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. Author manuscript; available in PMC: 2020 Jan 3.
Published in final edited form as: Methods Cell Biol. 2018 Sep 17;149:87–105. doi: 10.1016/bs.mcb.2018.08.007

Detection of misfolded rhodopsin aggregates in cells by Förster resonance energy transfer

Megan Gragg 1, Paul S–H Park 1,*
PMCID: PMC6941733  NIHMSID: NIHMS1064216  PMID: 30616829

Abstract

Rhodopsin is the light receptor in rod photoreceptor cells of the retina that plays a central role in phototransduction and rod photoreceptor cell health. Rhodopsin mutations are the leading known cause of autosomal dominant retinitis pigmentosa, a retinal degenerative disease. A majority of rhodopsin mutations cause misfolding and aggregation of the apoprotein opsin. The nature of aggregates formed by misfolded rhodopsin mutants and the associated cell toxicity is poorly understood. Misfolding rhodopsin mutants have been characterized biochemically, and categorized as either partial or complete misfolding mutants. This classification is incomplete and does not provide sufficient information to fully understand rhodopsin aggregation, disease pathogenesis, and evaluate therapeutic strategies. To better understand the aggregation of misfolded rhodopsin mutants, a Förster resonance energy transfer assay has been developed to monitor the aggregation of fluorescently tagged mutant rhodopsins expressed in live cells.

Keywords: retinitis pigmentosa, retinal degeneration, protein misfolding, protein aggregation, phototransduction, Förster resonance energy transfer

1. Introduction

Rhodopsin is the light receptor in rod photoreceptor cells of the retina and a member of the G protein-coupled receptor (GPCR) family of cell surface proteins. Rhodopsin consists of the apoprotein opsin covalently bound to the chromophore 11-cis retinal, which acts as an inverse agonist. Light causes the isomerization of 11-cis retinal to all-trans retinal, which activates rhodopsin and initiates the G protein-mediated signaling cascade called phototransduction (Park, 2014). Rhodopsin is highly concentrated in the rod outer segment disc membranes of photoreceptor cells where it forms oligomers organized as nanodomains (Liang et al., 2003, Whited and Park, 2015, Rakshit et al., 2015). Rhodopsin is a major constituent of the rod outer segment of photoreceptor cells and is required for the formation of this compartment and for the prevention of retinal degeneration (Lem et al., 1999).

Dysfunction in rhodopsin can cause a variety of retinal diseases. Over 100 mutations in rhodopsin have been determined to cause inherited retinal disease (Athanasiou et al., 2018), most causing retinitis pigmentosa (RP), a progressive retinal degenerative disease. A majority of the mutations in rhodopsin with known biochemical defects result in misfolding and aggregation of the receptor (Mendes et al., 2005, Athanasiou et al., 2018). The first mutation in rhodopsin discovered was the P23H mutation (Dryja et al., 1990), which causes autosomal dominant RP (adRP) and is the most common mutation in the United States. Since this initial discovery, the molecular defects caused by mutations in rhodopsin have been characterized biochemically and by cell biology (Garriga et al., 1996, Hwa et al., 1997, Kaushal and Khorana, 1994, Sung et al., 1991a, Sung et al., 1991b). The mutations that cause misfolding and aggregation have been characterized biochemically by examining the ability of the apoprotein opsin to bind 11-cis retinal using UV-Vis spectroscopy or examining the maturation of the synthesized protein by examining the migration patterns in SDS-PAGE gels. Misfolding mutants exhibit impairment in the binding of 11-cis retinal and in achieving a mature glycosylated form. The mutants have also been characterized by microscopy to determine the localization of the mutant receptors in the cell. Misfolding mutants are improperly trafficked and retained in the endoplasmic reticulum (ER).

