Temperature Dependence of CN and SCN IR Absorptions Facilitates

Nov 2, 2015 - Cyano and thiocyano groups have received attention as IR probes of local protein electrostatics or solvation, due to their strong absorp...
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Temperature Dependence of CN and SCN IR Absorptions Facilitates Their Interpretation and Use as Probes of Proteins Ramkrishna Adhikary, Jörg Zimmermann, Philip E. Dawson, and Floyd E. Romesberg* Department of Chemistry, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, California 92037, United States S Supporting Information *

ABSTRACT: Cyano and thiocyano groups have received attention as IR probes of local protein electrostatics or solvation, due to their strong absorptions and the ability to site specifically incorporate them within proteins. However, interpreting their spectra requires knowing whether they engage in hydrogen bonds (H-bonds). Existing methods for the detection of such H-bonding interactions are based on structural analysis or correlations between IR and NMR signals and are labor intensive and possibly ambiguous. Here, using model systems we show that the absorption frequency of both probes is linearly correlated with temperature and that the slope of the resulting line (frequency−temperature line slope or FTLS) reflects the nature of the probe’s microenvironment, including whether or not the probe is engaged in H-bonds. We then show that the same linear dependence is observed with p-cyano phenylalanine, cyanylated cysteine, or cyanylated homocysteine incorporated at different positions within the N-terminal Src homology 3 domain of the murine adapter protein Crk-II. The FTLSs indicate that p-cyano phenylalanine incorporated at two positions is engaged in strong H-bonding, while it is involved in weaker H-bonding at a third position. In contrast, the FTLS of the cyanylated cysteine or cyanylated homocysteine absorptions indicates that they do not engage in H-bonding at either a buried or surface exposed position. While the differences likely reflect side chain flexibility and the probe’s ability to avoid solvent, the data suggest that the temperature dependence of the absorption provides a simple method to gauge the probe’s environment, including the presence of H-bonding.

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correlation between absorption frequency and dielectric constant in protic solvents,4,6 as in this case hydrogen-bonding (H-bonding) with the (S)CN moiety introduces significant quadrupole-mediated contributions to the absorption frequency.21−24 Problematically, virtually all biological environments possess potential H-bond donors that may interact with the (S)CN probe and introduce such interactions. Thus, the use of (S)CN groups as probes of protein electrostatics at a given position requires knowing that they are free of H-bonding interactions. In contrast, the use of (S)CN groups as probes of solvation requires knowing that they engage in H-bonds with water. The determination of whether an (S)CN moiety incorporated at a specific position within a protein engages in Hbonding has previously been accomplished via structure elucidation6,7 or by comparing IR absorption frequencies and 13 C NMR chemical shifts.4,6 However, the detection of Hbonds via X-ray crystallography requires high resolution data, which aside from the experimental challenges is also impractical for the many sites of a protein that may be of interest, and NMR shifts are only qualitatively interpretable in terms of electric fields due to variable contributions of magnetic

he ability to characterize proteins with high spatial and temporal resolution is central to understanding their function. The most direct tool, vibrational spectroscopy, is complicated by spectral complexity, but this limitation may be overcome with the use of probes that absorb IR light between 1800−2600 cm−1, a spectral region that is free of obscuring protein absorptions. Carbon−deuterium (C−D) bonds were the first such probes to be incorporated into proteins,1 and they have the advantage of being nonperturbative but the disadvantage of being relatively weak chromophores. The incorporation of cyano (CN), thiocyanate (SCN), and azide (N3) probes into amino acids, peptides, or proteins has since been explored, in large part due to their stronger absorptions.2,3 Much effort has been focused on using CN and SCN (collectively here referred to as (S)CN) groups as site-specific probes of protein electrostatics, 4−12 or as probes of solvation.13−19 Their use as probes of electrostatics is based on the well-known linear relationship between the frequency of a chromophore’s absorption and its electric field strength as measured by Stark spectroscopy (electrochromism).4,6,20 Indeed, the stretching absorptions of model (S)CN probes exhibit a linear correlation with dielectric constant in aprotic solvents.4,6 However, the linear dependence (i.e., Stark-like behavior) requires a dominance of the solvatochromic dipoleelectric field interaction relative to higher multipolar contributions, such as those mediated by the solvatochromic quadrupole.21 Correspondingly, (S)CN probes show no © 2015 American Chemical Society

Received: September 9, 2015 Accepted: October 21, 2015 Published: November 2, 2015 11561

