MBP is a clamshell shaped protein that undergoes a significant closure of the clamshell upon binding its ligand, maltose (Figure 1A; Video 1). In addition, maltose binding leads to a smaller opening of the backside of the clamshell. By introducing one FRET pair at the top of the clamshell and a second FRET pair on the backside of the clamshell, we can address the accuracy and precision of ACCuRET to measure absolute distances as well as ligand-dependent increases and decreases in distance.

Our criteria for selection of FRET pairs were (1) the sites are solvent exposed, (2) the sites are on rigid secondary structural elements (α helices in MBP), (3) the distance between the sites is predicted to fall in the working range of tmFRET (~10–20 Å), and (4) the distance between sites undergoes a moderate change between apo and holo MBP. Our first FRET pair was at positions 295 (donor) and 237 (acceptor) on the outer lip of the clamshell (Figure 1A; Video 1). From β-carbon distances in the X-ray structures, these sites are 21 Å apart in the apo state and 13 Å apart in the holo state, a distance change of about −8 Å (shortening) with maltose binding. Our second FRET pair was at positions 322 (donor) and 309 (acceptor) on the backside of the clamshell (Figure 1A; Video 1). These sites are 13 Å apart in the apo state and 17 Å apart in the holo state, a distance change of +4 Å (lengthening) with maltose binding. In this study, we used these two FRET pairs to evaluate the accuracy and precision of ACCuRET with different acceptor metal-binding sites and different metals as well as for MBP in solution and bound to the membrane.

ACCuRET requires the protein to be site-specifically labeled with a donor fluorophore and an acceptor transition metal ion. To achieve specific labeling of MBP with a small donor fluorophore, we used amber codon suppression to introduce the fluorescent noncanonical amino acid L-Anap (Figure 2A) (Chatterjee et al., 2013; Kalstrup and Blunck, 2013). Our amber codon suppression method required the use of two plasmids (Zagotta et al., 2016; Aman et al., 2016). One plasmid encodes MBP with a TAG stop codon (the amber codon) engineered at the site for L-Anap incorporation. The second plasmid, pAnap, developed by Peter Schultz’s lab (Chatterjee et al., 2013), encodes an evolved amino acyl tRNA synthetase and 4 copies of tRNA, which the cell uses to produce L-Anap-loaded tRNA complementary to the TAG codon (Figure 2A). HEK293T/17 cells were co-transfected with the two plasmids and incubated in a cell-permeable methyl ester (ME) version of L-Anap.

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We achieved specific incorporation of L-Anap at donor positions 295 and 322 (Figure 1A). MBP-Anap was the primary band visible in gels of crude cell lysates using in-gel L-Anap fluorescence (Figure 2—figure supplement 1), indicating that L-Anap was not appreciably incorporated at other amber codon-containing sites. In addition, Western blots of FLAG-MBP containing the K295TAG or E322TAG mutation revealed full-length MBP only when the cells were incubated with L-Anap-ME (Figure 2B,C). However, even when L-Anap was present in the medium, a moderate amount of truncated product, in which translation terminated at the introduced stop codon, was observed for both MBP-295TAG and MBP-322TAG, as shown by the plots in Figure 2D,E (fraction of full-length MBP was 30% and 10%, respectively). These results indicate that L-Anap incorporation at positions 295 and 322 in MBP in HEK293T/17 cells was specific, but not particularly efficient.

To increase the efficiency of L-Anap incorporation, we co-expressed the MBP-TAG and pAnap plasmids with a third plasmid encoding a dominant negative form of eukaryotic Release Factor 1 (DN-eRF1). Previously, Jason Chin’s group showed that DN-eRF1 increases the efficiency of incorporation of noncanonical amino acids in mammalian cells (Schmied et al., 2014), presumably by delaying release of the translation product from the ribosome when encountering an amber stop codon. For both sites in MBP, coexpression with DN-eRF1 substantially increased the fraction of full-length product from cells incubated with L-Anap-ME (from 30% to 86% for MBP-295TAG and from 10% to 79% for MBP-322TAG) (Figure 2D,E). Coexpression with DN-eRF1 also increased the absolute amount of full-length protein relative to that observed with wild-type MBP, from 1.6% (±0.4%; n = 3) to 6.3% (±0.5%; n = 3) for MBP-295TAG and from 5.5% (±4%; n = 3) to 50% (±27%; n = 3) for MBP-322TAG. We observed little or no read-through (i.e. incorporation of a natural amino acid at the stop codon), even when co-expressing DN-eRF1, as little full length MBP-TAG protein was observed when L-Anap was omitted from the cell growth medium (Figure 2B,C). These results establish a method for achieving specific, high-efficiency fluorescent labeling of MBP for ACCuRET experiments.

L-Anap’s small size, short linker, and spectral properties makes it useful for tmFRET experiments (Zagotta et al., 2016; Aman et al., 2016; Dai et al., 2018; Dai and Zagotta, 2017). The peak absorption wavelength of L-Anap is 360 nm (Chatterjee et al., 2013). The emission of L-Anap is environmentally sensitive, blue shifting as the environment becomes less polar. The emission peak of L-Anap was 494 nm in our buffer and 475 nm in ethanol (see Materials and methods, Figure 13B). For the MBP-295Anap and MBP-322Anap in this study, the peak emissions were 491 nm and 478 nm, respectively (Figure 13A), suggesting that neither 295Anap nor 322Anap is fully solvent exposed and that the environment of 322Anap is more hydrophobic than that of 295Anap. We found that the quantum yield of L-Anap was also site dependent, measured to be 0.31 for MBP-295Anap and 0.47 for MBP-322Anap (see Materials and methods and Figure 13C). L-Anap’s low quantum yield, together with its low extinction coefficient (~20×10³ M⁻¹cm⁻¹; [Chatterjee et al., 2013; Zagotta et al., 2016]), makes L-Anap relatively dim compared to most visible light fluorophores; however, its fluorescence intensity is sufficient for most macroscopic fluorescence experiments. L-Anap’s emission spectrum overlaps the absorption spectra of many transition metal ions (such as Ni²⁺, Co²⁺, and Cu²⁺) making it a useful FRET donor for tmFRET (Figure 3C).

