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Probing the Conformation-Dependent Preferential Binding of Ethanol to Cationic Glycylalanylglycine in Water/Ethanol by Vibrational and NMR Spectroscopy David DiGuiseppi,† Bridget Milorey,† Gabrielle Lewis,‡ Nina Kubatova,§ Stefanie Farrell,† Harald Schwalbe,§ and Reinhard Schweitzer-Stenner*,† †

Department of Chemistry and ‡Department of Biology, Drexel University, Philadelphia, Pennsylvania 19104, United States § Institut für Organische Chemie und Chemische Biologie, Johann Wolfgang Goethe-Universität, 60438 Frankfurt am Main, Germany S Supporting Information *

ABSTRACT: The conformational propensity of amino acid residues is determined by an intricate balance of peptide−solvent and solvent−solvent interactions. To explore how the systematic replacement of water by a cosolvent affects the solvation of both the amino acid backbone and side chains, we performed a combined vibrational spectroscopy and NMR study of cationic glycylalanylglycine (GAG) in different ethanol/water mixtures of between 0 and 42 mol percent ethanol. Classical model peptide N′-methylacetamide was used as a reference system to probe solvent-induced spectroscopic changes. The alanine residue of GAG in water is known to exhibit a very high propensity for polyproline II (pPII). Adding up to 30 mol % ethanol at room temperature leads only to minor changes in the Ramachandran distribution of alanine, which mostly changes within the individual conformational subspaces. A further increase in the ethanol fractions leads to a destabilization of pPII and a stabilization of β-strand conformations. At higher temperatures, different degrees of enthalpy−entropy compensations lead to a much stronger influence of ethanol on the peptide’s conformational distribution. Ethanol-induced changes in chemical shifts and amide I wavenumbers strongly suggest that ethanol replaces water preferentially in the solvation shell of the polar C-terminal peptide group and of the alanine side chain, whereas the N-terminal group remains mostly hydrated. Furthermore, we found that ethanol interacts more strongly with the peptide if the latter adopts β-strand conformations. This leads to an unusual positive temperature coefficient for the chemical shift of the C-terminal amide proton. Our data suggests a picture in which GAG eventually accumulates at water−ethanol interfaces if the ethanol fractions exceed 0.3, which explains why the further addition of ethanol eventually causes self-aggregation and the subsequent formation of a hydrogel.

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INTRODUCTION Over the last 60 years, short peptides, particularly blocked dipeptides, have been used as model systems to explore the conformational sampling of individual amino acid residues in unfolded peptides and proteins.1−13 Early (mostly computational and theoretical) results of Ramachandran,1 Flory,2 and Scheraga14,15 suggested that all amino acids sample the entire accessible region of the Ramachandran space. With the exception of proline and glycine, the conformational distributions of amino acids were reported to be comparable. These results constitute the basis of the so-called random-coil model, which assumes that unfolded polypeptides behave like a polymer in a good solvent where the conformational sampling is solely restricted by the excluded volume effect.16 However, over the last 15 years, multiple experimental10,17−25 and computational studies6,11,26−36 as well as analyses of coil libraries37−41 have provided compelling evidence for the notion that amino acids sample a much more restricted Ramachandran space than predicted by the above studies. The conformational manifolds of amino acids depend on the steric and physical properties of their side chains as well as on the properties of their neighbors, in particular, i-1 residues.37,42,43 Alanine, which © 2017 American Chemical Society

has been generally used as the canonical representative for presenting the conformational heterogeneity of peptide backbone structures,1,2 was found to actually depart the most from the classical local random-coil behavior in that it samples a very high fraction of a polyproline II (pPII)-like conformation in unblocked tripeptides (GAG, AAA), 11,23,44 the alanine dipeptide,10,11 tetrapeptides,22,45,46 and longer polyalanines in aqueous solutions18,47,48 and even in nonregularly structured regions of proteins.37,49 The reason for this significant restriction of the conformational space of alanine has been a matter of debate. A vast majority of studies have identified water as the main stabilizer of pPII at room temperature, but the underlying mechanisms remain controversial. The preferred packing of water molecules,26 electrostatic screening,26,31 the interplay between peptide−water and water−water interactions in the hydration shell,50 solvent accessibility,51 and the formation of clathrate water structures32 around alanine side chains have been Received: March 27, 2017 Revised: May 17, 2017 Published: May 22, 2017 5744

DOI: 10.1021/acs.jpcb.7b02899 J. Phys. Chem. B 2017, 121, 5744−5758

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The Journal of Physical Chemistry B introduced as pPII stabilizers. Raines and co-workers proposed an nπ interpeptide interaction between the peptide oxygen and the subsequent carbonyl carbon as a pPII stabilizing force.52 However, the dominant role of the solvent has been underscored by thermodynamic comparisons of conformational distributions of a large series of GxG (x, guest amino acid) model peptides.53 If solvation water is indeed stabilizing the pPII conformation of alanine and other amino acid residues, then its replacement by other solvents should lead to the destabilization of this conformation. This has indeed been observed. Eker et al., for instance, showed that if the solvent was changed from water to dimethylsulfoxide (DMSO) then the preference for sampling the pPII conformation of AcAA−OH was practically eliminated.54 Liu et al. probed the pPII fraction of host−guest peptide AcGGAGGNH2 and an alanine based 11-mer in different alcohols and found that the pPII content of their peptides decreased with the increasing (aliphatic) chain length of the alcohol.55 To understand the effect of such solvent replacement on the structure of peptides, the current study is aimed at a more systematic investigation of how the replacement of water by a cosolvent alters the hydration shell of a peptide and thus affects its backbone. Alcohols are considered to be excellent tools for modifying the solvation of peptides and proteins in different ways. Glycerol, for instance, is known to cause preferred peptide/ protein hydration and thus a stabilization of folded states.56−60 The picture is more complex for primary alcohols such as methanol and ethanol. In principle, their interaction with peptide and water should be easier to understand than that of chemically more complicated glycerol. However, this is not the case. Ethanol as a cosolvent, for instance, can stabilize and destabilize proteins61−63 and produce a variety of effects in living organisms.64,65 Generally, it stabilizes unfolded over folded protein states, which indicates preferential binding to unfolded peptides/proteins.66 Several lines of evidence indeed suggest that it interacts with peptides via hydrophobic and hydrogen bonding.67−69 It has also been shown to reduce amyloid aggregation.62 These observations and the fact that it has a single site for hydrogen bonding and for hydrophobic interactions, respectively, make it an ideal candidate for systematically perturbing the hydration shell of peptides such as GAG. Some investigations of peptide−alcohol interactions have been performed in our laboratory. Toal et al. used UV-CD and proton NMR to probe conformational changes of cationic AAA in various ethanol/water mixtures.69 In this study, a highly nonlinear and nonmonotonous structural response to ethanol addition led to a slight stabilization of pPII at very low ethanol mole fractions (0.03) and a subsequent destabilization upon further increasing the ethanol fraction to 0.12. Spectroscopic evidence suggested the side chain−ethanol interaction to occur even at very low ethanol concentrations. More recently, Milorey et al. performed a more systematic study on cationic GAG by varying the ethanol mole fraction range from 0 to 0.5.70 To explore structural changes as a function of the ethanol fraction, the authors utilized the 3J(HNHα) coupling constant of the N-terminal amide group as a structural indicator. The results shown in Figure 1 reveal a highly nonmonotonous and nonlinear response. Systematic changes in 3J(HNHα) were observed in three regions indicated in the figure. On average, they observed an increase in the 3J(HNHα) value with increasing ethanol fraction. If this were due to only the

