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Bifunctional Spin Labeling of Muscle Proteins
CHAPTER FOUR
Bifunctional Spin Labeling of Muscle Proteins: Accurate Rotational Dynamics, Orientation, and Distance by EPR Andrew R. Thompson, Benjamin P. Binder, Jesse E. McCaffrey, Bengt Svensson, David D. Thomas1 Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota, Minneapolis, Minnesota, USA 1 Corresponding author: e-mail address: [email protected]
Contents 1. 2. 3. 4.
Introduction Methods of BSL Labeling Rotational Dynamics Orientation 4.1 Mechanically Aligned Systems 4.2 Magnetically Aligned Systems 5. Distance 5.1 Distance Measurements with DEER 5.2 The Problem of Orientation Selection 6. Discussion 6.1 Labeling Specificity and Protein Function 6.2 The BEER Technique References
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Abstract While EPR allows for the characterization of protein structure and function due to its exquisite sensitivity to spin label dynamics, orientation, and distance, these measurements are often limited in sensitivity due to the use of labels that are attached via flexible monofunctional bonds, incurring additional disorder and nanosecond dynamics. In this chapter, we present methods for using a bifunctional spin label (BSL) to measure muscle protein structure and dynamics. We demonstrate that bifunctional attachment eliminates nanosecond internal rotation of the spin label, thereby allowing the accurate measurement of protein backbone rotational dynamics, including microsecond-tomillisecond motions by saturation transfer EPR. BSL also allows for accurate determination of helix orientation and disorder in mechanically and magnetically aligned systems,
Methods in Enzymology, Volume 564 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2015.06.029
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due to the label's stereospecific attachment. Similarly, labeling with a pair of BSL greatly enhances the resolution and accuracy of distance measurements measured by double electron–electron resonance (DEER). Finally, when BSL is applied to a protein with high helical content in an assembly with high orientational order (e.g., muscle fiber or membrane), two-probe DEER experiments can be combined with single-probe EPR experiments on an oriented sample in a process we call BEER, which has the potential for ab initio high-resolution structure determination.
1. INTRODUCTION EPR has long been a mainstay technique in our study of muscle proteins due to its ability to make a diverse number of measurements for the characterization of protein structure and function including solvent accessibility (Surek & Thomas, 2008), nanosecond to millisecond dynamics ( James, McCaffrey, Torgersen, Karim, & Thomas, 2012; Karim, Zhang, Howard, Torgersen, & Thomas, 2006; Thompson, Naber, Wilson, Cooke, & Thomas, 2008), orientation and disorder (Mello & Thomas, 2012; Moen, Thomas, & Klein, 2013), and distance measurement from 0.5 to >6 nm (Agafonov et al., 2009; Lin, Prochniewicz, James, Svensson, & Thomas, 2011; Moen, Klein, & Thomas, 2014). While complementary measurements are possible using fluorescence methods (Agafonov et al., 2009), spin labels offer distinct advantages over fluorescence probes due to their stability during measurement (i.e., they do not “bleach”), and their small size, typically on the order of a large amino acid side chain. In the measurement of distances, spin labels offer an additional advantage over fluorescent labels as they do not require distinct donor and acceptor probes. EPR measurements have typically involved the use of monofunctionally attached labels targeted to cysteine residues, present either natively in the protein structure or introduced via protein mutation, a process commonly referred to as site-directed spin labeling (SDSL) (Altenbach, Marti, Khorana, & Hubbell, 1990). Due to the innate flexibility of such monofunctional attachments, the spectrum is influenced by both label and protein conformations, such that the sensitivity of EPR to changes in protein structure and function may be limited or masked entirely by spin label mobility (Fig. 1, top). While careful choice of spin label variety and labeling site may allow for restriction of spin label mobility, neither is a panacea for highresolution measurement of dynamics, orientation, and distance. Bifunctional spin labels, especially the label 3,4-bis(methanethiosulfonylmethyl)-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-1-
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Figure 1 Conventional EPR spectra of the monofunctional spin label MTSSL versus the bifunctional spin label BSL attached to the myosin motor protein at positions 707 or 697/707, respectively. Data from Thompson et al. (2008).
yloxy spin label (HO-1944 or BSL) (Kalai, Balog, Jeko, & Hideg, 1999), eliminate these problems by addition of a second sulfhydryl linker, rigidly tethering the label to the protein backbone, thus eliminating the nanosecond rotational dynamics found in most monofunctionally attached labels (Fig. 1, bottom). When attached to a helix with cysteine residues engineered at i and i + 4 (or in some cases i and i + 3), BSL attains a stereospecifically defined orientation with respect to the helix axis (Fleissner et al., 2011), allowing for precise orientation measurements in oriented systems. The conformational restriction afforded by the second cysteine linker also allows for more precise measurement of distance distributions in dipolar EPR experiments, eliminating contributions from spin label flexibility in the measured distance distributions. In this chapter, we demonstrate the power of BSL over traditional monofunctionally attached labels in the context of the characterization of muscle protein structural dynamics. We illustrate how the reduction of spin label mobility allows for measurement of microsecond-to-millisecond protein rotational dynamics, as explored by saturation transfer EPR (STEPR) studies of the myosin motor domain. We examine orientation measurements on several helices in the motor domain of myosin in mechanically aligned skinned muscle fibers, as well as on the transmembrane helix of phospholamban (PLB) in magnetically aligned lipid bicelles. In both cases, the precise coupling of BSL to the protein backbone allows for accurate measurement of helix orientation with respect to the symmetry axis of the ordered system, with resolution on the order of 1 degree. Finally, we show how distance measurements in myosin are further refined by BSL, and how such measurements can be used to constrain the computation of
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helix orientation and provide information about the relative motion of domains upon the addition of nucleotide, a process we call bifunctional electron–electron resonance (BEER).
2. METHODS OF BSL LABELING With appropriate choices for labeling site and labeling conditions, attaching BSL to a protein is just as straightforward as attaching a conventional monofunctional spin label. BSL should be attached to cysteines on a helix at residues i and i + 4 or i and i + 3, or on a β-sheet at residues i and i + 2 (Fleissner et al., 2011; Islam & Roux, 2015). The selection of specific residues for labeling is highly specific to the protein and its environment, and is beyond the scope of this chapter. In general, though, one should choose residues that avoid major steric hindrances by side chains and tertiary structure to ensure optimal labeling efficiency and minimize disruption of protein structure and function. When selecting helix labeling sites, one should aim to select residues at least a half turn away from the helix ends to limit the possibility that the addition of the nitroxide side chain will disrupt the native secondary structure. Additionally, labeling a helix at i and i +4 has been shown to be less disruptive of the native helical structure than at i and i + 3 (Islam & Roux, 2015). The general labeling procedure is as follows: 1. The target cysteines for labeling are reduced via the addition of a reducing agent such as dithiothreitol or tris(2-carboxyethyl)phosphine. This solution is kept stirring on ice for 1 h. 2. The reducing agent is removed by dialysis and/or by two sequential sizeexclusion spin columns (preferred) such as a Pierce Zeba Spin desalting columns (Thermo Scientific). 3. The protein is labeled by the addition of 2–10 equivalents (spin labels/ bifunctional labeling sites) of BSL (3,4-bis-(methanethiosulfonylmethyl)2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-1-yloxy spin label, Toronto Research Chemicals) for 1–3 h on ice. 4. Excess unbound spin label is rapidly removed with two sequential spin columns. In this step, either centrifugal filters or spin columns may be used, but dialysis is not recommended because long equilibration times may contribute to a significant loss in labeling efficiency. 5. Labeled protein is either used immediately or stored in organic solvent such as trifluoroethanol at 20 °C or below to prevent the slow
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accumulation of BSL released from the protein through disulfide exchange (see Section 6; Fava, Iliceto, & Camera, 1957). 6. Spin labeling efficiency is quantified by spin counting of labeled protein dissolved in pH neutral buffer or reconstituted in lipid or detergent, using a known standard such as TEMPO or TEMPOL (Eaton, Eaton, Barr, & Weber, 2010). Specific reconstitution conditions such as buffer configuration and spin label regeneration can be found in the literature, including Arata et al. (2003), Mello and Thomas (2012), Moen et al. (2013), and Thompson et al. (2008).
3. ROTATIONAL DYNAMICS EPR is sensitive to rotational motion in the picosecond-tomillisecond range (Fig. 2). Despite their small size in comparison to fluorescent labels, spin labels can possess significant internal dynamics, which often manifest on the nanosecond timescale (Columbus, Kalai, Jeko, Hideg, & Hubbell, 2001). These motions are essentially indistinguishable from internal protein dynamics (detected by conventional EPR) (Goldman, Bruno, & Freed, 1972) and cause fast spectral diffusion which destroys the microsecond sensitivity of saturation transfer EPR (Squier & Thomas, 1986). Therefore, a rigidly bound label is essential for both fast and slow dynamics measurements. Fortuitous placement of a monofunctional spin label on a protein can result in immobilization due to local steric and electrostatic interactions
Figure 2 Sensitivity of conventional (left) and saturation transfer (right) EPR to isotropic rotational diffusion, shown by spectral simulations (Thomas & McConnell, 1974). τR is the rotational correlation time. Conventional EPR lacks sensitivity to dynamics for τR > 107 s, necessitating the use of saturation transfer EPR for these measurements.
