Developmental expression and biophysical characterization of a Drosophila melanogaster aquaporin

Nancy Kaufmann,1 John C. Mathai,2 Warren G. Hill,2 Julian A. T. Dow,3 Mark L. Zeidel,2 and Jeffrey L. Brodsky1

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


    ABSTRACT
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Aquaporins (AQPs) accelerate the movement of water and other solutes across biological membranes, yet the molecular mechanisms of each AQP's transport function and the diverse physiological roles played by AQP family members are still being defined. We therefore have characterized an AQP in a model organism, Drosophila melanogaster, which is amenable to genetic manipulation and developmental analysis. To study the mechanism of Drosophila Malpighian tubule (MT)-facilitated water transport, we identified seven putative AQPs in the Drosophila genome and found that one of these, previously named DRIP, has the greatest sequence similarity to those vertebrate AQPs that exhibit the highest rates of water transport. In situ mRNA analyses showed that DRIP is expressed in both embryonic and adult MTs, as well as in other tissues in which fluid transport is essential. In addition, the pattern of DRIP expression was dynamic. To define DRIP-mediated water transport, the protein was expressed in Xenopus oocytes and in yeast secretory vesicles, and we found that significantly elevated rates of water transport correlated with DRIP expression. Moreover, the activation energy required for water transport in DRIP-expressing secretory vesicles was 4.9 kcal/mol. This low value is characteristic of AQP-mediated water transport, whereas the value in control vesicles was 16.4 kcal/mol. In contrast, glycerol, urea, ammonia, and proton transport were unaffected by DRIP expression, suggesting that DRIP is a highly selective water-specific channel. This result is consistent with the homology between DRIP and mammalian water-specific AQPs. Together, these data establish Drosophila as a new model system with which to investigate AQP function.

fluid homeostasis; osmosis; channel; membrane


IN ORDER FOR ORGANISMS to maintain fluid homeostasis between themselves and the external environment, they must be able to control water transport. Members of the aquaporin (AQP) family of water channels facilitate water transport and appear to play an essential role in regulating the flow of water between compartments and between the organism and its environment. For example, after hormonal signaling of dehydration, AQP2 levels in the apical membrane of kidney collecting duct cells rise, allowing more water to move into the cells and then out to the interstitium (via AQP3 and AQP4); a defect in AQP2 results in diabetes insipidus, which is characterized by a failure to concentrate urine (25). In contrast to the known function of AQP2, why seven different AQP family members need to be expressed in this organ is still not understood. In one hypothetical model, differences in substrate specificity explain the breadth of AQPs expressed in the kidney. It is also possible that AQP levels or localizations change during development and thus can be adjusted. It appears highly likely that many of the physiological roles of AQPs have yet to be discovered.

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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Fig. 1. DRIP shares sequence similarity with mammalian and insect aquaporins (AQPs). A: amino acid sequence alignment of DRIP with human AQP1 and AQP4, with Aedes aegypti AQP (AeaAQP), and with an incomplete but predicted AQP from Anopheles gambiae. The two Asn-Pro-Ala (NPA) motifs, a hallmark of many AQPs, are boxed. B: amino acid sequence alignment of human AQP4 and the seven Drosophila melanogaster putative AQPs. NH2- and COOH-terminal regions are not included. C and E loop regions are boxed, as are the NPA motifs. The additional length of the C loops in Aqp17664 Aqp17662 Aqp4019, and Aqp5398 may suggest aquaglyceroporin function (29). However, no pattern in the genomic structures among the Drosophila AQPs (DAQPs) was noted (http://flybase.bio.indiana.edu/cgi-bin/gbrowse_fb/dmel). C: Kyte-Doolittle hydropathy plot indicates six hydrophobic amino acid motifs of sufficient length in DRIP (closed bars) to form transmembrane domains. For comparison, the same plot is shown for AQP1.

 

    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
mRNA in situ hybridization. Antisense riboprobes were generated using the following cDNAs obtained from the Berkeley Drosophila Genome Project: RE60324, (DRIP), GH16993 (Aqp17664, RE34617 (Aqp7777), GH26134 (Aqp5398), and RE51883 (Aqp4019). After the indicated restriction enzymes were used to linearize the cDNA (XhoI: RE60324, RE34616; EcoRI: GH16993, RE51883; and BglII: GH26134), T3 (RE60324, RE51883, RE34617) or SP6 (GH16993, GH26134) RNA polymerase was added to generate digoxigenin-labeled probes using digoxigenin-11-UTP RNA labeling mix (Roche) as described previously (44). Protector RNase Inhibitor (Roche Diagnostics) was also included at a final concentration of 40 U/µl. Embryos from 20-h egg lays (25°C) of strain W118 adults were prepared, hybridized, developed, and staged as described previously (22, 44).