Figure 3

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The standard method to introduce a transition metal ion acceptor for tmFRET uses a dihistidine (HH) motif with histidines introduced at positions i and i + 4 on an α helix (Figure 3A) or at i and i + 2 on a βstrand (Taraska et al., 2009a; Taraska et al., 2009b; Richmond et al., 2000; Puljung and Zagotta, 2013; Arnold and Haymore, 1991; Suh et al., 1991). These minimal metal binding sites can bind Ni²⁺, Co²⁺, and Cu²⁺ with affinities in the 1 to 100 µM range, and binding can be reversed with EDTA. Although this method places the transition metal ion very close to the protein backbone, the metal concentrations required are high, so that they are not fully biocompatible and are subject to off-target binding.

To circumvent the limitations of standard HH sites, we developed a new, better method to introduce a specific, high-affinity metal-binding site in a protein by labeling a cysteine with 1-(2-pyridin-2-yldisulfanyl)ethyl)-1,4,7,10-tetraazacyclododecane (TETAC; Figure 3A,B). TETAC is a cysteine-reactive compound with a short linker to a cyclen ring that binds transition metal ions with very high-affinity (sub-nanomolar; [Kodama and Kimura, 1977]). Single cysteines can then be introduced in the protein at sites of interest and reacted with TETAC to create a mixed disulfide linkage to a cyclen metal-binding site.

ACCuRET uses TETAC to introduce minimal metal binding sites. TETAC has many advantages compared with standard HH sites for tmFRET: (1) it requires only nominal concentrations of free Cu²⁺ eliminating off-target binding of Cu²⁺ to endogenous binding sites on the protein; (2) it can be used with any type of secondary structural element; (3) its reaction with endogenous cysteines produces tmFRET only for sites that are very close (<20 Å), so endogenous cysteines need not be eliminated; (4) it can be easily removed with reducing agent, making its use compatible with physiological concentrations of other metals, for example Ca²⁺; and (5) it significantly blue shifts and increases the absorbance of Cu²⁺, extending the range of R₀ values available for tmFRET (Figure 3C). As shown in Figure 3D, the distance dependence of the FRET efficiency predicted by the Förster equation for each of our two L-Anap sites in MBP is shifter by almost 4 Å to longer distances for Cu²⁺-TETAC compared to Cu²⁺-HH. ACCuRET, therefore, offers a flexible and biocompatible method for measuring distances in the biological distance range.

For each of our two FRET pairs (Figure 1A), we made three constructs: one with no metal-binding site, one with a standard HH-binding site, and one with a Cys, for modification by TETAC. For MBP-295Anap, the HH was introduced at positions 233 and 237 (MBP-295Anap-HH) and the Cys was introduced at position 237 (MBP-295Anap-C). For MBP-322Anap, the HH was introduced at positions 305 and 309 (MBP-322Anap-HH) and the Cys was introduced at position 309 (MBP-322Anap-C). For both donor sites, control constructs were also made with no introduced metal-binding sites (MBP-295Anap and MBP-322Anap) to measure other forms of energy transfer and nonspecific decreases in fluorescence, and correct the FRET efficiency measurements accordingly (see Materials and methods). These constructs were all expressed in HEK293T/17 cells as amino-terminal FLAG fusions and purified using anti-FLAG beads for subsequent analysis (Figure 2—figure supplement 1).

ACCuRET was readily observed in both MBP-Anap-C constructs. We recorded emission spectra before and after application of 10 µM Cu²⁺-TETAC in a fluorometer (see Materials and methods and Approaches). As shown in Figure 4, Cu²⁺-TETAC produced a large decrease in fluorescence intensity for both MBP-295Anap-C (Figure 4A) and MBP-322Anap-C (Figure 4B). The emission spectra in the presence of Cu²⁺ were nearly identical in shape to the spectra without Cu²⁺-TETAC (Figure 4A–D, dashed traces) indicating the fluorescence quenching reflected a FRET mechanism as opposed to a change of environment of L-Anap or an inner filter effect (Lakowicz, 2006).

Figure 4

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Fluorescence quenching was strongly dependent on the presence of maltose. For MBP-295Anap-C, the quenching was greater with maltose indicating an increase in FRET efficiency and shorter distance (Figure 4C). For MBP-322Anap-C, the quenching was smaller with maltose indicating a decrease in FRET efficiency and longer distance (Figure 4D). Changes in tmFRET in MBP-295Anap-C therefore allowed us to visualize the closing of the top of the clamshell with ligand binding, and changes in tmFRET in MBP-322Anap-C allowed us to visualize the opening of the backside of the clamshell (Video 1). Importantly, maltose did not change the shape of the emission spectrum or shift the peak for either construct (Figure 4E,F), indicating the environment of L-Anap did not change upon maltose binding. However, because ACCuRET measures L-Anap fluorescence in the absence and presence of the acceptor for both the apo and ligand-bound states, it would account for any ligand-dependent changes in environment that would otherwise affect the calculation of FRET efficiency.