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( James et al., 2012; Thompson et al., 2008), but this is usually not the case. Often, labeling sites are at the far exterior of a protein, in order to avoid perturbations in internal structure or oligomeric interactions, and hence exert few local constraints on label dynamics. To reduce this ambiguity, it is desirable to use a spin label that makes two bonds with the protein. The synthetic spin-labeled amino acid 2,2,6,6-tetramethyl-piperidine-1-oxyl-4-amino4-carboxyl (TOAC) spin label incorporates directly into the protein backbone via amide bonds, with two carbon atoms in the nitroxide-containing piperidine ring bonded directly to the α-carbon, and thus directly reports backbone dynamics (Karim, Kirby, Zhang, Nesmelov, & Thomas, 2004; Karim et al., 2006; Karim, Zhang, & Thomas, 2007). While compact and immobilized, TOAC requires incorporation via peptide synthesis and thus is not compatible with proteins with molecular weights above about 6 kDa. BSL is an alternative to TOAC that also offers rigid attachment relative to the peptide backbone, but is compatible with SDSL by cysteine mutagenesis (Kalai et al., 1999). By reacting with two Cys residues, the probe’s internal dynamics are restricted, resulting in the immobilization necessary to measure both fast and slow protein dynamics by conventional and saturation transfer EPR. Figure 3 shows a comparison between the monofunctional methanethiosulfonate spin label (MTSSL) and BSL attached to myosin at two different sites. At both sites, MTSSL demonstrates significant nanosecond rotational mobility as evidenced by narrow splittings in the spectrum. In contrast, BSL is significantly more immobilized, as indicated by the near rigid-limit splitting value. Figure 4 shows the conventional EPR spectra
Figure 3 Conventional EPR spectra of MTSSL and BSL attached to myosin at several different locations. (A) 697 or 697/707, respectively. (B) 492 or 492/496, respectively.
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Figure 4 Singly and doubly attached BSL on the transmembrane helix of PLB at positions 32 and 32/36, respectively. Singly attached BSL (2Tjj0 ¼ 61.7 G) is only slightly less immobilized than doubly attached BSL (2Tjj0 ¼ 62.2 G).
Figure 5 Conventional and saturation transfer EPR spectra of BSL-labeled myosin free in solution and bound to actin filaments. Rotational correlation times are determined from L00 /L. Data adapted from Thompson et al. (2008).
of singly and doubly attached BSL on PLBs transmembrane helix. The outer splitting of singly attached BSL is slightly narrower, and the linewidth is slightly greater, indicating that it is only slightly more mobile than doubly attached BSL. This is consistent with previous studies that evaluated the effect of adding a bulky side chain to a monofunctional spin labels (Columbus et al., 2001). Figure 5 shows the conventional and saturation transfer EPR spectra of BSL–myosin, both free in solution and bound to actin. On the conventional
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EPR timescale (picoseconds to nanoseconds), the EPR spectra are quite similar and possess the characteristic powder lineshape, indicating strong label immobilization. In contrast, the saturation transfer EPR spectra are quite different, revealing a large decrease in myosin rotational mobility upon binding to actin, corresponding to an increase in rotational correlation time (τ R) from 500 ns to 600 μs. Had BSL not been immobilized on myosin as shown in Fig. 5, the protein’s microsecond global dynamics would not be detected, as the label’s nanosecond dynamics would dominate the saturation transfer measurement (Wilcox, Parce, Thomas, & Lyles, 1990).
4. ORIENTATION 4.1 Mechanically Aligned Systems The attachment of BSL is not only rigid on the conventional EPR time scale; it is also stereospecific with respect to the peptide backbone, permitting direct measurement of protein backbone orientation in a well-ordered, anisotropic system. The study of sarcomeric proteins stands to benefit greatly from this feature. Actin filaments within a muscle fiber can be oriented by positioning a fiber bundle within the cavity, thus setting up a biological scaffold upon which actin-binding proteins of interest may also be oriented (Fig. 6). Myosin is an excellent candidate for validating and exploiting this method, due to the high-binding affinity of the actomyosin complex and the abundance of mechanochemical coupling within its catalytic domain (CD) (Spudich, 2014; Thomas, Prochniewicz, & Roopnarine, 2002). Figure 7C shows EPR spectra of myosin labeled with either MTSSL or BSL at equivalent sites on the C-terminal end of the relay helix. Skinned muscle fiber bundles were incubated with the spin-labeled protein and oriented with the actin symmetry axis either parallel or perpendicular to the applied magnetic field. Spectra of oriented MTSSL–myosin show poor orientational resolution, evidenced by virtually indistinguishable parallel and perpendicular lineshapes, while spectra of BSL show great disparity between fiber orientations, indicating a highly ordered spin label ensemble. When these decorated fiber bundles are minced, removing all spectral contributions from orientation (Fig. 7C), the MTSSL ensemble also reveals significant motional narrowing, while the BSL spectrum exhibits a characteristic powder lineshape, indicating negligible nanosecond motion. The stereospecific attachment of BSL greatly simplifies the analysis of the resulting spectra and enables the derivation of label angular distributions. For the actomyosin data discussed above, the magnetic, hyperfine, and linewidth
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Figure 6 (A) Reference frame that defines the orientation dependence of nitroxide EPR (Earle & Budil, 2006). In the nitroxide frame, the applied magnetic field vector B forms the angles θNB and θNB relative to the nitroxide frame. (B) In oriented actomyosin samples, the actin filament forms the assembly axis vector A, which is usually aligned with B.
tensors can be determined from spectra of minced fibers, where there are no contributions from orientation. These values can then be fixed and applied to spectral simulations of the oriented samples, drastically reducing the parameter space of the subsequent fitting operation. With tensors thus predefined, the shape of the spectrum is entirely dependent on the ensemble spin label orientation, defined by θNB (axial orientation of the magnetic field in the nitroxide frame), ϕNB (azimuthal orientation in the nitroxide frame), and Δθ,ϕ (width of the angular distribution) (Fig. 6A). The extremely high-anisotropic sensitivity of EPR grants exceptional resolution to these parameters in the spectrum, such that localized changes in the orientation of individual structural elements are easily resolved; an example of this is given in Fig. 7E, where a nucleotide-induced structural change is detected on myosin’s relay helix. In samples with the symmetry axis parallel to the magnetic field, these angular parameters also describe label orientation relative to the symmetry axis (actin, in this case), and therefore θNB ¼ θNA and ϕNB ¼ ϕNA in Fig. 6B.
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Figure 7 (A) MTSSL and BSL attached to the myosin relay helix. (B) Myosin bound to actin within an oriented muscle fiber. (C) Conventional EPR spectra of actomyosin complexes labeled with MTSSL and BSL within oriented muscle fibers demonstrate orientation sensitivity. Minced (randomly oriented) fiber samples show nanosecond dynamics of MTSSL but not BSL. (D and E) Addition of ADP reveals a previously undetected rotation of the relay helix within actin-bound myosin.
From there, powerful structural constraints for the system can be developed by taking advantage of the stereospecific nature of BSL’s bifunctional attachment. For example, deriving the axial tilt of a myosin helix relative to actin requires two vectors in a common coordinate frame, one representing the actin vector, and the other representing the helix (Fig. 8). The actin vector can be accurately defined using the EPR-derived angle parameters (Fig. 8B), and the helix vector can be determined relative to the nitroxide frame (Fig. 8D) by geometric analysis of available crystal structures containing BSL (Fleissner et al., 2011). The helix tilt angle θAH is subsequently derived by finding the angle between these two vectors, thus generating a highresolution structural constraint independent of the spin label itself (Fig. 8F). Measurements obtained in this way for several sites across the myosin CD are in agreement with previous results from cryoelectron microscopy, in the absence of nucleotide (Holmes, Angert, Kull, Jahn, & Schroder, 2003). The real power of these derivations, though, lies in the ability to obtain constraints under a variety of biochemical conditions, potentially on the same protein sample. Figure 7D depicts the result of modeling the nucleotide-
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Figure 8 (A–D) Visualization of coordinate transformations used in the analysis of oriented muscle fiber EPR. Projection of the actin and helix vectors on the nitroxide frame yield the relative angle θAH, the axial tilt of a myosin helix relative to actin (E and F).
induced structural change on the relay helix, revealing a bending deformation that is similar to crystallographic results from myosin alone (Fisher et al., 1995; Smith & Rayment, 1996), but not previously observed for the actomyosin complex.
4.2 Magnetically Aligned Systems In addition to sarcomeric proteins such as myosin, BSL has great potential for structural elucidation of membrane proteins through orientation measurements. Membrane-bound proteins often exhibit nanosecond backbone dynamics due to their fluid environment, resulting in additional spectral effects detected by conventional EPR that complicate the measurement of orientation. While this motion can be resolved through spectral simulation, rigid label attachment is necessary to decouple label and protein dynamics, allowing determination of orientation. Indeed, a rigidly bound label such as BSL is absolutely essential for this type of measurement.
