Evidence for stabilization of aquaporin-2 folding mutants by N-linked glycosylation in endoplasmic reticulum

Teresa M. Buck, Joel Eledge, and William R. Skach

Molecular Medicine Division, Department of Medicine, Oregon Health Sciences University, Portland, Oregon 97201

Submitted 10 December 2003 ; accepted in final form 6 July 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Aquaporin-2 (AQP2) is the vasopressin-sensitive water channel that regulates water reabsorption in the distal nephron collecting duct. Inherited AQP2 mutations that disrupt folding lead to nephrogenic diabetes insipidus (NDI) by targeting newly synthesized protein for degradation in the endoplasmic reticulum (ER). During synthesis, a subset of wild-type (WT) AQP2 is covalently modified by N-linked glycosylation at residue Asn123. To investigate the affect of glycosylation, we expressed WT AQP2 and four NDI-related mutants in Xenopus laevis oocytes and compared stability of glycosylated and nonglycosylated isoforms. In all constructs, ~15–20% of newly synthesized AQP2 was covalently modified by N-linked glycosylation. At steady state, however, core glycosylated WT protein was nearly undetectable, whereas all mutants were found predominantly in the glycosylated form (60–70%). Pulse-chase metabolic labeling studies revealed that glycosylated isoforms of mutant AQP2 were significantly more stable than their nonglycosylated counterparts. For nonglycosylated isoforms, the half-life of WT AQP2 was significantly greater (>48 h) than that of mutant AQP2 (T126M 4.1 ± 1.0 h, A147T 4.2 ± 0.60 h, C181W 4.5 ± 0.50 h, R187C 6.8 ± 1.2 h). This is consistent with rapid turnover in the ER as previously reported. In contrast, the half-lives of mutant proteins containing N-linked glycans were similar to WT (~25 h), indicating that differences in steady-state glycosylation profiles are caused by increased stability of glycosylated mutant proteins. These results suggest that addition of a single N-linked oligosaccharide moiety can partially compensate for ER folding defects induced by disease-related mutations.

endoplasmic reticulum-associated degradation; nephrogenic diabetes insipidus; oocytes


NEPHROGENIC DIABETES INSIPIDUS (NDI) is characterized by the inability of the collecting duct to concentrate urine in response to stimulation by 8-arginine vasopressin (AVP). Most cases of congenital NDI are caused by mutations in one of two key proteins, the vasopressin (V2) receptor or aquaporin-2 (AQP2) (31, 52). AQP2, a member of the major intrinsic protein (MIP) family (13), is a hydrophobic protein of ~29 kDa that exists as a homotetramer in cell membranes. It contains six transmembrane segments and two inverted repeats (NPA motifs) that are thought to fold inward within the plane of the membrane to form the water-selective pore (12, 48). AQP2 synthesis, membrane insertion, folding, and maturation are facilitated by biosynthetic machinery in the endoplasmic reticulum (ER). Before its export from the ER, AQP2 must be properly folded and assembled into tetramers (20).

More than 26 AQP2 mutations have been causally linked to NDI, the majority of which are autosomal recessive and disrupt AQP2 function by altering stability of the immature protein (31). Four of these mutations, T126M, A147T, C181W, and R187C, have been extensively characterized in Xenopus oocyte, yeast, and mammalian systems and shown to exhibit defective intracellular trafficking and degradation by the ER-associated degradation (ERAD) pathway (4, 14, 21, 49, 53, 57). Although the precise mechanism of degradation remains unknown, it is believed that these mutations disrupt AQP2 folding and that the misfolded proteins are subsequently recognized by ER quality control machinery, ubiquitinated, and degraded by the 26S proteasome (9, 58). Certain mutations such as R187C and C181W that are located near the second NPA box disrupt the water-conducting pathway (5, 42, 49, 57), whereas other mutants, T126M and A147T, retain at least moderate water channel function (14, 29, 42, 49, 50). Thus channel function per se is not a criterion for recognition by ER quality control, raising the possibility that restored trafficking of mutant proteins may partially or completely correct the disease state.