The maltose affinity of wild-type MBP is about 2 µM (Martineau et al., 1990). We found that MBP purified from HEK293T/17 cells with L-Anap incorporated at position 295 or 322, and Cu²⁺ bound to HH or TETAC, appeared to be bound to an endogenous ligand, perhaps glycogen. These constructs exhibited FRET efficiencies expected for the maltose-bound states, and did not exhibit the maltose-dependent changes in tmFRET even after extensive washing and ion exchange chromatography (data not shown). Therefore, in all the MBP constructs in this paper we introduced a mutation into the maltose-binding site (W340A) that has previously been shown to reduce the maltose affinity (Martineau et al., 1990). This construct exhibited a maltose-dependent change in FRET (Figure 4) indicating that it did not copurify with an endogenous sugar.

We used ACCuRET to measure the maltose affinity of MBP-295Anap-C (containing W340A) modified with Cu²⁺-TETAC by measuring the fluorescence quenching as a function of maltose concentration. As shown in Figure 5, maltose caused a concentration-dependent decrease in fluorescence with an apparent affinity of 280 µM, similar to the value measured previously with tryptophan fluorescence (Martineau et al., 1990). No decrease in fluorescence was observed in MBP-295Anap lacking the introduced cysteine (Figure 5). These experiments illustrate the utility of ACCuRET for measuring both structural and functional properties of proteins.

Figure 5

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To quantify ACCuRET, we measured the time course of the fluorescence at a given wavelength before and after addition of Cu²⁺-TETAC and the reducing agent DTT. For both MBP-295Anap-C and MBP-322Anap-C, there was a rapid (<10 s) and substantial quenching of the fluorescence upon addition of 10 µM Cu²⁺-TETAC (Figure 6A,B). The quenching was nearly completely reversed upon addition of DTT (Figure 6A,B) and negligible in the absence of the introduced cysteine (Figure 6C,D), indicating that it resulted from a reaction of Cu²⁺-TETAC with the introduced cysteine. For MBP-295Anap-C the quenching was greater in the presence of maltose whereas for MBP-322Anap-C the quenching was less in the presence of maltose (Figure 6C,D), as expected from the predicted distance changes. These results establish that ACCuRET can serve as a useful alternative to tmFRET with standard HH sites to create a minimal metal-binding site for tmFRET with L-Anap and report intramolecular distance changes.

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We compared tmFRET efficiencies measured with ACCuRET to those measured with Cu²⁺ bound to standard HH sites, using the same donor-acceptor positions on MBP. Addition of 100 µM Cu²⁺ caused a rapid quenching of both MBP-295Anap-HH and MBP-322Anap-HH fluorescence (Figure 7A,B). The quenching was nearly completely reversed upon addition of EDTA (Figure 7A,B) and negligible in the absence of the introduced HH sites (Figure 7C,D), indicating that it resulted from binding of Cu²⁺ to the introduced HH sites. For MBP-295Anap-HH, the quenching was greater in the presence of maltose, whereas for MBP-322Anap-HH the quenching was less in the presence of maltose (Figure 7), as observed with Cu²⁺-TETAC (Figure 6).

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For both L-Anap sites, the quenching increased with increasing Cu²⁺ concentration, following a simple binding isotherm, with an apparent affinity corresponding to the affinity of the metal-binding site (Figure 7—figure supplement 1). For Cu²⁺ binding to MBP-295Anap-HH (in the presence of maltose), the affinity was 3 µM (Figure 7—figure supplement 1C), and for Cu²⁺ binding to MBP-322Anap-HH (in the absence of maltose) the affinity was 17 µM (Figure 7—figure supplement 1D). The quenching at saturating concentrations reflects the FRET efficiency between L-Anap and the bound metal. These data support the use of 100 µM Cu²⁺ as a saturating concentration for measuring tmFRET in these constructs.

Interestingly, at 1 mM Cu²⁺ there was some fluorescence quenching in MBP-295Anap, but little in MBP-322Anap (Figure 7—figure supplement 1). As these constructs lack the HH metal-binding site and 1 mM Cu²⁺ is a fairly high concentration, this quenching likely represents collisional quenching and suggests that 295Anap has a greater solvent accessibility compared to 322Anap, as also suggested by the emission spectra discussed above (Figure 13A). These data also indicate that there was negligible quenching by Cu²⁺ bound to endogenous sites in MBP at Cu²⁺ concentrations below 1 mM. Although not an issue with the constructs used here, it is worth noting that these potential problems caused by high concentrations of free Cu²⁺ are largely mitigated in ACCuRET.

One advantage of standard HH sites is the ability to use transition metals with different coordination chemistries and absorption profiles than Cu²⁺ (Figure 3C). These metals, including Co²⁺ and Ni²⁺, are expected to have different binding affinities for HH and endogenous sites as well as different R₀ values. Co²⁺ produced a similar FRET efficiency to Cu²⁺ but exhibited about a 10-fold lower binding affinity (Figure 7—figure supplement 1C, pink). Ni²⁺, however, produced a substantially lower FRET efficiency at saturating concentration (Figure 7—figure supplement 1C, green). This reflects the lower R₀ value predicted for Ni²⁺ and further supports a FRET mechanism for the energy transfer. More importantly, the lower R₀ value for Ni²⁺ expands the useable range of tmFRET measurements to shorter distances. With standard HH sites, metals with different R₀ values can be selected to match the distance of interest.

FRET efficiency is steeply dependent on distance between the donor and acceptor, making FRET a molecular ruler for measurements of distances (Stryer and Haugland, 1967). Accurate distance measurements, however, have historically been a challenge for FRET studies because they are highly dependent on the conformational heterogeneity and dynamics of the fluorophores, the relative orientation of the fluorophores, and the labeling specificity and efficiency (Best et al., 2007; Schuler et al., 2005; Sobakinskaya et al., 2018). ACCuRET was designed to address these problems by using small probes with short linkers, a metal ion acceptor that is isotropic, and labeling methods that are orthogonal, specific, and efficient.