Misfolding mutants of rhodopsin can be classified based on biochemical properties and have been subdivided as either complete or partial misfolding mutants (Kaushal and Khorana, 1994, Krebs et al., 2010, Sung et al., 1991a, Sung et al., 1993). Complete misfolding mutants bind little or no 11-cis retinal, are mostly retained in the ER, and cannot be chaperoned by retinoids. Partial misfolding mutants can bind some 11-cis retinal, are variably retained in the ER, can be chaperoned by retinoids, and can traffic properly when rescued by retinoid treatment. Secondary structure changes are observed in both partial and complete misfolding rhodopsin mutants (Miller et al., 2015, Liu et al., 1996). Variability is observed in the biochemical classification of misfolding mutants of rhodopsin (Krebs et al., 2010), and updates are required to include the aggregation properties of the mutants. Misfolding mutations are distributed throughout the structure of rhodopsin (Fig. 1). Complete misfolding mutations tend to be in close proximity to the covalently bound 11-cis retinal whereas partial misfolding mutations tend to be further in proximity to the bound chromophore (Fig. 1B).

Figure 1.

Figure 1.

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Partial and complete misfolding mutations. The secondary structure (A) and crystal structure (B) (PDB: 1U19) of rhodopsin are shown with residues that cause misfolding and adRP when mutated highlighted in blue (partial misfolding mutation), yellow (complete misfolding mutation), or green (partial or complete misfolding mutation, depending on mutation). Residues that cause misfolding and adRP when mutated in the crystal structure are depicted as spheres. 11-cis retinal is depicted as pink spheres. Classification is based on data in (Kaushal and Khorana, 1994, Krebs et al., 2010, Sung et al., 1991a, Sung et al., 1993).

While biochemical and microscopy methods have been useful for the characterization and classification of misfolding mutants of rhodopsin, none of these methods directly assess the aggregation of the misfolding mutants. While photoreceptor cell death is thought to occur, at least in part, because of toxic misfolded rhodopsin aggregates (Athanasiou et al., 2012, Vasireddy et al., 2011, Gorbatyuk et al., 2010, Price et al., 2011, Neri et al., 2010), little is known about the nature of the aggregate formed. Moreover, how misfolding mutants of rhodopsin interact with the wild-type receptor is unclear even though most reported cases of patients with adRP involving misfolding rhodopsin mutants are heterozygous for the mutation. The aggregation properties of misfolding rhodopsin mutants and the propensity of the mutants to physically interact with wild-type receptor must be considered to better understand the pathogenesis of adRP caused by misfolding rhodopsin mutants and to evaluate and formulate potential therapeutic strategies.

A Förster resonance energy transfer (FRET) method has been developed to directly probe the aggregation of rhodopsin mutants and can differentiate between oligomers and aggregates of rhodopsin in cells (Hovan et al., 2010, Miller et al., 2015, Gragg et al., 2016). FRET is non-radiative energy transfer between a donor and acceptor molecule that are less than 10 nm apart and have sufficient overlap between the emission and excitation spectra of the donor and acceptor molecules, respectively (Forster, 2012). The protocol outlined here has been used to characterize the aggregation properties of several misfolding mutants of rhodopsin causing adRP and provide additional molecular insights beyond those provided by biochemical and microscopy methods (Gragg et al., 2016, Gragg and Park, 2018). Studies utilizing this FRET method has revealed that there is variability in the aggregation properties of different misfolding mutants and that further subdivisions may be required among partial misfolding mutants classified biochemically. Furthermore, some misfolding mutants largely do not form physical interactions with the wild-type receptor and a pharmacological chaperone is predicted to be ineffective or even detrimental for some mutants. These studies further advance our molecular understanding about the impact of misfolding mutations in rhodopsin and should lead to a better understanding on the pathogenesis of the disease and how to combat the disease.

2. MATERIALS

Common reagents can be purchased from a variety of vendors. We only list those that have been used in our laboratory to conduct procedures outlined in this protocol.

2.1. GENERATION OF VECTORS FOR FRET

  1. pmTq-N1 (can be obtained at addgene.org)

  2. pYFP-N1 (can be obtained at addgene.org)

  3. Rhodopsin cDNA

  4. Thermal cycler

  5. Primers

  6. Restriction enzymes: EcoRI and BamHI (New England Biolabs, Ipswhich, MA)

  7. Heating block

  8. Alkaline phosphatase, calf intestinal (CIP) (New England Biolabs, Ipswhich MA)

  9. Agarose (VWR Life Science AMRESCO, Solon, OH)

  10. 2-Log DNA Ladder (New England Biolabs, Ipswhich MA)

  11. 6X Gel Loading Dye, Purple (New England Biolabs, Ipswhich MA)