DOI: 10.1021/acs.analchem.5b03437 Anal. Chem. 2015, 87, 11561−11567

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anisotropy. A more simple method to determine whether (S)CN IR absorptions are complicated by H-bonding, preferably one that requires minimal additional effort, would facilitate the use and further development of these probes. In aqueous buffer, the absorption spectrum of an (S)CN group consists of a single absorption or possibly overlapping absorptions, with the higher frequency component assigned to a strongly H-bonded species and a lower frequency component assigned to a less, or more weakly H-bonded species.25 Consistent with this interpretation, the absorptions of model (S)CN systems shift to higher frequency in more strongly Hbonding solvents.16 Importantly, the shifts are significantly larger than those observed with different aprotic solvents.4,6 Thus, we reasoned that under conditions that weaken Hbonding, such as increased temperature, (S)CN absorptions that are engaged in H-bonds should show large and diagnostic shifts to lower frequency. Here, we report a systematic study of the temperature dependence of the absorptions of (S)CN groups of model systems in solvents with varying physicochemical properties, including H-bond donor strength. In all cases, the absorption frequencies show a linear dependence on temperature with a unique and thus diagnostic slope (denoted here as the frequency−temperature line slope, FTLS), which is largest with the strongest H-bonding solvents. We then use these dependencies to analyze the local environments of the (S)CN probes of p-cyano phenylalanine ((CN)Phe), cyanylated cysteine ((SCN)Cys), or homocysteine ((SCN)HCys) incorporated at different positions of the N-terminal Src homology 3 domain of the murine adaptor protein Crk-II (nSH3) (Figure 1). The observed temperature-dependence of

Article

EXPERIMENTAL SECTION

Chemicals for solid phase peptide synthesis were obtained as described previously,26,27 except for Boc-Cys(Mbzl)−OH and Boc-Homocys(MBzl)−OH, which were purchased from BACHEM (King of Prussia, PA). 1-Cyano-4-dimethylaminopyridinium tetrafluoroborate (CDAP) was obtained from Sigma. p-Tolunitrile (98%) and methyl thiocyanate (>99%) were purchased from Sigma-Aldrich (St. Louis, MO) and TCI (Portland, OR), respectively. Cyclohexane (99.9%) and isopropyl alcohol (99.9%) were obtained from Fisher Chemicals, dimethyl sulfoxide (99.7%) and formamide (99%) were obtained from Acros Organics (New Jersey), and toluene (99.5%) was obtained from EMD Millipore (Billerica, MA). Synthesis and Purification of Cyanylated nSH3 Variants. Mutants of nSH3 with either a cysteine or a homocysteine at positions 159 or 181 were synthesized using Boc solid phase peptide synthesis. Synthesis was performed as described previously26,27 using p-methylbenzhydrylamine resin and the coupling reagent HCTU with Boc-Cys(Mbzl)−OH or Boc-Homocys(MBzl)−OH incorporated at the desired position. Peptides were cleaved from dry resin with HF and purified by preparative reversed-phase HPLC, which was performed with a C18 reversed-phase column using a linear gradient of solvent A (0.1% (v/v) aqueous TFA) and solvent B (90% acetonitrile (v/v), 10% water (v/v), 0.1% TFA (v/v)) of 30% B to 40% B over 60 min and a constant flow rate of 15 mL/min, while monitoring absorption at 230 nm. Fractions containing pure protein, confirmed by mass spectroscopy, were combined and lyophilized. The single sulfhydryl group of each nSH3 variant was cyanylated as described previously.28,29 Briefly, two stock solutions were prepared: a 500 μM protein solution in 100 mM citrate buffer (pH 3.0) containing 6 M guanidine-HCl and a 100 mM CDAP solution in 100 mM citrate buffer (pH 3.0). A 10-fold molar excess of the 100 mM CDAP solution was added to the protein solution, and the resulting mixture was incubated for 10 to 15 min at room temperature. Excess reagent was then removed by preparative reverse-phase HPLC, and pure fractions containing cyanylated protein were collected, confirmed by mass spectroscopy, combined, and lyophilized. Lyophilized nSH3 was washed and concentrated with 4 × 15 mL of sodium phosphate buffer (5 mM, pH 7.3) using a 3,000 MWCO centrifugal filter (Millipore). The concentration of protein was determined by its absorption at 280 nm using an extinction coefficient of 15,220 M−1cm−1.30 Aliquots of 1.0 μmol cyanylated protein were then lyophilized and stored at −20 °C, and the quality of each sample was assessed before use by analytical HPLC using a C12 reversed-phase column with a solvent gradient of 20% B to 50% B over 40 min, 1.5 mL/min flow rate, and monitoring absorption at 220 nm, and by mass spectrometry (Supporting Information). FT-IR Spectroscopy. Each aliquot of cyanylated protein was dissolved in 10 μL of 100 mM sodium phosphate buffer containing 200 mM NaCl (pH 7.3). Model compounds were added to the organic solvent of interest or to an identical aqueous solution. Eight μL of the sample was then loaded into a temperature controllable demountable liquid cell with CaF2 windows and a 75 μm Teflon spacer (Harrick Scientific Products, Inc.). Spectra were acquired on a Bruker Equinox 55 FT-IR spectrometer equipped with an MCT detector with a resolution of 2 cm−1. Spectra were acquired between 24 and 93 °C, in 5 °C intervals. At each temperature, samples were