Figure 1. 3J(HNHα) of the N-terminal amide proton of cationic GAG in different water/ethanol mixtures as determined from 1H NMR spectra taken at the indicated temperatures. Three notable regions are highlighted and labeled accordingly. Taken from ref 70.

redistribution between different conformations (i.e., pPII and the β-strand), then the data would be indicative of a decreasing pPII population (cf. right axis label). However, it is equally possible that the data reflect intrinsic structural changes in pPII or β-strand structures. In the current article, this issue is addressed more thoroughly by a comprehensive conformational analysis of GAG in three water−ethanol mixtures in the regions indicated in Figure 1. The nonlinear ethanol dependence of 3 J(H N H α ) is reminiscent of the fact that water−ethanol is a highly nontrivial, nonideal mixture that is not yet fully understood.71 The partial molar volume of ethanol exhibits a minimum at a mole fraction of 0.04, which lies in region 1.72 Ghosh et al. have performed simulations of ethanol/water mixtures in this region.73 They found that even at this very low concentration ethanol forms microaggregated clusters that are considerably stabilized with the lowering of temperature. The maximum in the negative excess enthalpy is close to region II, whereas excess of negative entropy occurs at the onset of region III.71 Mijaković et al. used Brillouin scattering to obtain the maximum in the sound velocity around a mole fraction of 0.1 at room temperature, close to the onset of region 2.74 Results of MD (molecular dynamics) simulations suggest the formation of organized clusters of ethanol at various levels of the ethanol mole fraction and a local maximum in the amount of water/ethanol microsegregation at a mole fraction of approximately 0.15, which lies right in region 2 in Milorey et al.70 This is accompanied by the formation of more rigid water clusters, which are suspected to cause the observed maximum in the sound velocity. The present study has multiple goals. First, we use canonical model peptide N′-methylacetamide (NMA) as a reference system to study the (IR) spectroscopic response of a blocked peptide group to the replacement of water by ethanol in the solvation shell. Second, we intend to determine the conformational distribution of GAG in the three regions indicated in Figure 1. Third, we use vibrational spectroscopy markers (amide I′) and changes in the chemical shifts of NH and CH3 protons to site specifically probe the replacement of water by ethanol. The obtained data shed some light on how the addition of ethanol prepares the system for the onset of largescale aggregation and gelation with a 0.55 fraction of ethanol, which we observed and reported recently.70,75 5745

DOI: 10.1021/acs.jpcb.7b02899 J. Phys. Chem. B 2017, 121, 5744−5758

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The Journal of Physical Chemistry B

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using 1H NMR. The spectra were recorded on a Varian 500 MHz FT-NMR with a 5 mm HCN triple-resonance probe, and the temperature was controlled using a Varian VT controller. Each sample was allowed to equilibrate at every temperature for 2 min before an experiment was started. The spectra were collected with 64 scans, and a PRESAT setting was used to suppress the water peak. The FIDs were analyzed using MestReC software, which was used to Fourier transform and phase the data. The raw data were then analyzed in Multifit. The signals were decomposed into Voigtian profiles with flexible half-widths on a frequency scale to obtain the positions. These frequency positions were then plotted in SigmaPlot as a function of temperature, and a linear regression was used to fit the data, as described by Toal et al.11 For heteronuclear NMR experiments, the GAG tripeptide was labeled at residue 1 with 13C in the carbonyl position, uniformly 13C- and 15N-labeled at residue 2 and 15N-labeled at residue 3. The C-terminal residue was manually attached to a chlorotrityl resin. The synthesis was manually carried out by standard Fmoc chemistry. Peptides were purified by reversedphase HPLC. Products were characterized using electrospray ionization mass spectrometry and analytical HPLC. 15N-FmocGlycine was purchased from Cambridge Isotope Laboratories, uniformly 13C- and 15N-labeled and 13C-carbonyl-labeled Fmoc-protected amino acids were purchased from SigmaAldrich, and 2-chlorotrityl chloride resin was purchased from MERK. All NMR samples were prepared by dissolving the peptide in water and deuterated ethanol containing 4,4dimethyl-4-silapentane-sulfonic acid (DSS) as an internal standard. The temperature was varied between 298 and 318 K. The spectra were recorded on a Bruker 600 MHz spectrometer equipped with a 5 mm HCN triple-resonance cryogenic probe with z gradients. Spectra were acquired and analyzed using the program TopSpin V3.5. 3 J(HN,Hα) coupling constants were obtained from a 13 C-,15N-decoupled 1H NMR spectrum using excitation sculpting to suppress the water solvent. The exact coupling constants were determined by fitting the proton signals with Voigtian profiles with flexible half-widths. The 3J(HN,C′) coupling constant was determined with E.COSY-type experiment soft HNCa-COSY, and the 1J(N,C⟨) coupling constant was determined through the J-modulated 1H,15N HSQC experiment.44,78 Conformational Analysis of GAG. Details of our spectroscopy-based conformational analysis of short peptides are described in detail in earlier publications.79 Briefly, we describe the conformational distribution of A as a superposition of twodimensional Gaussian functions centered at positions associated with different types of secondary structures

