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. 2009 Jun 11;150(9):4191–4202. doi: 10.1210/en.2009-0285

Soluble Corticotropin-Releasing Hormone Receptor 2α Splice Variant Is Efficiently Translated But Not Trafficked for Secretion

Ryan T Evans 1, Audrey F Seasholtz 1
PMCID: PMC2736082  PMID: 19520785

Abstract

CRH directs the physiological and behavioral responses to stress. Its activity is mediated by CRH receptors (CRH-R) 1 and 2 and modulated by the CRH-binding protein. Aberrant regulation of this system has been associated with anxiety disorders and major depression, demonstrating the importance of understanding the regulation of CRH activity. An mRNA splice variant of CRH-R2α (sCRH-R2α) was recently identified that encodes the receptor’s ligand-binding extracellular domain but terminates before the transmembrane domains. It was therefore predicted to serve as a secreted decoy receptor, mimicking the ability of CRH-binding protein to sequester free CRH. Although the splice variant contains a premature termination codon, predicting its degradation by nonsense-mediated RNA decay, cycloheximide experiments and polysome profiles demonstrated that sCRH-R2α mRNA escaped this regulation and was efficiently translated. However, the resulting protein was unable to serve as a decoy receptor because it failed to traffic for secretion because of an ineffective signal peptide and was ultimately subjected to proteosomal degradation. Several other truncated splice variants of G protein-coupled transmembrane receptors regulate the amount of full-length receptor expression through dimerization and misrouting; however, receptor binding assays and immunofluorescence of cells cotransfected with sCRH-R2α and CRH-R2α or CRH-R1 indicated that sCRH-R2α protein does not alter trafficking or binding of full-length CRH-R. Although sCRH-R2α protein does not appear to function as an intracellular or extracellular decoy receptor, the regulated unproductive splicing of CRH-R2α pre-mRNA to sCRH-R2α may selectively alter the cellular levels of full-length CRH-R2α mRNA and hence functional CRH-R2α receptor levels.

The soluble splice variant of corticotropin-releasing hormone receptor 2α is expressed in vivo, but is unable to function as a soluble decoy receptor because it is mistrafficked and degraded by the proteasome.

CRH is the primary hypothalamic mediator of the mammalian neuroendocrine stress response. In response to stress, CRH is released at the median eminence and stimulates corticotropes in the anterior pituitary to express and release ACTH. ACTH stimulates the adrenal glands to secrete glucocorticoids, which mediate many of the physiological responses to stress and negatively regulate the HPA axis to quell the response (1). CRH also acts as a neurotransmitter in numerous other sites in the central nervous system, mediating the metabolic, behavioral, autonomic, and immune responses to stress (1,2). In addition to CRH, several other CRH-like peptides have been identified, including urocortin (Ucn) I, II, and III (3,4,5,6). These CRH-like ligands have diverse expression patterns and contribute to a range of physiological functions, including energy balance and cardiovascular and intestinal function (reviewed in Refs. 7 and 8). Dysregulation of CRH and the Ucns has been correlated with a number of disorders including major depression, anxiety disorders, anorexia, and inflammatory and cardiac disease, demonstrating the significance of understanding the regulation of their activity (9,10,11,12,13).

CRH and Ucn mediate their effects through two G protein-coupled transmembrane receptors (GPCRs) of the class B1 subfamily, CRH receptor 1 (CRH-R1) and CRH-R2 (for review see Ref. 14). Expressed by separate genes, these receptors are detected in a few overlapping, but largely distinct, sites in both the central nervous system and periphery (15). Functional studies and knockout mice models suggest that CRH-R1 may initiate the stress response, whereas CRH-R2 modulates it (reviewed in Ref. 16). Although CRH-R1 and CRH-R2 share about 70% amino acid identity, they have differing pharmacologies due to lower similarity in their N-terminal ligand-binding domains. CRH and Ucn I bind specifically to both CRH-R1 and CRH-R2, whereas Ucn II and Ucn III preferentially or selectively bind CRH-R2 (14). Although several alternative splice forms of CRH-R1 (α, β, c-n) have been identified in rodents or humans, CRH-R1α is the predominantly expressed and functional form (14). Most other CRH-R1 splice variants contain truncations and deletions that disrupt ligand binding and/or signaling capabilities, and functional roles for these variants are still under investigation (17,18,19,20,21). CRH-R2 has two isoforms in rodents (α and β) and three in humans (α, β, and γ) that arise from separate promoters and 5′ exons that splice to a common set of downstream exons (22,23). In rodents, CRH-R2α is expressed primarily in the brain, whereas CRH-R2β is found mainly in the periphery, including the heart and skeletal muscle (15,23,24,25,26,27). In addition to the receptors, CRH activity is modulated by the evolutionarily conserved CRH-binding protein (CRH-BP), a 37-kDa secreted glycoprotein that binds CRH with equal or greater affinity than the receptors (28,29). CRH-BP appears to predominantly function to sequester CRH and inhibit its activity (28,29), although several lines of evidence suggest that CRH-BP may have other functions as well (29,30).

Recently, Chen and co-workers (31) identified an alternative splice variant of CRH-R2α in mouse in which exon 6 is deleted (called sCRH-R2α). Deletion of exon 6 causes a frameshift and premature termination codon (PTC) in exon 7, before sequences encoding the transmembrane domains. As a result, the sCRH-R2α sequence was predicted to encode the ligand-binding extracellular domain and a unique, hydrophilic, 38-amino-acid C-terminal tail. Lacking the anchoring transmembrane domains and C-terminal signal domains, sCRH-R2α was thought to produce a secreted decoy receptor that would inhibit CRH activity similar to the CRH-BP. Studies by Chen and co-workers (31) supported this hypothesis as recombinant sCRH-R2α protein (expressed in bacteria or from a eukaryotic expression vector with secretion tag) bound CRH at an affinity similar to the full-length receptor and inhibited the CRH/Ucn I-induced cAMP and ERK1/2-p42,p44 signaling pathways in cultured cells expressing CRH-R1 or CRH-R2.

