable change was not observed in phospholipids obtained from neonate larvae and the control mixture. 4. Discussion To understand the process of lipid utilization from the ingested vertebrate blood, we have purified the Lp of the malaria vector An. gambiae and examined the properties of

the lipid carrier using fluorescently labeled lipids. For the first time, we have directly observed evidence of endocytosis of Lp in live mosquito eggs, We have also examined how lipids are both imported in the developing oocyte and distributed in embryo and neonate larvae. Lps are the major lipoprotein complex of insect hemolymph. While mammals have a complex system of lipoproteins, all lipid types in insects can be transported by this single class of lipoprotein complex (Chino et al., 1981; Beenakkers et al., 1985). Insect Lp genes are primarily expressed in the fat body and the translated proprotein is

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Fig. 4. Yolk body in developing oocytes. (A) Confocal microscope image of a section of a developing oocyte. (B) Size distribution of the yolk bodies. Diamond shape in the Box and Whisker plot represents the median value and vertical line represents the mean value of the measurements. (C) and (D) show fatty acid (green) and phospholipid (red) contents in the yolk bodies of a portion of the oocyte. (E) shows differential interference contrast (DIC) image of the same frame represented in (C) and (D).

Fig. 5. Lipid distribution in embryonic larvae. The egg was dissected and the embryonic larvae were examined with a confocal microscope 36 h after being laid. (A) DIC image of a dissected embryonic larvae. (B) Localization of TRITC-labeled Phosphatidyl ethanolamine in the larvae shown in (A). (C) Localization of BODIPY-labeled fatty acid in the larvae. (D) Composite of panels (B) and (C).

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Fig. 6. Localization and modification of imported fatty acid and phospholipids in newly hatched larvae. (A) Composite image showing distribution of phosphatidyl ethanolamine (red) and fatty acid (green) in a neonate larvae. Immediately after hatching, the larvae were examined with a confocal microscope. (B) Enlarged view of a portion of a neonate larvae, showing vesicle-like droplet of lipids at each segment of the body. (C) Localization of imported fatty acid in the same portion of the larvae shown in (B). The fatty acid appeared to be accumulated in the lipid-like vesicles seen in panel (B). (D) Modification of imported fatty acid into neutral lipids. Lipids were extracted from larvae hatched from eggs laid by mosquitoes injected with labeled lipophorins (lane T), and from uninjected mosquito (lane UT). Extracted lipids were separated by thin-layer chromatography along with the lipid mixture used to label the lipophorin before injection (lane C). Fluorescence from the lipids was visualized with a UV light. A major portion of the fatty acid (FA) from the original mixture (lane C) was converted to neutral lipid in the neonate larvae (arrow in lane T).

cleaved by a serine protease and produce different forms of the mature Lp (Sundermeyer et al., 1996; Van Heusden et al., 1998; Bogerd et al., 2000; Smolenaars et al., 2004). An. gambiae Lp also contains two apolipoproteins of molecular masses similar to those of other insect species, such as the fruit fly D. melanogaster, the mosquito Ae. aegypti, the midges C. maximus, the black fly S. vittatum and the crane fly N. abbreviata (Pennington and Wells, 2002). Like these dipteran insects, no apoLpIII was detected in the Anopheles. In M. sexta and Locusta migratoria, the apoLpIII seems to be involved in the molecular changes of Lp during insect flight (Weers and Ryan, 2003). These changes are regulated by adipokinetic hormone (Gade and Beenakkers, 1977; Shapiro and Law, 1983). The composition of the An. gambiae lipophorin is similar to M. sexta (Pattnaik et al., 1979), Rhodnius prolixus (Gondim et al., 1989), Ae. aegypti (Ford and Van Heusden, 1994) and other insects (Chino, 1985). An. gambiae Lp

contains hydrocarbon, carbohydrate and a large percentage of lipids, which together represents nearly half the mass of the Lp. The density of An. gambiae Lp, 1.135 g/ml, identifies it as a HDLp. Unlike M. sexta (Ryan et al., 1986) and L. migratoria (Chino et al., 1986), LDLp was not found in An. gambiae. A possible reason for the observed differences in the density of Lp between insects may be related to their diet. In this respect, R. prolixus also feed on blood and have HDLp (Coelho et al., 1997). Despite the similarities between An. gambiae Lp and other insect Lps, lipid composition in the Anopheles Lp seems to be somewhat different. In Ae. aegypti, triacylglycerol is the most abundant (32% of the total) lipid (Ford and Van Heusden, 1994). On the other hand, hydrocarbons are the predominant lipid class in Periplaneta americana (Chino et al., 1981) and Leptinotarsa decemlineata (Katagiri et al., 1991). Interestingly, Lp from An. gambiae also showed a high percentage of hydrocarbon and cholesteryl