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To start, BSL-labeled membrane proteins must be reconstituted in an anisotropic environment to avoid orientational averaging. This can be accomplished through a mechanical bilayer such as a substrate-supported bilayer (Inbaraj, Laryukhin, & Lorigan, 2007), or a magnetically aligned bilayer such as bicelles (Cardon, Tiburu, & Lorigan, 2003). Bicelles offer a well-hydrated, homogeneous environment quite analogous to native vesicles, and isolated sarcoplasmic reticulum. They have seen extensive use in nuclear magnetic resonance (NMR) studies due to the high magnetic field available (De Angelis & Opella, 2007; Durr, Soong, & Ramamoorthy, 2013), but with lipid optimization and the addition of lanthanides, bicelles have seen increasing use in EPR experiments performed under physiological conditions and temperatures (Caporini, Padmanabhan, Cardon, & Lorigan, 2003; Cho, Dominick, & Spence, 2010; Garber, Lorigan, & Howard, 1999; Lu, Caporini, & Lorigan, 2004; McCaffrey, James, & Thomas, 2015). Figure 9 shows EPR spectra of PLB with BSL at positions 32/36 in aligned bicelles. The substantial differences in the spectra shown in Fig. 9A indicate a narrow, well-defined orientation distribution. Fitting by spectral simulation determines a label tilt angle of 90° 3°. Figure 9B shows complementary molecular dynamics simulations of BSL on PLB’s transmembrane helix, yielding an angle of 89° between the probe principal
Figure 9 (A) EPR spectra of BSL attached at positions 32 and 36 on the transmembrane helix of PLB. Spectral analysis of this data yields a label tilt angle of 90° 3°. (B) Molecular dynamics simulation of BSL on PLB finds a label tilt angle of 89° 5°, consistent with the experimental result from (A).
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axis and the membrane normal. These results are consistent with the analysis of Fig. 9A, as well as solid state NMR measurements of PLB (Traaseth, Buffy, Zamoon, & Veglia, 2006) along with the crystal structure of BSL as determined by Fleissner and coworkers (Fleissner et al., 2011). Advanced EPR spectral simulation software such as NLSL (Budil, Lee, Saxena, & Freed, 1996; Khairy, Fajer, & Budil, 2002) and MultiComponent (Altenbach, Flitsch, Khorana, & Hubbell, 1989) allow partial macroscopic order microscopic disorder models that can accommodate nanosecond rotational motion and anisotropic orientational distributions characteristic of aligned bicelle EPR. Because of the complexity of analyzing spectra affected by both rotational motion and orientational anisotropy, acquiring spectra of control samples is recommended to determine magnetic and dynamics parameters, such as frozen samples and isotropic vesicle/bicelle samples (Zhang et al., 2010). These allow for better determination of label orientation, which affects the spectral splitting as does rotational motion. Figure 10 shows a general scheme for analysis of EPR measurements on oriented samples. Magnetic tensors (electron g and hyperfine A) are determined from frozen or otherwise immobilized sample spectra, as they lack significant dynamics and orientational anisotropy. These tensor values are carried into the analysis of the randomly oriented sample (e.g., minced muscle fiber, vesicles, or isotropic bicelles) to determine dynamics parameters such as rotational correlation time (R or τR) and order parameter (S, realted to the simulated orienting potential coefficient c20). These values are then carried into the analysis of the aligned sample spectra to determine label orientation relative to the magnetic field (diffusion angles αD, βD, and γ D). To further restrict the fitting space during this step, multiple sample orientations (usually with the symmetry axis parallel and perpendicular to the applied field) can be globally analyzed. The final step (if applicable) uses previous atomic structures of the spin label in a similar environment to transform the label diffusion angles into helix angles that relate the helix orientation to the symmetry axis. A full explanation of EPR fitting parameters and notation in the context of the simulation program NLSL is available from Earle and Budil (2006).
5. DISTANCE 5.1 Distance Measurements with DEER Distance measurements using EPR have the distinct advantage over the complementary fluorescence technique FRET in that there is no need to
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Figure 10 Typical workflow for EPR spectral analysis in oriented measurements.
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Figure 11 (A) The conformational flexibility of monofunctional labels introduces uncertainty in distance measurements between labels that is eliminated by bifunctional attachment. (B) DEER waveforms of MSL at position 498 and 639 on myosin, and BSL at positions 494/498 and 639/643 on myosin. (C) MSL reveals a single, broad distance, while BSL reveals two distinct distance populations.
introduce distinct donor and acceptor pairs in the system of interest (Kast, Espinoza-Fonseca, Yi, & Thomas, 2010); identical spin labels can be used and the distance is measured via the dipolar interaction. The two techniques most frequently used are continuous wave dipolar EPR, sensitive to distances between 0.8 and 2 nm (Rabenstein & Shin, 1995), and double electron– electron resonance (DEER), which is capable of measuring distances from 2 to 6 nm in typical protein experiments (Pannier, Veit, Godt, Jeschke, & Spiess, 2000). DEER, in particular, is a powerful technique for the characterization of protein structure as the resultant waveforms are explicitly encoded with the distribution of distances between nearby labels ( Jeschke, 2012). The sensitivity of DEER to distance distributions is diminished by label flexibility, with monofunctionally attached labels typically producing broader, less defined distributions (Fig. 11C; Fleissner et al., 2011; Islam & Roux, 2015; Sahu et al., 2013). Label flexibility also reduces the reliability of detecting observed changes in distance distribution, as true structural changes may remain unresolved or, conversely, observed changes may be due merely to a change in label conformation. These ambiguities are eliminated through the use of the BSL.
5.2 The Problem of Orientation Selection While the restricted label mobility afforded by BSL improves the colocalization of the label to the protein backbone, the rigid coupling introduces a new challenge to accurately analyzing data recorded by DEER, namely it introduces the effect of orientation selection (Marko et al., 2009). The traditional analysis of the DEER waveform assumes that there
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is no orientational dependence in the dipolar interaction between the interacting spin labels, which, in the case of BSL, is clearly not true. Fortunately, in the two most common frequency domains under which DEER is performed, X- and Q-band, orientation selection can be largely suppressed by following the standard protocol of pumping at the maximum of the EPR absorption spectrum and choosing nonselective pulses ( Jeschke, 2012). Even under these conditions, though, the distribution width (but not center) extracted from the recorded waveforms is sensitive to choice of the position of the probe pulse, indicative of at least low levels of orientation selection, as shown in Fig. 12.
6. DISCUSSION 6.1 Labeling Specificity and Protein Function While BSL possesses superior sensitivity to rotational dynamics, orientation, and distance, there are several technical points to consider when handling BSL and analyzing the data. First and foremost, labeling specificity is an important consideration for any spin label, but BSL in particular due to its two sulfhydryl groups. If the target protein contains native Cys residues, there is always a risk of nonspecific labeling that should be assessed with appropriate assays. To evaluate nonspecific labeling, spin counting and mass spectrometry should be used to determine the label-to-protein ratio (Eaton et al., 2010; Karim et al., 2004). In addition to these routine measurements, conventional EPR on BSL-labeled proteins in oriented systems can also provide compelling evidence for nonspecific labeling. Indeed, measurements of dynamics alone are insufficient to identify nonspecific labeling, as singly attached BSL is often almost as immobilized as doubly attached BSL (Fig. 4). However, EPR spectra of oriented samples reveal the broader orientational distribution characteristic of singly attached spin labels (Fig. 13). In particular, singly attached labels tend to produce a powder-like spectrum, which is very distinct from the oriented components that typically arise from bifunctionally attached BSL. Thus, the presence of disordered spectral components from an oriented sample suggests nonspecific labeling. The distinct behavior of singly and doubly attached BSL ultimately represents another advantage over monofunctional spin labels, as the doubly attached component of a spectrum can be identified and treated separately from any additional artifacts. Spectral resolution with BSL is usually sufficient to analyze both ordered and disordered components independently,
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Figure 12 The presence of orientation selection was explored on myosin labeled at 492/496 and 639/643 with BSL via the acquisition of DEER at multiple pump-observe positions (A). The resultant DEER waveforms (B) were analyzed using LongDistances by Christian Altenbach, available for free download at http://www.chemistry.ucla.edu/ directory/hubbell-wayne-l. (C) Changes in the distance distributions obtained from the waveforms in (B) are due to orientation selection.
pulling out accurate orientation parameters even when a disordered component is present. While orientation measurements with BSL are thus significantly less hindered by specificity concerns than traditional techniques, potential for nonspecific labeling should still be minimized as much as possible to avoid problems in DEER analysis and functional perturbation by additional modification. If nonspecific labeling is apparent, it can be
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Figure 13 Orientational order is pronounced in BSL spectra of oriented systems when bifunctional attachment is present. (A) Conventional EPR spectra from myosin on oriented fibers, labeled monofunctionally and bifunctionally with BSL on the regulatory light chain at positions 122 and 122/126, respectively. (B) Conventional EPR spectra from myosin on oriented fibers, labeled monofunctionally with MTSSL or BSL at position 639, or bifunctionally with BSL at positions 639/643 on the myosin head. (C) Conventional EPR spectra from unaligned and aligned bicelles containing PLB labeled monofunctionally with BSL at position 32. See Fig. 9A for corresponding spectra with bifunctional BSL attachment.
addressed by careful substitution of native Cys with benign nonreactive analogs (such as Ser and Leu) (Shih, Gryczynski, Lakowicz, & Spudich, 2000). In protein systems with engineered helical attachments points, BSL favors bifunctional attachment, presumably due to the rapid kinetics associated with disulfide bond formation. With nonideal cysteine orientations, monofunctional double labeling of the protein by BSL is possible even with steric restrictions, but this has been shown to be an uncommon occurrence
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(Fleissner et al., 2011). While bifunctionally attached BSL is quite stable in practice, the release of label through disulfide exchange is apparent by the accumulation of unbound label over time. While we have not studied these effects in detail, potential mechanisms are the presence of reducing agent, or the formation of inter- or intra-disulfide bonds. As disulfide exchange is strongly temperature dependent, reversibility of BSL attachment can be minimized by maintaining samples at 4 °C or lower and is virtually eliminated by storage below 0 °C (Fava et al., 1957). As with any approach that employs site-directed spectroscopy, it is essential to perform appropriate functional assays for the protein in question to determine whether the introduction of labeling sites and/or bifunctional labeling perturbs function. Small changes in enzymatic activity and/or substrate binding affinity are expected even with well-chosen labeling sites, but care should be taken to assess the extent of perturbation, and to move labeling sites if necessary. In addition to the considerations applicable to conventional labels, such as the potential for steric hindrance, bifunctional labeling can also hyperstabilize α-helices, and so should not be deployed on α-helical segments for which high flexibility or helical unfolding is key to function (Fleissner et al., 2011).