However, it remained unclear whether the alternatively spliced sCRH-R2α transcript was efficiently translated in vivo and whether the protein was properly trafficked within the secretory pathway to function as a decoy receptor. First, transcripts containing PTCs, such as sCRH-R2α, are often regulated by nonsense-mediated RNA decay (NMD), which degrades aberrant mRNA transcripts to presumably prevent the expression of harmful truncated or mutated proteins (i.e. through dominant-negative activity or energy-expensive translation of inactive protein) (32,33). Although sCRH-R2α mRNA contains a PTC that would predict its regulation by NMD, it was unclear whether this occurs, because there are several examples of transcripts that fit this criterion yet escape NMD regulation (34,35,36,37). Second, studies by Rutz et al. (38) suggested that endoplasmic reticulum (ER) translocation and proper trafficking of the full-length CRH-R2α is driven by the first transmembrane domain. Because sCRH-R2α lacks any transmembrane domains, it was unclear whether sCRH-R2α protein would properly traffic for secretion as a decoy receptor in vivo. Several other alternatively spliced and truncated GPCRs (including members of the class B1 subfamily) show misrouted expression, and interestingly, instead of acting as decoy receptors, some of these truncated proteins serve to regulate functional expression of their full-length receptor counterparts through heterodimerization and coretention in a misrouted location (20,21,39,40,41,42,43,44,45).

To further elucidate the potential role of the sCRH-R2α splice variant in the CRH system, the regulation of sCRH-R2α mRNA and protein expression was examined. Studies presented here make use of NMD-inhibition and polysome profiles to determine whether the sCRH-R2α transcript is efficiently translated, and Western analysis and immunofluorescence confocal microscopy to examine whether the potentially synthesized sCRH-R2α protein is appropriately trafficked for secretion in vivo. Finally, based on these analyses, alternative (nondecoy receptor) roles for sCRH-R2α were examined.

Materials and Methods

Animals

Wild-type C57BL/6 male mice were given food and water ad libitum. Mice were killed under nonstressed conditions and tissues removed and immediately frozen at −80 C or extracted for analysis. All animal experiments were approved by the University of Michigan Committee on Use and Care of Animals and performed according to National Institutes of Health guidelines. Rat tissue was kindly provided by Dr. Robert Thompson, University of Michigan.

Cell culture

αT3-1, LβT2, CATH.a, Cos-1, and HEK293 cells were cultured as described in supplemental Materials and Methods (published as supplemental data on The Endocrine Society’s Journals Online web site at http://endo.endojournals.org).

RT-PCR and quantitative real-time PCR (qRT-PCR)

Cultured cells or tissues were harvested with Trizol (Invitrogen, Carlsbad, CA), and isolated RNA was treated with deoxyribonuclease (Turbo DNA-free; Ambion, Austin, TX) and used for cDNA synthesis as previously described (46). PCR and qRT-PCR analyses were performed on cDNA and −RT samples using Taq DNA polymerase (Invitrogen) and SYBR Green I Master Mix (SuperArray Bioscience, Frederick, MD), respectively, as described previously (47). All primer sequences, cycling conditions, and product sizes are provided in supplemental Materials and Methods. Relative gene expression was determined by R = (Etarget)ΔCttarget/(Eref)ΔCtref (48), where E is primer efficiency and ΔCt is the difference in cycle threshold between the control and sample for target or reference (ref) genes. For every sample, the average of at least duplicate qRT-PCR was used as the Ct value in the above calculation (in addition to biological replicates used for statistical analysis).

Cycloheximide (CHX) treatment

LβT2 or CATH.a cells were grown to 80% confluency on 10-cm plates, treated with 50 μg/ml CHX (Calbiochem, Gibbstown, NJ) (34), and harvested for qRT-PCR after various time points. For the relief condition, cells were treated for 3 h with CHX, washed twice with CHX-free medium, and cultured in CHX-free medium for 8 h before harvesting.

Polysome analysis

Polysome analysis was performed as previously described (49) with slight modifications. Briefly, 8 × 107 LβT2 or CATH.a cells were washed with cold Dulbecco’s PBS (PBS-D) containing 3 μm CHX, and lysed in 1 ml HNM buffer [20 mm HEPES (pH 7.5), 100 mm NaCl, 1.5 mm MgCl2, 5% Triton X-100, 3 μm CHX, and 10 U/ml RNAseOUT (Invitrogen)] by passing 10 times through an 18-gauge needle. Nuclei were removed by centrifugation at 10,000 × g for 5 min at 4 C. The lysate was split into two equal aliquots (one aliquot was adjusted to 20 mm EDTA) and separated across 5–50% sucrose gradients in HNM buffer lacking Triton X-100 (with or without 20 mm EDTA) by centrifugation at 38,000 rpm for 2 h at 4 C in a SW41Ti rotor (Beckman, Fullerton, CA). Gradients were manually fractionated (0.75 ml/fraction) from the top and measured for A260 to determine the polysome profile. RNA was isolated from select fractions using Trizol LS (Invitrogen) and used for qRT-PCR. Procedural modifications for analysis of brain lysates are in supplemental Materials and Methods.

Expression vectors

All plasmids used in this study (except eGFP constructs) were made by ligation of mouse cDNA sequences into pcDNA3.1D/V5-His-TOPO (Invitrogen) for expression with or without C-terminal V5-His tags. Details are provided in supplemental Materials and Methods.