Several AQPs (e.g., AQP1, AQP4, and AQP2) contain N-linked glycosylation consensus sites in their extracellular loops. Some of these sites are inefficiently recognized during protein synthesis by oligosaccharyltransferase (OST) to generate a mixture of glycosylated and nonglycosylated species (4, 11, 16, 17, 21, 29, 41, 46). Although N-linked glycosylation may be important for AQP2 trafficking through the Golgi (20), it has little effect on stability of wild-type (WT) AQP1 or AQP2, as both proteins exhibit normal function when glycosylation sites are removed or blocked by tunicamycin (2, 51). An unusual feature of AQP2 is that different glycosylation patterns have been reported for WT and mutant proteins (21, 29, 33). In membrane preparations derived from microinjected Xenopus oocytes, AQP2 mutants (T126M, A147T, and R187C) were found to be predominantly glycosylated, whereas WT AQP2 was present primarily in a nonglycosylated form (29). It was initially proposed that these differences were due to deglycosylation of WT protein as it passed through the Golgi, thus reducing the apparent glycosylation efficiency. However, it has recently been shown that WT and R187C AQP2 are both inefficiently glycosylated in oocytes (20). In addition, nonglycosylated forms of T126M AQP2 exhibited a shortened half-life compared with glycosylated forms when expressed in cultured hepatoma cells (21). In light of these findings we sought to determine whether N-linked glycosylation confers a general effect on AQP2 stability in the ER compartment. Using Xenopus oocytes and pulse-chase metabolic labeling, we examined four mutant AQP2 proteins that are known to cause NDI: T126M, A147T, C181W, and R187C. At early time points, all proteins were cotranslationally glycosylated to a similar extent as WT AQP2 (~20%). Interestingly, glycosylated forms of all mutant proteins were significantly more stable than their nonglycosylated counterparts, thus causing the percentage of glycosylated mutant protein to increase progressively over time. In contrast, N-linked oligosaccharides had little effect on the stability of the WT protein. These findings indicate that the presence of N-linked sugars exerts a generalized effect on AQP2 ERAD by delaying the recognition of misfolded substrate by ER quality control machinery.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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cDNA construction. Plasmid pSP64.myc-AQP2 was constructed by using a sense oligonucleotide (CCGGGTACATGTCTGAACAAAAACTTATTTCTGA AGAAGATCTGTGGGACCTCCGCTCCATA) to fuse the 10-residue myc epitope (EKQEEDL) (10) to the NH2 terminus of AQP2 (provided by A. Verkman, Univ. of California, San Francisco, CA) with standard cloning methods described previously (43). The missense mutants myc-AQP2 T126M, myc-AQP2 A147T, myc-AQP2 C181W, and myc-AQP2 R187C were constructed from pSP64.myc-AQP2 with site-directed mutagenesis (PCR overlap extension; Ref. 22). The chimeric clones thus encode N-terminally myc-tagged AQP2, AQP2T126M, AQP2A147T, AQP2C181W, and AQP2R187C. All cloned fragments were verified by DNA sequencing.

Truncated fusion proteins were generated by ligating the COOH-terminal 142 residues from bovine prolactin (43) to the indicated codons in AQP2. This was accomplished by using a sense oligonucleotide (oligo 242 base pairs upstream of the SP6 promoter) and antisense oligonucleotides encoding a BstEII restriction site at AQP2 codons Val131 (GAGCTCGGTCACCACCGCCTGGCCAGCCGT), Pro157 (AGCAGGGGTCACCGGGTTCTCTCCGCGGCG), Trp202 (GATCCCAGGTCACCCAGTGGTCATCAAAT), and Arg267 (GGCCTTGGTCACCCGTGGCAGGCTCTGC). Fragments were digested with NheI and BstEII and ligated 5' to a NheI/BstEII-digested vector, S.LST.gG.P, as described previously (43). The chimeric constructs thus encode the AQP2 coding sequence specified and the reporter domain (diagrammed in Fig. 6A). Importantly, this reporter contains no intrinsic topogenic information and faithfully follows the direction of upstream topogenic determinants (39, 44).



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Fig. 6. Cotranslational glycosylation is unaffected in AQP2 mutants. A: WT and mutant AQP2 cDNA was truncated after TM3, TM4, TM5, or TM6 as diagrammed and fused to a prolactin-derived COOH terminus reporter (P). B: oocytes injected with mRNA and [35S]methionine were incubated 3 h, immunoprecipitated with anti-prolactin antibody, and analyzed by SDS-PAGE. Predicted topology for the AQP2 protein at each truncation is diagrammed beneath sets of autoradiograms. Circle denotes attachment of N-linked sugars. ER, endoplasmic reticulum. C: oocytes injected with mRNA + [35S]methionine were incubated for time specified and immunoprecipitated with 9E10 antibody. D: data in C were quantitated to determine the % glycosylation at each time point. Signals of glycosylated protein at time points earlier than 30 min were not sufficient for quantitation.