Figure 1. (A) Structure of nSH3 (PDB 1CKA) showing the side chains of Phe143, Phe153, Leu159, Met181, and Tyr186 and (B) structures of the (S)CN probes used in this study (shown as free amino acids).

each (S)CN probe is again linear with temperature, and the FTLSs suggest that the (CN)Phe probe at each of the three positions examined is engaged in H-bonds, but of varying strength, while the SCN probes at either of the two positions, including a surface exposed position, are not H-bonded. The data are consistent with structure and stability studies and thus suggest that the simple acquisition of IR spectra at multiple temperatures provides the means to evaluate the probe’s environment, including the participation of H-bonding, which at the positions examined in nSH3 appears to be more common with (CN)Phe than the SCN probes. 11562

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Analytical Chemistry equilibrated for 10 min, and a total of 8,000 scans (corresponding to ∼50 min of data acquisition) were averaged. We note that experiments with model systems were performed at 5 mM for p-tolunitrile and 10 mM for methyl thiocyanate. Spectra were collected at 5 °C intervals between 6 and 93 °C for p-tolunitrile and between 24 and 93 °C for methyl thiocyanate or over the accessible temperature range as dictated by the solvent (Supporting Information). Background spectra (wild type nSH3 or neat solvent) measured under identical conditions were autosubtracted from the sample spectra using Bruker’s OPUS software from 1,900 cm−1 to 2,400 cm−1. The resulting difference spectra were further background corrected with a polynomial function and fit with pseudo-Voigt functions using Matlab (Mathworks, Inc.) as described previously.27 In order to determine the melting temperature (Tm) of cyanylated proteins, all spectra at intermediate temperatures were fit as a linear combination of folded and unfolded state spectra. The resulting folded state contributions were fit with a sigmoidal function as described previously27 and in the Supporting Information. All experiments were performed in triplicate with three independently prepared samples, where shown error bars represent standard deviation.

Figure 2. Background-corrected FT-IR spectra of (A) p-tolunitrile and (B) CH3SCN at 24 °C. Spectra of p-tolunitrile in solvents with increasing order of peak frequencies are DMSO, toluene, formamide, isopropyl alcohol, cylcohexane, and aqueous buffer. Spectra of CH3SCN in solvents with increasing order of peak frequencies are DMSO, formamide, toluene, isopropyl alcohol, buffer, and cyclohexane.

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RESULTS To establish references for understanding how the environment of the (S)CN probes impacts the temperature dependence of their IR absorption, we first characterized the spectra of the model compounds p-tolunitrile and methyl thiocyanate (CH3SCN) in aqueous buffer, isopropyl alcohol, formamide, dimethyl sulfoxide, toluene, and cyclohexane, which provide environments with varying H-bonding potential and dielectric constant. As expected, 4,6,16 at room temperature, the absorptions of both model compounds generally shifted to higher frequency with increased H-bonding potential of the solvent (Figure 2). Spectra were then collected as a function of temperature. With the exception of isopropyl alcohol, the spectrum of p-tolunitrile at each temperature was well fit by a single pseudo-Voigt function (Supporting Information). In isopropyl alcohol, the p-tolunitrile spectra required two pseudoVoigt functions, with the high frequency component tentatively assigned to an H-bonded population, and the low frequency component assigned to a weaker or less H-bonded species. Consistent with reports from the Londergan16,31 and Boxer laboratories,5 the spectrum of CH3SCN at each temperature was well fit by a single pseudo-Voigt function (Supporting Information). Plotting the frequency of each (S)CN absorption as a function of temperature revealed a linear correlation (Figure 3A and B and Supporting Information), and importantly, the FTLS with each solvent was generally unique. With both probes, the FTLS values were by far the largest for the absorptions in aqueous solvent, followed by isopropyl alcohol (the strongly Hbonded isopropyl alcohol absorption with p-tolunitrile) and formamide, and then by cyclohexane and DMSO, and finally by toluene and for p-tolunitrile, the non-H-bonded absorption in isopropyl alcohol. In fact, for CH3SCN in toluene and for the non-H-bonded absorption of p-tolunitrile in isopropyl alcohol, the shifts were actually to the blue (e.g., positive FTLSs). To characterize the behavior of the (S)CN probes when incorporated within nSH3, we first analyzed the temperature dependence of the previously reported spectra of (CN)Phe143 and (CN)Phe153 at 5 °C intervals between 6 and 93 °C and (CN)Phe186 at 5 °C intervals between 24 and 93 °C.26 To