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ester and a relatively low percentage of triacylglycerol (Fig. 1C). The cause and effect of such difference is not known; however, this indicates that a high triacylglycerol content may not be a general phenomenon for mosquito Lp as suggested by Ford and Van Heusden (1994). Lp picks up lipid from the insect gut and delivers it to different tissues (Canavoso et al., 2001). In most insects, lipid is stored in the fat body (Atella et al., 2000; Arrese et al., 2001; Yun et al., 2002). The location at which the maximum lipid deposition occurs may depend on the stage of the insect and its activity. In larvae , such as of M. sexta, most lipids are delivered to the fat body. In adult, lipids are translocated from the fat body to the flight muscle (Ryan and van der Horst, 2000; Canavoso et al., 2003), in a hormonally controlled process (Shapiro and Law, 1983; Van der Horst et al., 2001; Van der Horst, 2003). In An. gambiae at the peak of digestion, most vertebrate blood lipid is taken up by the ovary (Fig. 2A). Anautogenous mosquitoes require a blood meal to initiate egg development. Perhaps to meet the high demand of lipids by the activated oocytes, loaded Lp primarily deliver lipids to the eggs and excess lipids are stored in the fat body. Concurrently, the midgut also uptakes lipid from Lp. This perhaps reflects an exchange of lipids between Lp and the midgut during loading of the Lp. A small amount of lipid was also transferred to the muscles. This demonstrated that in addition to delivering lipids to the developing oocyte and fat body, Lp also serves its house-keeping function by delivering needed lipids to other tissues. Dantuma et al. (1997) demonstrated that in the fat body of L. migratoria, HDLp is resecreted after internalization and intracellular processing. This pathway known as retroendocytosis is not likely to occur in An. gambiae oocytes. During lipid transfer to An. gambiae oocytes, Lp is not recycled back to the hemolymph. The observation of a high level of FITC-labeled Lp associated with oocytes (Atella and Shahabuddin, 2002) supports this notion. The lipid accumulation occurs simultaneously with the deposition of the protein moiety inside yolk granules. This finding is not compatible with the general idea of Lp as a reusable shuttle for lipids. Bauerfeind and Komnick (1992) demonstrated endocytosis of HDLp by larval fat body cells and postulated that the function of this process could be either degradation of defective lipoproteins or lipid transport by a retroendocytic pathway. Lipoprotein-mediated lipid transfer has been studied in mammals and insects. The transfer occurs via receptormediated endocytosis (Dantuma et al., 1999; Van Hoof et al., 2003). In mammals, the LDL is released from the receptor in endosomes that mature to lysosomes and degrade the LDL (Stoorvogel et al., 1991; Innerarity, 2002). Mammalian LDL receptors are transmembrane proteins (Hussain et al., 1999). Genes for similar receptors have been cloned and sequenced from L. migratoria (Dantuma et al., 1999), Galleria mellonella (Lee et al., 2003) and Ae. aegypti (Cheon et al., 2001; Seo et al., 2003). Endocytic uptake of LP, however, has not been directly

