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Article Cite This: Inorg. Chem. XXXX, XXX, XXX−XXX

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Cu(II) Binding to the Peptide Ala-His-His, a Chimera of the Canonical Cu(II)-Binding Motifs Xxx-His and Xxx-Zzz-His Paulina Gonzalez,†,‡ Bertrand Vileno,†,§ Karolina Bossak,∥ Youssef El Khoury,⊥ Petra Hellwig,⊥ Wojciech Bal,∥ Christelle Hureau,*,‡,#,□ and Peter Faller*,†,‡ †

Institut de Chimie, UMR 7177, CNRS, Université de Strasbourg, 4 rue Blaise Pascal 67000, Strasbourg, France University of Strasbourg Institute for Advanced Study (USIAS) 67000, Strasbourg, France § French EPR Federation of Research (REseau NAtional de RPE interDisciplinaire (RENARD), Fédération IR-RPE CNRS #3443) 67081, Strasbourg, France ∥ Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Pawińskiego 5a 02-106, Warsaw, Poland ⊥ Laboratoire de bioélectrochimie et spectroscopie, UMR 7140, Université de Strasbourg, 4 Rue Blaise Pascal 67000, Strasbourg, France # LCC (Laboratoire de Chimie de Coordination), CNRS, 205, route de Narbonne F-31077, Toulouse, France □ UPS, INPT, LCC, Université de Toulouse F-31077, Toulouse, France ‡

S Supporting Information *

ABSTRACT: Peptides and proteins with the N-terminal motifs NH2-Xxx-His and NH2-XxxZzz-His form well-established Cu(II) complexes. The canonical peptides are Gly-His-Lys and Asp-Ala-His-Lys (from the wound healing factor and human serum albumin, respectively). Cu(II) is bound to NH2-Xxx-His via three nitrogens from the peptide and an external ligand in the equatorial plane (called 3N form here). In contrast, Cu(II) is bound to NH2-Xxx-ZzzHis via four nitrogens from the peptide in the equatorial plane (called 4N form here). These two motifs are not mutually exclusive, as the peptides with the sequence NH2-Xxx-His-His contain both of them. However, this chimera has never been fully explored. In this work, we use a multispectroscopic approach to analyze the Cu(II) binding to the chimeric peptide AlaHis-His (AHH). AHH is capable of forming the 3N- and 4N-type complexes in a pH dependent manner. The 3N form predominates at pH ∼ 4−6.5 and the 4N form at ∼ pH 6.5−10. NMR experiments showed that at pH 8.5, where Cu(II) is almost exclusively bound in the 4N form, the Cu(II)-exchange between AHH or the amidated AHH-NH2 is fast, in comparison to the nonchimeric 4N form (AAH). Together, the results show that the chimeric AHH can access both Cu(II) coordination types, that minor changes in the second (or further) coordination sphere can impact considerably the equilibrium between the forms, and that Cu kinetic exchange is fast even when Cu-AHH is mainly in the 4N form.

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INTRODUCTION

include a His close to the unprotected N-terminal of the peptide/protein, where the sequence is NH2-Xxx-Zzz-His (XZH) in which Xxx/Zzz is any amino acid and Zzz is any amino acid but Pro.9,10 The most representatives of the high affinity Cu(II) binding domain are found in human serum albumins, in which the sequence is Asp-Ala-His. Serum albumin is implicated in the Cu transport in the bloodstream via the complex with this motif.9 More peptides/proteins in the human body possess XZH N-terminal sequences, for example the saliva peptide Histatin-5,11,12 the copper transporter Ctr1,13 the truncated amyloid-β peptide,14 etc. The other motif is NH2-Xxx-His (XH), in which Xxx is any amino acid but Pro. The most classical example of this motif is the short peptide Gly-His-Lys (GHK).6 This peptide was