  12. Tris-acetate-EDTA (TAE) buffer: 40 mM Tris base, 20 mM acetic acid, 1 mM EDTA

  13. Ethidium bromide (Thermo Fisher Scientific, Waltham, MA)

  14. Agarose gel electrophoresis apparatus

  15. UV transilluminator

  16. QIAEX II Gel Extraction Kit (Qiagen, Germantown, MD)

  17. T4 DNA Ligase (New England Biolabs, Ipswhich, MA)

  18. 10-beta Competent E. coli – High Efficiency (New England Biolabs, Ipswhich MA)

  19. LB agar (VWR Life Science AMRESCO, Solon, OH)

  20. Kanamycin (MilliporeSigma, St. Louis, MO)

  21. 37 °C bacterial culture incubator

  22. LB Broth (VWR Life Science AMRESCO, Solon, OH)

  23. 14 mL round-bottom cell culture tube (Corning, Corning, NY)

  24. Orbital incubator shaker

  25. QIAprep Spin Miniprep kit (Qiagen, Germantown, MD)

  26. UV-Vis spectrophotometer

  27. UltraPure Distilled Water, nuclease-free (Invitrogen, Carlsbad, CA)

  28. QuickChange II Site-Directed Mutagenesis Kit (Agilent Technologies, Santa Clara, CA).

2.2. CELL CULTURE AND TRANSFECTION

  1. HEK293T/17 cells (American Type Culture Collection, Manassas, VA)

  2. 12-well cell culture plates (Corning, Corning, NY)

  3. Dulbecco’s Modified Eagle Medium, high glucose (DMEM-H), with L-glutamine and sodium pyruvate (Thermo Fisher Scientific, Waltham, MA)

  4. Fetal bovine serum (Thermo Fisher Scientific, Waltham, MA)

  5. Cell culture incubator (37 °C, 5% CO2)

  6. Lipofectamine 2000 Transfection Reagent (Invitrogen, Carlsbad, CA)

  7. Purified DNA vectors (e.g., pRho-YFP and pRho-mTq)

  8. 9-cis retinal (MilliporeSigma, St. Louis, MO)

  9. Dimethyl sulfoxide (DMSO) (Thermo Fisher Scientific, Waltham, MA)

  10. Aluminum foil

2.3. FRET EXPERIMENTS

  1. Spectrofluorometer with circulating water bath

  2. Quartz cuvette and stir bar

  3. 15 mL conical tube

  4. Vortexer

  5. 1X phosphate buffered saline (PBS) (Corning, Manassas, VA)

  6. n-dodecyl-β-D-maltoside (DM) (Anatrace, Maumee, OH)

  7. Sodium dodecyl sulfate (SDS) (Invitrogen, Carlsbad, CA)

  8. Prism (GraphPad Software, La Jolla, CA)

3. METHODS

Rhodopsin must be tagged with appropriate fluorescent proteins for FRET measurements in cells. Tagging rhodopsin at its C-terminus largely does not impact the folding or function of the receptor (Miller et al., 2015, Jin et al., 2003). Cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP), or their variants, are a widely used donor and acceptor pair for FRET studies. We have previously used tagged rhodopsin with the CFP variants SCFP3A, mTurquoise (mTq), and mTurquoise2 (mTq2) and YFP variant SYFP2 (Kremers et al., 2006, Goedhart et al., 2010, Goedhart et al., 2012, Miller et al., 2015, Gragg et al., 2016, Gragg and Park, 2018, Mishra et al., 2016). The protocol described here will focus on the receptor tagged with mTq or SYFP2, which is referred to as YFP for simplicity.

FRET is an indicator of whether or not the tagged rhodopsin molecules are in close enough proximity to form physical interactions (Figs. 2A and 2B). Significant overlap is present between the emission spectrum of mTq and excitation spectrum of YFP (Fig. 2C), making this donor and acceptor pair a good choice for FRET experiments. When mTq- and YFP-tagged rhodopsin are physically interacting (Fig. 2A), FRET can occur between the fluorescent proteins. Two indicators of FRET are observed in the emission spectra obtained after excitation of mTq. The fluorescence from mTq is quenched and a sensitized emission signal from YFP is observed when FRET occurs (Fig. 2D). The FRET efficiency can be determined by quantifying either the quenching of mTq or the sensitized emission signal.