MATERIALS AND METHODS Materials. Unblocked glycyl-alanyl-glycine (H-Gly-Ala-GlyOH) was purchased from Bachem, and N-methylacetamide was purchased from Sigma-Aldrich, both with >98% purity. No further purification was carried out on either. Deuterated solvents, D2O and ethan(ol)-d (EtOD), were used for vibrational spectroscopy studies to avoid the interference of the water bending mode with the amide I region. EtOD is the deuterated ethyl alcohol with the alcoholic hydrogen replaced by a deuterium. D2O and EtOD were purchased from SigmaAldrich in 99.9% purity. Ethanol (200 proof) was purchased from Pharmco-Aaper and utilized for nonvibrational spectroscopy studies. Methods. Fourier Transform Infrared Spectroscopy (FTIR). As described above, vibrational spectroscopy studies required the use of deuterated solvents. FTIR spectra of NMA were recorded on a PerkinElmer, Inc. (Shelton, CT) Spectrum One Fourier transform infrared absorption spectrometer, and samples were loaded onto a 59.5 μm CaF2 Biocell from BioTools (Jupiter, FL). The spectral range used was 450−4000 cm−1, the spectral resolution was 4 cm−1, and 25 scans were taken for each spectrum. Ultraviolet Circular Dichroism Spectroscopy. The UV-CD spectra of 200 mM GAG were recorded on a Jasco J-810 spectropolarimeter (model J-810-150S) purged with nitrogen. The temperature was controlled using a Peltier controller (model PTC-423S). Samples were loaded onto a 100 μm cell from International Crystal Laboratories. UVCD spectra were recorded between 190 and 250 nm with a 500 nm/min scan speed, 1 s response time, 0.05 data pitch, and 5 nm bandwidth. Five spectra were obtained and averaged for each temperature interval, and all spectra were corrected using appropriate background subtraction. Raman Spectroscopy. Polarized Raman spectra were obtained with 442 nm excitation with a He−Cd laser (model IK 4601R-E, Kimmon Electric, USA). The laser beam was directed onto an RM 100 Renishaw confocal Raman microscope focused onto a Fisherbrand microscope slide with a single concavity and a glass coverslip placed on top of the slide. The scattered light was filtered with a 442 nm notch filter, dispersed by a single-grating 2400 1 mm-1 grating, and imaged onto a back-thinned Wright Instrument CCD. The spectra of polarized Raman scattering were obtained using a linear polarizer for the parallel component (x polarization) and a combination of the linter polarizer and a λ/2 plate for the perpendicular component (y polarization). The x and y spectra were measured three and six times, respectively, and averaged to optimize their signal-to-noise ratios. The isotropic and anisotropic Raman intensities were calculated as described elsewhere.76 VCD/FTIR Spectroscopy. Infrared absorption and VCD spectra of GAG samples were recorded on a BioTools ChiralIR. Samples of 0.2 M were loaded onto a 59.5 μm CaF2 biocell (BioTools). A BioTools water-cooled temperature controller was used to maintain the temperature. Spectra were taken with 8 cm−1 resolution and a total integration time of 600 min (540 min for VCD and 60 min for IR) and collected with Grams/IR 7.00 (Thermo Galactic). The absorbance spectra were collected with the subtraction of the appropriate background using Multifit.77 NMR Spectroscopy. Amide proton 3J(HNHα) coupling constants and various chemical shift positions were determined

N

pA (ϕ , ψ ) =

∑ j=1

χA, j 2π |V̂ |

0 T

̂

0

e−0.5(ρA⃗ − ρA,⃗ j ) VA,j(ρA⃗ − ρA,⃗ j ) (1)

where ⎛ϕ ⎞ A ρA⃗ = ⎜⎜ ⎟⎟ ⎝ ψA ⎠

(2)

is the position vector of the jth Gaussian subdistribution in the Ramachandran space. The matrix 5746

DOI: 10.1021/acs.jpcb.7b02899 J. Phys. Chem. B 2017, 121, 5744−5758

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The Journal of Physical Chemistry B ⎛ σ 2 σ2 ⎞ ϕψ ,A ⎜ ϕA ⎟ VA,̂ j = ⎜ 2 ⎟ ⎜σ 2 ⎟ ⎝ ϕψ ,A σϕA ⎠

UV circular dichroism spectrum of the peptide to determine the enthalpic and entropic difference between peptide conformers for the investigated ethanol/water mixtures.11 Finally, we explore the temperature and solvent dependence of the chemical shifts of amide protons. IR Spectroscopy of N-Methylacetamide in Water/ Ethanol Mixtures. We measured the FTIR spectrum of NMAD in various water/ethanol mixtures. To this end, we started the series with pure D2O and subsequently increased the volume fraction of ethanol in increments of 0.1. The corresponding mole fractions are listed in Table 1. Figure 2

(3)

is the covariance matrix where the diagonal elements denote the half-widths of the distribution along the Ramachandran coordinates and the off-diagonal elements reflect correlations between the coordinates. χA,j denotes the statistical weight of the jth distribution, which together with the parameters of the Gaussian distribution were used as free parameters in our fits of conformational ensembles to amide I′ profiles and the 3 J(HNHα), 3J(HNCO), and 1J(NCα) coupling constants obtained from NMR experiments (vide supra). Amide I′ profiles and the coupling constants were calculated for 32 400 2° × 2° bins in Ramachandran space by using the excitonic coupling model by Schweitzer-Stenner80 and the most recent versions of the empirical Karplus equations.23 For the calculation of amide I′ profiles, we allowed for minor changes in earlier-determined transition dipole moments (for IR and VCD) and Raman tensor elements. The used values are listed in Table S1. Thermodynamic Analysis. The temperature dependence of 3 J(HNHα) was analyzed by utilizing a pseudo-two-state model23,53 which leads to the following equation

Table 1. List of Mole Fractions of Ethanol Corresponding to Volume Fractions of Ethanol in Ethanol/H2O (or D2O) Mixtures

⎡ ΔHβ − ΔGβ (TR ) ⎤ J pPII +3 Jβ exp⎢− ΔHβ /RT − RTR ⎣ ⎦⎥ J (T ) = (χpPII + χβ ) ⎡ ΔHβ − ΔGβ (TR ) ⎤ 1 + exp⎢− ΔHβ /RT − ⎥ RTR ⎣ ⎦

(

3

3

(

+

∑ JT ,i χT ,i i

)

)

volume fraction (%)

mole fraction

10 20 30 40 50 60 70 80 90 100

0.03 0.07 0.12 0.17 0.24 0.32 0.42 0.55 0.74 1.000

(4)

where 3Jj and χj are the J coupling constants and normalized mole fractions of the considered alanine conformations: j = pPII, β, and turn. ΔG(TR) is the corresponding Gibbs free energy difference between pPII and β at room temperature, TR. The values for ΔGR were calculated from earlier-reported pPII/ β mole fraction ratios.11 Reference 3Jj values were calculated as averages over the unique ϕ, ψ subdistributions of the jth conformation using the Karplus equation. Thus, only ΔHβ was used as a free parameter in the fit of eq 4 to experimental 3 J(HN,Hα)(T). Notably, we obtained very good fits to the experimental data using this approach, confirming the relative temperature-independence of turn-like conformations in the unfolded state.