Western analysis

Cos-1 or αT3-1 cells were transiently transfected with expression plasmids using Lipofectamine (Invitrogen) and harvested after 48 h in PBS and lysed in TNE-Triton [10 mm Tris-HCl (pH 7.5), 150 mm NaCl, 1 mm EDTA, 0.5% Triton X-100, and 1:50 protease inhibitor cocktail (Sigma, St. Louis, MO)]. Protein concentration and transfection efficiency were determined by Bradford method (50) (Bio-Rad, Hercules, CA), and β-galactosidase assay (51), respectively. For MG132 treatment, cells were treated with vehicle [dimethylsulfoxide (DMSO)] or 20 μm MG132 (Calbiochem) in cell medium for 3 h before harvesting. For glycosidase treatment, 20 μg protein was treated for 1 h at 37 C with 500 U peptide N-glycosidase F (PNGase F; New England Biolabs, Ipswich, MA). The 20-μg protein samples were separated by SDS-PAGE (10% acrylamide) and transferred to Immobilon-P (Millipore, Bedford, MA). Immunoblotting procedures are described in supplemental Materials and Methods.

Immunofluorescence

Cos-1 or αT3-1 cells were grown on poly-d-lysine-coated coverslips in six-well plates and cotransfected with DNA of both the indicated expression construct (1.2 μg) and eGFP-N2 (Clontech, Mountain View, CA) or CRH-BP-eGFP (1.2 μg) using 7.2 μl Lipofectamine (Invitrogen), following the manufacturer’s recommendations. After 48 h, cells were treated with MG132 or vehicle as described above, washed in PBS-D, fixed 10 min with 3.7% formaldehyde, permeabilized 8 min with 0.1% Triton X-100, and blocked in 1.5% normal goat serum. Cells transfected with V5-tagged constructs were incubated with mouse anti-V5 primary at 1:2500 in PBS-D with 0.05% Tween 20, followed by goat antimouse-AlexaFluor 568 (Invitrogen) secondary at 1:3000, whereas cells transfected with untagged sCRH-R2α were incubated with rabbit anti-sCRH-R2α (113–143) (31) serum at 1:2000 and goat antirabbit-AlexaFluor 555 (Invitrogen) secondary at 1:3000. Antibody incubations were each performed for 1 h at room temperature. All cells were stained with 300 nm 4′,6-diamidino-2-phenylindole (DAPI) (Invitrogen) for 1 min and washed before mounting coverslips on slides with Prolong Anti-fade Gold (Invitrogen). Confocal microscopy was performed as described in supplemental Materials and Methods.

Receptor binding assay

Receptor binding assays on intact HEK293 cells were performed 48 h after transient transfection using rat [125I]Ucn I (Phoenix Pharmaceuticals, Burlingame, CA) and binding methods similar to those described previously (52,53). See supplemental Materials and Methods for details.

Statistical analysis

Statistical methods (performed with Statview; SAS Institute, Cary, NC), P values, and sample sizes are included in figure legends.

Results

Alternatively spliced sCRH-R2α mRNA is detected in mouse and rat brain regions and in multiple cell lines

The expression of sCRH-R2α mRNA was detected by RT-PCR using primers in exon 3 and exon 7 (Fig. 1A, a and b), which amplify both CRH-R2α and sCRH-R2α fragments. sCRH-R2α mRNA was detected in mouse brain regions (consistent with previous studies) (31) and in several murine cell lines (αT3-1, LβT2, and CATH.a) known to express CRH-R2α (Fig. 1C) (23,54). Examination of mouse CRH-R2 splice sites revealed that both the splice acceptor and splice donor of exon 6 are of nonconsensus sequence (55,56), perhaps increasing alternative splicing for sCRH-R2α (Fig. 1B). Interestingly, the nonconsensus splice sites surrounding this exon are conserved across multiple species (exon 6 in rat, exon 7 in human), which suggested the alternative sCRH-R2α splice variant might be conserved as well (Fig. 1B). RT-PCR followed by DNA cloning and sequence confirmation revealed the presence of sCRH-R2α in numerous rat brain regions including thalamus, hypothalamus, hippocampus (Fig. 1C), midbrain, medulla/pons, cortex, and cerebellum (data not shown). However, we were unable to detect sCRH-R2α in human brain or SH-SY5Y cDNA samples, both of which express detectable CRH-R2α (data not shown). Instead, we identified fragments corresponding to other human CRH-R2α splice variants that have similar features to sCRH-R2α (exclusion of human exon 6 or exons 6–8 resulting in a frameshift and PTC, data not shown).

Figure 1.

Figure 1

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Genomic structure of CRH-R2 and expression of sCRH-R2α mRNA. A, Schematic of the CRH-R2 gene (top) and α-isoform splice variants, CRH-R2α (middle) and sCRH-R2α (bottom). Gray boxes denote translated sequence, and white boxes represent untranslated regions. CRH-R2α includes contiguous splicing of exons 3–14, whereas sCRH-R2α skips exon 6 (black highlighted box). Sequences encoding the ligand-binding extracellular domain and transmembrane domains in CRH-R2α are indicated. Primers used for RT-PCR and qRT-PCR are positioned schematically to represent their annealing sites (see supplemental Table 1 for PCR primer sequences and product sizes). B, Conservation of nonconsensus splice site sequences surrounding the exon skipped in sCRH-R2α (exon 6 in mouse and rat, exon 7 in human). CRH-R2 exon sequences (uppercase letters, 5′ and 3′ ends) with flanking intron sequence (lowercase letters) are shown for mouse exons 5–7. Rat and human CRH-R2 sequences are aligned below (•, conserved nucleotides when compared with mouse). The consensus sequence for splice donor and acceptor sites is shown in the middle row, and boxes indicate where CRH-R2 splice sites differ from consensus (y = c or t; r = a or g; n = a, g, c, or t; x = variable #). C, RT-PCR using dual primers (Fig. 1A, a and b) demonstrate expression of CRH-R2α and sCRH-R2α in several mouse cell lines (αT3-1, LβT2, and CATH.a) and in mouse and rat brain regions. Each RT-PCR was replicated at least twice using separate cell/tissue samples, and PCR products were confirmed by sequence analysis. Ctx, Cortex; Hpc, hippocampus; Hypo, hypothalamus; Thal, thalamus. +, PCR on cDNA, −, no-RT control.