Figure 3. Plot of frequency shift of (A) p-tolunitrile and (B) CH3SCN with temperature in various solvents. Solvent systems in panel (A) are designated as isopropyl alcohol (a, non-H-bonded), toluene (b), DMSO (c), cyclohexane (d), formamide (e), isopropyl alcohol (f, Hbonded), and aqueous buffer (g). Solvent systems in panel (B) are designated as toluene (a), DMSO (b), cyclohexane (c), isopropyl alcohol (d), formamide (e), and aqueous buffer (f). The open circles are experimental data points, and solid lines are best fitting straight lines. Spectral shifts at the lowest temperature are normalized to zero.

increase the range over which the folded state could be monitored, we acquired additional spectra for (CN)Phe153 at 2 °C and for (CN)Phe186 at 6, 11, 16, and 21 °C. Briefly, CN stretching absorptions obtained at temperatures where the protein was more than 80% folded were each fit to a single pseudo-Voigt function, and the normalized frequencies were plotted as a function of temperature (Figure 4A). The plots of frequency versus temperature for each (CN)Phe probe again yielded linear correlations. Interestingly, the FTLSs of (CN)11563

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(10.5 cm−1). Similarly, the spectrum of (SCN)HCys159 showed a single band that was well fit by a single pseudoVoigt function, with a peak maximum at 2159.7 cm−1 and a narrow line width of 6.1 cm−1. Each of these absorptions is assigned to the folded state of the protein. Fitting the spectrum of (SCN)Cys159 required two pseudo-Voigt functions, with peak frequencies at 2160.5 and 2162.4 cm−1 and widths of 5.9 and 14.2 cm−1, respectively. Based on the similarity of the more red-shifted absorption with those of the other folded proteins, the similarity of the more blue-shifted absorption with the other unfolded proteins (see below), and the thermal denaturation studies (see below), the red-shifted absorption is assigned to the folded state, and the blue-shifted absorption is assigned to the unfolded state. Spectra were then acquired at 5 °C intervals between 24 and 93 °C. With increasing temperature, a gradual decrease of the FT-IR absorption peak intensity was observed. For (SCN)Cys159 and (SCN)Cys181, the amplitude was reduced ∼10fold at 65 °C compared to the corresponding folded states and became unobservable at higher temperatures. A decrease was also observed for (SCN)HCys159 and (SCN)HCys181, but in these cases the decrease was only ∼2-fold at 93 °C relative to the intensity at 24 °C. However, in all cases the spectra of the folded state could not be fully recovered after a single heating− cooling cycle, and thus quenching of the absorption bands was irreversible. These observations are consistent with several previously reported studies of small molecules, peptides, and proteins with similar SCN moieties, all of which revealed that at elevated temperature the probes undergo either isomerization to the isothiocyanate, cyclization and peptide bond cleavage, or β-elimination.35−38 The FT-IR spectra acquired as a function of temperature were deconvoluted into linear combinations of folded (low temperature) and unfolded (high temperature) spectra for each nSH3 variant. The fractional concentrations of the folded state as a function of temperature were well fit by a simple sigmoidal function (Figure 5 and Supporting Information), confirming that the high temperature spectra correspond to unfolded protein. The melting temperatures or Tm values, which refer to the temperature at which the protein is 50% unfolded, are 56.3 ± 1.3 °C and 52.4 ± 0.3 °C for (SCN)HCys181 and

Figure 4. Plot of frequency shift of (CN)Phe143 (red circles), (CN)Phe153 (green circles), and (CN)Phe186 (blue circles) with temperature along with p-tolunitrile in the various solvents examined (gray solid lines, with individual data omitted for clarity and labeled as defined in Figure 3A). Frequency shifts of the folded and unfolded proteins are displayed in panels (A) and (B), respectively. Shifts of the folded proteins were normalized to zero at 6 °C and those of the unfolded proteins were normalized to that of p-tolunitrile in aqueous buffer at 93 °C. The data points for the model systems are omitted for clarity.

