1Department of Biological Sciences and 2Renal-Electrolyte Division, Department of Medicine, University of Pittsburgh, Pittsburgh, Pennsylvania; and 3IBLS Division of Molecular Genetics, University of Glasgow, Glasgow, United Kingdom
Submitted 16 December 2004 ; accepted in final form 25 March 2005
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ABSTRACT |
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fluid homeostasis; osmosis; channel; membrane
To date, most AQPs have been found to share certain properties regarding permeability: the movement of water or other solutes is driven by a concentration or osmotic gradient, and thus energy is not required. Moreover, the activation energy (Ea) required for water transport is much less than that needed for transport through the lipid bilayer (52). However, there are clear functional differences within the AQP family, the most notable being distinctions in transport rates and solute specificity. In particular, several AQPs are permeable to glycerol (aquaglyceroporins), and some AQPs are permeable to larger polyols or other solutes (25). Still other AQPs transport cations or anions, and some may move CO2 (7). Although many AQPs have been functionally characterized, it is still not possible to predict AQP solute specificity accurately on the basis of only the primary amino acid sequence. To improve predictive algorithms, it is essential that additional AQP family members be identified and biophysically characterized.
One organism that encodes several AQPs and is amenable to genetic and developmental analysis is Drosophila melanogaster. Drosophila AQPs (DAQPs) have been proposed to maintain fluid homeostasis, which is a particularly daunting task because flies are at constant risk of dehydration as a result of their high surface area-to-volume ratio (30). Furthermore, Drosophila undergo significant morphological changes during metamorphosis, so their fluid needs change considerably. The primary organ for fluid secretion in all insects is the Malpighian tubule (MT), which is a blind-ended tubule composed of a single layer of cells. In Drosophila, the two pairs of tubules float freely in the hemolymph and attach via ureters to the gut at the midgut-hindgut border (16). A V-type proton ATPase pump in the apical membranes of principal cells drives cation movement into the lumen, probably via apical Na+/H+ and K+/H+ exchangers (12). Cl follows passively, either paracellularly, as in the mosquito Aedes aegypti (46), or transcellularly through channels in the less numerous stellate cells (31). In both Drosophila and the malaria mosquito Anopheles gambiae, Cl shunt conductance is regulated by the neuropeptide leukokinin acting through intracellular Ca2+ (37, 38). After stimulation with hormones, fluid secretion rates across the MT are as high as 6 nl/min (14).
We previously reported the cloning of the first putative DAQP, DRIP, from an adult Drosophila MT cDNA library (15). At that time, the cellular localization of DRIP had not been determined, and it was not known whether DRIP was expressed in the tubule epithelium or in associated tissue, as had been found for the Aedes aegypti putative aquaporin AeaAQP (17, 33). Since the publication of our original report, five additional putative DAQPs have been cloned (39), and, on the basis of microarray studies, it appears that two of these are also highly expressed in the adult MT (45). Herein we propose to refer to the five new putative DAQPs using their database CG numbers, replacing "CG" with "Aqp" to acknowledge that their function as an AQP has not been determined. For historical continuity, however, we refer to DRIP (Aqp9023) and Big Brain (BIB; Aqp4722) as before. Notably, of the seven DAQPs, the DRIP sequence is the most similar to hAQP4, a water-specific AQP exhibiting the highest transport rates of any AQP (6). We know now that DRIP is most closely related to putative AQPs cloned from the yellow fever carrier Aedes aegypti and the malaria carrier Anopheles gambiae, suggesting that the pore properties determined for DRIP may be relevant to its dipteran relatives (Fig. 1A). This makes a detailed characterization of DRIP of great importance and may provide one means to fight the spread of disease. To this end, we report that DRIP is expressed in the stellate cells of the MTs and that it is a water-specific AQP. We also find that DRIP is expressed at many stages during development, suggesting that this protein plays important roles throughout the organism's life cycle.