6.2 The BEER Technique While the aforementioned techniques for measuring distance and orientation are powerful in their own right, BSL presents an unusual opportunity for parallel analysis, using the features of one method to constrain the other. Such analysis, here referred to collectively as BEER, represents a means to perform direct modeling of individual protein structural elements de novo, based on a small number of experiments. As discussed above, deployment of BSL in oriented systems can deliver accurate axial tilt angles for individual protein helices relative to the sample’s symmetry axis. However, axial tilt alone is not enough to model each helix within a protein’s tertiary structure, because the azimuthal angle with respect to the symmetry axis will always be degenerate. Furthermore, the intrinsic angular degeneracy of EPR allows for more than one potential helix tilt angle in most cases. These issues can often be resolved as long as a reference structure exists to contextualize the results, but for proteins without general established models, orientation measurements alone are not enough. In BEER, narrow intra-protein distance constraints obtained by DEER can significantly constrain the potential solutions given by orientation measurements, identifying the most likely
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conformer. Once the orientation of the given helices is thus established, the stereospecificity of BSL can be exploited to directly calculate the relative orientations of the labels used in DEER. This information may in turn be used to constrain the fitting of DEER waveforms by directly addressing orientation selection. BEER analysis may thus be performed iteratively to progressively constrain both results until an optimum solution is reached de novo, much like the model for analysis currently employed in structure determination by crystallography and NMR spectroscopy.
REFERENCES Agafonov, R. V., Negrashov, I. V., Tkachev, Y. V., Blakely, S. E., Titus, M. A., Thomas, D. D., et al. (2009). Structural dynamics of the myosin relay helix by timeresolved EPR and FRET. Proceedings of the National Academy of Sciences of the United States of America, 106(51), 21625–21630. Altenbach, C., Flitsch, S. L., Khorana, H. G., & Hubbell, W. L. (1989). Structural studies on transmembrane proteins. 2. Spin labeling of bacteriorhodopsin mutants at unique cysteines. Biochemistry, 28(19), 7806–7812. http://dx.doi.org/10.1021/Bi00445a042. Altenbach, C., Marti, T., Khorana, H. G., & Hubbell, W. L. (1990). Transmembrane protein structure: Spin labeling of bacteriorhodopsin mutants. Science, 248(4959), 1088–1092. Arata, T., Nakamura, M., Akahane, H., Aihara, T., Ueki, S., Sugata, K., et al. (2003). Orientation and motion of myosin light chain and troponin in reconstituted muscle fibers as detected by ESR with a new bifunctional spin label. Advances in Experimental Medicine and Biology, 538, 279–283. discussion 284. Budil, D. E., Lee, S., Saxena, S., & Freed, J. H. (1996). Nonlinear-least-squares analysis of slow-motion EPR spectra in one and two dimensions using a modified LevenbergMarquardt algorithm. Journal of Magnetic Resonance, Series A, 120(2), 155–189. http:// dx.doi.org/10.1006/jmra.1996.0113. Caporini, M. A., Padmanabhan, A., Cardon, T. B., & Lorigan, G. A. (2003). Investigating magnetically aligned phospholipid bilayers with various lanthanide ions for X-band spinlabel EPR studies. Biochimica et Biophysica Acta-Biomembranes, 1612(1), 52–58. http://dx. doi.org/10.1016/S0005-2736(03)00085-3. Cardon, T. B., Tiburu, E. K., & Lorigan, G. A. (2003). Magnetically aligned phospholipid bilayers in weak magnetic fields: Optimization, mechanism, and advantages for X-band EPR studies. Journal of Magnetic Resonance, 161(1), 77–90. Cho, H. S., Dominick, J. L., & Spence, M. M. (2010). Lipid domains in bicelles containing unsaturated lipids and cholesterol. Journal of Physical Chemistry B, 114(28), 9238–9245. http://dx.doi.org/10.1021/Jp100276u. Columbus, L., Kalai, T., Jeko, J., Hideg, K., & Hubbell, W. L. (2001). Molecular motion of spin labeled side chains in alpha-helices: Analysis by variation of side chain structure. Biochemistry, 40(13), 3828–3846. De Angelis, A. A., & Opella, S. J. (2007). Bicelle samples for solid-state NMR of membrane proteins. Nature Protocols, 2(10), 2332–2338. http://dx.doi.org/10.1038/ nprot.2007.329. Durr, U. H. N., Soong, R., & Ramamoorthy, A. (2013). When detergent meets bilayer: Birth and coming of age of lipid bicelles. Progress in Nuclear Magnetic Resonance Spectroscopy, 69, 1–22. http://dx.doi.org/10.1016/j.pnmrs.2013.01.001.
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Earle, K. A., & Budil, D. E. (2006). Calculating slow-motion ESR spectra of spin-labeled polymers. In S. Schlick (Ed.), Advanced ESR methods in polymer research (pp. 53–83). New York: John Wiley & Sons, Inc. Eaton, G. R., Eaton, S. S., Barr, D. P., & Weber, R. T. (2010). Quantitative EPR. Vienna: Springer. Fava, A., Iliceto, A., & Camera, E. (1957). Kinetics of the thiol-disulfide exchange. Journal of the American Chemical Society, 79(4), 833–838. http://dx.doi.org/10.1021/ja01561a014. Fisher, A. J., Smith, C. A., Thoden, J., Smith, R., Sutoh, K., Holden, H. M., et al. (1995). Structural studies of myosin:nucleotide complexes: A revised model for the molecular basis of muscle contraction. Biophysical Journal, 68(Suppl. 4), 19S–26S. discussion 27S–28S. Fleissner, M. R., Bridges, M. D., Brooks, E. K., Cascio, D., Kalai, T., Hideg, K., et al. (2011). Structure and dynamics of a conformationally constrained nitroxide side chain and applications in EPR spectroscopy. Proceedings of the National Academy of Sciences of the United States of America, 108(39), 16241–16246. http://dx.doi.org/10.1073/pnas.1111420108. Garber, S. M., Lorigan, G. A., & Howard, K. P. (1999). Magnetically oriented phospholipid bilayers for spin label EPR studies. Journal of the American Chemical Society, 121(13), 3240–3241. http://dx.doi.org/10.1021/Ja984371f. Goldman, S. A., Bruno, G. V., & Freed, J. H. (1972). Estimating slow-motional rotational correlation times for nitroxides by electron spin resonance. The Journal of Physical Chemistry, 76(13), 1858–1860. http://dx.doi.org/10.1021/j100657a013. Holmes, K. C., Angert, I., Kull, F. J., Jahn, W., & Schroder, R. R. (2003). Electron cryomicroscopy shows how strong binding of myosin to actin releases nucleotide. Nature, 425(6956), 423–427. Inbaraj, J. J., Laryukhin, M., & Lorigan, G. A. (2007). Determining the helical tilt angle of a transmembrane helix in mechanically aligned lipid bilayers using EPR spectroscopy. Journal of the American Chemical Society, 129(25), 7710–7711. http://dx.doi.org/10.1021/ ja071587l. Islam, S. M., & Roux, B. (2015). Simulating the distance distribution between spin-labels attached to proteins. The Journal of Physical Chemistry. B, 119(10), 3901–3911. http:// dx.doi.org/10.1021/jp510745d. James, Z. M., McCaffrey, J. E., Torgersen, K. D., Karim, C. B., & Thomas, D. D. (2012). Protein-protein interactions in calcium transport regulation probed by saturation transfer electron paramagnetic resonance. Biophysical Journal, 103(6), 1370–1378. http://dx.doi. org/10.1016/j.bpj.2012.08.032 (pii). Jeschke, G. (2012). DEER distance measurements on proteins. Annual Review of Physical Chemistry, 63, 419–446. http://dx.doi.org/10.1146/annurev-physchem-032511143716. Kalai, T., Balog, M., Jeko, J., & Hideg, K. (1999). Synthesis and reactions of a symmetric paramagnetic pyrrolidine diene. Synthesis, 6, 973–980. Karim, C. B., Kirby, T. L., Zhang, Z., Nesmelov, Y., & Thomas, D. D. (2004). Phospholamban structural dynamics in lipid bilayers probed by a spin label rigidly coupled to the peptide backbone. Proceedings of the National Academy of Sciences of the United States of America, 101(40), 14437–14442. Karim, C. B., Zhang, Z., Howard, E. C., Torgersen, K. D., & Thomas, D. D. (2006). Phosphorylation-dependent conformational switch in spin-labeled phospholamban bound to SERCA. Journal of Molecular Biology, 358(4), 1032–1040. Karim, C. B., Zhang, Z., & Thomas, D. D. (2007). Synthesis of TOAC spin-labeled proteins and reconstitution in lipid membranes. Nature Protocols, 2(1), 42–49. Kast, D., Espinoza-Fonseca, L. M., Yi, C., & Thomas, D. D. (2010). Phosphorylationinduced structural changes in smooth muscle myosin regulatory light chain. Proceedings of the National Academy of Sciences of the United States of America, 107(18), 8207–8212.