 
Xenopus laevis expression. mRNA was transcribed in vitro with SP6 RNA polymerase (New England Biolabs) using 2 µg of plasmid DNA in a 10-µl volume at 40°C for 1 h as previously described (39). Aliquots were used immediately or frozen in liquid nitrogen and stored at –80°C. Two microliters of transcript was mixed with fifty microcuries of 35S-labeled methionine (0.5 µl of a 10x concentrated Tran35S-label; ICN Pharmaceuticals, Irvine, CA) and injected into stage VI Xenopus oocytes (50 nl/oocyte). Oocytes were incubated at 18°C in modified Barth’s saline with HEPES (mBSH) [in mM: 88 NaCl, 1 KCl, 24 NaHCO3, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.41 CaCl2, and 10 HEPES pH 7.4 with 50 µg/ml gentamicin, 100 U/ml penicillin, and 100 mg/ml streptomycin sulfate]. For pulse labeling experiments, oocytes were homogenized at the indicated time points and radiolabeled protein was immunoprecipitated before SDS-PAGE. For pulse-chase experiments, medium was removed from oocytes 1.5 h after injection and replaced with mBSH supplemented with 2 mM unlabeled methionine. Incubation was then continued for an additional 30 min to equilibrate unlabeled methionine and complete translation of labeled AQP2 protein (56). This time was used as our initial time point for pulse-chase studies. At the times indicated, oocytes (5 per lane) were flash frozen in liquid nitrogen and stored at –80°C.

Immunoprecipitation. Groups of oocytes were thawed, solubilized in 100 µl of 0.1 M Tris-Cl, pH 8.0, 1% SDS in a 1.5-ml microcentrifuge tube and incubated at 37°C for 30 min. Homogenates were then diluted in 10 volumes of buffer A (1% Triton X-100, 100 mM Tris-Cl, pH 8.0, 100 mM NaCl, 5 mM EDTA) and incubated on ice for 30 min, and insoluble material was removed by centrifugation (16,000 g for 15 min at 4°C); 5.0 µl of protein A Affigel (Bio-Rad, Hercules, CA) and 0.75 µl of antibody Myc-9E10 (mouse ascites fluid; Ref. 10) were added. The sample was mixed at 4°C for 10 h before being washed three times with buffer A and twice with 0.1 M NaCl and 0.1 M Tris-Cl, pH 8.0. Samples were analyzed by SDS-PAGE, EN3HANCE (PerkinElmer Life Sciences, Boston, MA) fluorography, and autoradiography. Band signals were quantitated with a Bio-Rad personal Molecular PhosphorImager Fx (Kodak screens, Quantity-1 software).

PNGase F digest. After SDS solubilization at 37°C, samples were split and immunoprecipitated as described above. Protein A beads were washed, resuspended in 15 µl of 0.1 M Tris-Cl, pH 7.5, containing 0.3 µl of PNGase F (New England Biolabs, Beverly, MA), and incubated for 3 h at 37°C before SDS-PAGE.

Immunoblotting oocyte membranes. Oocytes were injected as above but without [35S]methionine, incubated at 18°C for the times indicated, and homogenized in 0.25 M sucrose, 50 mM potassium acetate (KOAc), 5 mM MgCl2, 1 mM DTT, and 50 mM Tris-Cl, pH 7.5 (5 µl/oocyte). Homogenates were centrifuged at 800 g for 5 min. The supernatant was layered onto 400 µl of 0.5 M sucrose, 50 mM KOAc, 5 mM MgCl2, and 50 mM Tris-Cl, pH 7.5, on top of 100 µl of 1.8 M sucrose in the same buffer. Samples were centrifuged at 186,000 g for 10 min. Total cellular membranes were collected at the 0.5 M–1.8 M sucrose interface, diluted with 400 µl of 50 mM KOAc, 5 mM MgCl2, and 50 mM Tris-Cl, pH 7.5, and repelleted through 0.5 M sucrose, 50 mM KOAc, 5 mM MgCl2, and 50 mM Tris-Cl, pH 7.5, by centrifugation at 186,000 g for 10 min. Membrane pellets were solubilized and separated by SDS-PAGE and transferred to nitrocellulose. Blots were blocked overnight with 5% nonfat dry milk, probed for 1 h with Myc 1-9E10 antibody (1:5,000, mouse ascites fluid), washed six times with 10 mM Tris-Cl, pH 8.0, 150 mM NaCl, and 0.1% Tween 20, probed with goat anti-mouse IgG horseradish peroxidase (HRP) antibody (1:10,000; Bio-Rad) for 1 h, and washed as before. Bands were detected by enhanced chemiluminescence (ECL) with pico west Supersignal (Pierce, Rockford, IL) per manufacturer's instructions.