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RESULTS This section is organized as follows. First, we present the results of our IR spectroscopy on NMA in different D2O−ethanol mixtures. Because both cosolvents are deuterated, the peptide maintains a state in which NH is replaced by ND (NMAD). This work is aimed at exploring spectral changes that reflect the replacement of D2O by ethanol in the peptide’s hydration shell. To this end, we focus on the amide I′ band. The eigenvector of the respective normal mode is dominated by the CO stretching mode.81 The corresponding wavenumber therefore depends on the strength of hydrogen bonding between the peptide and solvent and also on the polarity of the latter.82−85 In the second section, we first present a conformational analysis of GAG in three different water/ethanol mixtures that correspond to regions 1−3 of GAG/ethanol/water mixtures identified by Milorey et al.70 This is followed by a thermodynamic analysis that utilizes the temperature dependence of the 3J(HNHα) coupling constants of the N-terminal amide proton and of the

Figure 2. Amide II and I′/I regions of the IR spectra of 0.2 M NMA in the indicated binary mixtures of D2O and EtOH (in volume %).

shows the amide I′ region of these spectra. As observed earlier for NMAD in D2O−methanol mixtures,86 the addition of ethanol causes first a blue shift of amide I′, which is followed by a substantial broadening of the band profile at very high mole fractions of the cosolvent. Generally, this is indicative of the decreasing strength and increasing heterogeneity of hydrogen bonding between the solute and solvent.82,87,88 To gain more quantitative insights into solvent-induced changes in spectroscopic properties, we subjected all band profiles in Figure 2 to a spectral decomposition into Voigtian bands. An example is shown in Figure 3. This analysis revealed the following picture. Below a mole fraction of 0.2, the amide I′ profile is composed of a single band with its wavenumber shifting continuously to the blue; we term this band AI1′. The corresponding integrated intensity first increases, reaches a maximum at ca. 0.1, and then 5747

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al.70 It is in this region where these authors observed the onset of GAG aggregation and gelation (e.g., at χET = 0.55 for a peptide concentration of 200 mM). Moreover, a second amide I′ band appears at 1648 cm−1 (AI2′) (Figures 3,4), which exhibits a blue shift to 1652 cm−1 at very large ethanol fractions (>0.7). One is tempted to assign the occurrence of two amide I′ bands to the coexistence of NMAD monomers and oligomers, as observed for comparable concentrations of NMA in acetonitrile.90 Recent investigations of NMAD in methanol/ D2O mixtures, for which a similar coexistence of bands was observed (vide infra), have ruled out this option. To check whether aggregation affects NMA in D2O−ethanol, we measured the amide I′ band profile of samples with three different NMAD concentrations (50, 100, and 200 mM) in a mixture with an ethanol mole fraction of 0.316. As shown in Figure S1, the three spectra are practically identical, in contrast to the picture that similar measurements have revealed for NMA in acetonitrile.90 The absence of any NMAD aggregation is also supported by the absence of any measurable noncoincidence between the amide I′ bands in the IR and Raman spectra, as shown in Figure S2. In the case of NMAD oligomerization, transition dipole coupling between amide I′ modes would produce such a noncoincidence.90 As we argue in the Discussion, the two amide I′ subbands should instead be assigned to NMAD species hydrogen bonded (AI1) and nonhydrogen bonded (AI2) to functional groups of the solvent. The solvent dependence of AI1′ indicates that ethanol penetrates the carbonyl site of the hydration shell at rather low ethanol concentration, thus indicating some sort of preferential binding of the cosolvent. Structure Analysis of Cationic GAG. We measured the amide I′ band profile in the IR, isotropic Raman, anisotropic Raman, and VCD spectra of cationic GAG in ethanol/D2O mixtures with ethanol fractions of 0.03 (region 1), 0.12 (region 2), and 0.42 (region 3) at room temperature. The band profiles are shown in Figure 5, where the respective profiles for AI′ of GAG in D2O are also shown for comparison.44 Respective profiles observed for GAG in water and in the mixtures representing regions 1 and 2 are very similar. In all of them, the two amide I′ bands of the peptides are clearly resolved. The lower- and higher-wavenumber bands are predominantly assignable to the C-terminal and N-terminal peptide groups,21 respectively. We denote them as AIC and AIN in the following text. It must be noted that the two modes are subject to excitonic through-bond coupling that produces a mixing of their respective eigenvectors.20,21,76 Minor differences between the band intensity ratios are noticeable in the IR spectrum. The spectra observed for region 3 are clearly distinct from those of the other regions. Owing to a substantial red shift of AIC, a single band is now obtained with two underlying peptide bands. The VCD signal is reduced, which could in part be due to an increase in the spectral overlap between the negative and positive signals. Therefore, only a more detailed analysis of the profiles can reveal whether the observed changes in the region 3 spectrum reflect conformational changes in addition to spectral changes as recorded for NMA in ethanol/D2O (vide supra). As shown in numerous earlier papers, these band profiles reflect the conformational mixture sampled by the central alanine residue.76,79,91 To assess the conformational ensembles sampled at the investigated ethanol mole fractions, we additionally measured the homonuclear and heteronuclear

Figure 3. Spectral decomposition of the amide I′ region of NMA in 0.42 mol % EtOH/0.58 mol % D2O.

decreases upon further increases in the ethanol fraction. The maximum corresponds to region 1, where a microclustering of ethanol molecules is taking place.73,89 Above a mole fraction of 0.2, the wavenumber of AI′ levels off at ca. 1630 cm−1 and the intensity decreases with increasing ethanol content. The decrease is not monotonous; a plateau appears between 0.4 and 0.6 (Figure 4), which corresponds to region 3 in Milorey et

Figure 4. Wavenumber positions (upper panel) and integrated peak intensities (lower panel) of the amide I′ subbands of NMA plotted as a function of the ethanol mole fraction of binary D2O/ethanol mixtures: AI1 (closed circles) and AI2 (open circles). The spectral parameters were obtained by a decomposition of the amide I′ band profiles in Figure 1 using the program Multifit. 5748

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The Journal of Physical Chemistry B

Figure 5. Experimental (dotted line) and simulated (solid line) isotropic (top row) and anisotropic (second row) Raman, IR (third row), and VCD (bottom row) spectra of GAG in D2O (left) and in ethanol mole fractions of 0.03 (second from left), 0.12 (second from right), and 0.42 (right) plotted from 1600 to 1750 cm−1.

NMR J coupling constants 3J(HN,Hα) and 3J(HN,C′) as a probe of dihedral angle φ and 1J(N,Cα) as a probe of ψ. The φ and ψ dependences of these coupling parameters are empirically described by Karplus equations reported by Graf et al.23 To subject all of these spectroscopic data to a global fit, we constructed Ramachandran distributions as the superposition of two-dimensional Gaussian subdistributions associated with different secondary structures, as described in the Material and Methods section. For the simulations of the amide I′ band profiles, we used the algorithm of Schweitzer-Stenner, which reflects the conformational dependence of excitonic coupling between the N- and C-terminal amide I′ modes and the different conformational sensitivities of IR, Raman and VCD intensity distributions.79 To account for the experimental J coupling constants, constructed Ramachandran plots were used in combination with the respective Karplus equations. Table 2 lists the parameters and statistical weights of the twodimensional subdistributions that constitute the Ramachandran distributions with which we obtained the fits visualized by solid lines in Figure 5. The calculated J coupling constants are listed in Table 3 together with the respective experimental values. Generally, the agreement between simulation and experiment is very good. Figure 6 shows the Ramachandran plots of the distributions obtained for the peptide in water44 and in 0.42 mole fraction ethanol. The respective plots for GAG in 0.03 and 0.12 mole fraction ethanol are shown in Figure S3. The plots and the statistical weights listed in Table 2 reveal that conformational

Table 2. Parameters Used for the Simulations of the Vibrational Spectra of GAGa pPII