The RT-PCR experiments suggested that the sCRH-R2α might be expressed not only at different absolute levels across brain regions but also at varying levels relative to CRH-R2α, which could implicate regulated splicing control (Fig. 1C) (31). To accurately quantify sCRH-R2α and CRH-R2α mRNA, qRT-PCR was used to analyze cDNA from dissected mouse tissue using primers specific for each splice variant. The specificity of the CRH-R2α primer pair resulted from a 3′ primer targeted to exon 6, which is absent in sCRH-R2α (Fig. 1A, primer c). The sCRH-R2α-specific pair employed a 3′ primer complementary to the exon 5/7 boundary (Fig. 1A, primer e). The four nucleotides at the 3′ end of the sCRH-R2α-specific primer were complementary to the end of exon 5; critically, the three terminal nucleotides were not complementary to the end of exon 6, preventing annealing to the exon 6/7 boundary. The specificity of the primer pairs was confirmed by qRT-PCR analysis of RNA isolated from Cos-1 cells transfected with cDNA expression constructs for sCRH-R2α or CRH-R2α (Fig. 2A). Primers for CRH-R2α showed >1 × 105-fold specificity for CRH-R2α over sCRH-R2α cDNA, whereas sCRH-R2α-specific primers showed a >1 × 108-fold specificity for sCRH-R2α over CRH-R2α cDNA. Using these primers, splice variant-specific qRT-PCR of various mouse brain regions [normalized to TATA-binding protein (TBP) and scaled to 1.0] revealed relative mRNA levels for full-length CRH-R2α (Fig. 2B, black bars) consistent with previous findings (23,25). The mRNA expression of sCRH-R2α (Fig. 2B, gray bars) was lower than CRH-R2α in each region, yet still significant. The expression of sCRH-R2α relative to CRH-R2α was significantly different across brain regions, ranging from 4–40% of CRH-R2α mRNA levels (Fig. 2B). Strikingly, the ratio of sCRH-R2α to CRH-R2α was drastically reduced in peripheral tissues such as heart and muscle, where sCRH-R2α was 0.1 and 0.7% of CRH-R2α expression, respectively (data not shown).

Figure 2.

Figure 2

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Quantification of relative CRH-R2α and sCRH-R2α mRNA expression in mouse brain and pituitary. A, Demonstration of CRH-R2α and sCRH-R2α primer pair specificity in qRT-PCR. The table lists Ct (threshold cycle) values from qRT-PCR of Cos-1 cells transfected with cDNA expression vectors for each splice variant. B, Normalized mRNA expression of CRH-R2α and sCRH-R2α splice variants across various mouse tissue regions. Bars (sCRH-R2α, gray; CRH-R2α, black) represent the average expression determined by splice variant-specific qRT-PCR from three independent samples (error bars represent sem). mRNA expression was normalized to TBP and adjusted for an axis scale of 1. Numerical values above each data set represent the percentage of sCRH-R2α relative to CRH-R2α for that region ± sem (n = 3). Cere, Cerebellum; Ctx, cortex; Hpc, hippocampus; Hypo, hypothalamus; Med/pons, medulla/pons; Midbrn, midbrain; Pit, pituitary; Thal, thalamus.

sCRH-R2α mRNA escapes NMD and is efficiently translated on polysomes

Transcripts containing PTCs, like sCRH-R2α, are often regulated by NMD. In NMD, transcripts are targeted for degradation during a pioneering round of translation, during which a single processing ribosome removes exon junction complexes (EJCs) deposited by the spliceasome machinery until it terminates at a stop codon (32,57). If ribosome termination occurs upstream of an EJC (due to a PTC), proteins recruited to the termination site interact with the intact EJC to recruit various RNA degradation factors for NMD (32). Due to the requirement for a pioneering round of translation, NMD can be prevented by translational inhibitors, such as CHX (34,58). Transcripts normally subjected to NMD will readily increase with CHX treatment. To determine whether sCRH-R2α is degraded by NMD, mRNA levels were measured using splice variant-specific qRT-PCR in LβT2 cells treated with 50 μg/ml CHX. Ribosomal protein L3 splice variant a (RPL3a), which contains a PTC due to alternative splicing, was used as a positive control for NMD in these experiments. Although not previously identified in mouse, the RPL3a splice variant was shown to be regulated by NMD in human and rat cells with splice sites conserved in mouse (58). Indeed, the mouse RPL3a transcript (Fig. 3A, black bars) increased upon inhibition of NMD in LβT2 cells and decreased again upon CHX removal, demonstrating it as a target of NMD. Normalized sCRH-R2α mRNA levels (Fig. 3A, white bars), like CRH-R2α (Fig. 3A, gray bars), were unaffected by CHX treatment compared with control, suggesting sCRH-R2α is not degraded by NMD. Similar results were obtained in CATH.a cells (data not shown).

Figure 3.

Figure 3

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sCRH-R2α mRNA evades NMD regulation and is associated with polysomes. A, Inhibition of NMD by CHX. qRT-PCR was performed on LβT2 cells treated with 50 μg/ml CHX for the indicated times. For 3 CHX 8 Relief, cells were treated for 3 h with CHX followed by incubation in CHX-free medium for 8 h. Expression at each time point was normalized to TBP and expressed relative to the average of duplicate untreated controls (0 h) for each experiment. Bars represent the average expression from independent experiments ± sem (1 or 11 h CHX, n = 1; 3 h CHX, n = 5; 6 h CHX, n = 3; 3 h CHX 8 h Relief, n = 6). The 0-, 3-, and 6-h values for each mRNA were analyzed by ANOVA (RPL3a, P < 0.005) followed by Scheffé post hoc analysis; values statistically different from 0 h are indicated: *, P < 0.01. Results were replicated in CATH.a cells (data not shown). B, Polysome analysis of sCRH-R2α mRNA. Lysates from LβT2 cells were separated across 5–50% sucrose gradients without EDTA (−EDTA) for fractionation of monosomes and polysomes as measured by A260 (bottom panel, broad peak indicates polysome fractions; flanked by dashed vertical lines). Select fractions were processed for qRT-PCR of sCRH-R2α (top panel, closed circles/solid line), CRH-R2α (top panel, open squares/dashed line), RPL3a (middle panel, open circles/dashed line), and CypA (middle panel, closed squares/solid line). Relative values for each fraction were calculated by the equation R = (E)(Ctref−Ct), where Ctref is the lowest cycle threshold (highest mRNA) fraction for that gene. LβT2 lysates were also separated on gradients containing 20 mm EDTA (+EDTA), which dissociates ribosomes, to demonstrate that the observed mRNA profiles (−EDTA) were dependent on ribosome association. The representative results shown were replicated twice in LβT2 cells and once in CATH.a cells and in mouse whole brain lysates.