Phe153 and (CN)Phe186 are virtually identical to those of the model system in aqueous buffer, while that of (CN)Phe143 is indistinguishable from that of the model system in isopropyl alcohol (for the H-bonded absorption) or formamide. As a control, we also performed a similar analysis of the absorption spectra of each probe acquired at temperatures where the protein was at least 80% unfolded. A plot of peak frequency versus temperature again yielded a linear correlation for each protein (Figure 4B). However, in this case, all of FTLSs were indistinguishable, with values intermediate between aqueous buffer and formamide. The data suggest that the (CN)Phe probes are solvent exposed in the unfolded state and that the FTLS for each probe within the folded protein is the result of its specific folded microenvironment. To extend these studies to SCN probes, nSH3 was prepared with (SCN)Cys or (SCN)HCys replacing either Met181 or Leu159 (resulting in four different nSH3 variants). These probes were selected because they have been used as mimics of different amino acids, including methionine5,7,9,32−34 and leucine,8 and because they make it possible to examine the robustness of the results to small changes in probe positioning. In addition, incorporation at positions 181 or 159 enables an examination of the probe’s behavior at a surface exposed or buried position, respectively. Thus, proteins were synthesized with either cysteine or homocysteine at position 181 or 159 using Boc solid phase peptide synthesis. The single sulfhydryl was then cyanylated using 1-cyano-4-dimethylaminopyridinium tetrafluoroborate (CDAP)28,29 to produce the corresponding SCN-labeled variants of nSH3 ((SCN)Cys159, (SCN)HCys159, (SCN)Cys181, and (SCN)HCys181). The IR spectrum of both (SCN)HCys181 and (SCN)Cys181 at 24 °C showed a single absorption band that was well fit by a single pseudo-Voigt function, each with a center frequency of ∼2158.3 cm−1 (Supporting Information). However, the line width of the (SCN)HCys181 absorption (13.7 cm−1) is 3.2 cm−1 greater than that of (SCN)Cys181

Figure 5. Temperature-dependent fractional concentration of the folded state for (SCN)Cys159 (black circles), (SCN)HCys159 (red circles), (SCN)Cys181 (blue circles), and (SCN)HCys181 (green circles). Data was measured in triplicate, and lines are sigmoidal fits. 11564

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(SCN)Cys181, respectively, and 42.4 ± 0.7 °C and 15.2 ± 0.4 °C for (SC)HCys159 and (SCN)Cys159, respectively. The uniquely low Tm of (SCN)Cys159 is consistent with the assignment of the unique blue-shifted band observed in the 24 °C spectrum to the unfolded state. Clearly, the incorporation of SCN probes can be destabilizing; however, in each case the folded state spectra may be unambiguously identified. To characterize the temperature-dependent frequency shifts of the SCN probes within the folded state of each protein, spectra were analyzed as described above for the (CN)Phe probes (Supporting Information). The folded state of (SCN)Cys159 was not included as it was not sufficiently populated for analysis. The plots of absorption frequency as a function of temperature for (SCN)HCys159, (SCN)Cys181, and (SCN)HCys181 again yielded linear correlations (Figure 6A and

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DISCUSSION

While providing a strong and thus easily observed absorption, the use of (S)CN probes requires caution, due to destabilization of the protein, as is the case with any extrinsic probe, and to H-bonding. As shown here and previously,26 incorporation of (S)CN probes can significantly destabilize the native fold of the protein. Because a protein’s behavior is dependent on its native structure, the use of (S)CN probes at positions that result in significant perturbation should be avoided. Once sites of minimal perturbation are identified, the major challenge with the interpretation of (S)CN absorptions is determining whether they participate in H-bonding. The first attempts to identify (S)CN H-bonding interactions were based on X-ray crystallography;6,7 however, the high resolution structural analysis of every (S)CN-modified protein of interest is prohibitively labor intensive. Boxer, Hershlag, and co-workers have also established a procedure to determine whether a specific (S)CN probe engages in an H-bonding interaction that is based on comparisons of IR and NMR spectra.4,6 While conceptually elegant, this approach requires additional experiments, and quantitative interpretation of the data is complicated by anisotropic contributions to the NMR frequencies. Moreover, in cases where there is fast exchange between different species, such as with p-tolunitrile in isopropyl alcohol or benzonitrile in protic solvents,39,40 NMR cannot resolve the individual species, and thus an assessment of their individual shifts is not straightforward. Our approach to determine whether H-bonding contributes to the IR absorption frequency of a given (S)CN probe within a protein is more straightforward and is based on the larger blueshift induced by H-bonding, which are weakened or ablated at elevated temperatures. Importantly, the FTLS is dependent on the strength of the H-bond, as would be expected if weaker Hbonds induce smaller blue shifts in the first place. In addition, although the absorption frequency of each probe in DMSO is significantly lower than in cyclohexane (6 to 10 cm−1), it is interesting that both probes show similar FTLS values in the two solvents. This is likely a reflection of the disruption of only nonspecific interactions with increasing temperature in both solvents. Finally, the positive FTLS observed with CH3SCN in toluene (i.e., increasing temperature induces a blue-shift) likely reflects the disruption of η-type interactions with the (S)CN moiety, which Cho and co-workers have suggested induce a red-shift.21 Regardless of the detailed origins of the shifts, plotting the observed absorption frequency as a function of temperature revealed linear dependencies in all cases, with FTLS values that clearly reflect the probe environment, including the presence and strength of H-bonds. Interestingly, very different behaviors are observed with (CN)Phe and SCN probes incorporated at different positions within nSH3. When the (CN)Phe probe is incorporated at position 153 or 186 of nSH3, the FTLS is virtually identical to that observed for p-tolunitrile in aqueous buffer. This suggests that at these two positions, the CN moieties are engaged in strong H-bonds. Based on the shifts observed in formamide, the CN moieties are unlikely to involve backbone N−H donors, which result in smaller FTLSs. The strong H-bonding at (CN)Phe186 is consistent with its surface exposure. While the side chain at position 153 in the wild type protein is buried, the incorporation of the CN moiety at this position is by far the most destabilizing (Tm = 29.2 ± 0.3 °C),26 which is consistent