Figure 2.

Figure 2.

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Illustration of FRET. (A) FRET between tagged rhodopsin. When mTq-tagged rhodopsin and YFP-tagged rhodopsin form physical interactions, the fluorescent proteins are in close enough proximity for FRET to occur. Excitation of mTq will result in quenching of the emission signal from mTq and the sensitized emission from YFP. (B) No FRET between tagged rhodopsin. When mTq-tagged rhodopsin and YFP-tagged rhodopsin do not form physical interactions, the fluorescent proteins are not in close enough proximity for FRET to occur. Excitation of mTq will result only in emission from mTq. (C) Fluorescence spectra of mTq and YFP. The excitation (dashed lines) and emission (solid lines) spectra of mTq (blue) and YFP (yellow) are shown. (D) Example of emission spectra in the presence or absence of FRET. The emission spectra obtained after excitation of mTq is shown when FRET occurs (yellow) and when FRET does not occur (blue). When FRET occurs, the signal from mTq is quenched and the sensitized emission from YFP is detected.

The observed FRET efficiency is dependent on the acceptor:donor (A:D) ratio. As the A:D ratio is increased, the number of complexes containing both a donor and acceptor molecule will increase until all donor molecules are in complex with an acceptor molecule (Fig. 3A). For this reason, the observed FRET efficiency will increase with increasing A:D ratios (Fig. 3B). FRET experiments must be conducted at different ratios of mTq- and YFP-tagged rhodopsin in order to generate FRET curves and determine the maximal FRET efficiency (Emax) (Fig. 3C). Computing the Emax allows for quantitative comparisons of FRET among different receptors or mutant forms of the receptor.

Figure 3.

Figure 3.

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Relationship of apparent FRET efficiency and A:D ratio. (A) Cartoon illustrating the complement of complexes containing both an acceptor (yellow) and donor (blue) molecule as the A:D ratio is increased. (B) FRET spectra showing increased apparent FRET efficiency with increasing A:D ratio. The apparent FRET efficiency (E) can be computed from the quenching of the donor or sensitized emission from the acceptor. (C) FRET curves can be generated by plotting the relationship between the apparent E and the A:D ratio. The maximal E (Emax) can be obtained by fitting the data to a rectangular hyperbolic function.

The detection of FRET alone does not mean that tagged rhodopsins physically interact. Rhodopsin is embedded in the membrane of cells, which has a finite surface area. Chance encounters among freely diffusing rhodopsin molecules within a crowded membrane environment that bring the donor and acceptor fluorescent proteins in close enough proximity for FRET can result in non-specific FRET (Fig. 4A). Specific FRET, which derives from physical interactions (Fig. 4B), has to be distinguished from non-specific FRET. In FRET experiments described here, non-specific FRET is defined by control experiments where YFP-tagged rhodopsin is coexpressed with mTq-tagged m2 muscarinic receptor, an unrelated GPCR that should not physically interact with rhodopsin. Only FRET signals that exceed the level of non-specific FRET can be considered to be specific FRET indicative of physical interactions (Szalai et al., 2014, King et al., 2014, Lan et al., 2015).

Figure 4.

Figure 4.

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Non-specific versus specific FRET. (A) Non-specific FRET can occur even when no physical interactions occur between the donor and acceptor molecules, or the molecules they are attached to. Non-specific FRET occurs by chance encounters among freely diffusing donor and acceptor molecules, within a finite area like the cell membrane, that come in close enough proximity for FRET to occur. (B) Specific FRET occurs when the donor and acceptor molecules, or the molecules they are attached to, form physical interactions.