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 80–85°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{alpha}, ura3–52, leu2–3,112, his4–619, sec6–4, 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 {beta}-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 25–50 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 2–3 h at RT, cultures were incubated at 37°C for 2–3 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:

to determine the osmotic permeability coefficient (Pf). SAV refers to the surface area-to-volume ratio of the vesicle; Cin is the concentration inside the vesicle; Cout is the concentration outside the vesicle; V(t) is the relative volume of vesicle at time t (V and time t divided by V at time 0), and dV/dt is the ratio of the derivative V(t) to the derivative of time. The second rate was similar to control vesicles lacking DRIP. MATHCAD software (MathSoft) was used to perform the calculations as previously described (8). As a control, rapid mixing of vesicles with isotonic buffer showed no fluorescence change. For Ea calculations, the temperature of the water bath in the stopped-flow chamber was varied from 5 to 25°C and vesicles were allowed to adjust to the desired temperature for 5 min before permeability measurements were obtained.

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% {beta}-mercaptoethanol, 10% SDS, 25 mg/ml bromophenol blue, 325 mM Tris, pH 6.8, and 50% glycerol) and incubated at 65–75°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.


    RESULTS
 TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Dynamic expression of the Drosophila melanogaster AQP homolog, DRIP, during development. AQPs are traditionally classified into one of two groups on the basis of their transport properties: water-specific channels and aquaglyceroporins, which support the movement of small solutes such as glycerol and often water. A search of predicted AQPs in the Drosophila melanogaster proteome reveals seven putative homologs (1, 41) (Fig. 1B). Of these, DRIP is the most similar to mammalian water-specific AQPs [BLAST E values compared with hAQP1: DRIP = 5 x 10–50 vs. Aqp5398 = 8 x 10–9 (3)], and thus we chose to characterize it further. A Kyte-Doolittle hydrophobicity plot (26) (Fig. 1C) suggested that DRIP, like established AQP family members, contains six transmembrane domains (indicated by closed bars). Established AQP family members also contain two hemitransmembrane domains, which form much of the AQP pore (2), and two embedded Asn-Pro-Ala (NPA) motifs (boxed areas in Fig. 1, A and B). Notably, the alignment presented in Fig. 1B predicts that DRIP will contain two hemitransmembrane domains (amino acid residues 76–88 and 192–203) as well as the NPA motif. Only three of the other six DAQPs (Aqp4019, BIB, and Aqp7777) encode both NPAs (Fig. 1B).

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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Fig. 2. DRIP is expressed dynamically in developing Drosophila melanogaster embryos. A: DRIP riboprobe is specific. mRNA in situ analyses were performed with stage 17 embryos labeled with riboprobes specific for DRIP (RE60324), Aqp17664(GH16993), Aqp7777 (RE34617), and Aqp4019 (RE51883). Unique expression patterns for each riboprobe were observed. The embryos are oriented with the anterior portion to the left. All views are from the side, except a dorsal view is shown for RE51883 because otherwise total body wall muscle staining in the side view obscures the staining pattern (data not shown). B: DRIP is expressed throughout embryonic development. DRIP expression begins in the developing primordial gut (stage 8) and later (stage 14) demarcates the foregut-midgut border (closed arrowhead), where the proventriculus will form, and the midgut-hindgut border, where the Malpighian tubules (MTs) will connect. DRIP mRNA is also found in the developing pharynx (open arrowhead). In addition, DRIP mRNA localizes to the developing posterior spiracles (right box in stage 17, bottom image). This experiment was repeated three times with similar results.

 
Next, the DRIP riboprobe was hybridized to embryos from 20-h egg lay collections, allowing all stages during embryogenesis to be visualized (Fig. 2B). At the earliest stage examined, the blastoderm (stage 5) (22), we found DRIP mRNA localized under the newly formed embryonic cells, which provides evidence of maternal contribution in the oocyte. In contrast, DRIP was observed at stage 8 in the amnioproctodeal invagination, a tissue that is a combination of midgut and hindgut primordial (4). In stages 14 and 15 embryos, DRIP mRNA resides in cells that form the pharynx and in cells at the foregut-midgut boundary: A hint of future gut expression can even be traced back to the stage 12 embryo. In stage 16, and particularly in stage 17 embryos, these DRIP-expressing gut regions form the proventriculus at the foregut-hindgut border and a region of posterior midgut (also note DRIP expression in the posterior regions in Fig. 2A). At stage 17, a subset of cells in the posterior spiracles is also labeled (Fig. 2B, stage 17, inset). The formation of these structures, which regulate gas access to the trachea (see below), can be traced back to stage 12 embryos. Overall, the dynamic expression of DRIP observed throughout embryogenesis is consistent with an important role for this DAQP during the development of specific Drosophila organ systems.