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Khairy, K., Fajer, P., & Budil, D. (2002). Simulation of spin label motion in EPR spectra of muscle fibers. Biophysical Journal, 82(1), 479a. Lin, A. Y., Prochniewicz, E., James, Z., Svensson, B., & Thomas, D. D. (2011). Large-scale opening of utrophin’s tandem CH domains upon actin binding, by an induced-fit mechanism. Proceedings of the National Academy of Sciences of the United States of America, 108(31), 12729–12733. Lu, J. X., Caporini, M. A., & Lorigan, G. A. (2004). The effects of cholesterol on magnetically aligned phospholipid bilayers: A solid-state NMR and EPR spectroscopy study. Journal of Magnetic Resonance, 168(1), 18–30. http://dx.doi.org/10.1016/j. jmr.2004.01.013. Marko, A., Margraf, D., Yu, H., Mu, Y., Stock, G., & Prisner, T. (2009). Molecular orientation studies by pulsed electron–electron double resonance experiments. The Journal of Chemical Physics, 130(6), 064102. http://dx.doi.org/10.1063/1.3073040. McCaffrey, J. E., James, Z. M., & Thomas, D. D. (2015). Optimization of bicelle lipid composition and temperature for EPR spectroscopy of aligned membranes. Journal of Magnetic Resonance, 250, 71–75. http://dx.doi.org/10.1016/j.jmr.2014.09.026. Mello, R. N., & Thomas, D. D. (2012). Three distinct actin-attached structural states of myosin in muscle fibers. Biophysical Journal, 102(5), 1088–1096. http://dx.doi.org/ 10.1016/j.bpj.2011.11.4027 (pii). Moen, R. J., Klein, J. C., & Thomas, D. D. (2014). Electron paramagnetic resonance resolves effects of oxidative stress on muscle proteins. Exercise and Sport Sciences Reviews, 42(1), 30–36. http://dx.doi.org/10.1249/JES.0000000000000004. Moen, R. J., Thomas, D. D., & Klein, J. C. (2013). Conformationally trapping the actinbinding cleft of myosin with a bifunctional spin label. The Journal of Biological Chemistry, 288(5), 3016–3024. http://dx.doi.org/10.1074/jbc.M112.428565. Pannier, M., Veit, S., Godt, A., Jeschke, G., & Spiess, H. W. (2000). Dead-time free measurement of dipole-dipole interactions between electron spins. Journal of Magnetic Resonance, 142(2), 331–340. http://dx.doi.org/10.1006/jmre.1999.1944. Rabenstein, M. D., & Shin, Y. K. (1995). Determination of the distance between 2 spin labels attached to a macromolecule. Proceedings of the National Academy of Sciences of the United States of America, 92(18), 8239–8243. http://dx.doi.org/10.1073/pnas.92.18.8239. Sahu, I. D., McCarrick, R. M., Troxel, K. R., Zhang, R., Smith, H. J., Dunagan, M. M., et al. (2013). DEER EPR measurements for membrane protein structures via bifunctional spin labels and lipodisq nanoparticles. Biochemistry, 52(38), 6627–6632. http:// dx.doi.org/10.1021/bi4009984. Shih, W. M., Gryczynski, Z., Lakowicz, J. R., & Spudich, J. A. (2000). A FRET-based sensor reveals large ATP hydrolysis-induced conformational changes and three distinct states of the molecular motor myosin. Cell, 102(5), 683–694. Smith, C. A., & Rayment, I. (1996). X-ray structure of the magnesium(II).ADP.vanadate complex of the dictyostelium discoideum myosin motor domain to 1.9 A˚ resolution. Biochemistry, 35(17), 5404–5417. Spudich, J. A. (2014). Hypertrophic and dilated cardiomyopathy: Four decades of basic research on muscle lead to potential therapeutic approaches to these devastating genetic diseases. Biophysical Journal, 106(6), 1236–1249. http://dx.doi.org/10.1016/j. bpj.2014.02.011. Squier, T. C., & Thomas, D. D. (1986). Methodology for increased precision in saturation transfer electron paramagnetic resonance studies of rotational dynamics. Biophysical Journal, 49(4), 921–935. Surek, J. T., & Thomas, D. D. (2008). A paramagnetic molecular voltmeter. Journal of Magnetic Resonance, 190(1), 7–25. Thomas, D. D., & McConnell, H. M. (1974). Calculation of paramagnetic resonance spectra sensitive to very slow rotational motion. Chemical Physics Letters, 25, 470–475.
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Thomas, D. D., Prochniewicz, E., & Roopnarine, O. (2002). Changes in actin and myosin structural dynamics due to their weak and strong interactions. Results and Problems in Cell Differentiation, 36, 7–19. Thompson, A. R., Naber, N., Wilson, C., Cooke, R., & Thomas, D. D. (2008). Structural dynamics of the actomyosin complex probed by a bifunctional spin label that cross-links SH1 and SH2. Biophysical Journal, 95(11), 5238–5246. Traaseth, N. J., Buffy, J. J., Zamoon, J., & Veglia, G. (2006). Structural dynamics and topology of phospholamban in oriented lipid bilayers using multidimensional solid-state NMR. Biochemistry, 45(46), 13827–13834. http://dx.doi.org/10.1021/Bi0607610. Wilcox, M., Parce, J., Thomas, M., & Lyles, D. (1990). A new bifunctional spin-label suitable for saturation-transfer EPR studies of protein rotational motion. Biochemistry, 29(24), 5734–5743. Zhang, Z. W., Fleissner, M. R., Tipikin, D. S., Liang, Z. C., Moscicki, J. K., Earle, K. A., et al. (2010). Multifrequency electron spin resonance study of the dynamics of spin labeled T4 lysozyme. Journal of Physical Chemistry B, 114(16), 5503–5521. http://dx. doi.org/10.1021/Jp910606h.
Bifunctional Spin Labeling of Muscle Proteins: Accurate Rotational Dynamics, Orientation, and Distance by EPR Andrew R. Thompson, Benjamin P. Binder, Jesse E. McCaffrey, Bengt Svensson, David D. Thomas1 Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota, Minneapolis, Minnesota, USA 1 Corresponding author: e-mail address: [email protected]
Contents 1. 2. 3. 4.
Introduction Methods of BSL Labeling Rotational Dynamics Orientation 4.1 Mechanically Aligned Systems 4.2 Magnetically Aligned Systems 5. Distance 5.1 Distance Measurements with DEER 5.2 The Problem of Orientation Selection 6. Discussion 6.1 Labeling Specificity and Protein Function 6.2 The BEER Technique References
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Abstract While EPR allows for the characterization of protein structure and function due to its exquisite sensitivity to spin label dynamics, orientation, and distance, these measurements are often limited in sensitivity due to the use of labels that are attached via flexible monofunctional bonds, incurring additional disorder and nanosecond dynamics. In this chapter, we present methods for using a bifunctional spin label (BSL) to measure muscle protein structure and dynamics. We demonstrate that bifunctional attachment eliminates nanosecond internal rotation of the spin label, thereby allowing the accurate measurement of protein backbone rotational dynamics, including microsecond-tomillisecond motions by saturation transfer EPR. BSL also allows for accurate determination of helix orientation and disorder in mechanically and magnetically aligned systems,
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due to the label's stereospecific attachment. Similarly, labeling with a pair of BSL greatly enhances the resolution and accuracy of distance measurements measured by double electron–electron resonance (DEER). Finally, when BSL is applied to a protein with high helical content in an assembly with high orientational order (e.g., muscle fiber or membrane), two-probe DEER experiments can be combined with single-probe EPR experiments on an oriented sample in a process we call BEER, which has the potential for ab initio high-resolution structure determination.
