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Abstract

We have developed a novel reporter assay that leverages SNAP-epitope tag/near-infrared (NIR) imaging technology to monitor G protein-coupled receptor (GPCR) degradation in human cell lines. N-terminal SNAP-tagged GPCRs were subcloned and expressed in human embryonic kidney (HEK) 293 cells and then subjected to 24 h of cycloheximide (CHX)-chase degradation assays to quantify receptor degradation half-lives (t_1/2) using LICOR NIR imaging–polyacrylamide gel electrophoresis (PAGE) analysis. Thus far, we have used this method to quantify t_1/2 for all nine adrenergic (ADRA1A, ADRA1B, ADRA1D, ADRA2A, ADRA2B, ADRA2C, ADRB1, ADRB2, ADRB3), five somatostatin (SSTR1, SSTR2, SSTR3, SSTR4, SSTR5), four chemokine (CXCR1, CXCR2, CXCR3, CXCR5), and three 5-HT2 (5HT2A, 5HT2B, 5HT2C) receptor subtypes. SNAP-GPCR-CHX degradation t_1/2 values ranged from 0.52 h (ADRA1D) to 5.5 h (SSTR3). On the contrary, both the SNAP-tag alone and SNAP-tagged and endogenous β-actin were resistant to degradation with CHX treatment. Treatment with the proteasome inhibitor bortezomib produced significant but variable increases in SNAP-GPCR protein expression levels, indicating that SNAP-GPCR degradation primarily occurs through the proteasome. Remarkably, endogenous β2-adrenergic receptor/ADRB2 dynamic mass redistribution functional responses to norepinephrine were significantly decreased following CHX treatment, with a time course equivalent to that observed with the SNAP-ADRB2 degradation assay. We subsequently adapted this assay into a 96-well glass-bottom plate format to facilitate high-throughput GPCR degradation screening. t_1/2 values quantified for the α₁-adrenergic receptor subtypes (ADRA1A, ADRA1B, ADR1D) using the 96-well-plate format correlated with t_1/2 values quantified using NIR-PAGE imaging analysis. In summary, this novel assay permits precise quantitative analysis of GPCR degradation in human cells and can be readily adapted to quantify degradation for any membrane protein of interest.

Keywords

G protein-coupled receptor cycloheximide-chase label-free dynamic mass redistribution degradation SNAP

Introduction

It has been estimated that ~30%–40% of all approved medications target the G protein-coupled receptor (GPCR) family of membrane proteins.¹ Thus, understanding the biological basis of GPCR function is essential for human health, as each new revelation of a conserved GPCR process provides a potential interface to be modulated with novel small molecules. One such basic process that has been gathering increasing interest includes GPCR ubiquitin-dependent degradation.^2,3 With the discovery of small molecules that act as “molecular glues” to enhance the interaction affinity between target substrates and their cognate E3 ligases, the potential to enhance or block GPCR degradation with novel small molecules appears tractable.⁴ However, before this nascent field in GPCR drug discovery can be harvested, revealing the precise mechanisms regulating GPCR degradation in human cells is essential.