Half-life determination. To correct for variation in oocyte equilibration with the unlabeled methionine in the medium, the 1.5 h time point was normalized to 100% protein. Half-lives for the total and nonglycosylated protein were calculated assuming steady-state first-order elimination kinetics by fitting data to the equation A = Aoekt with Prism graphing software, where A is the protein remaining, Ao is protein at time 0, k is the rate constant, and t is time. Goodness of fit for the regression was analyzed by determining an F-statistic to determine the ability of the regression equation to fit the data (SigmaPlot). Data with P < 0.05 were eliminated from the calculations (2 of 26 experiments). All data are shown as means ± SE for a minimum of five experiments. Because of the stability of WT protein, an accurate half-life could not be calculated but was estimated to be >48 h. Data for the glycosylated protein did not follow first-order degradation kinetics, and so half-lives were estimated from Fig. 5B.



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Fig. 5. Glycosylation stabilizes mutant protein. Stability of nonglycosylated (A) and glycosylated (B) protein was determined as in Fig. 4 by quantifying 32-kDa and 29-kDa bands independently from separate pulse-chase experiments. Half-lives for mutants were determined as described in MATERIALS AND METHODS. WT half-life was estimated to be >48 h.

 

    RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Xenopus oocytes efficiently express a wide variety of AQP water channels and have been extensively used to study AQP2 function and trafficking (4, 5, 33, 40, 49). Importantly, oocytes also faithfully reconstitute quality control events at the level of the ER (56). Although this quality control is sometimes less stringent than in mammalian cells, oocytes maintain the ability to discriminate and degrade mutant proteins via the ERAD pathway and therefore provide a useful system for examining early events of protein biogenesis (56). We expressed myc-tagged versions of WT and four mutant AQP2 proteins (T126M, A147T, C181W, R187C) in microinjected oocytes to examine the role of N-linked glycosylation on AQP2 stability. These mutations were chosen because they exhibit similar trafficking defects in multiple expression systems including oocytes and because they reside in different regions of the protein, extracellular loop 2, intracellular loop 2, and extracellular loop 3 (see Fig. 1), and therefore likely disrupt different aspects of AQP2 folding.



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Fig. 1. Diagram of predicted aquaporin-2 (AQP2) topology and relative locations of 4 nephrogenic diabetes insipidus (NDI)-linked mutations, T126M, A147T, C181W, and R1817C. Topology and location of transmembrane (TM) segments was based on crystal structure of AQP1 and GlpF (6, 12, 34); branching structure represents high-mannose core oligosaccharide attached to Asn123.

 
WT and mutant AQP2 proteins exhibit different glycosylation profiles. AQP2 N-glycosylation status was initially examined by using total oocyte membranes that were collected either 2 h after injection, when the proteins are predominantly localized to the ER (56), or 48 h after injection, when WT protein should be maximally expressed at the plasma membrane (Refs. 14 and 32 and data not shown). Immunoblot analysis revealed that both WT and mutant proteins were present as two distinct isoforms, a major band of 29 kDa and a minor band of 32 kDa (Fig. 2A). PNGase F digestion confirmed that the difference in size was due to the covalent attachment of a single N-linked high-mannose core oligosaccharide (Fig. 2B). Initial glycosylation efficiency for both WT and mutant proteins was 10–20%. Thus the only N-linked AQP2 consensus site, residue Asn123, which is located in extracellular loop 2, is relatively inaccessible to OST. Forty-eight hours after injection a distinctly different glycosylation pattern was noted. WT protein remained predominantly in the nonglycosylated 29-kDa form, whereas mutant proteins were found mainly in the glycosylated form. These results were observed for all four mutants regardless of the site of mutation. Thus they appear to represent a general pattern independent of the specific folding defect.