βT/βα

type II βT

αl

γ

0.00

0.03

0.12

0.42

0.72 −69 155 0.18 −115 155 0.04 −60 −120 0.03 −60 −30 0.04 20 −60

0.70 −75 150 0.15 −122 160 0.05 −70 −30 0.05 −52 −60 0.05 82 −60

0.65 −73 150 0.20 −120 160 0.05 −80 −30 0.05 −43 −60 0.05 82 −60

0.55 −75 140 0.35 −105 105 0.02 −80 −30 0.03 −43 −60 0.05 82 −60

a

The parameters include mole fractions as well as the centers of their (φ, ψ) distributions (in degrees) in the upper and lower parts of the respective cells.

distributions of GAG in mixtures with 0.03 and 0.12 ethanol are not much different from that of GAG in D2O. There is a slight tendency toward a decreasing pPII fraction with increasing ethanol content. The β-strand is slightly lower at 0.03 and somewhat higher at 0.12, but these variations might be in the margin of error. Of greater significance is the shift in the φ 5749

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The Journal of Physical Chemistry B Table 3. Comparison of Experimental and Calculated 3 J(HNHα) (Top), 3J(HNCO) (Middle), and 1J(NCα) (Bottom) Coupling Constants for GAG in Varying Mole Fractions of Ethanol at Room Temperature 3

0.03 0.12 0.42

J(HNHα)

3

J(HNCO)

J(NCα)

1

exptl

calcd

exptl

calcd

exptl

calcd

6.28 6.32 6.47

6.28 6.29 6.71

0.96 1.02 0.86

0.97 1.03 0.84

10.99 10.99 10.97

11.23 11.24 11.07

coordinates of both the pPII and β-strand to more negative values. This shift rather than conformational redistributions gives rise to the increase in 3J(HNHα) reported by Milorey et al.70 (Figure 1). The situation is totally different for a mixture with an ethanol fraction of 0.42, where the pPII fraction drops from 0.72 (in pure D2O) to 0.55. Concomitantly, the β-strand fraction increases from 0.18 in pure D2O to 0.35. Hence, the conformational distribution of GAG now becomes very similar to those observed for other GxG peptides with aliphatic residues.44,92 The total fraction of different turnlike structures, which includes the sampling of right- and left-handed helical conformations, varies between 0.11 (in water) and 0.15 (for regions 1 and 2). Temperature Dependence of 3J(HNHα). Figure 7 depicts the temperature dependence of the 3J(HNHα) coupling of GAG in the above investigated ethanol−water mixtures. We subjected these and 3J(HNHα) values measured earlier for other mixtures of ethanol and water to the following thermodynamic analysis. First, we dealt with the data in Figure 7. From the conformational analysis of GAG, we obtained the Gibbs energy difference ΔGPβ between pPII and β-strand conformations at room temperature TR. This allows us to relate enthalpic and entropic differences ΔSPβ by ΔSPβ =

Figure 7. 3J(NHNα) coupling constant of the N-terminal amide proton of GAG in mixtures with ethanol mole fractions of 0.03 (black), 0.12 (red), and 0.42 (green) plotted as a function of temperature. The solid lines result from fits described in the text, and the reduced χ2 values are 0.01, 0.95, and 0.04, respectively.

Hence, we could fit 3J(HNHα)(T) with eq 4 by using only ΔHPβ as a free parameter. As shown in Figure 7, the thusobtained fits are very good. To analyze additional 3J(HNHα)(T) data reported by Milorey et al. for other ethanol/water mixtures, we first estimated the ΔGPβ(TR) values by interpolation from the respective values obtained from the above-described conformational analysis. Subsequently, we again used ΔHPβ solely as a free parameters for our fits, which again yielded a satisfactory reproduction of the experimental data (Figure S4), as documented by the respective regression coefficient. For a further check of our thermodynamic analysis, we measured the UV-CD spectra of GAG in ethanol−D2O mixtures with 0.03, 0.12, and 0.42 ethanol fractions as a function of temperature (data not shown). From these spectra, we obtained the respective temperature dependence of the dichroism Δε216 at 216 nm, as plotted in Figure S5. We fit these data with a two-state model (pPII/β) by using

ΔHPβ − ΔG Pβ (TR ) TR

(5)

Figure 6. Contour plots depicting the conformational Ramachandran distribution of GAG in water (left) obtained from ref 11 and χET = 0.42 (right) as obtained from the analysis of the amide I′ band profiles in Figure 4 and the corresponding J coupling constants in Table 3. The procedure is detailed in the text. 5750

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The Journal of Physical Chemistry B Δε216(T ) =

ΔHPβ = ΔHP0β + TcΔSPβ

ΔεP2 + Δεβ e−ΔGPβ(T )/RT 1+e

−ΔG Pβ (T )/RT

(6)

where Tc is the compensation temperature and ΔH0Pβ is the excess enthalpy. For these parameters, the fit yielded Tc = 312 ± 25 K and ΔH0Pβ = −2.45 ± 0.61 kJ/mol. Both values are lower than those that Toal et al. observed from a similar plot for cationic trialanine in ethanol and glycerol (321.5 K and −4.28 kJ/mol).69 It should be noted that the enthalpic and entropic differences observed by these authors are substantially larger, which suggests significant nearest-neighbor influences on the thermodynamics of the central residue in tripeptides.43,69 One might suspect that substituting cosolvent D2O by H2O for the NMR measurements might cause some structural rearrangements due to the isotope effect. However, earlier experiments in our laboratory where we compared the CD spectra of GAG in H2O and D2O did not reveal any significant differences (data not shown). Analysis of Spectral Indicators. To relate spectral changes observed for GAG in ethanol−water to the abovedescribed changes in NMA band profiles, we subjected a larger set of earlier-reported amide I′ band profiles70 to spectral decomposition with Multifit. The results are depicted in Figure S4. The wavenumber position of AIN is practically unaffected by the addition of ethanol, and the obtained wavenumber indicates just a slight blue shift. On the contrary, AIC shows a clear blue shift of ca. 10 cm−1, which starts right below the ethanol fraction of 0.2 and levels off around 0.3. Thus, the shift of AIC correlates nicely with the above-reported shift of AI1 in the spectrum of NMA. Our results indicate a preferred ethanol solvation in the vicinity of the C-terminal peptide group. Chemical shifts are excellent indicators of peptide/protein structure and peptide−solvent interactions. The notion particularly applies to the signal of amide protons, which are frequently involved in hydrogen bonding interactions. For peptides such as GAG in ethanol−water, one can expect that the amide groups donate hydrogen bonds to both cosolvents. Generally, an upfield shift of a chemical shift position is interpreted as indicating a weakening of hydrogen bonding. Because thermal fluctuation (e.g., the occupation of higher vibrational levels of the highly anharmonic potential of hydrogen bonds) leads to a weakening of hydrogen bonds, it is generally expected that the NMR signal of a solvent-exposed amide proton moves upfield with increasing temperature. Temperature coefficients below e.g −6 ppb/Co are indicative of fully exposed amide groups that might be hydrogen bonded to the solvent.93 Significantly higher (i.e., less negative) values indicate some involvement of intramolecular hydrogen bonding. Figure 9 exhibits the chemical shift of the N-terminal amide proton of GAG in ethanol−water as a function of ethanol fraction for five different temperatures. At low ethanol fractions (region I), the NH signal moves upfield before it continuously increases with increasing ethanol content until it reaches a value in region III that exceeds that of GAG in water. These data indicate that upon moving across regions II and III the strength of peptide−solvent hydrogen bonding involving the amide proton increases continuously until the effect is leveling off above χET = 0.4.94 This is a somewhat surprising result in that it indicates that for ethanol fractions above this value NH− solvent hydrogen bonding is stronger than it is in pure water. For all ethanol fractions, the NH signal moves downfield with increasing temperature, reflecting the expected weakening of hydrogen bonding. Figure S7 depicts the corresponding