To date, significant effort has been put forth to identify E3 ubiquitin ligases that govern trafficking and/or degradation of specific GPCR cohorts. Potential GPCR:E3 ligase combinations that have been proposed include MDM2,⁵ Nedd4,⁶ and MARCH2⁷ for β2-AR, AIP4 for CXCR4⁸ and the delta-opioid receptor,⁹ SMURF2 for the mu-opioid receptor,¹⁰ Nedd4-2 for the PAR1 and P2Y1 receptors,¹¹ Cbl for the PAR2 receptor,¹² and Siah1A for metabotropic glutamate receptors 1 and 5.¹³ Given that the human body encodes >800 GPCRs,¹⁴ much work remains to be performed if we are to categorize GPCRs into modules of E3 ubiquitin ligase nodes. Unfortunately, matching GPCRs with their cognate E3 ligases has proven to be experimentally challenging, primarily due to the inherent technical issues associated with performing biochemical assays on membrane proteins. Commercially available anti-GPCR antibodies are often of minimal experimental utility,¹⁵ and as a result, the GPCR field has had to rely heavily on fusion epitope tags to monitor GPCR degradation in cell culture using standard biochemical (i.e., co-immunoprecipitation) and imaging (i.e., confocal fluorescence microscopy) assays. Although useful, these approaches have limited quantitative accuracy and often produce numerous nonspecific artifacts that can reduce the reliability of data sets. Thus, to achieve the goal of pairing all human GPCRs with the E3 ubiquitin ligases that facilitate their degradation, assays that can be performed with relatively high throughput, can produce reliable, reproducible data sets, and are sufficiently sensitive to detect low GPCR expression levels are essential.

Herein, we report the development of a novel cycloheximide (CHX)-chase degradation assay we termed SNAP-GPCR-CHX. This assay leverages the self-labeling SNAP-epitope reporter tag to facilitate the monitoring of SNAP-tagged GPCR protein expression levels using LICOR near-infrared (NIR) imaging–polyacrylamide gel electrophoresis (PAGE) assays. SNAP-GPCR-CHX was validated using a series of essential control experiments that (1) compared endogenous and SNAP-tagged protein degradation rates, (2) determined the effect of altering SNAP-tagged GPCR expression levels on degradation rate, and (3) examined the effects of the proteasome inhibitor bortezomib. Next, we used SNAP-GPCR-CHX to calculate t_1/2 degradation rates for 21 human GPCRs expressed transiently in human embryonic kidney (HEK) cells. SNAP-GPCR degradation rates were subsequently correlated with degradation rates of endogenous GPCRs expressed in cultured cells by monitoring agonist-stimulated label-free dynamic mass redistribution (DMR) functional responses. Finally, we repurposed SNAP-GPCR-CHX into 96-well plates to facilitate high-throughput screening of GPCR degradation. In doing so, we demonstrate that SNAP-GPCR-CHX permits rapid, quantitative, and reproducible determination of GPCR protein half-life in cultured human cells with the potential for high-throughput screening and is thus amenable to identifying degradative mechanisms targeting GPCRs.

Materials and Methods

Chemicals, Constructs, and Cell Lines

HEK293 and SW480 human colon carcinoma cells were purchased from ATCC (Manassas, VA) and grown in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum and 2 mM l-glutamine. Cells were plated at 3 million cells per 10 cm³ dish and transfected with 10 µg of cDNA of select GPCRs expressed in the pSNAP vector using 1 mg/mL polyethyleneimine (PEI). Twenty-four hours posttransfection, cells were trypsinized and replated onto six-well plates for SNAP-CHX-chase assays. Theoretical cDNA constructs were created with aPe-A Plasmid Editor v2.0 software. All pSNAP-GPCR cDNA constructs were created using Takara inFusion molecular cloning kits and sequenced for nucleotide accuracy by Eurofins. pSNAP vector cDNA and SNAP substrate BG-782 were obtained from New England Biolabs (Ipswich, MA). PageRuler Prestained NIR Protein, goat anti-rabbit DyLight 680 antibody, and ladders used for all NIR-PAGE analyses were from Thermo Fisher Scientific (Waltham, MA). Norepinephrine and phenylephrine were obtained from Sigma Inc. (St. Louis, MO)