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Fig. 2. Glycosylation profiles for wild-type (WT) and mutant AQP2. A: Xenopus oocytes were injected with WT or mutant, myc-tagged AQP2 mRNA as indicated. Total oocyte membranes were collected 2 and 48 h after RNA injection and immunoblotted with anti-myc (9E10) antibody. Glycosylated (+g) and nonglycosylated (–g) AQP2 bands are indicated. Exposure times were adjusted to best demonstrate the glycosylation profile. B: oocytes were coinjected with mRNA and 35S-labeled methionine, incubated 8 h, and homogenized. Immunoprecipitated samples (myc, 9E10 antibody) were analyzed directly (–) or after digestion with PNGase F (+) by SDS-PAGE and autoradiography.

 
Pulse-chase analysis of AQP2 synthesis in Xenopus oocytes. To determine why mutant and WT AQP2 exhibited different glycosylation profiles at later time points, pulse-chase metabolic labeling studies were performed. Oocytes were coinjected with mRNA and [35S]methionine to label newly synthesized protein. Previous studies demonstrated that [35S]methionine is efficiently taken up into the aminoacyl (aa)-tRNAmet pool and incorporated into protein (Ref. 56 and data not shown). Ninety minutes after injection, oocytes were placed in fresh medium containing 2 mM unlabeled methionine and incubated for an additional 30 min (prechase period) to allow uptake and equilibration of cold methionine and to complete synthesis of labeled AQP2 polypeptides. Groups of oocytes were then homogenized at indicated time points, immunoprecipitated with anti-myc antibody, and analyzed by SDS-PAGE and autoradiography. As expected, initial glycosylation efficiency was low; ~20% of AQP2 was recovered as the 32-kDa glycosylated species for all constructs tested (Fig. 3A). Nonglycosylated WT AQP2 was remarkably stable throughout the chase period, whereas its glycosylated form was stable for 10 h and then gradually decreased in intensity. This late disappearance likely represents Golgi processing of high-mannose core carbohydrates, although it has been difficult to consistently detect Golgi processed forms in oocytes because of their low abundance and heterogeneous appearance (32). It is also possible that some WT AQP2 may pass through the Golgi without further modification. For mutant proteins, nonglycosylated forms were rapidly degraded (after a brief lag period), with little protein detected after 24 h. Interestingly, glycosylated isoforms of mutant proteins were significantly more stable than their nonglycosylated counterparts, suggesting that the increase in AQP2 glycosylation observed at late time points primarily resulted from increased stability of mutant AQP2 induced by N-linked glycosylation.



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Fig. 3. Pulse-chase analysis of AQP2 synthesis. A: Xenopus oocytes were coinjected with myc-tagged AQP2 mRNA and [35S]methionine, labeled for 1.5 h, and chased in fresh medium containing unlabeled methionine. Time 0 was taken as 30 min after medium change to allow equilibration of cold methionine. At the times indicated oocytes (5 oocytes/lane) were homogenized and immunoprecipitated with Myc-9E10 antibody and analyzed by SDS-PAGE and autoradiography. Glycosylated (+g) and nonglycosylated (–g) AQP2 is indicated. B: data were quantitated by phosphorimaging, and values were normalized to the 1.5 h time point. The data represent the total protein, 29-kDa band + 32-kDa band. Results show means ± SE for ≥5 separate experiments.

 
AQP2 trafficking mutants display reduced half-life in Xenopus oocytes. Data from multiple pulse-chase experiments were quantitated by phosphorimaging to determine the half-lives of WT and mutant AQP2 proteins. The half-life of each protein was then determined by fitting a standard curve for first-order degradation kinetics as described in MATERIALS AND METHODS. Data shown in Fig. 3B clearly demonstrate a decrease in stability of mutant proteins compared with WT AQP2. WT protein had a half-life of >48 h, whereas the half-lives for mutant proteins were ~7.5 ± 1.4 h (T126M), 7.1 ± 1.6 h (A147T), 8.0 ± 1.0 h (C181W), and 12 ± 2.2 h (R187C).

We next examined the fraction of glycosylated proteins as a function of time (Fig. 4). Two hours after injection, WT and mutant proteins were all glycosylated to the same extent, consistent with Fig. 2. All mutants showed a progressive increase in percent glycosylation from ~20% to 65–70%, whereas the fraction of glycosylated WT protein remained nearly constant. Together, these results further demonstrate that N-linked glycans have a markedly different effect on mutant and WT AQP2.