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EXPERIMENTAL PROCEDURES |
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For adult MT in situ mRNA hybridization studies, a protocol kindly provided by Dr. Edward Blumenthal (Marquette University, Milwaukee, WI) was adapted. Oregon R flies were reared on standard medium at room temperature (RT) and humidity, and 7- to 10-day-old adults were briefly anesthetized with CO2, incubated on ice, and dissected under cold 1x PBS (in mM: 175 NaCl, 1.86 NaH2PO4, and 8.41 Na2HPO4, pH 7.2). Dissected tubules with guts attached were collected in fixative [4% paraformaldehyde (PFA), 50 mM EGTA, and 1x PBS] in a glass scintillation vial. After 20 min, the vial was adjusted to RT and fixation continued for 2 h. Tubules were washed four times with 100% methanol, five times for 5 min with 100% ethanol, and fixed for 30 min in 5% PFA in PBS with 0.01% Tween 20 (PBT). The fixed material was then washed five times for 2 min each time in PBT and treated with proteinase K (Roche Diagnostics) at a final concentration of 5.7 µg/ml in PBT for 10 min at RT. Five additional washes in PBT on ice were followed by 30-min postfixation in 5% PFA in PBT. After five 5-min washes in PBT, the tissue was incubated at 57°C overnight in hybridization solution {50% formamide, 5x SSC (0.75 M NaCl and 0.075 M Na3 citrate), 1x Denhardt's solution (0.02% Ficoll 400; 0.02% polyvinylpyrrolidone; 0.02% bovine serum albumin), 0.1% Triton X-100, 0.1% 3-([3-cholamidopropyl]dimethylammonio)-1-propanesulfonate (CHAPS), 1 mg/ml transfer RNA, 5 mM EDTA, and 50 µg/ml heparin}. The next day, riboprobe at a final concentration of 2.5 µg/ml was denatured for 15 min at 8085°C in hybridization solution and incubated with MTs overnight at 57°C. Tubules were washed three times for 20 min each (2x SSC, 0.1% CHAPS, and 50% formamide) at 57°C, three times for 20 min each (0.2x SSC, 0.1% CHAPS, and 50% formamide) at 57°C, and two times for 10 min each in KTBT (50 mM Tris, pH 7.5, 150 mM NaCl, 10 mM KCl, and 1% Triton X-100) at RT. After being blocked for nonspecific binding in 20% normal goat serum in KTBT (KTBTN) for 2.5 h at 4°C, tubules were incubated with anti-digoxigenin antibody (Roche Diagnostics) diluted 1:2,000 in KTBTN overnight at 4°C. The labeled tubules were then washed five times for 10 min each in KTBT at RT followed by two 10-min washes in NTMT (100 mM Tris, pH 9.5, 50 mM MgCl2, 100 mM NaCl, 0.1% Triton X-100, and 1 mM levamisole). Finally, labeling was visualized using 4.5 ng/ml 4-nitro blue tetrazolium chloride and 1.75 ng/ml 5-bromo-4-chloro-3-indoyl-phosphate in NTMT.
Xenopus plasmid construction and sense RNA preparation. DRIP was amplified by performing PCR from RE60324 (see above) using primers (5') CCTGAATTCATGGTCGAGAAAACAG AAATGTCG and (3') GTCCTCGAGTTAGAAGTCGTACGAGTCGG cloned into the Xenopus expression vector pXT7 between the EcoRI and XhoI sites, and the sequence of the insert was confirmed using automated sequencing (DNA Sequencing Facility, University of Pittsburgh, Pittsburgh, PA). The resulting plasmid was linearized using XbaI, and the gene encoding human AQP1 in a Xenopus expression vector (American Type Culture Collection, Manassas, VA) was linearized using SmaI. mMessage mMachine (Ambion, Austin, TX) was used to generate methyl-G-capped sense RNA using T7 RNA polymerase for DRIP and T3 RNA polymerase for AQP1. After DNase1 digestion, the RNA was precipitated, phenol-chloroform was extracted, and the RNA was resuspended in RNase-free water (Ambion). The final concentrations of the message were 10 ng/50 nl DRIP and 1 ng/50 nl AQP1. RNA size and integrity were checked on a denaturing formaldehyde agarose gel according to standard protocols.
Xenopus oocyte injection and swelling assay. Oocytes were collected and treated with collagenase. Stage 6 oocytes were sorted and allowed to recover overnight at 18°C before 50 nl of DRIP and AQP1 sense RNA were injected into them. Injected oocytes were incubated for 3 days at 18°C with daily changes of 1x modified Barth's saline [MBS; in mM: 88 NaCl, 1 KCl, 2.4 NaHCO3, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.42 CaCl2, and 10 HEPES, pH 7.9]. To assay for water transport, oocytes were digitally photographed using Simple PCI image-capturing software (Compix), with images captured every 5 s. After 2 min, the buffer was exchanged with a perfusion apparatus to x MBS. Two-dimensional oocyte images were converted to black-and-white binary images, and oocyte areas were measured using NIH ImageJ software (http://rsb.info.nih.gov/ij/). Oocyte areas were plotted against time, and the swelling rate was determined on the basis of a linear curve fit.