To determine whether sCRH-R2α mRNA is efficiently translated, polysome profiles were performed. Efficiently translated transcripts are associated with polysomes, whereas those degraded by NMD are generally associated with monosomes because they are degraded during the pioneering round of translation, before loading of additional ribosomes (57). Lysates from LβT2 cells were separated over a 5–50% sucrose gradient (with or without EDTA), and isolated fractions were measured for an A260 polysome profile followed by qRT-PCR for sCRH-R2α, CRH-R2α, and controls. A duplicate gradient containing 20 mm EDTA, which causes ribosomes to dissociate, was used as a control to demonstrate that the normal mRNA profiles (without EDTA) were caused by ribosome association. Figure 3B shows the resulting profile with sCRH-R2α mRNA (closed circles/solid line) most abundant in polysome fractions, albeit at lower fraction numbers than CRH-R2α (open squares/dashed line), suggesting it is efficiently translated. It should be noted that the lower peak number for sCRH-R2α (fractions 8 and 9) compared with CRH-R2α (fraction 10) resulted from a shorter translational length (5′-untranslated region to stop codon) for sCRH-R2α (∼0.68 kb) compared with CRH-R2α (∼1.4 kb). Cyclophilin A (CypA) (Fig. 3B, closed squares/solid line), which is efficiently translated and has a similar translational length (∼0.54 kb) to sCRH-R2α, closely matched the sCRH-R2α peak in fractions 8 and 9. Again, RPL3a was used as a NMD-regulated positive control (Fig. 3B, open circles/dashed line), and although it has a similar translational length to sCRH-R2α and CypA at about 0.54 kb, it was found predominantly in monosome fractions. These results were replicated in CATH.a cells and total mouse brain (data not shown) and indicate that sCRH-R2α mRNA is not degraded by NMD and is associated with translating ribosomes, suggesting efficient production of sCRH-R2α protein in both cell lines and in vivo.

sCRH-R2α protein does not traffic to the secretory pathway and is degraded by the proteasome

For translated sCRH-R2α to function as a soluble decoy receptor, it must traffic through the secretory pathway and be secreted from the cell. Lacking any confirmed ER translocation motifs, it was unclear whether sCRH-R2α protein would be appropriately trafficked for secretion. Initial trials to detect sCRH-R2α in concentrated media or lysates from cells transfected with sCRH-R2αV5 (sCRH-R2α with C-terminal V5-His fusion tag) were unsuccessful at detecting protein (Fig. 4A, lanes 3 and 7), even though abundant mRNA levels were confirmed by qRT-PCR and CRH-BP expressed from the same vector (CRH-BPV5) was readily detected in lysates and cell media (Fig. 4A, lanes 1 and 5). We therefore hypothesized that the sCRH-R2αV5 protein might be excluded from the ER, which could cause misfolding and targeting for degradation. Upon inhibition of the proteasome with MG132, sCRH-R2αV5 protein levels increased in lysates (Fig. 4A, lanes 3 and 4, and Fig. 4B, lanes 2 and 3), whereas those of CRH-BPV5 (Fig. 4A, lanes 1 and 2, and Fig. 4B, lanes 5 and 6) and CRH-R2αV5 (Fig. 4B, lanes 8 and 9) did not (Fig. 4D shows quantified protein levels from Western analysis). These results were replicated several times in both Cos-1 and αT3-1 cell lines and with the nonpeptide proteasome inhibitor, lactacystin (data not shown). Although sCRH-R2α levels increased with proteasome inhibition in cell lysates, importantly, sCRH-R2αV5 remained undetected in concentrated media (Fig. 4A, lanes 7 and 8). Also, treatment of lysates with PNGase F, an N-linked glycosylase, demonstrated that, unlike CRH-R2αV5 and CRH-BPV5, sCRH-R2αV5 was unglycosylated (Fig. 4B). Because known N-linked glycosylation sites in the CRH-R2α N-terminal region are also encoded by sCRH-R2α, the lack of sCRH-R2α glycosylation suggested that sCRH-R2α protein was not maintained in the ER where these modifications occur.

Figure 4.