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Fig. 4. Percentage of glycosylated mutant AQP2 protein increases over time. The percentage of glycosylated protein as a function of total protein at each time point was determined as in Fig. 3. Results show means ± SE of ≥5 separate experiments.

 
Glycosylation stabilizes mutant protein. We next determined the half-lives of the 29-kDa and 32-kDa isoforms independently. Nonglycosylated WT AQP2 has a significantly longer half-life (>48 h) than any of the nonglycosylated mutant proteins (T126M 4.1 ± 1.0 h, A147T 4.2 ± 0.60 h, C181W 4.5 ± 0.50 h, R187C 6.8 ± 1.2 h; Fig. 5A). In contrast, the half-life of glycosylated mutant proteins (15–26 h) was very similar to that of WT protein (20 h), and in each case, the glycosylated isoform was more stable than its nonglycosylated counterpart (Fig. 5B). We also noted that for R187C there was an absolute increase in intensity of the 32-kDa band during early time points. The reason for this remains unclear, but it suggests that oligosaccharides may continue to be added to R187C polypeptide after synthesis is complete (discussed below).

AQP2 mutations do not affect cotranslational glycosylation and early folding events. Although our data support a model in which N-linked glycosylation confers increased stability on mutant (and misfolded) AQP2 proteins, an alternative possibility is that these mutations might influence glycosylation on a subset of proteins that had already achieved a more stable conformation at the time of glycosylation. AQP topology is directed by topogenic determinants (e.g., signal anchor and stop transfer sequences) that target the ribosome nascent chain complex to the ER membrane, initiate and terminate translocation of peptide loops, and integrate transmembrane (TM) segments into the lipid bilayer (8, 11, 27, 44). Because N-linked glycosylation normally occurs cotranslationally as peptide loops translocate into the ER lumen, we tested whether accessibility of Asn123 by OST might affected by NDI-related mutations that altered AQP2 folding.

Plasmids encoding WT and mutant AQP2 proteins were truncated at codons Val131, Pro157, Trp202, and Arg267 and ligated to a COOH terminus reporter to generate fusion proteins containing 3, 4, 5, or 6 of the AQP2 TM segments (Fig. 6A). In vitro transcribed mRNA was expressed in Xenopus oocytes, and reporter-containing polypeptides were recovered by immunoprecipitation and analyzed by autoradiography. These experiments revealed that WT and mutant constructs at any given truncation site were cotranslationally glycosylated to the same extent even though constructs encoded only a portion of the AQP2 protein. Thus synthesis of TMs 4, 5, and 6 is not required for glycosylation of Asn123 to occur. Moreover, glycosylation of Asn123 occurs even before mutant residues are synthesized, confirming that AQP2 misfolding must be a late event relative to the timing of initial oligosaccharide attachment. Of note, fusion proteins truncated at residue Val131, just before TM4, were more efficiently glycosylated (~50%; Fig. 6B). This occurs because removal of TM4 increases the accessibility of Asn123 to OST by increasing its tethered distance from the ER membrane (see Fig. 1 and Ref. 11). Given that the fusion proteins lack one or more TM segments (TMs 5 and/or 6), the glycosylation observed here is not dependent on acquisition of tertiary folded structure.

Finally, to confirm that the extent of glycosylation observed (20%) reflects cotranslational attachment of oligosaccharide rather than posttranslational addition, we compared the timing of AQP2 glycosylation relative to protein synthesis. For these experiments, oocytes were harvested at very short time intervals after injection of mRNA and [35S]methionine. Full-length radiolabeled AQP2 first appeared 10–20 min after microinjection, consistent with the time required for [35S]methionine incorporation into the aa-tRNA pool and synthesis of the 29-kDa protein (Fig. 6, C and D). Importantly, appearance of N-linked glycosylated species coincided precisely with the completion of synthesis, and levels of glycosylation (12–14%) are in good agreement with those observed at the initiation of the pulse-chase experiments (see Fig. 2). Because of technical constraints and the low abundance of protein present at these short time points, it was not possible to determine whether partial-length translation intermediates were also glycosylated. However, together our data support a model in which cotranslational attachment of N-linked sugars is primarily dictated by steric accessibility of the consensus site as extracellular loop 2 topology is established and not by the presence of specific mutations.