SNAP-CHX-Chase Assays

Forty-eight hours posttransfection, 50 µg/mL CHX was added to cultured cells. Cells were treated for 0–6 and 24 h. Cells were then counted with the Corning cytosmart, diluted to ensure uniformity across samples, and immediately lysed in 50 mM Tris, pH 7.4, 150 mM NaCl 0.1% Tween-20, 1% NP40 buffer at 4 °C. Cell lysates were subsequently treated with 0.5 µM SNAP substrate BG-782 and subjected to PAGE. Next, GPCR monomer band fluorescent intensities at 800 nm were quantified with the LI-COR Clx NIR imaging system (Lincoln, NE). PAGE gels were Coomassie stained to quantify total protein levels per gel lane in order to normalize each data point by total protein per sample. For β-actin SNAP-CHX assays, polyacrylamide gels were transferred to nitrocellulose blots, blocked with 5% bovine serum albumin, and incubated overnight at 4 °C with 1:1000 rabbit anti-β-actin monoclonal antibody from Cell Signaling Technologies (Danvers, MA). Nitrocellulose was then washed 3× with PBST and incubated for 1 h with 1:5000 goat anti-rabbit DyLight 680 antibody.

96-Well-Plate SNAP-CHX-Chase Assays

Following transient transfection using the conditions described in “Chemicals, Constructs, and Cell Lines,” HEK293 cells were trypsinized and plated at ~100,000 cells/well into 96-well glass-bottom plates. CHX (50 µg/mL) was added to each well and incubated for 0–6 h. Cells were then fixed with 4% paraformaldehyde for 30 min. Next, cells were labeled with 0.5 µM SNAP–substrate BG-782 for 30 min at 37 °C in darkness. Cell nuclei were stained with To-Pro-3. Plates were then imaged with LI-COR Odyssey Clx NIR imaging in the 800 nm channel to measure SNAP-GPCR protein expression. To-Pro-3 nuclear stain signals were measured in the 700 nm channel and used as a normalization control for each well.

Label-Free DMR Assays

Cells were counted with the Corning cytosmart counter and plated uniformly at ~50,000 cells/well in 384-well Corning Epic microsensor plates. Cells were cultured for 24 h and then mounted on the Corning Epic BT reader at 37 °C. Baseline DMR measurements were recorded for 1 h. Compounds were added with the Sorenson Biosciences 96-well Benchtop Pipettor and agonist DMR responses were recorded for 1 h. Data were exported to Microsoft Excel using Epic Analyzer Software for analysis with GraphPad Prism software (La Jolla, CA).

Data Analysis

All PAGE SNAP-CHX-chase data were analyzed with GraphPad Prism software and are expressed as percent band intensity per total protein as measured by Coomassie staining. All 96-well-plate data are expressed as %800 nm/%700 nm signal values in each individual well.

Results and Discussion

The SNAP-tag is a 20 kDa mutant of the DNA repair protein O⁶-alkylguanine-DNA alkyltransferase that can be covalently bound by benzylguanine derivatives linked to an array of synthetic probes. We and others have utilized SNAP-tag technology to monitor GPCR protein expression and localization in vitro.^16–25 Here, we investigated the utility of incorporating SNAP-tag technology into traditional CHX-chase assays to permit calculation of GPCR protein half-lives when expressed in cultured human cells. Our experimental workflow involved fusing the SNAP-tag to the N-terminal domain of select GPCRs, transiently transfecting SNAP-GPCR cDNA constructs into HEK293 cells, and then treating cells with 50 µg/mL CHX for 0–24 h. Cells were then lysed and treated with the irreversible SNAP substrate BG-782, subjected to PAGE, and analyzed with LICOR Odyssey NIR quantitative analysis.