Figure 4

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sCRH-R2α fails to be secreted and is degraded by the proteasome. A, SDS-PAGE and Western blot of lysates or concentrated media from Cos-1 cells transfected with the indicated expression constructs and treated with 20 μm MG132 (or vehicle, DMSO) to inhibit the proteasome. Anti-V5 blots (top) were reprobed for β-tubulin (bottom) as a protein loading control. Approximate molecular masses are as follows: CRH-BPV5, 45 kDa; sCRH-R2α, 20 kDa; β-tubulin, 55 kDa. B, Anti-V5 Western blot analysis of lysates from αT3-1 cells transfected with the indicated expression constructs and treated with 20 μm MG132 (or vehicle, DMSO) and/or the N-linked glycosidase PNGase F to determine proteasome sensitivity and glycosylation state, respectively. sCRH-R2αV5 was detected at about 20 kDa in an unglycosylated (u) state, CRH-BPV5 was about 45 kDa glycosylated (g) and about 41 kDa unglycosylated, and CRH-R2αV5 was about 80 kDa glycosylated and about 50 kDa unglycosylated. Blots were reprobed for β-tubulin as a loading control. C, Anti-sCRH-R2α (113–143) Western blot of lysates or concentrated media from Cos-1 cells transfected with untagged sCRH-R2α or a N13A mutant of sCRH-R2αV5 and treated with 20 μm MG132 (or vehicle, DMSO). N13A-sCRH-R2αV5 was about 30 kDa glycosylated and about 20 kDa unglycosylated and untagged sCRH-R2α was about 16 kDa in an unglycosylated state only. Blots were reprobed for β-tubulin as a loading control. D, Effect of proteasome inhibition on protein expression. Protein levels from B and C were quantified with ImageJ and normalized to β-tubulin and transfection efficiency to show relative protein expression with and without proteasome inhibition (±MG132). Results are representative of replicated trials and consistent with experiments using an alternate proteasome inhibitor, lactacystin (data not shown). BP, CRH-BP; R2α, CRH-R2α; sR2α, sCRH-R2α.

Supporting the Western analysis, immunofluorescence of Cos-1 cells cotransfected with eGFP and sCRH-R2αV5 showed few detectable sCRH-R2αV5-positive cells without proteasome inhibition. Under MG132 treatment, the number of detectable sCRH-R2αV5-positive cells increased by about 30-fold (data not shown) and sCRH-R2αV5 showed colocalization with eGFP in the cytoplasm and nucleus but not ER or Golgi (Fig. 5, first row). Cytoplasmic and nuclear localization is consistent with exclusion of sCRH-R2αV5 from the secretory pathway and free diffusion of sCRH-R2αV5 through the nuclear pore due to its small size (59). This localization was not the result of MG132 treatment because the few sCRH-R2αV5-positive cells present without MG132 treatment mimicked this localization pattern (data not shown). In contrast, CRH-R2αV5 (membrane receptor) and CRH-BP-eGFP (marker for secreted protein) showed localization to ER and Golgi within the secretory pathway (Fig. 5, second row), and the localization and number of signal-positive cells was unchanged by MG132 treatment (data not shown). In addition to ER and Golgi, CRH-R2αV5 showed plasma membrane expression (Fig. 5, second row), which was not detected for sCRH-R2α or CRH-BP-eGFP.

Figure 5.

Figure 5

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sCRH-R2α is not localized to the secretory vesicles but rather to the cytoplasm and nucleus. Cos-1 cells were cotransfected with sCRH-R2αV5, CRH-R2αV5, sCRH-R2α, or N13A-sCRH-R2αV5 and eGFP or CRH-BP-eGFP and processed for immunofluorescence as described in Materials and Methods. Images were sequentially recorded for DAPI (blue, first column), eGFP (green, second column), and AlexaFluor 568 or 555 (red, third column) in the same field. Labels above each green or red panel indicate the protein responsible for the signal. Merged images are shown for each row in the fourth column (yellow indicates colocolization of green and red signals). sCRH-R2α with or without the V5-His tag showed localization to the cytoplasm and nucleus, similar to eGFP (rows 1 and 3, respectively), whereas CRH-R2αV5 and the N13A-sCRH-R2αV5 mutant localized to ER and Golgi within the secretory pathway, similar to CRH-BP-eGFP (rows 2 and 4, respectively). Cells in rows 1 and 3 were treated with 20 μm MG132 for 3 h before processing to increase sCRH-R2α protein expression; however, MG132 treatment did not alter localization (data not shown). Scale bars in each DAPI panel represent approximately 20 μm. The representative images shown were replicated through at least two independent experiments in Cos-1 and αT3-1 cells.

Although the C-terminal V5-His tag had no effect on the proper trafficking of CRH-BPV5, CRH-R1V5, or CRH-R2αV5 or the CRH-induced increase in cAMP signaling by CRH-R1V5 or CRH-R2αV5 (data not shown), it was possible that the V5-His tag differentially affected sCRH-R2α expression, folding, and stability. To examine whether the V5-His tag caused the mistrafficked and proteasome-degraded phenotype of sCRH-R2αV5, two approaches were used. First, the expression of an untagged version of sCRH-R2α was examined using an antibody specific to its unique C-terminal tail (31) (provided by W. Vale, The Salk Institute, La Jolla, CA). In transiently transfected cells, untagged sCRH-R2α mimicked the V5-His-tagged version and showed sensitivity to proteasome degradation (Fig. 4C, lanes 1 and 2), a lack of detectable secretion (Fig. 4C, lanes 7 and 8) or glycosylation (data not shown), and localization to the cytoplasm and nucleus (Fig. 5, third row). As a second approach, an N13A mutation, previously shown to enhance the activity of CRH-R2α’s pseudo-signal peptide (38), was introduced into the V5-His-tagged sCRH-R2α (N13A-sCRH-R2αV5). The N13A- sCRH-R2αV5 protein was secreted in a glycosylated state (confirmed by PNGase F treatment, data not shown) in concentrated media (Fig. 4C, lanes 5 and 6), confirming that the native signal peptide was insufficient for proper trafficking. The glycosylated N13A-sCRH-R2αV5 was insensitive to proteasome inhibition; however, an unglycosylated form increased upon proteasome inhibition (Fig. 4C, lanes 3 and 4), indicating that a portion was routed for degradation. Supporting this, immunofluorescence showed predominant localization of N13A-sCRH-R2α to secretory vesicles (Fig. 5 fourth row), with only a slight nuclear/cytoplasmic signal that, unlike the signal in secretory vesicles, increased in propensity and intensity upon MG132 treatment (data not shown). Together, these results indicate that sCRH-R2α protein fails to traffic through the secretory pathway due to an ineffective signal peptide and lack of transmembrane domain and, as a result, is targeted for degradation by the proteasome.