    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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The current study examines the effect of N-linked glycosylation on the early ER quality control of WT and mutant forms of the vasopressin-regulated water channel AQP2. Because AQP2 is inefficiently glycosylated in most cell types including mammalian cells, it affords a unique opportunity to directly compare, in the same cell, the fate of identical polypeptides that differ only in the presence of a single N-linked oligosaccharide. In the case of WT protein, nonglycosylated isoforms have relatively long half-lives that likely reflect productive processing and trafficking out of the ER compartment. This finding is consistent with previous studies demonstrating that glycosylation is not required for AQP function; some AQPs lack N-linked consensus sites (AQP6), whereas others have extracellular sites that are not utilized (AQP4; Ref. 41). Still others, such as AQP1, AQP2, and AQP3, have sites that are utilized in only a subset of synthesized proteins (1, 2, 46), and removal or blockage of N-linked glycosylation has no discernable effect on function (2, 51). In contrast, our studies of NDI mutants provide evidence that the effects of AQP glycosylation can be both subtle and complex. In Xenopus oocytes, each of the four NDI mutations studied had a dramatic effect on the stability of AQP2 in the ER, decreasing the half-life of mutant proteins by more than fourfold compared with WT. This was expected given that all mutants were chosen on the basis of previous reports showing defective trafficking and rapid degradation in the ER (33, 49, 50). When stability of glycosylated and nonglycosylated species was examined separately, however, we found that the presence of N-linked glycans markedly and selectively stabilized mutant proteins, with the half-life (in the ER) approaching that of WT. These results thus extend the previous findings of Hirano et al. (21) and demonstrate a general role for N-linked glycans in early AQP2 folding and ER quality control. It should be noted that glycosylated and nonglycosylated monomers of WT AQP2 readily formed heterotetramers. In contrast, recessive mutations that disrupt AQP2 folding and ER trafficking fail to tetramerize (5, 20, 24). Therefore, it seems likely that the differential stability of mutant isoforms reflects that of monomeric proteins.

N-linked glycans have long been known to facilitate protein folding. Glycosylation inhibitors cause generalized protein misfolding in the ER lumen, as demonstrated by the induction of a strong unfolded protein response (15, 19, 25, 28, 3638, 45, 54). N-linked glycans are also involved in targeting misfolded substrates to the ERAD pathway. Cleavage of a single mannose residue by ER mannosidase I generates a Man8 intermediate and provides a degradation signal recognized by the ER protein EDEM (23). This process may provide a timing mechanism for monitoring folding, because if folding is completed before Man9 trimming, the substrate is exported from the ER. In contrast, if mannose trimming occurs before export, the substrate may become a degradation target. Our findings that glycosylation specifically improves the stability of trafficking mutants, but not WT AQP2, suggests a novel physiological role for N-linked glycans, namely, that they provide a selective advantage for proteins slightly below the threshold for efficient folding. Such a finding is particularly intriguing given the large number of disease-related mutations that direct proteins into the ERAD pathway.

AQP2 is a highly hydrophobic protein with a significant fraction of its mass located within the lipid bilayer and only a small portion of luminally exposed polypeptide. It is therefore unlikely that glycosylation would effect solubility or aggregation of AQP2 as has been proposed for soluble proteins. Consistent with this notion, none of the mutations examined affected protein translocation or early folding events that establish cotranslational topology of individual TM segments (Fig. 6 and data not shown), and addition of N-linked sugars occurs cotranslationally even before the mutant residues have been synthesized. This eliminates the possibility that a cohort of more stable mutant proteins is preferentially glycosylated. It therefore seems likely that the presence of the oligosaccharide moiety either directly facilitates late folding events, possibly within the lipid bilayer, or in some manner masks the misfolded protein from recognition by quality control machinery (30).

One possibility is that interactions with calnexin (CNX) or calreticulin (CRT) may stabilize glycosylated AQP2 by binding to partially trimmed glucose residues. For example, when a truncated form of ribophorin I (RI), RI332, was pharmacologically prevented from binding to CNX/CRT, its half-life was reduced (7). Similarly, RI mutants lacking glycosylation consensus sites exhibited decreased half-lives. Major histocompatibility complex (MHC) class I heavy chain was also degraded more rapidly when interactions with CNX were blocked, but, in contrast to RI332, removal of the N-linked consensus site had a stabilizing effect (55), suggesting a complex relationship between carbohydrate effects on folding and the CNX/CRT pathway. In our case, we were unable to detect a significant difference in the stability of either WT or mutant glycosylated AQP2 after treatment of oocytes with the glucosidase inhibitors castanospermine or deoxynorjirimycin (data not shown). However, interpretation of these experiments was difficult because of the incubation times necessary to observe stability differences (10–20 h) and the toxicity of long-term exposure to glucosidase inhibitors.