Figure 6. Plot of frequency shift of (SCN)Cys181 (green circles), (SCN)HCys181 (red circles), (SCN)Cys159 (blue circles), and (SCN)HCys159 (violet circles) with temperature along with CH3SCN in the various solvents examined (gray solid lines, with individual data omitted for clarity and labeled as defined in Figure 3B). Frequency shifts of folded and unfolded nSH3 are displayed in panels (A) and (B), respectively, except for the unfolded state of (SCN)Cys159 (blue circles), which is included in panel (A) because it could only be characterized over the corresponding temperature range (see text). Spectral shifts of folded proteins (and unfolded (SCN)Cys159) were normalized to zero at 24 °C, and the shifts of unfolded (SCN)HCys159 and (SCN)HCys181 were normalized to that of CH3SCN at 93 °C in formamide.

Supporting Information). However, in these cases, all of the FTLS values were similar and close to that observed with CH3SCN in cyclohexane. As a control, we again performed an analysis of the absorption spectra of the unfolded proteins. The unfolded state of (SCN)Cys181 could not be analyzed due to signal loss (see above), but it was possible to analyze the unfolded state of (SCN)Cys159 because it is populated at lower temperatures. The frequency shift of (SCN)Cys159 and of both (SCN)HCys probes in the unfolded state were again linear with temperature with FTLS values similar to that of CH3SCN in isopropyl alcohol or formamide for (SCN)HCys181 and slightly larger for (SCN)HCys159 and (SCN)Cys159 (but still significantly less than for CH3SCN in aqueous buffer) (Figure 6B). Clearly the cyclohexane-like FTLS values observed in the folded state result from the probes specific folded microenvironments. 11565

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We note that the data should not be interpreted as suggesting that all aliphatic SCN probes will generally be free from H-bonding when incorporated within a protein. Indeed, Boxer has reported clear structural evidence that several SCN probes incorporated within ketosteroid isomerase are involved in H-bonding interactions.6,7 Moreover, the significant temperature dependence of the absorptions of (SCN)Cys13 of RNase S5 suggests H-bonding by the standards of the current work. Clearly, (S)CN moieties are capable of experiencing different environments, including those involving H-bonding, and in fact it is this diversity of environments that should make the proposed method useful.