Energy transfer from donor to acceptor causes quenching of donor fluorescence upon the addition of acceptor. With tmFRET, the FRET efficiency, E, can be easily calculated from the decrease in L-Anap fluorescence upon addition of metal acceptor. Other sources of energy transfer (e.g. solution quenching and off-target metal binding) and nonspecific decreases in fluorescence (e.g. bleaching and loss of protein) can confound calculations of FRET efficiency. The extent of this background quenching can be determined using constructs without an introduced metal-binding site and the FRET efficiencies corrected for as described in Materials and methods (Taraska et al., 2009a; Dai and Zagotta, 2017; Puljung and Zagotta, 2013).

From the FRET efficiencies, we calculated the distances between the donors and acceptors using the Förster equation r = R₀(1/E− 1)^1/6 where R₀ is the distance producing 50% FRET efficiency. We calculated R₀ using our measurements of the emission spectrum and quantum yield of L-Anap at each of the two positions in MBP, 295 and 322 (Figure 13), and the absorption spectrum of Cu²⁺ bound to either a HH motif or cyclen (Figure 3C). We assume random orientations of the donor and acceptor (κ² = 2/3), a reasonable assumption when one member of the FRET pair is a metal ion (Selvin, 2002).

ACCuRET accurately measured intramolecular distances and distance changes, comparable to tmFRET using standard HH sites. Figure 8 compares the tmFRET distance measurements for all our MBP constructs, with either Cu²⁺-TETAC or Cu²⁺-HH, in the absence (open circles) or presence (closed circles) of maltose. Also shown are the β-carbon distances predicted from the X-ray crystal structures of MBP in the absence and presence of ligand. Overall, the distances were highly accurate, with a root-mean-square deviation (RMSD) between measured distances and crystal structure distances of 1.5 Å for ACCuRET and 1.8 Å for tmFRET with standard HH sites. There was little variability in our measurements (Figure 6C,D, and Figure 7C,D), suggesting that the errors were more likely to be in our estimates of the probe distances from the β-carbon positions, or in assumptions of the Förster equation (see below), than in experimental variability due to the method. In addition, the changes in distances with maltose were similar in direction and magnitude to the changes in β-carbon distances predicted from the crystal structures. Finally, we did not detect a systematic difference between the distances measured with Cu²⁺-HH and Cu²⁺-TETAC (Figure 8). In conclusion, ACCuRET accurately determined the absolute distances and maltose-dependent distance changes for the FRET pair at the top of the clamshell, for which the distance decreased with maltose, and the FRET pair on the back of the clamshell, for which the distance increased with maltose.

Figure 8

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Membrane proteins present a major challenge for measuring protein structure and conformational dynamics. Compared to soluble proteins, membrane proteins typically express at much lower levels, are difficult to purify in their native state, and often require their native membrane environment to exhibit physiological structural and functional properties. ACCuRET is well suited to address these challenges because it leverages the exquisite sensitivity of fluorescence measurements, the efficient incorporation of L-Anap into proteins in mammalian cells, the ability to work on unpurified proteins, and the ability to record from proteins in their native membranes. We therefore set out to validate and optimize ACCuRET for measuring conformational dynamics of proteins in the membrane.

To record fluorescence from native membranes we utilized a method called cell unroofing. Our implementation of this method utilizes a probe sonicator mounted on the stage of an inverted microscope to shear off the dorsal surface of cells and dislodge all soluble cellular contents and organelles (Figure 9; [Zagotta et al., 2016]). Cell unroofing leaves the ventral surface of cells intact as plasma membrane sheets attached to the coverslip for simple epifluorescence imaging. Cell unroofing also provides access to the intracellular surface of the membrane for application of transition metals and intracellular ligands. tmFRET measurements can then be made by recording the fluorescence from the unroofed cell before and after the application of Cu²⁺-TETAC or free transition metal.

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To validate and compare ACCuRET in unroofed cells, we made membrane-bound MBP constructs by adding a carboxy-terminal CAAX motif, which is then farnesylated in the cell (Farnsworth et al., 1989; Anderegg et al., 1988). These MBP-295TAG-CAAX and MBP-322TAG-CAAX constructs were each cotransfected into HEK293T/17 cells with pANAP and DN-eRF1 and incubated with L-Anap-ME in the medium, as described above. Both MBP-295Anap-CAAX and MBP-322Anap-CAAX expressed robustly and localized to the intracellular surface of the plasma membrane (Figure 9—figure supplement 1). Upon unroofing, the cells were no longer easily visible with bright field illumination. However, epifluorescence illumination showed plasma membrane sheets with relatively uniform L-Anap fluorescence (Figure 9) which were stable for >30 min (data not shown). Previously, we showed that these images look similar with total internal reflection fluorescence (TIRF) microscopy, confirming that most or all the fluorescence is from the plasma membrane (Zagotta et al., 2016). These results indicate that MBP-295Anap-CAAX and MBP-322Anap-CAAX proteins were stably associated with the membrane.