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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Fig. 3. DRIP is expressed in the MTs. A: DRIP mRNA in situ of late stage 17 embryo shows DRIP expression in the MT. The elongated stage 17 embryo (compare with stage 17 in Fig. 2B) will soon hatch as a first instar larva. B: DRIP mRNA in situ of a dissected adult MT. Note that DRIP expression is restricted to the stellate cells. Four experiments were performed with similar results.

 
To explore whether other DAQPs that may reside in the MT are also present in the stellate cells, we performed in situ studies with three other DAQPs. On the basis of microarray analysis, two of these (Aqp4019 and Aqp 17664), like DRIP, are enriched in the MT (compared with whole adult). Aqp5398 is absent from the MT and was chosen as a negative control (45). Consistent with the microarray data, both Aqp17664and Aqp4019 showed strong expression in the MT, while Aqp5398 was not detected (Fig. 4, AD). In contrast to DRIP, Aqp4019 and AQP17664clearly localized to the principal cells and we were unable to discern staining in the stellate cells. If Aqp4019 and Aqp17664are in the stellate cells, they are clearly present at significantly lower levels than DRIP.



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Fig. 4. Distinct expression patterns of DAQPs in the MT. A: DRIP mRNA in the adult MT, shown by in situ analysis, is restricted to the stellate cells. Staining is absent in the adjacent principal cells. B: Aqp4019 is expressed in the principal cells as determined by mRNA in situ. Two attached tubules are shown. Experimental and photographic conditions are identical to those in A. C: Aqp17664mRNA in situ shows expression restricted to the principal cells of the main segment of the tubule. D: Aqp5398 is absent from both the stellate and principal cells as shown using in situ analysis. Tubules treated identically but without probe also show absence of staining (data not shown).

 
DRIP facilitates water transport across both the Xenopus oocyte plasma membrane and yeast secretory vesicle membranes. To examine the solute specificity of DRIP, we expressed DRIP in Xenopus oocytes because water channel expression accelerates oocyte swelling in hyposmotic saline (36). We first showed that oocytes injected with mRNA encoding the mammalian water channel, AQP1, swelled more rapidly than controls after incubation in hyposmotic saline (Fig. 5; compare rates for hAQP1 and control in the three oocytes shown). Under the same conditions, oocytes injected with DRIP mRNA also swelled quickly and leaked their yolk proteins (data not shown) as previously reported for hAQP1-injected oocytes (36).



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Fig. 5. In hypotonic solution, Xenopus oocytes injected with mRNA encoding DRIP swell faster than control oocytes. Each bar represents the initial swelling rate from a single experiment in which an individual oocyte was injected with the same amount of a methyl-G-capped sense RNA (human AQP1 or DRIP) or water (negative control).

 
To confirm these results and to further characterize DRIP water and solute permeabilities, we used a yeast AQP expression system studied previously to examine the pore specificities of human AQP1 and AQP2 (8). DRIP cDNA containing a COOH-terminal His tag was cloned into the yeast galactose-inducible expression vector pYES2, and both the DRIP-containing vector and the expression vector lacking the insert were transformed into a sec6 S. cerevisiae strain (see EXPERIMENTAL PROCEDURES). This strain accumulates late secretory vesicles of uniform size at the restrictive temperature of 37°C (35). AQP-containing secretory vesicles can then be loaded with the concentration-specific probe, CF, and isolated. Under hyperosmotic conditions, the change in CF fluorescence is linearly related to the change in volume, allowing for measurement of Pf (52).

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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Fig. 6. DRIP-expressing vesicles exhibit elevated water permeability (Pf). A: expression of DRIP in yeast. Protein lysates from yeast containing either a DRIP-His-tagged expression vector or the vector lacking an insert were chromatographed on a nickel nitrilotriacetate agarose bead (Qiagen) column, bound protein was eluted with imidazole, and the His-tagged protein was detected using immunoblot analysis as described in EXPERIMENTAL PROCEDURES. The band at ~29 kDa represents the DRIP monomer, and the higher-molecular-weight band (asterisks) may be a DRIP multimer. B: DRIP increases water permeability across yeast secretory (sec) vesicle membranes. A stopped-flow trace of secretory vesicles either containing or lacking DRIP is shown after rapid mixing of each vesicle population in hypertonic solution at 7°C. DRIP-expressing vesicles shrink faster than control vesicles, which results in self-quenching of 5,6-carboxyfluorescein (CF). In this experiment, Pf (DRIP) = 0.014 cm/s and Pf (Control) = 0.0008 cm/s. The average Pf relative difference (~15-fold) is shown in the inset. Error bars represent SD.