Yeast strain and plasmid construction.
DRIP was amplified from RE60324, and eight histidines were appended at the COOH terminus using PCR primers (5') CCTGAATTCATG GTCGAGAAAACAGAAATGTCG and (3') GTCCTCGAGCTAATGATGATGATGA TGATGATGATGGAAGTCGTACGAGTC. The PCR product was cloned into the EcoRI and XhoI sites in the yeast galactose-regulated expression vector pYES2 (Invitrogen). The insert was verified by performing DNA sequence analysis, and the expression vector either containing or lacking the insert was transformed into a sec6 temperature-sensitive Saccharomyces cerevisiae mutant strain SY1 (MAT, ura352, leu23,112, his4619, sec64, GAL+) (28, 35) using a standard lithium acetate technique. Transformants were selected on synthetic complete medium lacking uracil (SC-ura) but supplemented with glucose to a final concentration of 2%. A frozen glycerol stock was made from a single colony and used for all subsequent studies.
To check DRIP expression, a single colony of yeast transformed with either vector lacking insert or containing the DRIP-His-tagged construct was used to inoculate 50 ml of SC-ura medium containing 2% raffinose (wt/vol). A total of 10 ml of this culture was used to inoculate 2 L of SC-ura containing 2% galactose. After 20 h at RT, the yeast were collected using centrifugation and washed in 0.7 M sorbitol (4,400 OD600 of cells). After recentrifugation, cells were resuspended in
1 ml of 0.7 M sorbitol and frozen at 80°C. Cell wall digestion was performed in the same manner used for vesicle preparation (see below), and spheroplasts were resuspended in lysis buffer (LB; 0.8 M sorbitol, 10 mM triethanolamine, 1 mM EDTA, 0.25 mM phenylmethylsulfonyl fluoride, 1 µg/ml pepstatin A, and 1 µg/ml leupeptin, pH 7.4) at 1 ml/400 OD600 and lysed by vigorous agitation with a one-half volume of glass beads four times for 1 min each time. After centrifugation at 5,000 rpm for 5 min (SS34; Sorvall), the supernatant was removed and spun at 15,000 rpm in the same rotor for 20 min. The membrane pellet was resuspended in
1 ml of LB and refrozen at 80°C. After being allowed to thaw, the lysate was spun again and the pellet was resuspended in 3 ml of solubilization buffer (SB; 50 mM Tris, pH 8.0, 300 mM NaCl, 20% glycerol, 2 mM
-mercaptoethanol, 1.5% octyl glucoside, 0.25 mM phenylmethylsulfonyl fluoride, 1 µg/ml pepstatin A, and 1 µg/ml leupeptin). After 1-h incubation at 4°C, the solubilized protein extract was incubated with 1 ml of a 1:1 slurry of nickel nitrilotriacetate agarose beads (Qiagen) containing 5 mM imidazole and was rotated overnight at 4°C. The resin was used to form a column and was washed with 15 ml of SB containing 5 mM imidazole, 5 ml of SB containing 50 mM imidazole, and then 5 ml of SB containing 150 mM imidazole. The bound protein was eluted with SB containing 500 mM imidazole and visualized using silver staining after electrophoresis was performed on a 15% SDS-polyacrylamide gel. Alternatively, the resolved protein was transferred to nitrocellulose membranes, and consistent protein loading was verified using Ponceau S staining. Nonspecific antibody binding sites were blocked for 1 h at RT with 2% bovine serum albumin fraction V (A-7888; Sigma) in TBST (50 mM Tris, pH 7.4, 150 mM NaCl, and 2% Tween 20). The blots were then incubated with anti-pentahistidine antibody (Qiagen) at a 1:5,000 dilution overnight at 4°C. After being washed in TBST and incubated with horseradish peroxidase-conjugated sheep anti-mouse antibody (Amersham) at a 1:5,000 dilution, bound antibody was visualized using SuperSignal West Pico enhanced chemiluminescence (Pierce) on a Kodak 440 CF Image Station.
Yeast vesicle preparation.