Copper is an essential metal for most living organisms. Copper exists mainly in the two redox states +1 and +2 in vivo and is generally bound to proteins.1,2 Several classes of such proteins can then be conceptually distinguished: (i) enzymes with one or more Cu centers that are essential for catalytic activity such as electron transfer and oxygen reduction in cytochrome c oxidase or redox reaction with superoxide in SOD, etc; (ii) Cu transporters, which bring the essential copper to aforementioned proteins and can be subdivided into soluble Cu transporters, designated as Cu chaperons that transport Cu from one protein to another,3 and transmembrane proteins such as Ctr1 or ATP7a/b that transport Cu across the membrane;4,5 and (iii) small molecules, often peptides such as glutathione or the wound-healing factor GHK.6 In addition, there are two well-known, strong, and very small Cu(II)-binding motifs found in peptides and proteins.7,8 Both © XXXX American Chemical Society

Received: August 16, 2017

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DOI: 10.1021/acs.inorgchem.7b01996 Inorg. Chem. XXXX, XXX, XXX−XXX

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Inorganic Chemistry

Figure 1. Top: Scheme of the structure of the complexes Cu(II)-Xxx-His (3N coordination) and Cu(II)-Xxx-Zzz-His (4N coordination). L depicts an external ligand such as H2O. Bottom: 3N and 4N form of the Cu(II)-Ala-His-His. The applied nomenclature for the His imidazole ring is shown at the bottom left.

of the metal ion slows with the number of amide ligands due to the slow protonation or deprotonation needed to release or bind metal ions, respectively.31 Thus, XH and XZH motifs are of fundamental interest in coordination chemistry. These motifs are naturally present in several peptides and proteins and have been widely studied for more than five decades.7−9,32−34 Despite this interest, reports on the Cu coordination of a chimera containing the two motifs, i.e. Xxx-His-His (XHH), have been very limited,35 and a systematic study of the pure motif has not been reported. This motif has the possibility to bind Cu(II) in either coordination mode (Figure 1), i.e. as in Xxx-His (e.g., GHK) or Xxx-Zzz-His (e.g., DAHK). The two binding sites are mutually exclusive in terms of binding Cu(II) ions because they share potential ligands (N-terminal amine and first amide). However, an intramolecular pH dependent rearrangement is possible (switching from XH to XZH and vice versa).

described as a wound-healing factor and was shown to participate in tissue repair. Moreover, GHK stimulated the growth and improved viability of several types of cultured cells and organisms.15 Both peptide types bind Cu(II) with the N-terminal amine and the Nπ of the His side chain (see Figure 1, bottom left for the nomenclature of His). The difference between them lies in the number of amide ligands coordinated. For XH, there is only one amide binding between Xxx and His, whereas in XZH, both amides in between the N-terminal amine and His bind the metal ion. It is interesting to note that the affinities of the two most representative peptides, i.e. GHK and DAHK (from human serum albumin), for Cu(II) at pH 7.4 are almost the same.16−26 Both have quite strong affinity with a Kd around 10−13 M for GHK, while the DAHK is 2−3 times stronger. Moreover, the simplest representative of the XZH family, GGH, has a Cu(II) affinity about 10-fold higher than that of GH at pH 7.4.7,27 This means that the advantage from having more coordination bonds from one peptide, i.e. a chelate effect by going from a tridentate to a tetradentate ligand, is mostly but not totally compensated by the high energy that requires the deprotonation of the amide (which has a high pKa of around 15).28 Despite this very similar affinity at pH 7.4, the two motifs have quite different kinetic properties. NMR experiments showed a very fast self-exchange of Cu-GHK (ms or faster), whereas DAHK did not exchange Cu(II) on the NMR time scale. This difference was assigned to the different coordination sphere, to the number of coordinating amidate ligands, and in particular to the free equatorial coordination site in Cu-GHK that is occupied in solution by water or other small molecules.29,30 It has been known that binding and releasing

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RESULTS AND DISCUSSION Cu Binding to AHH. The pH dependence of the Cu(II) coordination of XH and XZH motifs has been widely studied.35−38 Cu(II) binds to the XH motif via three nitrogens (Figure 1; N-terminal amine: NH2; amide: N−; imidazole: Nim) and to XZH motifs via four nitrogens (NH2, N−, N−, and Nim). Such complexes are often denoted as 3N and 4N, with respect to the amount of nitrogens being coordinated. Cu(II) binding occurs over a wide range of pH values with a full complex formation for 5 < pH < 11. The typical absorption maxima for d−d transitions are around 610 and 520 nm for 3N and 4N, respectively.7 B