The detection of specific FRET indicates that tagged rhodopsins are physically interacting; however, whether those interactions represent oligomers formed by properly folded rhodopsin or aggregates of misfolded rhodopsin is unknown. Oligomers and aggregates of rhodopsin can be distinguished by treatment with the mild detergent n-dodecyl-β-D-maltoside (DM) (Miller et al., 2015, Gragg et al., 2016). DM has been shown previously to disrupt the oligomers of rhodopsin in photoreceptor cell membranes into monomers (Jastrzebska et al., 2004). As expected, the FRET signal from tagged WT rhodopsin expressed in HEK293 cells is disrupted by DM treatment (Fig. 5A), which indicates that oligomers of tagged WT rhodopsin are disrupted by treating cells with DM similar to rhodopsin from native photoreceptor cells (Fig. 5C). In contrast, the FRET signal from tagged misfolding mutants of rhodopsin cannot be disrupted by DM (Fig. 5B), which indicates that the mutants form aggregates that are resistant to disaggregation by DM (Fig. 5C). The harsher detergent SDS is required to disrupt aggregation and eliminate the FRET signal (Fig. 5B). Thus, the FRET signal that is DM-sensitive can be assigned to oligomers and the FRET signal that is DM-insensitive can be assigned to aggregates. This assignment is supported by SDS-PAGE and confocal microscopy studies (Gragg et al., 2016).

Figure 5.

Figure 5.

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Distinguishing oligomers and aggregates by detergent treatment. (A, B) Emission spectra obtained after excitation at 425 nm of cells coexpressing mTq- and YFP-tagged wild-type rhodopsins (A) or mutant rhodopsins (B). Spectra are obtained on untreated cells (blue), cells treated with DM (red) and then SDS (green). The emission signal at 476 nm is used to compute the various FRET efficiency values. These spectra were adapted from those presented in (Gragg et al., 2016). (C) Detergent treatment of cells expressing wild-type or mutant rhodopsins. Wild-type rhodopsin forms oligomers in untreated cells that become disrupted after DM treatment. Mutant rhodopsin form aggregates intracellularly. The aggregates are not disrupted by DM treatment but become disrupted after SDS treatment.

An overview of FRET experiments described in detail below is provided in Fig. 6. A typical FRET experiment requires the generation of vectors encoding tagged rhodopsin, transfection of cells, and measurement of FRET using a spectrofluorometer.

Figure 6.

Figure 6.

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Overview of collecting FRET data. (A) Vector creation. Vectors containing mTq or YFP preceded with a multiple cloning site allow for insertion of the sequence for rhodopsin to generate a fusion construct of rhodopsin tagged with the fluorescent proteins at its C-terminus. The vectors pmTq-N1 or pYFP-N1 are digested with restriction enzymes and the PCR-amplified sequence for rhodopsin is inserted by ligation procedures. The resulting vectors pRho-mTq and pRho-YFP are used for transfection in cells. (B) FRET experiments. Cells are co-transfected with pRho-mTq and pRho-YFP. 24–28 hours post transfection, cells are harvested and resuspended in PBS. The cell suspension is placed in a cuvette and the fluorescence spectra obtained and analyzed for FRET.

3.1. GENERATION OF VECTORS FOR FRET

  1. The vectors pmTq-N1 and pYFP-N1 are similar to the commercially available vector pECFP-N1 (Clontech Laboratories, Mountain View, CA), except the sequence for ECFP is replaced by either mTq or YFP.

  2. Amplify the sequence for rhodopsin by polymerase chain reaction (PCR) using the appropriate forward and reverse primers and cDNA template for rhodopsin. The forward primer should include an EcoRI restriction endonuclease site (GAATTC) and Kozak consensus sequence (GCCACC). The reverse primer should include a BamHI restriction endonuclease site (GGATCC). Primer sequences used to amplify rhodopsin can be found in previously published work (Gragg and Park, 2018).

  3. Digest the PCR-amplified rhodopsin product and pmTq-N1 and pYFP-N1 vectors with the restriction enzymes EcoRI and BamHI for 15 minutes at 37 °C, according to the manufacturer’s protocol. After digestion, heat-inactivate the restriction enzymes for 20 minutes at 65 °C.

  4. Dephosphorylate the digested ends of the vector with calf intestinal phosphatase for 1 hour at 37 °C, according to manufacturer’s protocol.

  5. Run the digested vector and PCR product on a 1% agarose gel in TAE buffer containing 0.5 μg/mL ethidium bromide to separate out the digested material.

  6. Visualize DNA in the gel on a UV transilluminator. Excise bands in the gel corresponding to the digested vector backbone and rhodopsin PCR product. Purify the DNA from the excised bands using a gel purification kit, according to the manufacturer’s protocol.