With the use of a method similar to one previously published (9), a single colony of SY1 yeast containing either the vector control or the DRIP expression vector was used to inoculate 2550 ml of SC-ura medium containing 2% raffinose. This culture was diluted into 1 L of identical medium at an initial OD600 of 0.025 and the culture was grown overnight using shaking at RT to an OD600 of
0.5. Yeast were collected using centrifugation at 5,000 rpm in a GS3 rotor (PTI) and resuspended in 2 L of yeast extractpeptone (YP) medium (1% yeast extract and 2% bactopeptone) containing 2% galactose at an initial OD600 of 0.25 to induce DRIP expression. After 23 h at RT, cultures were incubated at 37°C for 23 h to induce secretory vesicle accumulation. The cells were harvested by performing centrifugation and washed twice with ice-cold 0.7 M sorbitol before the pellet was resuspended in 1 ml of 0.7 M sorbitol and frozen at 80°C. Vesicles were prepared as described previously (8), except that the vesicles were incubated in 20 mM 5,6-carboxyfluorescein (CF; Molecular Probes).
Measurement of water transport in sec vesicles.
Unincorporated, extravesicular CF was quenched using anti-CF antibody (Molecular Probes) after titrating the amount of anti-CF that failed to further reduce fluorescence at 520 nm on an Aminco Bowman series 2 spectrometer (SLM Aminco). To measure the rate of water transport in a SF.17 MV Applied Photophysics stopped-flow device, vesicles were mixed rapidly with a 2.4 M sorbitol solution, approximately doubling the extravesicular osmolality. CF self-quenching was measured (490 nm excitation/520 nm emission) as vesicles shrank, and the rate of fluorescence change was determined from the curve fit using a nonlinear regression algorithm and Applied Photophysics software. The rate of fluorescence change is directly proportionate to the vesicle size change at this concentration of CF. Osmolality was measured using freezing point depression in an osmometer (Osmette A; Precision Systems), and vesicle size was determined using dynamic light scattering (LSR; DynaPro). To calculate osmotic permeability, fluorescence change was fit to a double exponential algorithm and the first obtained rate was used in the following formula:
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To verify DRIP-His protein expression in vesicles after water transport assays, 12 µl of vesicles were mixed with 3 µl of 5x SDS sample buffer (5% -mercaptoethanol, 10% SDS, 25 mg/ml bromophenol blue, 325 mM Tris, pH 6.8, and 50% glycerol) and incubated at 6575°C for 10 min before electrophoresis was performed on a 15% SDS-polyacrylamide gel. Immunoblots were obtained as described above using anti-pentahistidine antibody (Qiagen).
Measurements of transport of other solutes in sec vesicles. For glycerol and urea permeability measurements, vesicles were incubated for 30 min on ice in 0.8 M glycerol or urea in LB. The vesicles were mixed rapidly with isosmotic 0.4 M NaCl in LB in the stopped flow (see above). The vesicle shrinkage rate was determined as described above and as described previously elsewhere (9). For proton and ammonia permeability measurements, vesicles were prepared in 2 mM CF, a concentration at which fluorescence is pH sensitive. To measure the rate of proton movement, vesicles in LB at pH 7.4 were rapidly mixed in the stopped flow with an equal volume of isosmotic LB at pH 5.0. Under these conditions, protons entering the vesicles quenched the fluorescence. The buffering capacity of the vesicles was established by measuring the fluorescence change upon addition of 10 mM sodium acetate. The relationship between pH and fluorescence was established by titrating LB with HCl on a pH meter and vesicles on the luminescence spectrometer. A similar method was used to determine NH3 permeability, with the following changes. Vesicles were equilibrated for 30 min on ice in LB at pH 6.8 before analysis with LB at pH 6.8 and supplemented with 40 mM NH4Cl, such that NH3 entering the vesicles increased CF fluorescence. Water transport was measured in all yeast preparations used for solute studies.
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RESULTS |
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One key to understanding DRIP function is to determine the site of DRIP expression. Furthermore, very little is known about AQP expression during development in most organisms. Because the developmental program has been studied extensively in Drosophila melanogaster embryos, it is an ideal system in which to search for evidence of developmental roles for DRIP. To this end, we performed mRNA hybridization studies using a digoxigenin-labeled DRIP riboprobe. As controls, antisense riboprobes were made for four other putative DAQPs (Aqp17664 Aqp7777, Aqp4019, and Aqp5398) and hybridized to embryos at the same developmental stage (stage 17). As shown in Fig. 2A, we found four distinct staining patterns using the Aqp17664(GH16993), Aqp7777 (RE34617), Aqp4019 (RE51883), and DRIP riboprobes, whereas no embryonic expression was observed for Aqp5398 (data not shown). Of interest, Aqp7777 (RE34617) appeared to be expressed strongly in the brain and in the segmental ganglia, Aqp17664was expressed in the salivary glands and a section of the gut, and Aqp4019 was expressed in the body wall and visceral muscles. These data demonstrate that the DRIP riboprobe is specific and can be used to assess the developmental pattern of DRIP expression.