As for soluble MBP, Cu²⁺-TETAC was an effective tmFRET acceptor for membrane-bound MBP-Anap in unroofed cells. We unroofed the MBP-Anap-C-CAAX constructs and recorded the L-Anap fluorescence before and after addition of Cu²⁺-TETAC and DTT. As shown in Figure 10A,C, Cu²⁺-TETAC produced a rapid decrease in L-Anap fluorescence of MBP-295Anap-C-CAAX that was reversed by the addition of DTT. The quenching was nearly absent in MBP-295Anap-CAAX without the introduced Cys (Figure 10E), indicating it arose from Cu²⁺-TETAC modification of this Cys. The quenching was greater in the presence of maltose (Figure 10C,E) as expected from the decrease in distance between the donor and acceptor with maltose for the MBP-295Anap FRET pair (Video 1). Similarly, for MBP-322Anap-C-CAAX, Cu²⁺-TETAC produced a rapid and reversible decrease in L-Anap fluorescence that was absent in MBP-322Anap-CAAX (Figure 10B,D,F). With MBP-322Anap-C-CAAX, however, the quenching was less in the presence of maltose (Figure 10D,F), as expected from the maltose-induced increase in distance between the donor and acceptor for the MBP-322Anap FRET pair (Video 1). The fractional fluorescence quenching was highly reproducible from cell to cell and was similar to the results obtained for soluble MBP in a cuvette (Figure 6). These results establish that ACCuRET can be used to measure tmFRET in membrane proteins.

Figure 10

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We compared tmFRET efficiencies measured with ACCuRET to those measured with Cu²⁺ bound to standard HH sites, using the same donor-acceptor positions. Addition of 100 µM Cu²⁺ to unroofed cells expressing MBP-295Anap-HH-CAAX and MBP-322Anap-HH-CAAX produced a rapid decrease in fluorescence that was reversed by EDTA and nearly absent without the introduced HH sites (Figure 11). Furthermore, the quenching with Cu²⁺ was greater with maltose than without maltose in MBP-295Anap-HH-CAAX and less with maltose than without maltose in MBP-322Anap-HH-CAAX. The fractional fluorescence quenching was highly reproducible from cell to cell and closely mirrored the results obtained for soluble MBP in a cuvette.

Figure 11

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From the FRET efficiencies, we calculated the distances between the donors and acceptors using the Förster equation as described above. Comparing our determination of distances from membrane-bound MBP to those predicted from the X-ray crystal structures of MBP (Figure 8) gives RMSD values of 2.9 Å for ACCuRET and 1.3 Å for tmFRET with standard HH sites, comparable to the values for MBP in solution. However, distances calculated from membrane-bound MBP-Anap constructs were systematically longer than those determined from MBP-Anap constructs in solution (see Discussion). Together, our data demonstrate that ACCuRET provides accurate determinations of distances and distance changes in both soluble proteins and membrane proteins.

The Förster equation predicts a steep distance dependence for FRET efficiency (Figure 12A, black curve) and assumes that the donor and acceptor probes are a fixed distance apart (Figure 12B, black). However, the linkers between the probes and the protein backbone produce substantial heterogeneity and mobility of the probes that effect the measured FRET efficiency (Best et al., 2007; Schuler et al., 2005; Taraska et al., 2009b). This heterogeneity in distance, together with the nonlinear dependence of the FRET efficiency on distance, cause the distances closer to the R₀ distance to be weighted more heavily. The distance dependence of FRET efficiency, therefore, becomes shallower than predicted by the Förster equation. Our approach to reduce this effect has been to minimize these linkers by using a fluorescent amino acid and minimal metal binding sites closely associate with the backbone (HH or TETAC). Still, we observed a small, systematic bias in our tmFRET measurements where the distances greater than R₀ have somewhat greater FRET efficiency and the distances less than R₀ have somewhat lower FRET efficiency than predicted by the Förster equation (Figure 12A). This manifests as an apparent shallowing of the distance dependence and an underestimation of the changes in distance with maltose.

Figure 12

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To correct for this heterogeneity, we developed a method we call FCG (Förster convolved with Gaussian) analysis. FCG analysis assumes a Gaussian distribution of donor-acceptor distances in the sample (Figure 12B). This distribution is typical of the distance distributions between two spin labels on rigid proteins measured using double electron-electron resonance (DEER) spectroscopy (Jeschke, 2012). When the Förster equation was convolved with a Gaussian distribution, the apparent FRET efficiency has a similar (but not identical) R₀ but a more shallow distance dependence than the Förster equation (Figure 12A). The greater the width of the Gaussian distribution, the shallower the distance dependence. Using a Gaussian distribution with a full width at half maximum (FWHM) of 8 Å, we found that, in most cases, the distances and changes in distance calculated from the FCG model (Figure 8, asterisks) more closely matched the distances predicted from the crystal structures than the distances calculated with the Förster equation alone. Since all proteins at physiological temperature possess some structural heterogeneity (Frauenfelder et al., 1991), we propose that this correction may be useful for many FRET studies of distance.

All constructs were made in the pcDNA3.1 mammalian expression vector (Invitrogen, Carlsbad, CA), except as noted below. The amino acid sequence of MBP was based on the pMAL-c5x vector (New England Biolabs, Ipswich, MA), and was codon optimized for mammalian cell expression. To reduce the affinity of MBP for endogenous ligands and allow subsequent testing of maltose binding, we introduced a point mutation, W340A (Martineau et al., 1990), in all constructs. In addition, a FLAG epitope was added to the N-terminus of MBP. FLAG-MBP cDNA was synthesized by Bio Basic (Amherst, NY). The FLAG epitope is present in all constructs but is omitted from the construct names except in the discussion of western blot analysis. For membrane localization, the C-terminus of FLAG-MBP was fused to a CAAX domain with the following sequence: KMSKDGKKKKKKSKTKCVIM. The pAnap vector ([Chatterjee et al., 2013]; purchased from Addgene, Cambridge, MA) was used as previously described (Zagotta et al., 2016). The dominant negative eukaryotic release factor one construct (DN-eRF1) was kindly provided by Jason Chin (Cambridge, UK) and used in the provided pcDNATM5/FRT/TO vector (Schmied et al., 2014).