with structural distortions that allow solvent access to the side chain. The absorption of (CN)Phe at position 143 again shifted linearly with temperature, but with a FTLS that was more similar to the H-bonded absorption of the model compound in isopropyl alcohol or to the absorption in formamide. This suggests that its environment involves a weaker H-bond. This is consistent with the predicted position of the side chain being at the interface between the protein surface and solvent, where it is at least partially shielded from solvation by the side chains of Asp150 and Leu151. The behavior with the SCN probes was distinctly different. In all three cases where analysis was possible in the folded state ((SCN)HCys159, (SCN)HCys181, and (SCN)Cys181), and regardless of whether the probe was incorporated at a surface exposed or buried position, the probes exhibited similar FTLSs that are decidedly smaller than that observed with CH3SCN in H-bonding solvents (aqueous buffer or even isopropyl alcohol or formamide) and very similar to that observed with cyclohexane. This suggests that the SCN probes at these positions do not engage in H-bonds. While the H-bond basicity of the (CN)Phe probes is slightly greater than that of the SCN probes,41 it seems insufficient to explain the decidedly increased solvation of the former. Instead, we tentatively attribute the difference to increased flexibility afforded the SCN probes by their aliphatic side chains, which likely allows them to access more stabilizing environments that also shield them from water. Indeed, position 159 is buried with many predominantly hydrophobic environments within reach of the (SCN)HCys side chain and position 181 is adjacent to a hydrophobic pocket formed by the side chain of Phe143 and Trp169. That the FTLS values for the SCN probes in the unfolded state still suggest nonaqueous environments, unlike for the (CN)Phe probes, is further evidence that the flexibility of the alkyl side chains allows these probes to access environments that shield them from water. Regardless of the physical underpinnings, the H-bonding-free environment of the SCN probes suggests that their absorption frequencies may be interpreted in terms of local electrostatics. While a quantitative analysis requires knowledge of the projection of the electric field onto the transition dipole, the availability of interpretable SCN absorption frequencies at multiple positions within the protein affords several interesting observations. The absorption frequencies for (SCN)HCys159, (SCN)HCys181, and (SCN)Cys181 were similar (2158.2 to 2159.7 cm−1). This implies that the electrostatic environment experienced by the SCN group is not altered by the additional methylene that differentiates the probes, at least when incorporated at position 181. Perhaps more surprisingly, the data also suggest that the (SCN)HCys probe experiences a similar environment when positioned within the core of the protein or on its surface, which likely again results from the flexibility of the aliphatic side chain allowing the SCN moiety to access any available hydrophobic microenvironments, which exist even on the surface of the protein. The folded absorption of (SCN)Cys159 is somewhat more blue-shifted than the other proteins (2160.5 cm−1), suggesting that it experiences a slightly less polar environment.6,16 This may result from a combination of the shorter side chain (relative to (SCN)HCys159) and the more buried positions (relative to (SCN)Cys181 and (SCN)HCys181) leading to a more sterically restricted environment, which is also consistent with the probe’s uniquely narrow line width (5.9 cm−1) and destabilizing effect on the protein.

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CONCLUSIONS We have developed a simple approach to fuller interpretation of the absorption spectrum of (S)CN probes incorporated at different positions within a protein that only requires the characterization of the absorption spectrum at several temperatures. Importantly, the approach appears capable of identifying environments in which the probes are free of H-bonding interactions, and thus where their shifts in absorption frequency are likely linearly dependent on local electric field. The collection of H-bond-free shifts at different positions of a protein should make it possible to determine not only the range of electrostatic environments available within a protein but also, when supplemented with structural and computational data, the absolute value of the electric field. Conversely, the ability to straightforwardly demonstrate that (S)CN probes are Hbonded should facilitate the interpretation of their spectra in terms of solvation. More generally, a method to characterize the ability of different sites within a protein to accommodate water or engage the extrinsic probe with complementary H-bond donors has important implications for our understanding of protein dynamics, engineering, and evolution.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.5b03437. Characterization of nSH3 variants, details of FT-IR data acquisition and analysis, additional figures and tables (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Phone: 858-784-7290. Fax: 858-784-7472. E-mail: floyd@ scripps.edu. Notes

The authors declare no competing financial interest.

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REFERENCES

(1) Chin, J. K.; Jimenez, R.; Romesberg, F. E. J. Am. Chem. Soc. 2001, 123, 2426−2427. (2) Waegele, M. M.; Culik, R. M.; Gai, F. J. Phys. Chem. Lett. 2011, 2, 2598−2609. (3) Ma, J.; Pazos, I. M.; Zhang, W.; Culik, R. M.; Gai, F. Annu. Rev. Phys. Chem. 2015, 66, 357−377. (4) Bagchi, S.; Fried, S. D.; Boxer, S. G. J. Am. Chem. Soc. 2012, 134, 10373−10376. (5) Fafarman, A. T.; Boxer, S. G. J. Phys. Chem. B 2010, 114, 13536− 13544. 11566