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The localization of DRIP in gut regions immediately before larval eclosion (stage 17) also suggests a role for DRIP in the adult. The insect gut is known to be involved in osmoregulatory water reabsorption, although traditionally most reabsorption occurs in the hindgut (11, 47). Water transport in the midgut may also aid in nutrient uptake, while water may be secreted in the foregut to help lubricate food passage. Finally, late expression of DRIP in the posterior spiracles is also consistent with an osmoregulatory role for DRIP. The posterior spiracles sit at the opening to the trachea, where gases, including water vapor, are exchanged. By regulating the opening and closing of the spiracles, the insect can balance water loss with respiration (27).
Because mammalian AQPs regulate fluid homeostasis in the kidney (25), we examined DRIP expression in the embryonic and adult MTs. Indeed, DRIP was expressed in the MTs of late stage 17 embryos that are about to eclose as first instar larvae (Fig. 3A). We also found DRIP expression in the adult MTs, although expression is clearly restricted to stellate cells (Fig. 3B), including the bar-shaped stellate cells in the distal MT (data not shown). This expression pattern is different from the pattern exhibited by an Aedes aegypti DRIP homolog; in this organism, the message resides in the tracheoles that attach to the MTs but is not found in the MTs themselves (17, 33). In contrast, our data suggest that DRIP plays a role in fluid secretion and osmotic balance akin to that exhibited by the kidney.
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We first established that the His-tagged DRIP protein was expressed in yeast by examining the protein bound to Ni affinity columns loaded with detergent-solubilized membranes. As shown in Fig. 6A , a diffuse band migrating at 29 kDa eluted from the column with 500 mM imidazole only when lysates from yeast containing the DRIP expression vector were examined. Slower migrating bands were also observed, suggesting that DRIP, like other AQPs (40), forms higher-order oligomers; however, we cannot rule out the possibility that the His tag may have played a role in DRIP oligomerization.
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The temperature dependence of Pf can be used to determine the Ea for water movement and to provide evidence for channel-mediated transport. A hallmark of protein water channels is their low Ea for water movement (2). We therefore examined the Pf as a function of temperature and obtained the Arrhenius plot in Fig. 7, which shows that DRIP-containing sec vesicles have an Ea of 4.9 kcal/mol for water transport, whereas vesicles lacking DRIP have an Ea of 16.4 kcal/mol. These data provide further evidence that DRIP functions as a water channel.
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Because many AQP superfamily members (e.g., aquaglyceroporins AQP3, AQP7, AQP9) transport small solutes (23, 24, 43), we investigated whether DRIP facilitates glycerol, urea, and ammonia transport. After equilibrating vesicles in glycerol- or urea-containing buffer, the rate of solute efflux was followed by rapid mixing in isosmotic buffer devoid of the solute (see EXPERIMENTAL PROCEDURES). Under these conditions, we found that the rate of glycerol and urea efflux is the same in secretory vesicles either containing or lacking DRIP (Fig. 8, B and C). Furthermore, we determined that DRIP-containing vesicles conduct ammonia at rates similar to those of vesicles from yeast transformed with a vector lacking the DRIP insert (data not shown). For all solutes tested, the same vesicle preparations showed osmotic permeability differences as high as 25-fold between DRIP and controls (Fig. 8A). From these collective data, we conclude that DRIP is not permeable to glycerol, urea, ammonia, or protons and is a water-selective AQP.
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DISCUSSION |
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DRIP is expressed at very early stages during Drosophila embryogenesis, and, surprisingly, the pattern of expression was quite dynamic throughout development. Importantly, the DRIP message is highest in several organs in which water transport is likely to play an important role: pharynx, gut, and posterior spiracles. DRIP was also expressed in adult MTs, which filter the hemolymph and thus function like the kidney in higher organisms. Consistent with these data, microarray analysis indicated that DRIP mRNA is enriched 3.6-fold in adult MTs relative to the adult fly (45). Specifically, in the present study, we found that MT expression was restricted to the stellate cells. Because the stellate cells compose only 20% of the tubule main segment, DRIP protein is likely to be highly enriched in this cell type. Together, these data suggest that DRIP is critical for fluid homeostasis in both the developing embryo and in adult flies, a hypothesis that will be tested in future studies.