To begin, we examined the utility of performing SNAP-GPCR-CHX studies on the adrenergic family of GPCRs, which are clinical targets for numerous cardiovascular²⁶ and central nervous system disorders.²⁷ Our previous studies have demonstrated the utility of using SNAP-tag labeling to monitor adrenergic receptor (AR) protein expression and agonist-stimulated functional responses in label-free DMR assays.¹⁶ Shown in Figure 1 are representative SNAP PAGE images of CHX-chase assays performed in HEK293 cells transiently expressing the α₁-AR (α_1A-AR/ADRA1A [ Fig. 1A ], α_1B-AR/ADRA1B [ Fig. 1B ], α_1D-AR/ADRA1D [ Fig. 1C ]), α₂-AR (α_2A-AR/ADRA2A [ Fig. 1E ], α_2B-AR/ADRA2B [ Fig. 1F ], α_2C-AR/ADRA2C [ Fig. 1G ]), and β-AR (β1-AR/ADRB1 [ Fig. 1I ], β2-AR/ADRB2 [ Fig. 1J ], β3-AR/ADRB3 [ Fig. 1K ]) receptor subtypes. Each SNAP-GPCR-CHX-chase assay was performed in triplicate, and data were analyzed with GraphPad Prism software to calculate the protein half-life (t_1/2) with nonlinear regression one-phase exponential decay curve fitting ( Fig. 1D,H,L ). Degradation values and the total GPCR degraded at 24 h of CHX treatment are displayed in Table 1 . As shown, the α_1A/ADRA1A subtype degraded the most rapidly with t_1/2 = 0.69 ± 0.12 h, whereas β3/ADRB3 degraded the least rapidly, with t_1/2 = 2.71 ± 0.51 h. Total receptors degraded ranged from 67.9% to 85.8%, with a mean of 74.8% ± 9.0%. To gain further insight into the potential variation of GPCR degradation rates, we subjected members of the 5HT2 serotonin (HTR2A HTR2B, HTR2C) ( Suppl. Fig. S1 ), CXCR chemokine (CXCR1, CXCR2, CXCR3, CXCR5) ( Suppl. Fig. S2 ), and SSTR somatostatin (SSTR1, SSTR2, SSTR3, SSTR4, SSTR5) ( Suppl. Fig. S3 ) receptor families to SNAP-GPCR-CHX assays ( Table 1 ). Similar to the adrenergic family, GPCR degradation t_1/2 values ranged from 0.87 h (SSTR5, CXCR5) to 2.83 h (HTR2A), with maximal 24 h CHX degradation ranging from 75.7% to 92%.

Table 1.

SNAP-GPCR Degradation Half-Lives (t_1/2) as Quantified with SNAP-CHX-Chase Assays (n = 3 or 4).

GPCR	t_1/2 (h), mean ± SEM	% Degraded^a	× BTZ,^b mean ± SEM
ADRA1A	0.69 ± 0.12	77.2	55.6 ± 3.4
ADRA1B	1.15 ± 0.23	85.8	82.9 ± 4.9
ADRA1D	2.14 ± 0.47	78.8	303.0 ± 41.8
ADRA2A	1.84 ± 0.21	67.9	78.6 ± 5.3
ADRA2B	2.55 ± 0.35	83.5	316.9 ± 21.2
ADRA2C	1.69 ± 0.17	55.6	81.1 ± 13.1
ADRB1	2.23 ± 0.46	71.7	397 ± 38.7
ADRB2	0.87 ± 0.18	76.7	136.7 ± 22.4
ADRB3	2.71 ± 0.51	76.3	192.9 ± 34.6
HTR2A	2.83 ± ±0.41	92.0	ND
HTR2B	2.32 ± 0.38	79.0	ND
HTR2C	2.54 ± 0.52	80.2	ND
SSTR1	0.77 ± 0.23	81.1	ND
SSTR2	0.89 ± 0.19	80.2	ND
SSTR3	1.03 ± 0.27	77.7	ND
SSTR4	1.09 ± 0.19	82.3	ND
SSTR5	0.87 ± 0.24	80.4	ND
CXCR1	1.20 ± 0.39	77.8	ND
CXCR2	1.26 ± 0.31	78.7	ND
CXCR3	1.99 ± 0.26	75.7	ND
CXCR5	0.87 ± 0.22	79.3	ND

ND, not determined.

Percent SNAP-GPCR degraded following 24 h of treatment with 50 µg/mL CHX.

Fold increase in SNAP-GPCR expression following 24 h of treatment with 1 µg/mL bortezomib (× BTZ).