DOI: 10.1021/acs.analchem.5b03437 Anal. Chem. 2015, 87, 11561−11567

Article

Analytical Chemistry (6) Fafarman, A. T.; Sigala, P. A.; Herschlag, D.; Boxer, S. G. J. Am. Chem. Soc. 2010, 132, 12811−12813. (7) Fafarman, A. T.; Sigala, P. A.; Schwans, J. P.; Fenn, T. D.; Herschlag, D.; Boxer, S. G. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, E299−308. (8) Liu, C. T.; Layfield, J. P.; Stewart, R. J., 3rd; French, J. B.; Hanoian, P.; Asbury, J. B.; Hammes-Schiffer, S.; Benkovic, S. J. J. Am. Chem. Soc. 2014, 136, 10349−10360. (9) Stafford, A. J.; Ensign, D. L.; Webb, L. J. J. Phys. Chem. B 2010, 114, 15331−15344. (10) Hu, W.; Webb, L. J. J. Phys. Chem. Lett. 2011, 2, 1925−1930. (11) Stafford, A. J.; Walker, D. M.; Webb, L. J. Biochemistry 2012, 51, 2757−2767. (12) Walker, D. M.; Wang, R.; Webb, L. J. Phys. Chem. Chem. Phys. 2014, 16, 20047−20060. (13) Johnson, M. N.; Londergan, C. H.; Charkoudian, L. K. J. Am. Chem. Soc. 2014, 136, 11240−11243. (14) Getahun, Z.; Huang, C. Y.; Wang, T.; De Leon, B.; DeGrado, W. F.; Gai, F. J. Am. Chem. Soc. 2003, 125, 405−411. (15) Ma, J.; Pazos, I. M.; Gai, F. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 8476−8481. (16) Maienschein-Cline, M. G.; Londergan, C. H. J. Phys. Chem. A 2007, 111, 10020−10025. (17) Mukherjee, S.; Chowdhury, P.; DeGrado, W. F.; Gai, F. Langmuir 2007, 23, 11174−11179. (18) Tucker, M. J.; Getahun, Z.; Nanda, V.; DeGrado, W. F.; Gai, F. J. Am. Chem. Soc. 2004, 126, 5078−5079. (19) Waegele, M. M.; Tucker, M. J.; Gai, F. Chem. Phys. Lett. 2009, 478, 249−253. (20) Levinson, N. M.; Fried, S. D.; Boxer, S. G. J. Phys. Chem. B 2012, 116, 10470−10476. (21) Kim, H.; Cho, M. Chem. Rev. 2013, 113, 5817−5847. (22) Cho, M. J. Chem. Phys. 2009, 130, 094505. (23) Lee, H.; Choi, J. H.; Cho, M. Phys. Chem. Chem. Phys. 2010, 12, 12658−12669. (24) Lee, H.; Choi, J. H.; Cho, M. J. Chem. Phys. 2012, 137, 114307. (25) Huang, C.-Y.; Wang, T.; Gai, F. Chem. Phys. Lett. 2003, 371, 731−738. (26) Adhikary, R.; Zimmermann, J.; Dawson, P. E.; Romesberg, F. E. ChemPhysChem 2014, 15, 849−853. (27) Adhikary, R.; Zimmermann, J.; Liu, J.; Dawson, P. E.; Romesberg, F. E. J. Phys. Chem. B 2013, 117, 13082−13089. (28) Wakselman, M.; Guibe-Jampel, E.; Raoult, A.; Busse, W. D. J. Chem. Soc., Chem. Commun. 1976, 21−22. (29) Wu, J.; Watson, J. T. In Posttranslational Modifications of Proteins; Kannicht, C., Ed.; Humana Press: Totowa, NJ, Vol. 194, pp 1−22. (30) Cremeens, M. E.; Zimmermann, J.; Yu, W.; Dawson, P. E.; Romesberg, F. E. J. Am. Chem. Soc. 2009, 131, 5726−5727. (31) McMahon, H. A.; Alfieri, K. N.; Clark, K. A. A.; Londergan, C. H. J. Phys. Chem. Lett. 2010, 1, 850−855. (32) Fafarman, A. T.; Webb, L. J.; Chuang, J. I.; Boxer, S. G. J. Am. Chem. Soc. 2006, 128, 13356−13357. (33) Jha, S. K.; Ji, M.; Gaffney, K. J.; Boxer, S. G. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 16612−16617. (34) Sigala, P. A.; Fafarman, A. T.; Bogard, P. E.; Boxer, S. G.; Herschlag, D. J. Am. Chem. Soc. 2007, 129, 12104−12105. (35) Catsimpoolas, N.; Wood, J. L. J. Biol. Chem. 1966, 241, 1790− 1796. (36) Degani, Y.; Patchornik, A. Biochemistry 1974, 13, 1−11. (37) Doherty, G. M.; Motherway, R.; Mayhew, S. G.; Malthouse, J. P. Biochemistry 1992, 31, 7922−7930. (38) Smith, P. A. S.; Emerson, D. W. J. Am. Chem. Soc. 1960, 82, 3076−3082. (39) Aschaffenburg, D. J.; Moog, R. S. J. Phys. Chem. B 2009, 113, 12736−12743. (40) Ghosh, A.; Remorino, A.; Tucker, M. J.; Hochstrasser, R. M. Chem. Phys. Lett. 2009, 469, 325−330.

(41) Abraham, M. H.; Grellier, P. L.; Prior, D. V.; Morris, J. J.; Taylor, P. J. J. Chem. Soc., Perkin Trans. 2 1990, 521−529.

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DOI: 10.1021/acs.analchem.5b03437 Anal. Chem. 2015, 87, 11561−11567