sCRH-R2α protein fails to regulate full-length CRH-R trafficking

CRH-Rs have been implicated in both homo- and heterodimerization (21,60,61,62). Also, truncated and misrouted versions of other GPCRs have been shown to decrease the membrane expression of their full-length counterparts by dimerization and misrouting of the full-length receptor (39,40,41,42,43,44,45). To determine whether sCRH-R2α protein expression could decrease the membrane levels of CRH-R2α, a receptor binding assay on intact HEK293 cells cotransfected with CRH-R2α and equimolar sCRH-R2α or empty vector was performed using [125I]Ucn I as ligand. Figure 6 shows that coexpression of sCRH-R2α did not alter the amount of [125I]Ucn I bound by the CRH-R2α-transfected cells. Binding was also unaffected by transfection of CRH-R2α with 10 molar equivalents of sCRH-R2α or upon treatment of cotransfected cells with 20 μm MG132 to increase sCRH-R2α protein (data not shown). Cells transfected with only sCRH-R2α showed no specific binding (data not shown). Because there is some evidence of interaction between certain CRH-R2 and CRH-R1 proteins (21), the effect of sCRH-R2α coexpression on CRH-R1 membrane binding was also examined. However, no effect was observed in receptor binding assays on HEK293 cells transfected with an equal (Fig. 6) or 10× molar ratio of sCRH-R2α to CRH-R1 (data not shown). Consistent with these results, sCRH-R2α coexpression caused no observable change in the subcellular localization of CRH-R2αV5 or CRH-R1V5 as determined by immunofluorescence (supplemental Fig. 1) and had no effect on Ucn I-induced cAMP signaling through either CRH-R2α or CRH-R1 (supplemental Fig. 2).

Figure 6.

Figure 6

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sCRH-R2α coexpression does not affect CRH-R2α or CRH-R1 membrane binding of [125I]Ucn I. Intact HEK293 cells cotransfected with CRH-R2α or CRH-R1 and sCRH-R2α or empty vector at equimolar ratios were subjected to receptor binding assays with 200 pm rat [125I]Ucn I. Excess unlabeled Ucn-I was used as competitor to determine nonspecific binding. After normalizing for transfection efficiency, the percent maximal binding was determined by dividing the specific binding by the average CRH-R plus vector value (n = 3) in each trial. Bars show the average percent maximal binding ± sem across several independent experiments for cells expressing CRH-R with empty vector (black bars) or sCRH-R2α (gray bars). CRH-R2α + vector and CRH-R2α + sCRH-R2α, n = 9; CRH-R1 + vector, n = 3; CRH-R1 + sCRH-R2α, n = 6. Student’s t test was used to confirm the lack of significant difference between empty vector- and sCRH-R2α-transfected samples (P > 0.8).

Discussion

The alternatively spliced sCRH-R2α transcript, originally identified in mouse (31) and later detected in rat brain (Fig. 1C), esophagus (63), and pituitary (64), encodes the CRH-R2α ligand-binding domain without any transmembrane domains and was therefore predicted to serve as a soluble decoy receptor or alternative binding protein for CRH and Ucn (31). However, legitimate concerns remained: 1) whether sCRH-R2α was efficiently translated in vivo because its mRNA contains a PTC that could target it for NMD (32,33), and 2) whether sCRH-R2α protein trafficked properly for secretion because the effectiveness of its putative signal peptide has been disputed (38). Somewhat surprisingly, inhibition of NMD with CHX (Fig. 3A) and polysome analysis (Fig. 3B) indicated that although sCRH-R2α mRNA contains a PTC, it escapes NMD and is poised for efficient translation through association with polysomes. Several other mRNA transcripts containing a PTC have been identified that also escape NMD (34,35,36,37), and various mechanisms have been proposed for how this occurs. However, as of yet, these mechanisms are insufficient to explain every circumstance, and how sCRH-R2α escapes NMD remains unclear, highlighting the complexity of the NMD pathway.

With indications that sCRH-R2α was efficiently translated in vivo, we evaluated whether sCRH-R2α protein was properly trafficked for secretion. Western analysis (Fig. 4) and immunofluorescence (Fig. 5) of exogenously expressed V5-His-tagged or untagged sCRH-R2α demonstrated that sCRH-R2α protein is not localized to secretory vesicles or secreted. Immunofluorescence data and lack of glycosylation suggest that sCRH-R2α is excluded from the secretory pathway at the point of ER translocation. The sCRH-R2α protein is also highly sensitive to proteasome degradation, possibly because it is misfolded outside of the ER lumen environment. Exclusion of sCRH-R2α from the ER appears to result from an insufficient signal peptide. Rutz et al. (38) demonstrated that although the putative signal peptide of CRH-R2α (and of sCRH-R2α) was predicted at a 0.98 probability, it was insufficient for ER translocation of CRH-R2α, and instead this process was mediated by the first transmembrane domain, which sCRH-R2α lacks. In the same study, an N13A mutation in the pseudo-signal peptide partially rescued CRH-R2α signal peptide function (38), consistent with our findings that the N13A mutation causes sCRH-R2α to become glycosylated, localized to secretory vesicles (Fig. 5), and secreted from the cell (Fig. 4). The improper trafficking and ultimate degradation of sCRH-R2α is consistent with the lack of sCRH-R2α immunohistochemical signal in CRH-R2α-expressing cells in mouse brain (31). The presence of sCRH-R2α immunohistochemical signal in major neuronal sites of CRH-R1 expression (31) may represent cross-reactivity with an alternative splice variant of CRH-R1 (18).