Figure 1.

CHX-chase of SNAP-tagged ARs in HEK293 cells. Representative images of polyacrylamide gels displaying SNAP-GPCR protein levels in HEK293 cell lysates expressing the α₁-ARs: (A) ADRA1A, (B) ADRA1B, (C) ADRA1D; the α₂-ARs: (E) ADRA2A, (F) ADRA2B, (G) ADRA2C; or the β-ARs: (I) ADRB1, (J) ADRB2, (K) ADRB3. Each receptor is displayed by gene name. For each gel, the GPCR monomer band intensity was used to quantify protein expression (denoted with an asterisk). The duration of 50 µg/mL CHX treatment is shown in hours. Data were quantified as mean ± SEM to calculate degradation t_1/2. Graphs represent data examining the (D) α₁-AR, (H) α₂-AR, and (L) β-AR subfamilies. All data are the mean ± SEM of three or four experiments.

Next, we tested if the observed GPCR degradation rates were affected by nonspecific artifacts induced by the presence of the SNAP-tag. First, CHX-chase degradation assays were performed on HEK293 cells expressing empty pSNAP vector. As shown in Figure 2A,B , SNAP is remarkably stable in HEK293 cells, with no significant degradation occurring with 24 h of CHX treatment. As a further control, we tested the effects of fusing the SNAP-tag to β-actin, a protein reported to be relatively stable in human cells, and thus often used as a loading control for Western blotting experiments. NIR-PAGE analysis revealed that SNAP-tagged β-actin was stable with CHX treatment ( Fig. 2C , green bands). By leveraging the stringent specificity of commercial antibodies to β-actin, we simultaneously co-monitored CHX-chase degradation of endogenous β-actin. This was achieved by transferring PAGE samples to nitrocellulose after imaging with the LICOR Odyssey to detect SNAP β-actin and then performing Western blotting with an anti-β-actin primary antibody and Alexa-Fluor 700 secondary antibody. As shown, endogenous β-actin ( Fig. 2C , red bands) stability in CHX-chase assays is equivalent to that observed with SNAP β-actin ( Fig. 2C , green bands) (quantification shown in Fig. 2D ). Taken together, these preliminary findings indicate that the SNAP-tag does not affect protein stability and that degradation rates of SNAP-tagged proteins are likely similar to those observed with their cognate endogenous proteins.

Figure 2.

CHX-chase of pSNAP and β-actin in HEK293 cells. (A) Representative image of polyacrylamide gel for HEK293 cell lysates expressing empty pSNAP vector. (B) Degradation data are quantified as mean ± SEM. (C) Image of nitrocellulose blot stained for SNAP β-actin with SNAP substrate BG-782 and anti-β-actin antibody. (D) Data are quantified as mean ± SEM. All data are the mean of three experiments.