Amber stop codons (TAG), histidines and cysteines were introduced using standard PCR and oligonucleotide-based mutagenesis. All sequences were confirmed using automated DNA sequencing (Eurofin/Operon, Louisville, KY). The stop codons were introduced into FLAG-MGP at positions 295 (MBP-295TAG) and 322 (MBP-322TAG). For each stop codon, three constructs were produced: no metal-binding site, a di-histidine, and a cysteine. For MBP-295TAG, the di-histidines were introduced at positions 233 and 237 (MBP-295TAG-HH), and the cysteine was introduced at position 237 (MBP-295TAG-C). MBP-295TAG-C also contained the 233H mutation. For MBP-322TAG, the di-histidines were introduced at positions 305 and 309 and the cysteine at position 309.

HEK293T/17 cells were obtained from ATCC (Manassas, VA; #CRL-11268; RRID: CVCL_1926), and were expanded only once immediately upon receipt. Cells were used for no more than 35 passages, and replenished from frozen aliquots of the originally expanded stock. Aliquots of the originally expanded stock were used for no more than 8 years, at which point a new stock was purchased. No additional authentication was performed. Testing for mycoplasma contamination was performed using the MicroFluor Mycoplasma Detection kit (catalog #M7006; ThermoFisher Scientific, Waltham, MA) at the end of the study and cells were found to be contaminated.

HEK293T/17 cells were plated in 6-well trays on glass coverslips treated with poly-lysine. Cells were transfected at ≈25% confluency with a total of 1.6 µg of DNA and 10 µL of Lipofectamine 2000 (Invitrogen, Carlsbad, CA) per well. The 1.6 µg of DNA consisted of 0.9 µg of FLAG-MBP, 0.3 µg of pANAP and 0.4 µg of DN-eRF1. For experiments in which DN-eRF1 was not used, it was replaced with 0.4 µg pcDNA3. The DNA/Lipofectamine mix was prepared in 300 µL Opti-MEM (Invitrogen, Carlsbad, CA) per well. For transfection, cells were incubated in growth medium without antibiotics for 4–6 hr at 37°C with 5% CO₂. After incubation, the medium was replaced with one including antibiotics and supplemented with 20 µM L-Anap-ME (AssisChem, Waltham, MA). The L-Anap-ME was made as a 10 mM stock in ethanol and stored at −20°C. Trays were wrapped with aluminum foil to block any light and incubated at 37°C until use.

For fluorometry or western blot experiments, cells were harvested approximately 24 hr after transfection. Cells were washed twice with PBS, and cell pellets, collected from 9 wells each, were stored at −20°C until use. For imaging experiments, the day after transfection, medium was removed from the wells and replaced with HEPES buffered Ringers (HBR) solution (in mM: NaCl 140; KCl 4; CaCl₂ 1.8; glucose 5; HEPES 10; pH 7.4). Cells were then placed on the bench top at room temperature for the remainder of the same day.

For fluorometry experiments, between 1 and 4 frozen pellets were thawed and resuspended in 0.5–0.8 mL Stabilization Buffer Tris (SBT) (in mM: KCl 70; MgCl₂ 1; Trizma Base 30; pH 7.4) supplemented with cOmplete mini EDTA-free protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO). The suspension was sonicated using a Sonifier 450 with MicroTip (Branson, Danbury, CT) with settings of power = 4 and duty cycle = 50% for a total of 10 pulses. Lysed cells were then spun in a benchtop centrifuge at 13,000 rpm for 20 min at 4°C, and the cleared lysate was moved to a new tube.

Anti-FLAG M2 affinity gel (Sigma-Aldrich, St. Louis, MO) was prepared by rinsing 100 µL of slurry per sample five times with 1 mL SBT. Cleared lysate was then added to the rinsed gel and nutated at 4°C for 1 to 2 hr. Tubes were wrapped in aluminum foil to prevent photodamage. The gel was rinsed five times with 1 mL of SBT. Between 0.25 to 0.5 mL of a 200 ng/mL solution of FLAG peptide (Sigma-Aldrich, St. Louis, MO) in SBT was added to the rinsed beads, which were then nutated at 4°C for between 90 min and 12 hr to elute the protein. The purified protein was harvested by spinning the beads and collecting the supernatant. The protein was stored in tubes wrapped in aluminum foil at 4°C for up to 1 week.

For western blot analysis, 25 µL of lysis buffer per 10 mg cells were used to normalize the protein concentration for each experiment. Lysis buffer was made as follows: to 35 ml H₂O, 185.7 mg triethanolamine, 876.6 mg NaCl, and 10 mL glycerol were added, then the pH was adjusted to 8.0 with NaOH. After bringing the volume up to 50 mL, 500 mg digitonin, 5 mM EDTA, and Halt protease inhibitor cocktail (ThermoFisher Scientific, Waltham, MA) were added. Cleared cell lysates were then were run on NuPage 10% bis-tris gels (ThermoFisher Scientific, Waltham, MA) with MOPS SDS running buffer (50 mM MOPS, 50 mM Tris, 3.5 mM SDS and 1 mM EDTA). Proteins were transferred to PVDF membranes using a BioRad Transblot SD (Hercules, CA) transfer cell with Bjerrum/Schafer-Nielsen transfer buffer with SDS (Bjerrum and Schafer-Nielsen, 1986). Membranes were blocked in 5% Milkman Instant Low Fat Dry Milk (Marron Foods, Harrison, NY) in TBS-T (20 mM Tris, 137 mM NaCl, 0.1% Tween20, pH 7.6) for either 1 hr at room temperature or overnight at 4°C. Anti-FLAG primary antibody (Sigma-Aldrich Cat F3165, St. Louis, MO) was used at a dilution of 1:20,000 and secondary antibody (Amersham Cat NA931, Pittsburgh, PA) HRP-linked anti mouse IgG was used at 1:30,000 dilution in TBS-T. Secondary antibodies were visualized with Super Signal West Femto Substrate (ThermoFisher, Waltham, MA) and imaged with a Proteinsimple gel imager (San Jose, CA). Densitometry was performed using ImageJ (Schneider et al., 2012). Regions of interest were drawn around each band and paired with adjacent background regions of the same size. The mean gray value within each background region was subtracted from the mean gray value within its corresponding band. We attempted to achieve similar intensities for wild-type and L-Anap incorporating MBP by loading different amounts of lysate for the different constructs. In all cases, the dilution of samples was taking into account after densitometry to allow comparisons to be made among lanes.