where we calculated the Gibbs energy difference from the thermodynamic parameters obtained from the fit to the corresponding 3J coupling data (vide supra). Thus, only dichroism values ΔεP2 and Δεβ were used as free parameters. This led to a good reproduction of the experimental data. The thermodynamic parameters obtained from this analysis are listed in Table 4; the corresponding spectroscopic parameters Table 4. Thermodynamic Values for GAG Obtained from Fitting the Vibrational Spectra and Various Coupling Constants ethanol mole fraction

ΔH (kJ/mol)

ΔS (J/mol·K)

0.00 0.03 0.07 0.12 0.17 0.24 0.32 0.42

−10 −5.2 −12 −11 −14 −7.2 −7.1 −8.8

−23 −6.7 −29 −26 −38 −16 −16 −24

(7)

can be found in Table S2. The existence of an isodichroic point suggests that we can neglect the most likely mutually neutralizing contributions from different turnlike structures. The behavior of the thermodynamic parameters can be correlated with regions 1−3 identified by Milorey et al.70 In region 1 (χET = 0.03), the enthalpy and entropy drop to lower values. In region 2 (χET = 0.1−0.2), the values are slightly though systematically higher (more negative) than they are for GAG in pure water. For χET > 0.2 (the transition region between regions 2 and 3), the values drop before they increase at 0.42 (region 3). The correlated changes in ΔHPβ and ΔSPβ suggest enthalpy−entropy compensation. This notion is confirmed by the enthalpy−entropy plot in Figure 8, to which we could fit the equation

Figure 8. Plot of ΔH versus ΔS values of cationic GAG in various ethanol/water mixtures obtained from a thermodynamic analysis of the temperature dependence of 3J(NHNα). The solid line results from a linear regression analysis that resulted in ΔH = −2.45 ± 0.61 kJ/mol and Tc = 312 ± 25 K. 5751

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switch that practically inverts the temperature dependence, thus yielding a positive temperature coefficient. Hence, for ethanol fractions of between 0.1 and 0.3 encompassing region 2 and the area between regions 2 and 3, the NH signal first shifts upfield and then shifts significantly downfield, indicating stronger hydrogen bonding with the solvent at high temperatures. In region 3, the temperature dependence is inverted in that the temperature coefficient is now positive over the entire temperature region investigated. Even though nonlinear temperature dependencies of chemical shifts are not unknown, their occurrence in the spectra of such a simple peptide is very surprising and indicates that the influence of the solvent on this tripeptide is far from being trivial. Although the chemical shifts of amide protons reflect the status of solute−solvent hydrogen bonding, changes in the chemical shift of alanine methyl protons could be indicative of hydrophobic alanine−ethanol interactions. Figure 12 depicts the apparent chemical shift position of the observed triplet as a function of ethanol fraction. At room temperature, there are not many changes below χET = 0.10, but the signal moves significantly upfield at 0.42. This is very reminiscent of what we observed for the C-terminal amide proton signal and indicates a common cause.

Figure 9. Chemical shift values of the N-terminal amide proton of GAG plotted as a function of the ethanol mole fraction of ethanol/ water mixtures for the indicated temperatures.

temperature coefficients as a function of ethanol fraction. Interestingly, the addition of only small amounts of ethanol (region 1) leads to a sharp increase in the temperature coefficient, which does not change much upon increasing the temperature further. A totally different picture emerges if one inspects the corresponding chemical shift behavior of the C-terminal proton displayed in Figure 10. At 30 °C, the signal position does not

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DISCUSSION Preferred Solvation of NMA by Ethanol. The spectroscopic composition of the amide I′ band profile indicates the coexistence of two NMA-solvent configurations for high fractions of ethanol and even in pure ethanol. Our experiments rule out the possibility that any of the two amide I′ bands reflect the formation of NMA-oligomers. To interpret our result, we refer to a spectroscopic study that Woutersen et al.86 conducted on NMAD in methanol. They observed two different bands amide I′ bands at 1631 and 1651 cm−1 that are rather close to the positions of corresponding bands in our spectra. On the basis of the results of 2D-IR experiments as well as MD simulations, they arrived at the conclusion that the lower-wavenumber band reflects a configuration with ethanol hydrogen bonded to the carbonyl group, whereas the higherwavenumber band probes an environment in which the carbonyl group is not hydrogen bonded. Their data do not provide any information about hydrogen bonding involving the amide group. In view of the similarity of methanol and ethanol, we conclude that the same scenario can be applied to our data: the AI1 band reflects hydrogen bonding between CO and ethanol, and AI2 indicates its absence. As described in the Results section, AI1 continuously blue shifts with increasing ethanol fraction and levels off after the latter becomes larger than 0.2. The shift itself indicates a replacement of D2O by ethanol as a hydrogen-bonding donor. This process certainly involves the formation of intermediates where both D2O and ethanol are hydrogen bonded to the carbonyl group, which could produce a wavenumber just in between the values for pure D2O and ethanol hydrogen bonding. Because the different configurations do not lead to discernible bands, the fluctuation between the configurations must be fast compared to the inverse of the difference between frequencies of individual vibrations.95 From the changes in the AI1′ wavenumber displayed in Figure 4, we thus estimate that these fluctuations should proceed on a subpicosecond time scale. The above data and the occurrence of AI2 at ethanol fractions larger than 0.2 are clearly indicative of preferred solvation by

Figure 10. Chemical shift values of the C-terminal amide proton of GAG plotted as a function of the ethanol mole fraction of ethanol/ water mixtures for the indicated temperatures.

change much until the mole fraction of ethanol exceeds 0.3 (entering region 3), where it shows a significant upfield shift, indicating a weakening of hydrogen bonding. Interestingly, this is the region where the wavenumber shifts of both AIC and AI1 are leveling off, thus indicating solvent exchange (Figures 4 and S4). Increasing the temperature causes rather dramatic qualitative changes in the ethanol dependence of the chemical shift, which is also visualized in Figure 11. It depicts the chemical shift as a function of temperature for different ethanol fractions. At low ethanol fractions (region 1), the behavior is quite normal in that the data indicate an upfield shift indicative of decreasing hydrogen bonding strength. In the transition region between 1 and 2 (close to χET = 0.1), we observe a 5752

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Figure 11. Chemical shift values for the C-terminal of GAG in pure water (top left) and in mixtures of ethanol/water with 0.03 (top right), 0.12 (bottom left), and 0.42 (bottom right) ethanol plotted as a function of temperature.