GPCRs are degraded by both E1/E2/E3 ubiquitin/proteasome-dependent and -independent (i.e., lysosomal) mechanisms.²⁸ To delineate the contribution of ubiquitin- and non-ubiquitin-dependent mechanisms to GPCR degradation, HEK293 cells expressing SNAP-GPCRs were treated with bortezomib, a selective inhibitor of the 26S proteasome that is FDA approved to treat multiple myeloma and lymphoma. If specific SNAP-tagged GPCRs are degraded by the ubiquitin/proteasome pathway, we would expect bortezomib treatment to enhance SNAP-GPCR protein expression levels following 24 h treatment. Figure 3 compares the effects of 1 µM bortezomib on HEK293 cells expressing the α₁-AR (α_1A-AR/ADRA1A [ Fig. 3A ], α_1B-AR/ADRA1B [ Fig. 3B ], α_1D-AR/ADRA1D [ Fig. 3C ]), α₂-AR (α_2A-AR/ADRA2A [ Fig. 3e ], α_2B-AR/ADRA2B [ Fig. 3F ], α_2C-AR/ADRA2C [ Fig. 3G ]), and β-AR (β1-AR/ADRB1 [ Fig. 3I ], β2-AR/ADRB2 [ Fig. 3J ], β3-AR/ADRB3 [ Fig. 3K ]) receptor subtypes. Remarkably, bortezomib enhanced protein expression levels of all nine AR subtypes with rank order ADRB1 > ADRA2B > ADRA1D > ADRB3 > ADRB2 > ADRA1B > ADRA2C > ADRA2A > ADRA1A (Fig. 3D,H,L, Table 1). To assess the potential contribution of lysosomal degradation pathways, HEK293 cells expressing SNAP-ADRA1D were treated with the lysosomal degradation inhibitors NH₄Cl, chloroquine, leupeptin, or all three combined (ALL) ( Suppl. Fig. S4 ). As shown, none of the lysosomal inhibitors administered alone or in combination produced any discernable increase in ADRA1D protein expression levels. Taken together, these findings indicate that the AR family of GPCRs is primarily degraded by ubiquitin/ERAD/proteasomal-dependent mechanisms in HEK293 cells.

Figure 3.

Effects of the proteasome inhibitor bortezomib on SNAP-GPCR protein expression in HEK293 cells. Representative images of polyacrylamide gels displaying SNAP-GPCR protein levels in HEK293 cell lysates expressing the α₁-ARs: (A) ADRA1A, (B) ADRA1B, (C) ADRA1D; the α₂-ARs: (E) ADRA2A, (F) ADRA2B, (G) ADRA2C; or the β-ARs: (I) ADRB1, (J) ADRB2, (K) ADRB3. Each receptor is displayed by gene name. For each gel, GPCR monomer band intensity was used to quantify protein expression (denoted with an asterisk). The duration of 1 µm bortezomib treatment is shown in hours. Data were quantified as mean ± SEM to calculated degradation t_1/2 values. Graphs represent data examining the (D) α₁-AR, (H) α₂-AR, and (L) β-AR subfamilies. All data are the mean of three experiments.

One potential limitation of this novel approach is the reliance on ectopic SNAP-GPCRs, which may produce results that are not entirely indicative of the cellular or molecular mechanisms degrading endogenous GPCRs in vivo. Thus, we examined the effects of CHX-induced degradation of endogenously expressed GPCRs by measuring agonist-stimulated label-free DMR responses in cultured cells. In previous studies, we determined that HEK293 cells endogenously express functional β-ARs using label-free DMR Schild regression analysis.¹⁶ Twenty-four-hour CHX treatment reduced protein expression levels of β-ARs by >70% in SNAP-GPCR-CHX assays ( Fig. 1L ). If these findings are indicative of endogenous β-AR degradation rates, we predict that 24 h CHX treatment will produce significant decreases in agonist efficacy by decreasing β-AR receptor reserve. This was tested by quantifying label-free DMR responses for the endogenous AR neurotransmitter norepinephrine in cultured HEK293 cells in the absence or presence of 24 h of CHX treatment. Figure 4A demonstrates that CHX treatment reduces both norepinephrine potency by ~4× (untreated EC₅₀ = 1.78 ± 0.06 µM vs CHX EC₅₀ = 7.18 ± 0.27 µM) and maximal responses by ~68%. Next, we examined CHX degradation in SW480 human colon carcinoma cells, which we previously determined to express primarily the α_1B-AR subtype, with minimal to no expression of the α_1A and α_1D subtypes.¹⁷ SNAP-CHX assays reveal that SNAP-α_1B-AR degrades with t_1/2 = 0.77 ± 0.23 h and is ~75% degraded at 24 h ( Fig. 4B,C ). Twenty-four hours of CHX treatment of untransfected SW480 cells expressing endogenous α_1B-ARs decreased the potency of the selective α₁-AR agonist phenylephrine (untreated EC₅₀ = 15.8 ± 0.02 mM vs CHX EC₅₀ = not determined) and maximal responses by ~99% ( Fig. 4D ). Thus, these findings suggest that SNAP-GPCR-CHX assays are indicative of endogenous GPCR degradation rates.