Starna (Atascadero, CA) sub-micro fluorometer cells (100 µL) were used for both fluorometry and spectrophotometry. Fluorometry experiments were performed using a Jobin Yvon Horiba FluoroMax-3 spectrofluorometer (Edison, NJ). For emission spectra of L-Anap, we used an excitation wavelength of 370 nm and 5 nm slits for excitation and emission, except for experiments to measure quantum yield in which 1 nm slits were used. For time course measurements, we excited samples at 370 nm and recorded the emission at 480 nm every 10 s using anti-photobleaching mode of the instrument. Reagents (transition metals, Cu²⁺-TETAC, and DTT) were added manually as 100x stocks during the interval between measurements by pipetting up and down in the cuvette without removing it from the instrument.

Protein samples were diluted 1:10 to 1:200 in SBT to keep the fluorescence intensity within the linear range of the spectrofluorometer. We found that it was essential to use Tris buffered solutions for experiments with Cu²⁺ and Cu²⁺-TETAC, as Cu²⁺ appeared to precipitate in HEPES-based solutions (data not shown). Transition metal ions (Cu²⁺, Co²⁺ and Ni²⁺) were prepared from sulfate (CuSO₄) or chloride (CoCl₂ and NiCl₂) salts as 100 mM stocks in water or SBT, with stocks of lower concentrations diluted from this stock in SBT. We noted no difference between CuSO₄ and CuCl₂ binding to TETAC, but chose to use CuSO₄. TETAC (Toronto Research Chemicals, Toronto, Canada) was prepared as a 100 mM stock in DMSO and stored at −20°C until the day of use. The TETAC stock appeared stable over a time scale of months and tolerated multiple freeze-thaw cycles. To prepare Cu²⁺-TETAC, 1 µL each of 100 mM TETAC stock and 110 mM CuSO₄ stock were mixed together and allowed to incubate for one minute as the solution turned a deeper shade of blue, reflecting binding of Cu²⁺ to the cyclen ring. To this mixture, 98 µL of SBT was added, giving a solution of 1.1 mM Cu²⁺ and 1 mM TETAC. The high concentrations in the binding reaction and 10% over-abundance of Cu²⁺ ensured that all the TETAC was bound with Cu²⁺. This stock solution was then diluted 1:100 when added to the cuvette for fluorometer experiments, giving a final concentration of 10 µM Cu²⁺-TETAC. Co²⁺ did not appear to be compatible with TETAC, eliminating its cysteine reactivity. 1,4-Dithiothreitol (DTT) (Sigma-Aldrich, St. Louis, MO) was prepared as a 100 mM stock in water.

Absorption measurements were made using a Beckman Coulter DU 800 (Brea, CA). To measure the absorption of Cu²⁺ and Co²⁺ bound to HH, we used an α-helical peptide with the following sequence: Ac-ACAAKHAAKHAAAAKA-NH₂, which was custom synthesized by Sigma-Aldrich (St. Louis, MO) (Taraska et al., 2009b). The peptide was dissolved in SBT to a final concentration of 1 mM. N-ethyl maleimide was prepared as a 200 mM stock in DMSO and added to the peptide solution at a final concentration of 2 mM to prevent the peptide from precipitating upon addition of Cu²⁺. To determine the extinction coefficient of Cu²⁺-TETAC, we measured the absorption spectrum of 2 mM Cu²⁺-cyclen.

Because the emission spectrum of L-Anap is sensitive to its environment, we measured the emission spectrum and quantum yield for L-Anap incorporated into each of our two sites in MBP. The emission spectrum of MBP-322Anap was blue shifted relative to that of MBP-295Anap by about 20 nm, suggesting that the L-Anap in MBP-322 is in a somewhat more hydrophobic environment (Figure 13A). To measure the quantum yield for each L-Anap position in MBP, we identified a solvent condition for free L-Anap that would mimic the environment of the fluorophores at each of the two sites. We measured the emission spectra of free L-Anap in SBT/ethanol mixtures that ranged from 0% to 100% ethanol. As shown in Figure 13B, both the peak wavelength and amplitude of the emission spectra were sensitive to ethanol. We found that the emission spectrum of MBP-295TAG matched most closely to the emissions spectrum of free L-Anap in 20% ethanol (Figure 13A, red), whereas the emission spectrum of MBP-322Anap matched most closely to free L-Anap in 85% ethanol (Figure 13A, blue).

Figure 13

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Next, we measure the quantum yield of L-Anap in the different SBT/ethanol mixtures. Samples of L-Anap were prepared in SBT, 20% ethanol in SBT, 85% ethanol in SBT, and 100% ethanol. The fluorescence intensity (excitation at 370 nm and emission at 480 nm) and absorption (at 370 nm) of each sample were measured as described above and plotted against each other in Figure 13C. The quantum yield of L-ANAP in each of the mixtures was calculated relative to the reference quantum yield of L-ANAP in ethanol (0.48) (Chatterjee et al., 2013) using the following equation (31):

where Q is the quantum yield of L-Anap, slope refers to the slope of the linear fits to the data, and η is the refractive index. Quantum yield values were 0.23, 0.31, and 0.47 for 0%, 20%, and 85% ethanol, respectively. We therefore estimated the quantum yield for MBP-295Anap of 0.31 and for MBP-322Anap of 0.47. These estimates assume that EtOH:SBT mixtures mimic the L-Anap environment in MBP and that there are no endogenous quenchers within MBP.