1. INTRODUCTION EPR has long been a mainstay technique in our study of muscle proteins due to its ability to make a diverse number of measurements for the characterization of protein structure and function including solvent accessibility (Surek & Thomas, 2008), nanosecond to millisecond dynamics ( James, McCaffrey, Torgersen, Karim, & Thomas, 2012; Karim, Zhang, Howard, Torgersen, & Thomas, 2006; Thompson, Naber, Wilson, Cooke, & Thomas, 2008), orientation and disorder (Mello & Thomas, 2012; Moen, Thomas, & Klein, 2013), and distance measurement from 0.5 to >6 nm (Agafonov et al., 2009; Lin, Prochniewicz, James, Svensson, & Thomas, 2011; Moen, Klein, & Thomas, 2014). While complementary measurements are possible using fluorescence methods (Agafonov et al., 2009), spin labels offer distinct advantages over fluorescence probes due to their stability during measurement (i.e., they do not “bleach”), and their small size, typically on the order of a large amino acid side chain. In the measurement of distances, spin labels offer an additional advantage over fluorescent labels as they do not require distinct donor and acceptor probes. EPR measurements have typically involved the use of monofunctionally attached labels targeted to cysteine residues, present either natively in the protein structure or introduced via protein mutation, a process commonly referred to as site-directed spin labeling (SDSL) (Altenbach, Marti, Khorana, & Hubbell, 1990). Due to the innate flexibility of such monofunctional attachments, the spectrum is influenced by both label and protein conformations, such that the sensitivity of EPR to changes in protein structure and function may be limited or masked entirely by spin label mobility (Fig. 1, top). While careful choice of spin label variety and labeling site may allow for restriction of spin label mobility, neither is a panacea for highresolution measurement of dynamics, orientation, and distance. Bifunctional spin labels, especially the label 3,4-bis(methanethiosulfonylmethyl)-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-1-
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Figure 1 Conventional EPR spectra of the monofunctional spin label MTSSL versus the bifunctional spin label BSL attached to the myosin motor protein at positions 707 or 697/707, respectively. Data from Thompson et al. (2008).
yloxy spin label (HO-1944 or BSL) (Kalai, Balog, Jeko, & Hideg, 1999), eliminate these problems by addition of a second sulfhydryl linker, rigidly tethering the label to the protein backbone, thus eliminating the nanosecond rotational dynamics found in most monofunctionally attached labels (Fig. 1, bottom). When attached to a helix with cysteine residues engineered at i and i + 4 (or in some cases i and i + 3), BSL attains a stereospecifically defined orientation with respect to the helix axis (Fleissner et al., 2011), allowing for precise orientation measurements in oriented systems. The conformational restriction afforded by the second cysteine linker also allows for more precise measurement of distance distributions in dipolar EPR experiments, eliminating contributions from spin label flexibility in the measured distance distributions. In this chapter, we demonstrate the power of BSL over traditional monofunctionally attached labels in the context of the characterization of muscle protein structural dynamics. We illustrate how the reduction of spin label mobility allows for measurement of microsecond-to-millisecond protein rotational dynamics, as explored by saturation transfer EPR (STEPR) studies of the myosin motor domain. We examine orientation measurements on several helices in the motor domain of myosin in mechanically aligned skinned muscle fibers, as well as on the transmembrane helix of phospholamban (PLB) in magnetically aligned lipid bicelles. In both cases, the precise coupling of BSL to the protein backbone allows for accurate measurement of helix orientation with respect to the symmetry axis of the ordered system, with resolution on the order of 1 degree. Finally, we show how distance measurements in myosin are further refined by BSL, and how such measurements can be used to constrain the computation of
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helix orientation and provide information about the relative motion of domains upon the addition of nucleotide, a process we call bifunctional electron–electron resonance (BEER).
2. METHODS OF BSL LABELING With appropriate choices for labeling site and labeling conditions, attaching BSL to a protein is just as straightforward as attaching a conventional monofunctional spin label. BSL should be attached to cysteines on a helix at residues i and i + 4 or i and i + 3, or on a β-sheet at residues i and i + 2 (Fleissner et al., 2011; Islam & Roux, 2015). The selection of specific residues for labeling is highly specific to the protein and its environment, and is beyond the scope of this chapter. In general, though, one should choose residues that avoid major steric hindrances by side chains and tertiary structure to ensure optimal labeling efficiency and minimize disruption of protein structure and function. When selecting helix labeling sites, one should aim to select residues at least a half turn away from the helix ends to limit the possibility that the addition of the nitroxide side chain will disrupt the native secondary structure. Additionally, labeling a helix at i and i +4 has been shown to be less disruptive of the native helical structure than at i and i + 3 (Islam & Roux, 2015). The general labeling procedure is as follows: 1. The target cysteines for labeling are reduced via the addition of a reducing agent such as dithiothreitol or tris(2-carboxyethyl)phosphine. This solution is kept stirring on ice for 1 h. 2. The reducing agent is removed by dialysis and/or by two sequential sizeexclusion spin columns (preferred) such as a Pierce Zeba Spin desalting columns (Thermo Scientific). 3. The protein is labeled by the addition of 2–10 equivalents (spin labels/ bifunctional labeling sites) of BSL (3,4-bis-(methanethiosulfonylmethyl)2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-1-yloxy spin label, Toronto Research Chemicals) for 1–3 h on ice. 4. Excess unbound spin label is rapidly removed with two sequential spin columns. In this step, either centrifugal filters or spin columns may be used, but dialysis is not recommended because long equilibration times may contribute to a significant loss in labeling efficiency. 5. Labeled protein is either used immediately or stored in organic solvent such as trifluoroethanol at 20 °C or below to prevent the slow
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accumulation of BSL released from the protein through disulfide exchange (see Section 6; Fava, Iliceto, & Camera, 1957). 6. Spin labeling efficiency is quantified by spin counting of labeled protein dissolved in pH neutral buffer or reconstituted in lipid or detergent, using a known standard such as TEMPO or TEMPOL (Eaton, Eaton, Barr, & Weber, 2010). Specific reconstitution conditions such as buffer configuration and spin label regeneration can be found in the literature, including Arata et al. (2003), Mello and Thomas (2012), Moen et al. (2013), and Thompson et al. (2008).
3. ROTATIONAL DYNAMICS EPR is sensitive to rotational motion in the picosecond-tomillisecond range (Fig. 2). Despite their small size in comparison to fluorescent labels, spin labels can possess significant internal dynamics, which often manifest on the nanosecond timescale (Columbus, Kalai, Jeko, Hideg, & Hubbell, 2001). These motions are essentially indistinguishable from internal protein dynamics (detected by conventional EPR) (Goldman, Bruno, & Freed, 1972) and cause fast spectral diffusion which destroys the microsecond sensitivity of saturation transfer EPR (Squier & Thomas, 1986). Therefore, a rigidly bound label is essential for both fast and slow dynamics measurements. Fortuitous placement of a monofunctional spin label on a protein can result in immobilization due to local steric and electrostatic interactions
Figure 2 Sensitivity of conventional (left) and saturation transfer (right) EPR to isotropic rotational diffusion, shown by spectral simulations (Thomas & McConnell, 1974). τR is the rotational correlation time. Conventional EPR lacks sensitivity to dynamics for τR > 107 s, necessitating the use of saturation transfer EPR for these measurements.
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( James et al., 2012; Thompson et al., 2008), but this is usually not the case. Often, labeling sites are at the far exterior of a protein, in order to avoid perturbations in internal structure or oligomeric interactions, and hence exert few local constraints on label dynamics. To reduce this ambiguity, it is desirable to use a spin label that makes two bonds with the protein. The synthetic spin-labeled amino acid 2,2,6,6-tetramethyl-piperidine-1-oxyl-4-amino4-carboxyl (TOAC) spin label incorporates directly into the protein backbone via amide bonds, with two carbon atoms in the nitroxide-containing piperidine ring bonded directly to the α-carbon, and thus directly reports backbone dynamics (Karim, Kirby, Zhang, Nesmelov, & Thomas, 2004; Karim et al., 2006; Karim, Zhang, & Thomas, 2007). While compact and immobilized, TOAC requires incorporation via peptide synthesis and thus is not compatible with proteins with molecular weights above about 6 kDa. BSL is an alternative to TOAC that also offers rigid attachment relative to the peptide backbone, but is compatible with SDSL by cysteine mutagenesis (Kalai et al., 1999). By reacting with two Cys residues, the probe’s internal dynamics are restricted, resulting in the immobilization necessary to measure both fast and slow protein dynamics by conventional and saturation transfer EPR. Figure 3 shows a comparison between the monofunctional methanethiosulfonate spin label (MTSSL) and BSL attached to myosin at two different sites. At both sites, MTSSL demonstrates significant nanosecond rotational mobility as evidenced by narrow splittings in the spectrum. In contrast, BSL is significantly more immobilized, as indicated by the near rigid-limit splitting value. Figure 4 shows the conventional EPR spectra
Figure 3 Conventional EPR spectra of MTSSL and BSL attached to myosin at several different locations. (A) 697 or 697/707, respectively. (B) 492 or 492/496, respectively.
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Figure 4 Singly and doubly attached BSL on the transmembrane helix of PLB at positions 32 and 32/36, respectively. Singly attached BSL (2Tjj0 ¼ 61.7 G) is only slightly less immobilized than doubly attached BSL (2Tjj0 ¼ 62.2 G).