Figure 4.

CHX treatment reduces GPCR functional responses in cultured human cells. (A) Label-free DMR concentration–response curves for norepinephrine in untreated cells and HEK293 cells treated with 50 µg/mL CHX for 24 h. DMR responses obtained at 30 min of norepinephrine stimulation were used. (B) Representative image of polyacrylamide gel for SW480 cell lysates expressing SNAP-ADRA1B. (C) Degradation data are quantified as mean ± SEM (n = 3). (D) Label-free DMR concentration–response curves for the selective α₁-AR agonist phenylephrine in untreated cells and SW480 cells treated with 50 µg/mL CHX for 24 h. DMR responses obtained at 30 min of phenylephrine stimulation were used. All label-free DMR data are the mean ± SEM of four experiments performed in quadruplicate.

Finally, we adapted the SNAP-GPCR-CHX assay to a 96-well-plate imaging, high-throughput format. In brief, HEK293 cells transfected with SNAP-GPCR subtypes were seeded in glass-bottom 96-well plates and treated with 50 µg/mL CHX for 0–24 h. Cells were then fixed with paraformaldehyde and stained with BG-782 to label SNAP-GPCRs or with TO-PRO-3 to label cell nuclei, as a normalization control. Plates were then imaged and analyzed with LICOR NIR. Shown in Figure 5A is a representative 96-well-plate SNAP-GPCR-CHX assay performed on HEK293 cells expressing the α₁AR subtypes; Figure 5B displays the calculated degradation rates for each subtype, which are quantified as the ratio of 800 nm fluorescence (SNAP substrate BG-782, green) divided by the 700 nm signal (TO-PRO-3, red). In comparison with SNAP-GPCR-CHX degradation rates determined using NIR-PAGE, the 96-well-plate format produced similar half-life values (ADRA1A t_1/2 = 0.83 ± 0.24 h; ADRA1B = 2.48 ± 0.76 h; ADRA1D t_1/2 = 1.33 ± 0.45 h), with a discrepancy in rank order of ADRA1D > ADRA1B, which we suspect is a result of the 96-well plate capturing all ADRA1D bands labeled with BG-782, including N-terminal cleavage products that are resistant to degradation ( Fig. 1C ). Taken together, these findings demonstrate that SNAP-GPCR-CHX assays have the potential to be optimized for high-throughput screens that require GPCR protein expression as a functional output.

Figure 5.

Ninety-six-well-plate CHX-chase assays quantifying SNAP-GPCR protein expression level in HEK293 cells. (A) Representative 96-well plate of HEK293 cells expressing SNAP-ADRA1A, SNAP-ADRA1B, or SNAP-ADRA1D stained with SNAP substrate BG-782 in an 800 nm channel (left plate, green) or To-Pro-3 to stain HEK293 cell nuclei (right plate, red). Cells were treated with 50 µg/mL CHX for the indicated times. (B) Data are quantified as mean ± SEM (n = 3, each with four replicates). Data were normalized for cell number by dividing SNAP-GPCR band intensity (800 nm channel) by To-Pro 3 stain intensity (700 nm channel).

Significant effort has been put forth to decipher the cellular and molecular mechanisms governing GPCR ubiquitin-dependent proteasomal degradation. Unfortunately, progress in the field has been hindered by technical difficulties in performing the currently available biochemical assays on integral membrane proteins like GPCRs. Previous studies have used a wide variety of epitope tags to circumvent these issues, such as HA,^29,30 V5,¹⁰ FLAG,⁵ and Myc–.^30,31 In most approaches, to obtain sufficient GPCR protein signal-to-noise ratios, epitope-tagged GPCRs must first be detergent solubilized and then immunoprecipitated with an affinity resin specific to the fused epitope tag. Standard PAGE and nitrocellulose transfer are performed, followed by immunoblotting with primary and secondary antibodies. Although informative, these biochemical approaches have limited quantitative accuracy, are costly, and are laboriously time-consuming. Alternatively, radiolabeled small-molecule agonists or antagonists can be used to monitor GPCR protein expression levels,^32,33 but require significant exposure time to produce detectable signals and are potentially hazardous.