  7. Ligate the PCR product insert into the vector backbone with T4 DNA ligase, according to the manufacturer’s protocol. Mix the insert with the vector backbone so that the former is in excess of the latter. Start with a ratio of 3:1 for insert and vector, and test higher ratios if unsuccessful. Incubate the reaction mixture with the ligase for 10 minutes at room temperature.

  8. Transform competent cells with up to 10 μL of the ligation mixture, according to the manufacturer’s high efficiency transformation protocol.

  9. Plate the transformed competent cells on LB agar plates containing 50 μg/mL kanamycin. The pmTq-N1 and pYFP-N1 vectors contain a kanamycin resistance gene and therefore cells that have taken up the ligated vector will be resistant to the antibiotic.

  10. Incubate the plates overnight at 37 °C. The next day, pick several colonies from the plate using sterile pipette tips. Transfer the pipette tips to individual cell culture tubes containing 5 mL of LB media with 50 μg/mL kanamycin.

  11. Incubate the inoculated cultures overnight at 37 °C with shaking at 250 rpm in an incubated orbital shaker.

  12. The next day, isolate and purify the vector DNA from each culture with a miniprep kit, according to the manufacturer’s protocol.

  13. Digest a small amount of the purified vector with EcoRI and BamHI restriction enzymes. Run the digested material on a 1 % agarose gel in TAE buffer containing 0.5 μg/mL ethidium bromide and visualize the separated bands. Determine which of the samples exhibit bands corresponding to both the vector backbone and rhodopsin insert using the DNA ladder as a reference.

  14. Sequence the purified DNA that contains the rhodopsin insert to confirm that the sequence for rhodopsin is correct and inserted as expected. The resulting vectors, pRho-mTq and pRho-YFP, will be used for FRET experiments or as a template for mutagenesis to introduce mutations that cause misfolding.

  15. Generate stocks of the vectors pRho-mTq and pRho-YFP by transformation and DNA purification procedures described above. Quantify the purified DNA by UV-Vis spectroscopy. Prior to transfection, dilute the DNA vectors to 100 ng/μL in nuclease-free water.

  16. Point mutations can be introduced in the rhodopsin sequence of pRho-mTq and pRho-YFP using the QuickChange II Site-Directed Mutagenesis Kit (Agilent Technologies, Santa Clara, CA). Primer sequences used to generate mutations can be found in previously published works (Miller et al., 2015, Gragg and Park, 2018)

3.2. CELL CULTURE AND TRANSFECTION

  1. Seed HEK293T cells at 150,000 cells per well in a 12-well cell culture plate in DMEM-H media supplemented with 10% FBS. Keep HEK293T cells below passage number 30 for good protein expression. Other transfectable cell lines can be used as well.

  2. Incubate cells at 37 °C in a 5 % CO2 incubator. 24 hours after seeding, cells should be 60–80 % confluent and ready for transfection.

  3. Transfection procedures using Lipofectamine 2000 outlined here are adapted from the manufacturer’s protocol for transfection in 12-well plates.

  4. Aspirate the media in each well and then add to the cells 0.8 mL of DMEM-H media supplemented with 10% FBS preheated to 37 °C.

  5. Dilute Lipofectamine 2000 in DMEM-H and incubate 5 minutes at room temperature. For 12-well plates, mix 4 μL of Lipofectamine 2000 with 96 μL DMEM-H per well.

  6. To generate FRET curves, cells transfected with only pRho-YFP and cells co-transfected with pRho-mTq and pRho-YFP at different ratios is required. Cells transfected with only pRho-YFP will serve as the background control when measuring spectra. The amount of DNA from a 100 ng/μL solution of purified vector that should be added in the diluted DNA mixture is given in Table 1. The total amount of DNA in the diluted DNA mixture containing both pRho-mTq and pRho-YFP should be 400 ng. The ratios of pRho-mTq and pRho-YFP presented in the table are a starting point and can be adjusted to cover the full range of A:D ratios required to generate FRET curves.

  7. Mix 100 μL of each diluted DNA mixture with 100 μL of diluted Lipofectamine 2000. Incubate for 20 minutes at room temperature.

  8. Add each 200 μL mixture dropwise down the well of a 12-well cell culture plate. Gently pipet the media up and down to ensure adequate mixing. The final volume in the well will be 1 mL.