 
Next, we isolated secretory vesicles from DRIP-expressing and control yeast after growth in galactose, loaded the vesicles with CF, and obtained water permeability measurements (see EXPERIMENTAL PROCEDURES). Figure 6B shows typical stopped-flow traces for vesicles containing or lacking DRIP after they were mixed rapidly into a hyperosmotic solution. The rate of shrinkage is clearly faster for the DRIP-containing vesicles, indicating that DRIP increases water permeability across the sec vesicle membrane. The average Pf (7°C) for DRIP-containing vesicles was ~10-fold higher than that for control vesicles (DRIP = 0.0148 cm/s, control = 0.0014 cm/s; n = 5) (Fig. 6B, inset). This value is nearly identical to the difference (10-fold) reported when the Aedes DRIP homolog was expressed in Xenopus oocytes and compared with controls, but it is significantly higher than the twofold difference reported when the Pf of the Rhodnius prolixus major intrinsic protein (Rp-MIP) was assessed (17, 18).

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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Fig. 7. Activation energy (Ea) of water transport decreases in DRIP-expressing vesicles. An Arrhenius plot of the natural log of the osmotic permeabilities of DRIP and control vesicles was determined from stopped-flow measurements at five different temperatures (7.2, 11.4, 15.0, 18.0, and 22.8°C) using curves fitted to single exponentials. Closed squares indicate data obtained for DRIP-containing vesicles, whereas open squares represent data obtained with vesicles lacking the DAQP. Data were fit using Microsoft Excel software [DRIP: y = –2.5073x + 12.245 (R2 = 0.9082); Control: y = –8.2272x + 30.989 (R2 = 0.9986)], and Ea was determined by calculating {kappa} = Ae–Ea/RT. This result was confirmed in a second experiment.

 
DRIP does not facilitate proton, glycerol, urea, or ammonia transport. Having demonstrated that DRIP functions as a water channel, we next investigated whether the path of water movement through the pore is similar to that of other AQP family members. In the AQP channel, H+ bonds between adjacent water molecules are interrupted and proton transport is not supported (25). In contrast, the gramicidin channel conducts an unbroken chain of water molecules through its pore and thus also transports protons (34). The mechanism by which the water chain in the AQP channel is broken is still disputed, but the NPA asparagines in the center of the pore likely play a key role in this process, as may a conserved arginine. To determine how water moves through the DRIP pore, we measured the proton permeabilities of DRIP-expressing and control vesicles. We observed identical rates of proton transport, even though a difference in osmotic permeability was demonstrated in the DRIP-expressing vesicles (data not shown). This finding suggests that water moves through DRIP by a mechanism similar to that used by other AQPs and is consistent with the high degree of identity in the predicted pore region in DRIP (see DISCUSSION).

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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Fig. 8. DRIP does not increase glycerol or urea transport. Stopped-flow traces of secretory vesicles either containing or lacking DRIP are shown after rapid mixing of each vesicle population in 2.4 M sorbitol (A) 0.8 M isosmotic glycerol (B), or 0.8 M urea gradients (C). Open circles indicate data obtained for DRIP-containing vesicles, whereas closed circles represent data obtained with vesicles lacking the DAQP. Vesicles used in B were also used in A, vesicles used in C were also used in Fig. 5B. Both sets of DRIP vesicles clearly show evidence of rapid water transport but no evidence of solute transport. Glycerol, n = 4 (B); urea, n = 2 (C).

 

    DISCUSSION
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 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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 REFERENCES
 
This report is the first biophysical characterization of a water channel from Drosophila melanogaster, an organism that is amenable to genetic, developmental, and cellular analysis. The characterized AQP, DRIP, is most closely related and is 44% identical to human AQP4, a channel that exhibits the highest measured rate of water transport (48, 49). To establish DRIP's solute specificity, we used two heterologous AQP expression systems. Data obtained from examining DRIP expression in both Xenopus oocytes and yeast sec vesicles indicate that DRIP facilitates water transport, which is consistent with the similarity between DRIP and AQP4. Like AQP4, DRIP appears to be a water-specific channel because DRIP-containing vesicles did not exhibit increased conductance of glycerol, urea, protons, or ammonia.

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.


    GRANTS
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 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) Grant DK-43955 (to M. L. Zeidel). N. Kaufmann is the recipient of a National Research Service Award from the NIDDK (DK-065443).


    ACKNOWLEDGMENTS
 
We thank Dr. Ed Blumenthal for the in situ protocol, Dr. Gerard Campbell and laboratory for help with technical issues and fly maintenance, Dr. Beth Stronach for valuable discussions, Dr. Jeff Hildebrand for the use of his microscope, the Kleyman Laboratory for advice and help with the oocyte system, and Laura Kean and Nicole Southern for technical assistance.


    FOOTNOTES
 

Address for reprint requests and other correspondence: J. L. Brodsky, Dept. of Biological Sciences, Univ. of Pittsburgh, 274 Crawford Hall, Pittsburgh, PA 15260 (e-mail: jbrodsky{at}pitt.edu)

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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