One motivation for this study was to discern how water moves across the MT. The path followed by osmotically driven water into the MT lumen has not been determined; whether water moves trans- or paracellularly has aroused considerable debate (5, 16, 29, 31, 32, 42). To drive fluid secretion, a proton gradient is established by a V-ATPase in principal cells. Protons move back into the cell in exchange for K+/Na+, and Cl moves through the stellate cells into the tubule lumen, which in turn drives water uptake (13). Because we observed DRIP expression within stellate cells of the adult MT (Figs. 3 and 4), we suggest that water moves transcellularly through DRIP. This model is consistent with the finding in Drosophila that leukokinin, which stimulates Cl transport in stellate cells, also stimulates fluid transport (31) and with the finding that slower fluid transport is observed across MTs in a Drosophila mutant that expresses fewer stellate cells (10). However, our data do not rule out the possibility of paracellular transport through septate junctions associated with stellate cells. Specifically, one other AQP, AQP0, has been shown to reside at cell-cell junctions in the lens of the mammalian eye (21). If DRIP similarly resides only at the junctions between stellate cells, it could mediate paracellular water transport (although we note that AQP0 in the junctions appears to be closed; see Ref. 21). In addition, water may move through the principal cells. We did not detect Aqp4019 or Aqp17664in the stellate cells, but message corresponding to both is clearly abundant in the principal cells of the adult MT main segment. Although the principal cells have not been implicated in directly facilitating water movement, the MTs move fluid faster than any other epithelium on a per cell basis (15, 16), and it would be an efficient use of surface area to employ these cells as well. Future studies will no doubt resolve this issue.
Because much still is not understood about the selectivity and mechanism of solute transport through AQPs, it is important to characterize AQPs biophysically from a diverse number of species and tissues. Only by performing such an analysis can improved algorithms be developed to predict AQP substrate specificity. Nevertheless, on the basis of sequence alignments comparing water-specific and solute-transporting AQP family members, it was proposed that particular residues could be used to predict specificity (20). Accordingly, it was suggested that water specificity may be predicted on the basis of the presence of two small, uncharged residues just after the NPA in the second hemitransmembrane domain (corresponding to positions 196 and 200 in DRIP) (Fig. 1B) (20). In addition, two residues (positions 212 and 213 in DRIP) in transmembrane domain 6 are aromatic in the water-specific AQPs, but a proline and a nonaromatic residue occupy these positions in the solute transporters (20). In support of this hypothesis, DRIP contains residues consistent with its being a water-specific channel (S196, A200, Y212, and W213). However, the analogous residues in the Drosophila AQP-like protein BIB (S242, S246, Y258, and W259) also predict a water-specific channel; yet BIB instead appears to be a monovalent cation channel (50, 51). These data emphasize limitations in existing predictions of AQP specificity, but like many water-specific family members, DRIP lacks the inserted amino acids in the C loop frequently found in the aquaglyceroporins (19). They are found, however, in Aqp4019 and Aqp17664(Fig. 1B). Thus it will be interesting in the future to determine the transport properties of Aqp4019 and Aqp17664and relate these characteristics to their principal cell expression.
Our results may also be applicable to an examination of AQPs in other Diptera, such as the malaria vector Anopheles gambiae and the yellow fever vector Aedes aegypti. An amino acid sequence alignment with these mosquito species indicates that DRIP is 64% identical to a predicted protein in Anopheles (EnSANGP00000016718; Anopheles Genome Sequencing Consortium) and 65% identical to Aedes AQP (AeaAQP), which has been characterized as a water-specific AQP (17). If AQPs control fluid homeostasis in dipteran insects, they could provide a specific molecular target to block the spread of mosquito-borne diseases. In adult flies and mosquitoes, tight regulation of water transport systems is essential because extra fluid carried in flight is energetically costly, yet the high surface-to-volume ratio puts the insect at risk of rapid desiccation (5). Moreover, the fluid volumes of Diptera change considerably at metamorphosis, and it is likely that AQPs play a role in this process. For example, before becoming airborne, Drosophila larvae survive in the moist environment of rotting fruit and shed waste as a green fluid called the meconium. Similarly, mosquitoes must emerge from their larval lives in water before commencing flight (5). Thus maintenance of fluid homeostasis throughout the life cycle presents multiple targets for insect population control. On the basis of the results of our study in Drosophila, we can now take advantage of this genetically tractable organism to address the roles of DRIP function in development and of fluid secretion in the adult.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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