GAG in Different Ethanol−Water Mixtures. Though still not providing a unifying and consistent picture, experimental and computational work on ethanol−water mixtures have revealed a high degree of nonideality that starts at very low ethanol fractions. Milorey et al., on the basis of the ethanol dependence of the N-terminal J(HNHα) of cationic GAG, identified three regions in which the addition of ethanol to water caused nonlinear changes that could be correlated in part with changes in the IR spectrum of different ethanol−water mixtures (Figure 1).70 Their results strongly indicate that processes in the bulk are transduced to structural changes in the peptide. Below, we now discuss the solvent-induced changes in GAG structure and solvation in more detail based on the present experimental data. Region 1 of Milorey et al. lies between χET values of 0 and 0.05. Here, the N-terminal J(HNHα) coupling constant shows a local maximum at ca. 0.03 over a temperature region of 30−50 °C (Figure 1). Region 2, in which J(HNHα) exhibits a sudden increase, lies between 0.12 and 0.18. Region 1 and the area in between regions 1 and 2 (1−2) corresponds to a large number of phenomena that are diagnostic of the nonideality of ethanol−water mixtures with regard to their mean molar volume, the density of water, and a local maximum in the chemical shift of aqueous OH groups.71,72,89,94 MD simulations indicate that these anomalies reflect phase-separation ethanol aggregation on a more microscopic scale.89 The observed NMR data (chemical shifts and N-terminal J coupling constants) suggest a weakening of hydrogen bonding between the Nterminal NH group and the solvent and some changes in the

Figure 12. Chemical shift values of the methyl protons of the alanine side chain of GAG plotted as a function of the ethanol mole fraction of ethanol/water mixtures for the indicated temperatures.

ethanol. This is not totally unexpected because experimental data indicate that peptides with hydrophobic residues in particular attract ethanol even if the fraction of the cosolvent is rather low.69,70 However, other data on, for example, tetrapeptides suggest that the preference for ethanol over water is minimal.96 In the case of NMA, the two methyl groups are certainly strong attractors for the hydrophobic side of ethanol, but our data indicate an accumulation at the carbonyl side as well. 5753

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The Journal of Physical Chemistry B backbone structure of GAG, but our detailed structural analysis of the peptide reveals only minor changes that mostly involve the φ coordinate of the alanine residue. Interestingly, H NMR data on ethanol−water mixtures indicate that water−water interactions increase in this region until they level off when the ethanol fraction reaches 0.1.94 Changes at the CO sites of both peptide groups and at the site of the N-terminal amide group are negligible at room temperature (NMR and IR data). With increasing temperature, the respective chemical shift changes in the 1−2 region, and the shift is upfield at lower and surprisingly downfield at higher temperatures. The most pronounced effect can be inferred from the thermodynamics obtained from the N-terminal J(HNHα) coupling constant: Upon the addition of a 0.03 fraction of ethanol, the enthalpic and entropic differences between pPII and the β-strand decreases by factor of 2 and 3, respectively. Although that does not significantly affect the populations of both states at room temperature owing to the close proximity of room temperature to the compensation temperature Tc (vise supra), a structural redistribution occurs at high temperatures (i.e., 59% pPII and 31% β-strands for D2O, 65% pPII and 25% at 0.03 ethanol). In other words, pPII becomes stabilized in region 1 at high temperatures, which is a very surprising result. This finding underscores the notion that just probing Gibbs energy landscapes of peptides at room temperatures does not yield a complete picture. Region 2 lies between χET values of 0.12 and 0.18, followed by the 2−3 transition region between 0.18 and 0.32. There are three remarkable observations for region 2. First, the (excess) enthalpy of ethanol−water mixing exhibits a minimum.71 Second, the velocity of sound depicts a local maximum.74 Third, the N-terminal 3J(HN,Hα) jumps from 6.18 to 6.26 Hz. Generally, it can be said that the ethanol clusters increase in this region and, as a consequence, hydrogen bonding between ethanol molecules and between ethanol and water increases at the expense of water−water bonding. Mijakovic et al., by means of MD simulations, arrived at a picture in which the percolation network in the remaining water clusters stiffens and thus affects the sound velocity.74 Our structural analysis suggests a slight increase in the β-strand population at the expense of pPII, in line with what the 3J(HH,Hα) case suggests. The N-terminal amide proton chemical shifts move continuously toward lower field values at all temperatures. This is indicative of a decrease in the hydrogen bonding between NH and H2O. Again, the behavior of the C-terminal amide proton is different. It remains unaffected at room temperature. The temperature dependence is peculiar again; the signal first moves upfield and subsequently (>T = 35 °C) downfield. The AIc wavenumber starts to blue shift in this region. The change of the thermodynamic parameter inferred from 3 J(HH,Hα)(T) is again remarkable in that both enthalpy and entropy jump from the very low values in region 1 to what one could characterize as a local maximum in region 2 (Table 4). This indicates a larger destabilization of pPII at high temperatures (ca. 350 K) in the latter region (61% pPII at 0.03 and 46% at a 0.17 ethanol fraction). Altogether, these data indicate an increased interaction of ethanol with the peptide at higher temperatures, which seems to occur more on the side of the C-terminal amide proton. Interestingly, the data of Mijaković et al. indicate that it is in region 2 where the temperature dependence of the sound velocity is inverted from a positive to a decreasing temperature coefficient, which could indicate a loosening of the water hydrogen-bonding network.74