Because water channel function could not be detected for any of the mutant proteins (data not shown), it is unlikely that their proper trafficking ever occurs in oocytes. We therefore favor the possibility that recognition of mutant AQP2 by quality control machinery is somehow delayed by the presence of the glycan moiety. This implies that folding defects in different mutants are impacted by events in the ER lumen. AQP2 stabilization also appears to be generalized and independent of the folding defect because it is shared by missense mutations located in different physical and functional regions of the protein (Fig. 1). Importantly, this effect is not limited to the Xenopus expression system because similar observations were made for the T126M mutant expressed in mammalian hepatocytes (21).

Most consensus sites in polytopic proteins must be located at least 12–14 residues from the ends of TM segments for OST to efficiently attach the carbohydrate moiety (26, 35). On the basis of AQP1 and Glpf crystal structures (6, 12, 34), we estimate that Asn123 is only 6–9 residues from the N-terminus of TM4 (see Fig. 1). This is likely responsible for the inefficient glycosylation observed for AQP2 because increasing this distance to ~16 residues (i.e., truncation before TM4) substantially improved cotranslational glycosylation efficiency (Fig. 6). AQP2 glycosylation also appears to be primarily cotranslational, as it takes place during translation and before AQP synthesis is completed. However, isolated reports have indicated that prolonged ER residence time may increase potential exposure of N-linked consensus sites to ER-localized OST and result in posttranslational glycosylation (3, 18). To some extent this may occur for R187C, in which the absolute amount of glycosylated material continued to increase after synthesis of radiolabeled protein was completed. ER retention per se, however, did not correlate well with the extent of posttranslational glycosylation, because T126M, A147T, and C181W mutants were all ER retained but did not exhibit the increase of total glycosylated protein observed for R187C. In addition, glycosylated proteins persisted after the nonglycosylated protein had been largely degraded, making it unlikely that increased stability was due solely to posttranslational glycosylation. Thus our data support a model in which AQP glycosylation is primarily cotranslational and limited by steric accessibility of the consensus site. Unfortunately, attempts to define the precise extent of posttranslational glycosylation with tunicamycin were inconclusive. Oocytes stores of dolichol-bound high-mannose oligosaccharides were only depleted after 8 h of tunicamycin exposure (data not shown), and stability of both glycosylated and nonglycosylated AQP2 isoforms was decreased during subsequent incubation, consistent with a general toxic effect (data not shown).

In conclusion, we have shown that the N-linked glycosylated forms of four naturally occurring disease-related AQP2 mutants, T126M, A147T, C181W, and R187C, persist longer in the ER than their nonglycosylated counterparts. It was previously proposed that high steady-state levels of glycosylated mutant AQP2 were caused by poor folding of the mutant proteins that interfered with the removal of the high-mannose carbohydrates in the early cis-Golgi complex (29). In contrast, our results are most consistent with the interpretation that addition of N-linked glycans results in a generalized increase in mutant AQP2 half-life, stabilizing the small pool of glycosylated (relative to nonglycosylated) species that accumulates and predominates over time. This does not occur for WT protein in which the nonglycosylated form is already very stable. We cannot completely rule out, however, the possibility that delayed glycosylation may contribute to the apparent half-life for some mutants, particularly R187C. Although glycosylation of some proteins is necessary for them to obtain their native structure, WT AQP2 does not require addition of N-linked glycosylation for proper folding or water channel function. Thus it is surprising that addition of N-linked sugars to AQP2 mutant proteins has such an effect. These studies raise the possibility that maneuvers to manipulate attachment of N-linked carbohydrates may provide one strategy for improving stability and possibly trafficking of mutant proteins in human disease.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by National Institutes of Health Grants GM-53457 and DK-51818. W. R. Skach is an established investigator of the American Heart Association.


    ACKNOWLEDGMENTS
 
We thank Dr. A. Verkman for providing AQP2 cDNA and Dr. D. Koop for helpful discussion regarding degradation kinetics.


    FOOTNOTES
 

Address for reprint requests and other correspondence: W. R. Skach, Div. of Molecular Medicine, Oregon Health Sciences Univ., 3181 SW Sam Jackson Park Rd., Portland, OR 97239 (E-mail: skachw{at}ohsu.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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