Many GPCRs are capable of both homo- and heterodimerization and, as seen for the class B1 subfamily, show extensive alternative splicing (65). As a result of these features, several examples have been described of protein interaction between separate alternative splice variants of a single gene. Interestingly, these interactions often affect expression or function of either interacting partner (20,21,39,40,41,42,43,44,45). For example, truncated splice variants of LH, GnRH, CRH-R1, and CRH-R2β with altered trafficking were able to misroute their respective canonical receptor (20,39,42,66), prompting us to consider whether sCRH-R2α protein was functioning in a similar manner to regulate the amount of full-length CRH-R2α expressed on the cell membrane, perhaps by recruiting CRH-R2α to the cytoplasm or proteasome. However, receptor binding assays (Fig. 6), immunofluorescence (supplemental Fig. 1), and cAMP signaling assays (supplemental Fig. 2) of cells cotransfected with sCRH-R2α and CRH-R2α demonstrated that sCRH-R2α protein expression did not affect CRH-R2α binding, trafficking, or signaling via cAMP. Similarly, sCRH-R2α had no affect on CRH-R1 signaling or trafficking, even though recent findings (21) showed interaction between a CRH-R1 splice variant (CRH-R1d) and CRH-R2β, suggesting sCRH-R2α might interact with CRH-R1. Interestingly, CRH-R1d, which is normally retained in the cytoplasm, was rescued to the membrane by its interaction with CRH-R2β (21). However, coexpression with either CRH-R2αV5 or CRH-R1V5 was unable to induce membrane expression of sCRH-R2α as determined by immunofluorescence (supplemental Fig. 1). Hence, we have been unable to detect a role for the cytosolic sCRH-R2α protein in modulating CRH-R function.

However, it remains likely that regulated alternative splicing of sCRH-R2α could be functioning to modify full-length CRH-R2α transcript levels, because splicing to sCRH-R2α reduces the pre-mRNA pool available for CRH-R2α transcript production (56). Interestingly, regulated alternative splicing has been suggested for CRH-R1 and CRH-R2β. CRH-R1 splice variants show differential regulation and expression in myometrium during pregnancy and the onset of labor (67,68) as well as preferential production in human skin upon environmental stimuli, such as UV exposure (18,20). For CRH-R2β, chronic variable stress increases mRNA expression of an alternative splice variant while decreasing the canonical form (66). Examinations to date of physiological stimuli known to regulate CRH-R2α mRNA expression, such as glucocorticoids (i.e. dexamethasone) treatment (23,69), showed no change in the relative expression of sCRH-R2α vs. CRH-R2α mRNA (R. Evans, unpublished data). Instead, both splice variants were equally affected by dexamethasone in CATH.a cells, suggesting the change in CRH-R2α mRNA expression induced by dexamethasone is due only to transcriptional regulation (23) and not alterations in splicing control. However, the variations in relative expression of sCRH-R2α to CRH-R2α mRNA across brain regions as determined by qRT-PCR in this study (Fig. 2B) lend initial support for regulated alternative splicing of sCRH-R2α, and it remains likely that regulated splicing control exists for sCRH-R2α under other unexplored conditions.

In conclusion, these studies extended the identification of sCRH-R2α mRNA expression to several murine cell lines and rat brain regions and quantified sCRH-R2α expression across mouse brain regions. Studies also demonstrated that the sCRH-R2α transcript escapes NMD and is efficiently translated, regardless of containing a PTC. However, due to an ineffective signal peptide, the protein is not trafficked for secretion and is largely degraded by the proteasome. Unlike several other truncated receptors, sCRH-R2α protein does not appear to alter trafficking, membrane binding, or signaling of the full-length receptors. Instead, regulation of alternative splicing in different cellular environments or under varying regulatory or developmental conditions may allow splicing of the alternative transcript to alter functional levels of the full-length CRH-R2α mRNA and subsequent protein.

Supplementary Material

[Supplemental Data]
en.2009-0285_index.html (2.9KB, html)

Acknowledgments

We thank Adina Williams for assistance with human sCRH-R2α and mouse sCRH-R2β RT-PCRs, Stephanie Garbern and Ruchira Srinivasakrishnan for assistance with cloning of expression vectors, Travis Williams for production of the CRH-BP-eGFP plasmid, Linda Gates for assistance with cell culture, Shirley Huang and James Beals for technical consultation, and Gwen Stinnett for discussions. We thank Dr. Wylie Vale (The Salk Institute, La Jolla, CA) for the generous gift of anti-sCRH-R2α (113–143) rabbit serum, Dr. Pamela Mellon (University of California, San Diego, CA) for kindly providing αT3-1 and LβT2 cell lines, and Dr. Robert Thompson (University of Michigan, Ann Arbor, MI) for kindly providing rat brain tissue and human brain cDNA samples.

Footnotes

This work was supported by National Institutes of Health (NIH) Grant T32 GM07544 and NIH T32 HD07048 (R.T.E.), NIH DK42730 and DK57660 (A.F.S.), and Michigan Diabetes Research and Training Center Cell and Molecular Biology Core NIH DK20572.

The NMD regulation of sCRH-R2α and protein expression analysis of sCRH-R2αV5 were presented in poster format at The Endocrine Society Meeting 2008 in San Francisco, CA, and at the American Neuroendocrine Society Meeting 2008 in San Rafael, CA.

Disclosure Summary: The authors have nothing to disclose.

First Published Online June 11, 2009

Abbreviations: CHX, Cycloheximide; CRH-BP, CRH-binding protein; CRH-R1, CRH receptor 1; CypA, cyclophilin A; DMSO, dimethylsulfoxide; EJC, exon junction complex; ER, endoplasmic reticulum; GPCR, G protein-coupled transmembrane receptor; NMD, nonsense-mediated RNA decay; PBS-D, Dulbecco’s PBS; PNGase F, peptide N-glycosidase F; PTC, premature termination codon; qRT-PCR, quantitative real-time PCR; RPL3a, ribosomal protein L3 splice variant a; TBP, TATA-binding protein; Ucn, urocortin.

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Supplementary Materials

[Supplemental Data]
en.2009-0285_index.html (2.9KB, html)
en.2009-0285_1.pdf (88.5KB, pdf)
en.2009-0285_2.pdf (164.6KB, pdf)
en.2009-0285_3.pdf (192.3KB, pdf)
en.2009-0285_4.pdf (122.1KB, pdf)

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