Thus, the primary goal of this study was to develop a simple, reproducible approach that can be used to quantify degradation half-lives of GPCRs (and possibly other membrane proteins) in high throughput relative to traditional immunoprecipitation/immunoblotting. Previous studies in which the SNAP-tag was leveraged to identify and characterize the α_1D-AR:Scribble:dystrophin-associated protein complex,³⁴ determined the importance of type I PDZ ligands for GPCR functional expression,¹⁵ or facilitated the serendipitous discovery that the α_1D-AR N-terminal domain is endogenously cleaved¹⁹ guided the decision to incorporate the SNAP-tag into CHX-chase assays. Herein, we demonstrated that the SNAP-epitope tag is remarkably stable and does not induce degradation of β-actin, and thus is not likely enhancing GPCR degradation rates. Additional support for this conclusion comes from our previous studies investigating SNAP-tagged GPCR label-free DMR responses, which clearly demonstrated agonist efficacy, and in turn GPCR functional expression is not hindered by the presence of SNAP-tags located in the N-terminal domain.¹⁵ In future studies, leveraging CRISPR gene editing technology to fuse SNAP-tags to endogenous GPCRs should provide significant information about these essential cellular processes.

Incorporating the SNAP-tag into GPCR degradation studies provides significant experimental advantages. Importantly, SNAP-tagged GPCRs are labeled in cell lysates prior to PAGE, permitting direct imaging of GPCR protein bands in acrylamide gels using LICOR NIR imaging, thereby reducing experiment duration, reagent cost, and nonspecific artifact and enhancing data set resolution. The SNAP-tag can also be repurposed for imaging analysis with cellular localization specificity through the use of cell-permeable and -impermeable SNAP substrates,^15,20 or for affinity purification/mass spectrometry of GPCR signaling complexes.¹⁸ Thus, we propose that SNAP-GPCR-CHX assays are amenable for high-throughput monitoring of GPCR protein expression, and thus applicable for studies examining mechanisms governing GPCR degradation, and potentially to screen for small molecules that can enhance or inhibit GPCR degradation. Future experiments to be performed using our de novo 96-well-plate SNAP-GPCR-CHX assay will include testing the effects of proteasome inhibitors (i.e., bortezomib) on GPCR degradation and quantifying GPCR resynthesis rates following CHX washout. These experiments will facilitate the long-term goal of adapting SNAP-NIR CHX assays to identify and characterize the degradation mechanisms of non-GPCR membrane proteins, such as tyrosine kinase receptors, ion channels, adhesion molecules, and others.

Supplemental Material

sj-pdf-1-jbx-10.1177_2472555220979793 – Supplemental material for Development of a Novel SNAP-Epitope Tag/Near-Infrared Imaging Assay to Quantify G Protein-Coupled Receptor Degradation in Human Cells

Supplemental material, sj-pdf-1-jbx-10.1177_2472555220979793 for Development of a Novel SNAP-Epitope Tag/Near-Infrared Imaging Assay to Quantify G Protein-Coupled Receptor Degradation in Human Cells by Kyung-Soon Lee, Edelmar Navaluna, Nicole M. Marsh, Eric M. Janezic and Chris Hague in SLAS Discovery

Footnotes

Supplemental material is available online with this article.

Declaration of Conflicting Interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: Funding for this research was provided by NIGMS T32GM007750 (E.M.J.) and R01GM100893 (C.H.).

ORCID iD

Nicole M. Marsh

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