For each time course experiment in the fluorometer, an averaged background trace from six to eight experiments that did not contain protein, but to which Cu²⁺-TETAC and DTT, or Cu²⁺ and EDTA, were added, was subtracted from the protein-containing trace. The fraction of fluorescence quenching (F) was defined as follows:

To determine the FRET efficiency, E, we corrected for other sources of energy transfer (e.g. solution quenching) using the following equations:

where F_Cys and F_{no Cys} are the fractional quenching by Cu²⁺-TETAC in protein with and without cysteines respectively, and F_HH and F_{no HH} are the fraction of quenching by Cu²⁺ in protein with and without HH sites, respectively. We also analyzed our data using simplified equations, which would account for nonspecific decreases in fluorescence (e.g. bleaching or loss of protein) but not energy transfer:

The values of E produced by the two analysis methods were similar because of the low degree of background quenching. For clarity, only the results of analysis with the latter equations are shown. To calculate the mean and standard error of the mean for E, we used the mean and standard error of the mean for our F measurements (i.e. F_HH, F_{No HH}, F_Cys, and F_{No Cys}) in Monte Carlo resampling (1 × 10⁶ cycles; NIST Uncertainty Machine v1.3.4; [Lafarge and Possolo, 2015]).

Cells were imaged and unroofed as previously described (Gordon et al., 2016; Zagotta et al., 2016). Briefly, coverslips were mounted in a homemade chamber on a microscope stage and perfused for several minutes with HBR via gravity-flow perfusion. Poly-lysine solution (0.1 mg/mL 30,000–70,000 MW in PBS) was then added to the chamber and the cells were incubated for 15 s. A 1:3 dilution of SBT was then added to the chamber, and the cells were incubated for 30 s. The solution was then replaced with SBT and the cells immediately exposed to a single pulse from a Branson Sonifier 450 with MicroTip with power = 2 and duty cycle = 50%. Unroofed cells were then perfused with SBT for at least 2 min to remove debris from the chamber before images were collected.

Experiments were performed using a Nikon Ti-E inverted microscope (Melville, NY) and a Nikon CFI Apo TIRF 60x oil immersion objective. Excitation light was provided by a Xenon arc lamp (model Lambda LS, Sutter Instrument Co., Novato, CA). For L-Anap excitation we used a 375/28 nm excitation filter and a 480/50 nm emission filter. Each experiment commenced with a pre-bleaching step, comprised of a 7-s exposure of the coverslip to L-Anap excitation light. Pre-bleaching has been previously shown to reduce small, non-specific L-Anap dependent background fluorescence (Gordon et al., 2016). Images for analysis were collected using exposures ranging from 100 ms to 500 ms, depending on sample intensity, using a QuantEM EMCCD camera (Photometrics, Tuscon, AZ) with readout speed of 5 MHz and the multiplier set to 20. The illumination and camera settings were the same for all fluorescence images acquired for a given field of view. Different metal concentrations and EDTA were applied via gravity perfusion. Cu²⁺-TETAC and DTT were applied directly to the chamber via a transfer pipette. Five times the chamber volume was added in each case to ensure complete changeover of the solution.

Images were imported into ImageJ (Schneider et al., 2012) for analysis. Regions of interest were drawn by hand to include most of a given cell, but excluding any regions that appeared only partially unroofed, which tended to occur at the cell edges. For each cell, a background region was selected nearby that did not contain cells. The mean gray value of the background region of interest was subtracted from the mean gray value of the region of interest of the corresponding cell. This background subtraction was repeated for each image collected in an experiment using the same regions of interest. The mean and standard error of the mean for the FRET efficiency were calculated as described above.

The R₀ values for each L-Anap site paired with each type of bound metal (i.e. Cu²⁺ or Cu²⁺-TETAC) were calculated using the measured emission spectrum and quantum yield of L-Anap at each site and the measured absorption spectra for each type of bound metal using the following equation (Lakowicz, 2006):

where C is a scaling factor, J is the normalized spectral overlap of the emission of the donor and absorption of the acceptor, Q is the quantum yield of L-Anap at the given site (see above), η is the index of refraction (1.33 in our case), and κ² is the orientation factor, assumed to be 2/3, a reasonable assumption for an isotropic acceptor (15). Distances were calculated from the FRET measurements using the Förster equation:

FRET efficiencies assuming a Gaussian distribution of distances between donor and acceptor, with FWHM = 8 Å (i.e. σ=3.4) were determined by numerically convolving the Förster equation with the Gaussian function in Microsoft (Redmond, WA) Excel 2016. The corrected distances were then determined from plots of the FRET efficiency vs. the mean distance of the Gaussian distribution.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Research reported in this publication was supported by the National Eye Institute of the National Institutes of Health under award numbers R01EY017564 (to SEG) and R01EY010329 (to WNZ), by the National Institute of Mental Health of the National Institutes of Health under award number R01MH102378 (to WNZ), by the National Institute of General Medical Sciences of the National Institutes of Health under award numbers R01GM100718 and R01GM125351 (to SEG and WNZ), and by the following additional awards from the National Institutes of Health: S10RR025429, P30DK017047, and P30EY001730. The authors declare no competing financial interests.

1. Received: April 4, 2018
2. Accepted: June 5, 2018
3. Accepted Manuscript published: June 20, 2018 (version 1)
4. Version of Record published: July 23, 2018 (version 2)
5. Version of Record updated: November 30, 2018 (version 3)

© 2018, Gordon et al.

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