observed in insect fat body or oocyte. Recently, Van Hoof et al. (2005) demonstrated that an endocytic pathway for Lp uptake does exist in insects by transforming human LDL receptor gene into insect cells. These authors used fixed transformed cells and antibodies to follow the Lp uptake. Here, we have provided direct evidence of endocytic uptake of Lp in live oocytes of An. gambiae. Clearly, Lp accumulated on the vitellin membrane and formed globular structures, which appeared as a punctate pattern around the oocyte. During the endocytosis of Lp, a similar pattern was observed for the LDL receptor transformed insect cells (Van Hoof et al., 2005). This observation together with temperature dependence of the process along with competition with unlabeled Lp indicated the endocytic uptake of Lp in the mosquito oocyte. Identification of the Anopheles Lp receptor and use of markers for endocytic vesicles would help to better understand the initial stage of the Lp uptake. The vesicles appear to move towards the oocyte cortex and fuse with existing yolk body over time. Size distribution of the yolk bodies showed that more yolk bodies are smaller rather than larger than the mean diameter of 1.8 mm. This indicated that yolk bodies grow until reaching an optimal size, but what determines the size of the yolk body is presently not clear. However, it can be speculated that maintaining a small yolk body would facilitate a larger total surface area compared to a single yolk. This may allow the egg to process the nutrients faster for rapid development. Processing of up-taken nutrients may occur inside the yolk body, which probably acts as a warehouse for structural components and energy for the developing embryo. Particulate round bodies were also present in the bodies of dissected embryo around 36 h (Fig. 5). Whether these are residual yolk bodies or newly formed vesicles with different chemical compositions remains to be determined. Immediately after hatching, however, vesicles with different consistency were observed. The neonate larvae vesicles along the body wall contain only the fatty acid component of the injected Lp and are completely devoid of the phospholipids. This observation is in contrast to the identification of phosopholipids in the gastric cecae and upper intestine of the neonate larvae (this study and (Atella and Shahabuddin, 2002)). Although the mechanism of such separation of the lipid types is not clear, it is evident that processing of the proteins and lipids occurs in developing eggs in the mosquito and during embryonic development after the egg is laid. Along with vitellogenin, apolipoproteins also serve as a yolk protein precursor (Sun et al., 2000). We extracted lipids from neonate larvae and separated by thin layer chromatography with an acidic solvent to examine the fate of injected fatty acid and phospholipids. Most of the fattyacid-associated fluorescence remained on the origin, suggesting that at least a large portion of the injected fatty acids converted to neutral lipid. Although the composition

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of the converted neutral lipid is unknown, this is in agreement with the lipid composition in adult mosquito (Fig. 1C) where free fatty acids are rare and most fatty acids are found as mono-, di- or triglycerides. In animals, most lipids stored in fat cells are composed primarily of triglycerides; some monoglycerides and diglycerides are mixed in, produced by incomplete esterification. In the mosquito, however, triglycerides are a minor component (Fig. 1C). Although it remains to be examined, these glycerides may release fatty acids on demand, which undergo b-oxidation and provide energy to the larval muscle. Storage of fatty acids along the body wall where the muscle for larval movement is located suggests that these are used in rapid movement of larvae after hatching. Lipid uptake from the ingested vertebrate blood and its utilization in egg development clearly impact the successful laying of eggs by malaria vector mosquitoes. Lp-mediated lipid uptake by the developing oocyte, therefore, plays an important role in the vector density in malaria endemic areas. Also, targeting of fatty acids along the body wall— which may provide the muscle needed energy for the neonate larval movement—and phospholipids in the intestine—which may serve as a emulsifying agent for the ingested food—play important role in the survival of the newly hatched larvae. Farther understanding the mechanism of vertebrate lipids metabolism in mosquitoes may lead to identifying ways to control the density of mosquito and impact the spreading of malaria. Acknowledgments We thank Drs. Luis Miller and Jose Ribeiro for support and encouragements. We are indebted to Andre Laughinghouse and Kevin Lee for maintaining the mosquito colony. Dr. Silva-Neto M.A.C. is recipient of a fellowship from Fundac- a˜o Coordenac- a˜o de Aperfeic- oamento de Pessoal de Nı´ vel Superior, CAPES, MEC, Brazil. G.C. Atella’s travel was supported by CAPES. References Arrese, E.L., Canavoso, L.E., Jouni, Z.E., Pennington, J.E., Tsuchida, K., Wells, M.A., 2001. Lipid storage and mobilization in insects: current status and future directions. Insect Biochem. Mol. Biol. 31 (1), 7–17. Atella, G.C., Gondim, K.C., Masuda, H., 1992. Transfer of phospholipids from fat body to lipophorin in Rhodnius prolixus. Arch. Insect Biochem. Physiol. 19 (2), 133–144. Atella, G.C., Arruda, M.A., Masuda, H., Gondim, K.C., 2000. Fatty acid incorporation by Rhodnius prolixus midgut. Arch. Insect Biochem. Physiol. 43 (3), 99–107. Atella, G.C., Shahabuddin, M., 2002. Differential partitioning of maternal fatty acid and phospholipid in neonate mosquito larvae. J. Exp. Biol. 205 (23), 3623–3630. Bauerfeind, R., Komnick, F.P., 1992. Immunocytochemical localization of lipophorin in the fat body of dragonfly larvae (Aesna cyanea). J. Insect Physiol. 38, 185–198. Beenakkers, A.M., Van der Horst, D.J., Van Marrewijk, W.J., 1985. Insect lipids and lipoproteins, and their role in physiological processes. Prog. Lipid Res. 24 (1), 19–67.

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