If this happens, then ethanol clusters might absorb a lot of water and thus dehydrate the solvation shell of the peptide. We now move to region 3 (χET values of between 0.32 and 0.44), in which the 3J(HH,Hα) of the N-terminal shows its largest increase (6.34 to 6.55 Hz) before leveling off again for higher ethanol fractions where the peptide is known to selfaggregate and gel.70,75 Here, the solvent excess entropy exhibits its minimum.71 This is the region where ethanol clustering becomes very dominant for NMA97 and our data indicate preferred solvation with ethanol. However, for GAG this occurs predominantly on the C-terminal side, as indicated by the substantial upshift of the low-wavenumber amide I′ and the abrupt high-field shift of the C-terminal amide proton signal (Figure 10). The temperature coefficient of this signal is now entirely positive. The structural analysis reveals a substantial destabilization of pPII even at room temperature, with corresponding gains for the β-strand. Interestingly, the enthalpy and entropy values moved back to lower values, which lie somewhere between water and 0.03 ethanol values. As a consequence, the pPII (0.48) and β-strand mole fractions (0.42) at 350 K are comparable with corresponding values obtained for region 2 (0.46 and 0.39, respectively). We derive the following picture of peptide−solvent interactions from our data. In line with the findings of Milorey et al.,70 our data show that the peculiar properties of water− ethanol have an impact on how a solvent and GAG interact. With increasing ethanol concentration, water in the hydration shell is replaced by ethanol. This occurs predominantly on the C-terminal side of the peptide. The preferred solvation with ethanol also affects the alanine side chain. Interestingly, our data reveal that the preferred solvation with ethanol starts earlier (i.e., at lower ethanol fractions) at higher temperatures. We assign this behavior to a higher preference of ethanol binding for the β-strand conformation over pPII. This model explains first why ethanol solvation stabilizes the β-strand. Second, it explains the positive temperature coefficient of the C-terminal chemical shift in region 3. Third, it explains why there is not much difference between the conformational distributions of GAG in regions 2 and 3 at high temperature (353 K). The enthalpy−entropy compensation displayed in Figure 9 deserves some further consideration. Its very existence suggests that the considered thermodynamic systems have very similar compensation temperatures at which the total enthalpic difference between two states is being neutralized by the respective entropy difference.98 A recent comparison of GAG with other GxG-type peptides (with x representing the host amino acid residues) in water revealed that GAG is peculiar in that it does not share isoequilibria exhibited by most of the other investigated GxG (10).53 The enthalpy−entropy plot for GAG in different water−ethanol mixtures reveals the reasons. First, the negative value of the non-compensated enthalpy ΔH0Pβ is much larger for GAG (−2.46 kJ/mol) than the average value observed for GxG (−0.77 kJ/mol). Second, compensation temperature Tc is higher for the former (312 K) than it is for the latter (295 K). Because the total Gibbs energy reads as ΔG Pβ = ΔHP0β + (Tc − T )ΔSPβ

(8)

both effects increase the negative amount of ΔGPβ below Tc, thus increasing the pPII fraction. The respective values for AAA (−4.28 kJ/mol and 321 K) are even more pronounced, reflecting the even higher pPII propensity of the central alanine. 5754

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The Journal of Physical Chemistry B In spite of inducing substantial changes in ΔHPβ and ΔSPβ, the addition of ethanol to ΔGPβ remains modest in regions 1 and 2 at room temperature, owing to enthalpy−entropy compensation and the proximity to the compensation temperature. The situation is different only in region 3, which might reflect a (slightly) lower ΔHPβ value or a lower compensation temperature than the average value that emerged from the fit to the data in Figure 9. If one combines the site-specific ethanol solvation of GAG suggested by our data with the suggested existence of large ethanol clusters in region 3,97 then a very interesting scenario emerges. Our data seem to indicate that GAG accumulates preferentially at the surface of ethanol clusters, which it penetrates with its polar C-terminal side while the N-terminal side sticks into the aqueous hydration shell of clusters. Egashira and Nishi recently suggested that ethanol clusters in water might form a bilayer that depicts the OH group at the ethanol− water interface and the carbon−hydrogen chain packed in the interior.97 Such a coexistence of water and primary alcohol clusters has recently been identified to form in mixtures of 70 mol % methanol/30 mol % water, where one would expect a preponderance of isolated water molecules in a bath of methanol if the solution was behaving ideally.99 As illustrated in Figure 13, our data could be understood as indicating that

solvent-induced spectroscopic shifts. Our results indicate a preferred ethanol solvation of NMA above an ethanol mole fraction of 0.3. For GAG, preferred solvation is restricted to the C-terminal half and the alanine side chain of the peptide. We propose that GAG accumulates on the interface between ethanol clusters and water where the C-terminal and alanine side chain penetrate the former. It is obvious that this would cause an increase in the local peptide concentration, thus facilitating aggregation and even fibrilization. Our structural analysis reveals that adding ethanol up to mole fractions below 0.4 at room temperature causes some minor structural rearrangements within pPII and β-strand subdistributions but only minor changes in their statistical weight. However, owing to rather significant changes in the enthalpic and entropic differences between these conformations, the addition of ethanol causes much more pronounced changes in the pPII/ β-strand equilibrium at high temperatures (350 K). The situation changes above an ethanol mole fraction of 0.4, where ethanol shifts the equilibrium significantly toward the βstrand at room temperature and even more strongly at 350 K. The abnormal temperature dependence of the chemical shift of the C-terminal amide proton and of the alanine side chain strongly suggests that for fractions above 0.3 ethanol stabilizes the β-strand over pPII by a stronger interaction (hydrogen bonding) with the solvent. The thermodynamic analysis of our data reveals again a strong enthalpy−entropy compensation at physiological temperature, which substantially reduces the influence of the ethanol cosolvent on GAG conformation.

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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.jpcb.7b02899. Fitting, IR, and Raman parameters; IR spectra of different concentrations of NMA in ethanol/water; a comparison of IR and Raman spectra of NMA in the same cosolvent mixture; Ramachandran plots of GAG in two different ethanol/D2O mixtures; plots of the amide I′ wavenumbers of GAG as a function of ethanol−D2O mixtures; J coupling constants and Δε216 plotted as a function of temperature for different ethanol−D2O mixtures; and temperature coefficients of chemical shifts plotted as a function of ethanol−D2O mixtures (PDF)

Figure 13. Pictorial representation of how GAG would assemble itself between the ethanol bilayers and water.

GAG peptides penetrate these bilayers to some extent so that the C-terminal and the alanine side chain predominantly probe the less-polar environment in the interior of the micelles. The N-terminal is still hydrated and to some extent reflects the hydration shell of the micelles. The water−ethanol interface could involve hydrogen bonding between ethanol and water. This model nicely explains the sharp decrease in the peptide’s solubility at higher ethanol concentration and at neutral pH where the C-terminal is charged (data not shown). Moreover, it could explain why GAG eventually aggregates if the ethanol fraction is increased beyond region 3. The proposed interaction with ethanol clusters would increase the local density and also cause some type of local order, which could facilitate interpeptide hydrogen bonding.

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AUTHOR INFORMATION

Corresponding Author

*Phone: 215-895-2268. E-mail: rschweitzer-stenner@drexel. edu.

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ORCID

SUMMARY It has been previously shown that 200 mM GAG aggregates into long fibrils and forms a gel once the ethanol mole fraction reaches the value of 0.55.70,75 The current study was aimed at exploring how ethanol solvation and solvent−peptide interactions prepare the system for aggregation and fibrilization. We utilized amide I′ band profiles, J coupling constants, and chemical shifts of amide and methyl protons of GAG in different ethanol−water/D2O mixtures. Concomitant vibrational spectroscopy measurements were carried out with NMAD, which was used as a reference system for identifying

Harald Schwalbe: 0000-0001-5693-7909 Reinhard Schweitzer-Stenner: 0000-0001-5616-0722 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We appreciate some financial support for this project from the Department of Chemistry at Drexel University (for R.S.-S.). We also thank Dr. Lynn Penn for sharing reference 99 with us. 5755

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