  9. Incubate cells at 37 °C in a 5 % CO2 incubator for 24–28 hours.

  10. To test the effect of a pharmacological chaperone, such as 11-cis retinal or 9-cis retinal, the pharmacological chaperone can be added shortly after transfection. Both 11-cis retinal or 9-cis retinal are inverse agonists that can bind the apoprotein opsin to form a functional receptor (Hubbard and Wald, 1952). 9-cis retinal is more stable and therefore more practical for experiments. All procedures involving 9-cis retinal must be conducted under dim red light conditions to avoid bleaching of the chromophore. 4–6 hours post-transfection, add 1.5 μL of 9-cis retinal from a 100 mM stock solution in DMSO into each well containing 1 mL of media, which results in a final concentration of 15 μM. Cover plates in foil and return cells to the 5 % CO2 incubator until they are ready for FRET experiments.

  11. Non-specific FRET can be defined by coexpressing rhodopsin with an unrelated GPCR. We have previously cotransfected the vector pFLAG-m2-mTq-1D4 (Gragg et al., 2016), which codes for the m2 muscarinic receptor tagged with mTq, and pRho-YFP for non-specific controls. 2500 ng of pFLAG-m2-mTq-1D4 is required to achieve comparable expression levels of the mTq-tagged m2 muscarinic receptor as that of tagged rhodopsin. 50–800 ng of pRho-YFP is cotransfected with 2500 ng of pFLAG-m2-mTq-1D4 to achieve a range of A:D ratios required for generation of FRET curves.

Table 1.

Example of composition of diluted DNA mixture used for transfection

pRho-YFPa 2 μL (200 ng) 1 μL (100 ng) 2 μL (200 ng) 2.5 μL (250 ng) 3 μL (300 ng) 3.5 μL (350 ng)
pRho-mTqa 0 μL 3 μL (300 ng) 2 μL (200 ng) 1.5 μL (150 ng) 1 μL (100 ng) 0.5 μL (50 ng)
DMEM-H 98 μL 96 μL 96 μL 96 μL 96 μL 96 μL
Total DNA 200 ng 400 ng 400 ng 400 ng 400 ng 400 ng
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a

DNA is added from a 100 ng/μL solution of purified vector

3.3. MEASURING FRET SPECTRA

  1. 24–28 hours post transfection, aspirate media in each well of the 12-well plate of transfected cells. Resuspend cells with 1 mL PBS by gentle pipetting. Repeat 2 more times and combine resuspended cells in a 15 mL conical tube for a total volume of 3 mL.

  2. Perform procedures below starting with cells transfected with only pRho-YFP, which will serve as the background control, and then on cells cotransfected with pRho-YFP and pRho-mTq.

  3. Gently mix the cell suspension on the lowest setting of a vortexer and transfer the 3 mL cell suspension into a cuvette containing a stir bar.

  4. The protocol here describes settings for a FluoroMax-4 spectrofluorometer (Horiba Scientific, Edison, NJ) to detect fluorescence from mTq and YFP. The optimal excitation wavelengths and slit widths should be determined for the spectrofluorometer and the donor acceptor pair used. Samples are maintained at 25 °C in the spectrofluorometer by a circulating water bath.

  5. Place the cuvette containing a cell suspension sample and load into the spectrofluorometer. Obtain spectra on the untreated cells. The emission spectra are obtained for both mTq and YFP excitation. For YFP excitation, samples are excited at 485 nm with a 5 nm slit width and the emission spectrum measured from 505 nm to 650 nm with a 10 nm slit width. For mTq excitation, samples are excited at 425 nm with a 5 nm slit width and the emission spectrum measured from 450 nm to 600 nm. Measurements are taken every nanometer in the emission spectra.

  6. Add 20 μL of 0.2 M DM in water to the cell suspension in the cuvette to achieve a final concentration of 1.3 mM DM. Let the stir bar stir for 5 minutes to thoroughly mix the sample. After 5 minutes, obtain the emission spectrum for mTq excitation. This spectrum is the DM-treated spectrum.

  7. Add 20 μL of 0.5 M SDS to the DM-treated cell suspension to achieve a final concentration of 3.3 mM SDS. Let the stir bar stir for 5 minutes to thoroughly mix the sample. After 5 minutes, obtain the emission spectrum for mTq excitation. This spectrum is the SDS-treated spectrum.