Figure 5 Conventional and saturation transfer EPR spectra of BSL-labeled myosin free in solution and bound to actin filaments. Rotational correlation times are determined from L00 /L. Data adapted from Thompson et al. (2008).
of singly and doubly attached BSL on PLBs transmembrane helix. The outer splitting of singly attached BSL is slightly narrower, and the linewidth is slightly greater, indicating that it is only slightly more mobile than doubly attached BSL. This is consistent with previous studies that evaluated the effect of adding a bulky side chain to a monofunctional spin labels (Columbus et al., 2001). Figure 5 shows the conventional and saturation transfer EPR spectra of BSL–myosin, both free in solution and bound to actin. On the conventional
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EPR timescale (picoseconds to nanoseconds), the EPR spectra are quite similar and possess the characteristic powder lineshape, indicating strong label immobilization. In contrast, the saturation transfer EPR spectra are quite different, revealing a large decrease in myosin rotational mobility upon binding to actin, corresponding to an increase in rotational correlation time (τ R) from 500 ns to 600 μs. Had BSL not been immobilized on myosin as shown in Fig. 5, the protein’s microsecond global dynamics would not be detected, as the label’s nanosecond dynamics would dominate the saturation transfer measurement (Wilcox, Parce, Thomas, & Lyles, 1990).
4. ORIENTATION 4.1 Mechanically Aligned Systems The attachment of BSL is not only rigid on the conventional EPR time scale; it is also stereospecific with respect to the peptide backbone, permitting direct measurement of protein backbone orientation in a well-ordered, anisotropic system. The study of sarcomeric proteins stands to benefit greatly from this feature. Actin filaments within a muscle fiber can be oriented by positioning a fiber bundle within the cavity, thus setting up a biological scaffold upon which actin-binding proteins of interest may also be oriented (Fig. 6). Myosin is an excellent candidate for validating and exploiting this method, due to the high-binding affinity of the actomyosin complex and the abundance of mechanochemical coupling within its catalytic domain (CD) (Spudich, 2014; Thomas, Prochniewicz, & Roopnarine, 2002). Figure 7C shows EPR spectra of myosin labeled with either MTSSL or BSL at equivalent sites on the C-terminal end of the relay helix. Skinned muscle fiber bundles were incubated with the spin-labeled protein and oriented with the actin symmetry axis either parallel or perpendicular to the applied magnetic field. Spectra of oriented MTSSL–myosin show poor orientational resolution, evidenced by virtually indistinguishable parallel and perpendicular lineshapes, while spectra of BSL show great disparity between fiber orientations, indicating a highly ordered spin label ensemble. When these decorated fiber bundles are minced, removing all spectral contributions from orientation (Fig. 7C), the MTSSL ensemble also reveals significant motional narrowing, while the BSL spectrum exhibits a characteristic powder lineshape, indicating negligible nanosecond motion. The stereospecific attachment of BSL greatly simplifies the analysis of the resulting spectra and enables the derivation of label angular distributions. For the actomyosin data discussed above, the magnetic, hyperfine, and linewidth
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Figure 6 (A) Reference frame that defines the orientation dependence of nitroxide EPR (Earle & Budil, 2006). In the nitroxide frame, the applied magnetic field vector B forms the angles θNB and θNB relative to the nitroxide frame. (B) In oriented actomyosin samples, the actin filament forms the assembly axis vector A, which is usually aligned with B.
tensors can be determined from spectra of minced fibers, where there are no contributions from orientation. These values can then be fixed and applied to spectral simulations of the oriented samples, drastically reducing the parameter space of the subsequent fitting operation. With tensors thus predefined, the shape of the spectrum is entirely dependent on the ensemble spin label orientation, defined by θNB (axial orientation of the magnetic field in the nitroxide frame), ϕNB (azimuthal orientation in the nitroxide frame), and Δθ,ϕ (width of the angular distribution) (Fig. 6A). The extremely high-anisotropic sensitivity of EPR grants exceptional resolution to these parameters in the spectrum, such that localized changes in the orientation of individual structural elements are easily resolved; an example of this is given in Fig. 7E, where a nucleotide-induced structural change is detected on myosin’s relay helix. In samples with the symmetry axis parallel to the magnetic field, these angular parameters also describe label orientation relative to the symmetry axis (actin, in this case), and therefore θNB ¼ θNA and ϕNB ¼ ϕNA in Fig. 6B.
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Figure 7 (A) MTSSL and BSL attached to the myosin relay helix. (B) Myosin bound to actin within an oriented muscle fiber. (C) Conventional EPR spectra of actomyosin complexes labeled with MTSSL and BSL within oriented muscle fibers demonstrate orientation sensitivity. Minced (randomly oriented) fiber samples show nanosecond dynamics of MTSSL but not BSL. (D and E) Addition of ADP reveals a previously undetected rotation of the relay helix within actin-bound myosin.
From there, powerful structural constraints for the system can be developed by taking advantage of the stereospecific nature of BSL’s bifunctional attachment. For example, deriving the axial tilt of a myosin helix relative to actin requires two vectors in a common coordinate frame, one representing the actin vector, and the other representing the helix (Fig. 8). The actin vector can be accurately defined using the EPR-derived angle parameters (Fig. 8B), and the helix vector can be determined relative to the nitroxide frame (Fig. 8D) by geometric analysis of available crystal structures containing BSL (Fleissner et al., 2011). The helix tilt angle θAH is subsequently derived by finding the angle between these two vectors, thus generating a highresolution structural constraint independent of the spin label itself (Fig. 8F). Measurements obtained in this way for several sites across the myosin CD are in agreement with previous results from cryoelectron microscopy, in the absence of nucleotide (Holmes, Angert, Kull, Jahn, & Schroder, 2003). The real power of these derivations, though, lies in the ability to obtain constraints under a variety of biochemical conditions, potentially on the same protein sample. Figure 7D depicts the result of modeling the nucleotide-
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Figure 8 (A–D) Visualization of coordinate transformations used in the analysis of oriented muscle fiber EPR. Projection of the actin and helix vectors on the nitroxide frame yield the relative angle θAH, the axial tilt of a myosin helix relative to actin (E and F).
induced structural change on the relay helix, revealing a bending deformation that is similar to crystallographic results from myosin alone (Fisher et al., 1995; Smith & Rayment, 1996), but not previously observed for the actomyosin complex.
4.2 Magnetically Aligned Systems In addition to sarcomeric proteins such as myosin, BSL has great potential for structural elucidation of membrane proteins through orientation measurements. Membrane-bound proteins often exhibit nanosecond backbone dynamics due to their fluid environment, resulting in additional spectral effects detected by conventional EPR that complicate the measurement of orientation. While this motion can be resolved through spectral simulation, rigid label attachment is necessary to decouple label and protein dynamics, allowing determination of orientation. Indeed, a rigidly bound label such as BSL is absolutely essential for this type of measurement.
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To start, BSL-labeled membrane proteins must be reconstituted in an anisotropic environment to avoid orientational averaging. This can be accomplished through a mechanical bilayer such as a substrate-supported bilayer (Inbaraj, Laryukhin, & Lorigan, 2007), or a magnetically aligned bilayer such as bicelles (Cardon, Tiburu, & Lorigan, 2003). Bicelles offer a well-hydrated, homogeneous environment quite analogous to native vesicles, and isolated sarcoplasmic reticulum. They have seen extensive use in nuclear magnetic resonance (NMR) studies due to the high magnetic field available (De Angelis & Opella, 2007; Durr, Soong, & Ramamoorthy, 2013), but with lipid optimization and the addition of lanthanides, bicelles have seen increasing use in EPR experiments performed under physiological conditions and temperatures (Caporini, Padmanabhan, Cardon, & Lorigan, 2003; Cho, Dominick, & Spence, 2010; Garber, Lorigan, & Howard, 1999; Lu, Caporini, & Lorigan, 2004; McCaffrey, James, & Thomas, 2015). Figure 9 shows EPR spectra of PLB with BSL at positions 32/36 in aligned bicelles. The substantial differences in the spectra shown in Fig. 9A indicate a narrow, well-defined orientation distribution. Fitting by spectral simulation determines a label tilt angle of 90° 3°. Figure 9B shows complementary molecular dynamics simulations of BSL on PLB’s transmembrane helix, yielding an angle of 89° between the probe principal
Figure 9 (A) EPR spectra of BSL attached at positions 32 and 36 on the transmembrane helix of PLB. Spectral analysis of this data yields a label tilt angle of 90° 3°. (B) Molecular dynamics simulation of BSL on PLB finds a label tilt angle of 89° 5°, consistent with the experimental result from (A).
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axis and the membrane normal. These results are consistent with the analysis of Fig. 9A, as well as solid state NMR measurements of PLB (Traaseth, Buffy, Zamoon, & Veglia, 2006) along with the crystal structure of BSL as determined by Fleissner and coworkers (Fleissner et al., 2011). Advanced EPR spectral simulation software such as NLSL (Budil, Lee, Saxena, & Freed, 1996; Khairy, Fajer, & Budil, 2002) and MultiComponent (Altenbach, Flitsch, Khorana, & Hubbell, 1989) allow partial macroscopic order microscopic disorder models that can accommodate nanosecond rotational motion and anisotropic orientational distributions characteristic of aligned bicelle EPR. Because of the complexity of analyzing spectra affected by both rotational motion and orientational anisotropy, acquiring spectra of control samples is recommended to determine magnetic and dynamics parameters, such as frozen samples and isotropic vesicle/bicelle samples (Zhang et al., 2010). These allow for better determination of label orientation, which affects the spectral splitting as does rotational motion. Figure 10 shows a general scheme for analysis of EPR measurements on oriented samples. Magnetic tensors (electron g and hyperfine A) are determined from frozen or otherwise immobilized sample spectra, as they lack significant dynamics and orientational anisotropy. These tensor values are carried into the analysis of the randomly oriented sample (e.g., minced muscle fiber, vesicles, or isotropic bicelles) to determine dynamics parameters such as rotational correlation time (R or τR) and order parameter (S, realted to the simulated orienting potential coefficient c20). These values are then carried into the analysis of the aligned sample spectra to determine label orientation relative to the magnetic field (diffusion angles αD, βD, and γ D). To further restrict the fitting space during this step, multiple sample orientations (usually with the symmetry axis parallel and perpendicular to the applied field) can be globally analyzed. The final step (if applicable) uses previous atomic structures of the spin label in a similar environment to transform the label diffusion angles into helix angles that relate the helix orientation to the symmetry axis. A full explanation of EPR fitting parameters and notation in the context of the simulation program NLSL is available from Earle and Budil (2006).