3.4. FRET ANALYSIS

  1. Analyze emission spectra obtained from cells coexpressing YFP- and mTq-tagged rhodopsin (e.g., Figs. 5A and 5B).

  2. For emission spectra obtained after mTq excitation on untreated, DM-treated, and SDS-treated cells, subtract the background signal and adjust the baseline. The emission spectrum after mTq excitation on cells expressing only YFP-tagged rhodopsin is used as the background control. Subtract this spectrum from the spectra obtained from cells coexpressing YFP- and mTq-tagged rhodopsin. Adjust the baseline of the background-subtracted spectra by subtracting the average of the last 15 fluorescence measurements from these spectra so that the baseline is equal to zero. Record the emission signal at 476 nm from untreated (Em476untreated), DM-treated (Em476DM-treated), and SDS-treated (Em476SDS-treated) cells (Figs. 5A and 5B). For A:D ratio computations, Em476SDS-treated is used since the fluorescence signal from mTq is fully unquenched only after SDS treatment.

  3. For emission spectra obtained after YFP excitation on untreated cells, adjust the baseline as described above. Record the emission signal at 527 nm (Em527untreated). YFP fluorescence is stable in DM but unstable in SDS (Gragg et al., 2016). Thus, only the signal from untreated or DM-treated cells can be used to determine the A:D ratio. The A:D ratio is computed as follows: A:D ratio = Em527untreated/ Em476SDS-treated. The computed A:D ratio is in arbitrary units and does not represent the absolute stoichiometry.

  4. The FRET efficiency (E) is calculated from the quenching of the mTq signal at 476 nm (Figs. 5A and 5B). The E for three types of FRET are calculated.

  5. Total FRET corresponds to the FRET signal from untreated cells and is a combination of DM-sensitive and DM-insensitive FRET (Fig. 5). Etotal = 1 − (Em476untreated /Em476SDS-treated).

  6. DM-sensitive FRET corresponds to the fraction of the total FRET signal originating from rhodopsin oligomers that are disrupted by DM treatment (Fig. 5). EDM-sensitive = (Em476DM-treated – Em476untreated))/Em476SDS-treated.

  7. DM-insensitive FRET corresponds to the fraction of the total FRET signal originating from rhodopsin aggregates that are resistant to DM treatment and requires SDS treatment to disrupt FRET (Fig. 5). EDM-insensitive = 1 − (Em476DM-treated/Em476SDS-treated).

  8. To generate FRET curves for total, DM-sensitive, and DM-insensitive FRET, plot the computed values of E versus the computed values of the A:D ratio (Fig. 3C). The maximal FRET efficiency (Emax) is determined by fitting the data by non-linear regression, using graphing software such as Prism, to the following rectangular hyperbolic function: E = (Emax × A:D) / (EC50 + A:D).

  9. Examples of FRET curves for a non-specific control, WT rhodopsin, and misfolded mutant rhodopsin is shown in Fig. 7. The Emax for total, DM-sensitive, and DM-insensitive FRET of the non-specific control must be exceeded to indicate specific FRET representing physical interactions (Figs. 4 and 7A). The total FRET signal from WT rhodopsin originates predominantly from DM-sensitive FRET (Fig. 7B), indicative of oligomers. The total FRET signal from misfolded mutant rhodopsin originates predominantly from DM-insensitive FRET (Fig. 7C), indicative of aggregates.

Figure 7.

Figure 7.

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Examples of FRET curves. FRET curves for the non-specific control (A), wild-type rhodopsin (B), and mutant rhodopsin (C) are shown. FRET curves for total FRET (blue), DM-sensitive FRET (red), and DM-insensitive FRET (green) are shown. The non-specific control shown is from cells coexpressing mTq-tagged m2 muscarinic receptor and YFP-tagged wild-type rhodopsin. The Emax from FRET curves from the non-specific control is shown as dashed lines. The FRET curves are adapted from those presented in (Gragg et al., 2016).

Acknowledgments

This work was funded by grants from the National Institutes of Health (R01EY021731) and Research to Prevent Blindness (Unrestricted Grant).

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