5. DISTANCE 5.1 Distance Measurements with DEER Distance measurements using EPR have the distinct advantage over the complementary fluorescence technique FRET in that there is no need to
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Figure 10 Typical workflow for EPR spectral analysis in oriented measurements.
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Figure 11 (A) The conformational flexibility of monofunctional labels introduces uncertainty in distance measurements between labels that is eliminated by bifunctional attachment. (B) DEER waveforms of MSL at position 498 and 639 on myosin, and BSL at positions 494/498 and 639/643 on myosin. (C) MSL reveals a single, broad distance, while BSL reveals two distinct distance populations.
introduce distinct donor and acceptor pairs in the system of interest (Kast, Espinoza-Fonseca, Yi, & Thomas, 2010); identical spin labels can be used and the distance is measured via the dipolar interaction. The two techniques most frequently used are continuous wave dipolar EPR, sensitive to distances between 0.8 and 2 nm (Rabenstein & Shin, 1995), and double electron– electron resonance (DEER), which is capable of measuring distances from 2 to 6 nm in typical protein experiments (Pannier, Veit, Godt, Jeschke, & Spiess, 2000). DEER, in particular, is a powerful technique for the characterization of protein structure as the resultant waveforms are explicitly encoded with the distribution of distances between nearby labels ( Jeschke, 2012). The sensitivity of DEER to distance distributions is diminished by label flexibility, with monofunctionally attached labels typically producing broader, less defined distributions (Fig. 11C; Fleissner et al., 2011; Islam & Roux, 2015; Sahu et al., 2013). Label flexibility also reduces the reliability of detecting observed changes in distance distribution, as true structural changes may remain unresolved or, conversely, observed changes may be due merely to a change in label conformation. These ambiguities are eliminated through the use of the BSL.
5.2 The Problem of Orientation Selection While the restricted label mobility afforded by BSL improves the colocalization of the label to the protein backbone, the rigid coupling introduces a new challenge to accurately analyzing data recorded by DEER, namely it introduces the effect of orientation selection (Marko et al., 2009). The traditional analysis of the DEER waveform assumes that there
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is no orientational dependence in the dipolar interaction between the interacting spin labels, which, in the case of BSL, is clearly not true. Fortunately, in the two most common frequency domains under which DEER is performed, X- and Q-band, orientation selection can be largely suppressed by following the standard protocol of pumping at the maximum of the EPR absorption spectrum and choosing nonselective pulses ( Jeschke, 2012). Even under these conditions, though, the distribution width (but not center) extracted from the recorded waveforms is sensitive to choice of the position of the probe pulse, indicative of at least low levels of orientation selection, as shown in Fig. 12.
6. DISCUSSION 6.1 Labeling Specificity and Protein Function While BSL possesses superior sensitivity to rotational dynamics, orientation, and distance, there are several technical points to consider when handling BSL and analyzing the data. First and foremost, labeling specificity is an important consideration for any spin label, but BSL in particular due to its two sulfhydryl groups. If the target protein contains native Cys residues, there is always a risk of nonspecific labeling that should be assessed with appropriate assays. To evaluate nonspecific labeling, spin counting and mass spectrometry should be used to determine the label-to-protein ratio (Eaton et al., 2010; Karim et al., 2004). In addition to these routine measurements, conventional EPR on BSL-labeled proteins in oriented systems can also provide compelling evidence for nonspecific labeling. Indeed, measurements of dynamics alone are insufficient to identify nonspecific labeling, as singly attached BSL is often almost as immobilized as doubly attached BSL (Fig. 4). However, EPR spectra of oriented samples reveal the broader orientational distribution characteristic of singly attached spin labels (Fig. 13). In particular, singly attached labels tend to produce a powder-like spectrum, which is very distinct from the oriented components that typically arise from bifunctionally attached BSL. Thus, the presence of disordered spectral components from an oriented sample suggests nonspecific labeling. The distinct behavior of singly and doubly attached BSL ultimately represents another advantage over monofunctional spin labels, as the doubly attached component of a spectrum can be identified and treated separately from any additional artifacts. Spectral resolution with BSL is usually sufficient to analyze both ordered and disordered components independently,
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Figure 12 The presence of orientation selection was explored on myosin labeled at 492/496 and 639/643 with BSL via the acquisition of DEER at multiple pump-observe positions (A). The resultant DEER waveforms (B) were analyzed using LongDistances by Christian Altenbach, available for free download at http://www.chemistry.ucla.edu/ directory/hubbell-wayne-l. (C) Changes in the distance distributions obtained from the waveforms in (B) are due to orientation selection.
pulling out accurate orientation parameters even when a disordered component is present. While orientation measurements with BSL are thus significantly less hindered by specificity concerns than traditional techniques, potential for nonspecific labeling should still be minimized as much as possible to avoid problems in DEER analysis and functional perturbation by additional modification. If nonspecific labeling is apparent, it can be
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Figure 13 Orientational order is pronounced in BSL spectra of oriented systems when bifunctional attachment is present. (A) Conventional EPR spectra from myosin on oriented fibers, labeled monofunctionally and bifunctionally with BSL on the regulatory light chain at positions 122 and 122/126, respectively. (B) Conventional EPR spectra from myosin on oriented fibers, labeled monofunctionally with MTSSL or BSL at position 639, or bifunctionally with BSL at positions 639/643 on the myosin head. (C) Conventional EPR spectra from unaligned and aligned bicelles containing PLB labeled monofunctionally with BSL at position 32. See Fig. 9A for corresponding spectra with bifunctional BSL attachment.
addressed by careful substitution of native Cys with benign nonreactive analogs (such as Ser and Leu) (Shih, Gryczynski, Lakowicz, & Spudich, 2000). In protein systems with engineered helical attachments points, BSL favors bifunctional attachment, presumably due to the rapid kinetics associated with disulfide bond formation. With nonideal cysteine orientations, monofunctional double labeling of the protein by BSL is possible even with steric restrictions, but this has been shown to be an uncommon occurrence
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(Fleissner et al., 2011). While bifunctionally attached BSL is quite stable in practice, the release of label through disulfide exchange is apparent by the accumulation of unbound label over time. While we have not studied these effects in detail, potential mechanisms are the presence of reducing agent, or the formation of inter- or intra-disulfide bonds. As disulfide exchange is strongly temperature dependent, reversibility of BSL attachment can be minimized by maintaining samples at 4 °C or lower and is virtually eliminated by storage below 0 °C (Fava et al., 1957). As with any approach that employs site-directed spectroscopy, it is essential to perform appropriate functional assays for the protein in question to determine whether the introduction of labeling sites and/or bifunctional labeling perturbs function. Small changes in enzymatic activity and/or substrate binding affinity are expected even with well-chosen labeling sites, but care should be taken to assess the extent of perturbation, and to move labeling sites if necessary. In addition to the considerations applicable to conventional labels, such as the potential for steric hindrance, bifunctional labeling can also hyperstabilize α-helices, and so should not be deployed on α-helical segments for which high flexibility or helical unfolding is key to function (Fleissner et al., 2011).
6.2 The BEER Technique While the aforementioned techniques for measuring distance and orientation are powerful in their own right, BSL presents an unusual opportunity for parallel analysis, using the features of one method to constrain the other. Such analysis, here referred to collectively as BEER, represents a means to perform direct modeling of individual protein structural elements de novo, based on a small number of experiments. As discussed above, deployment of BSL in oriented systems can deliver accurate axial tilt angles for individual protein helices relative to the sample’s symmetry axis. However, axial tilt alone is not enough to model each helix within a protein’s tertiary structure, because the azimuthal angle with respect to the symmetry axis will always be degenerate. Furthermore, the intrinsic angular degeneracy of EPR allows for more than one potential helix tilt angle in most cases. These issues can often be resolved as long as a reference structure exists to contextualize the results, but for proteins without general established models, orientation measurements alone are not enough. In BEER, narrow intra-protein distance constraints obtained by DEER can significantly constrain the potential solutions given by orientation measurements, identifying the most likely
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conformer. Once the orientation of the given helices is thus established, the stereospecificity of BSL can be exploited to directly calculate the relative orientations of the labels used in DEER. This information may in turn be used to constrain the fitting of DEER waveforms by directly addressing orientation selection. BEER analysis may thus be performed iteratively to progressively constrain both results until an optimum solution is reached de novo, much like the model for analysis currently employed in structure determination by crystallography and NMR spectroscopy.
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