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Inorganic Chemistry

Figure 2. UV−vis (left) and circular dichroism (right) spectra of Cu-AHH at different pH values (where the dashed arrow in the UV−vis spectra indicates increasing absorption). Conditions for the UV and CD experiments: final concentrations of 2 mM AHH and 1.8 mM CuCl2 were titrated with 1 μL aliquots of 5 M NaOH solution, and the reaction was left to equilibrate for ∼10 min at each NaOH addition.

Before studying AHH, a double-histidine model peptide, we analyzed individual motifs within simple AH and AAH sequences. As expected, at pH 7.4, Cu(II) binds to AH and AAH in a 1:1 stoichiometry with the d−d transition bands at 610 and 520 nm, respectively (Figure S1; an overview of the reported spectroscopic characterization is given in Table S1). Afterward, pH dependent absorption spectra of Cu-AHH were recorded for 0.9:1 Cu(II) to peptide stoichiometry, making sure there was no excess of copper over peptide, thus preventing a copper hydroxide precipitation in neutral and alkaline pH (Figure 2). The spectra are characterized by at least three distinctive species. At low pH, the λmax of the d−d bands are around 800 nm, which correlates with a band for a Cu(II) aqueous ion. With the increase in pH, first a band at ∼610 nm specific to 3N coordination started to appear, followed by a blue shift of the band maximum to 520 nm, thus indicating the formation of a 4N complex. With further increase of pH, again, a small blue shift from 520 to 510 nm occurs. The final formed species with this d−d band maximum we call 4N′. The origin of this species is not clear, but it does not change the first equatorial coordination sphere (see below). The circular dichroism (CD) data are in a good agreement with the pH dependent UV−vis spectra (Figure 2, right). The pattern of the CD spectrum at pH ∼ 4 is very similar to the one of Cu-GHK29 with two positive bands around 600 and 330 nm and two negative ones around 510 and 300 nm. At higher pH values, the Cu/AHH spectrum presents major changes, where previous characteristic bands are substituted with two new positive bands at 490 and 310 nm and just one negative band around 550 nm. This resembles the sign pattern and position of the spectra for the Cu-DAHK complexes.29 As shown in Figure 3, the coordination of Cu-AHH complexes as a function of pH was also investigated by means of low temperature electron paramagnetic resonance (EPR) (100 K). In agreement with the UV−vis and CD spectra, the EPR fingerprints (g parallel values and corresponding hyperfine splitting constants) show the presence of the 3N form (g// = 2.232, g⊥ = 2.048, A// = 562 MHz = 187 × 10−4 cm−1 at pH 3.6), which shifts with the increase of pH toward the 4N conformation (g// = 2.177, g⊥ = 2.043, A// = 614 MHz = 204 × 10−4 cm−1 at pH 7.8)29 (EPR parameters are assembled in Tables S1 and S2). It is important to note that the changes occurred at pH lower than that obtained in the absorption and CD spectra. These variations

Figure 3. EPR spectra of Cu-AHH at pH 3.6 (purple), 4.9 (red), 6.2 (green), 6.9 (blue), and 7.8 (orange): conditions: C[Cu] = 1.8 mM [AHH] = 2 mM, no buffer, 30% glycerol at 100 K. Dotted lines a and b indicate the lowest field peak of the Cu(II) parallel hyperfine splitting constant of the 3N and 4N complexes, respectively. EPR parameters as follows: g// = 2.232, g⊥ = 2.048, and A// = 562 MHz = 187 × 10−4 cm−1) at pH 3.6 and g// = 2.177, g⊥ = 2.043, and A// = 614 MHz (= 204 × 10−4 cm−1) at pH 7.8 (EPR experimental parameters: microwave power, 0.1 mW; modulation amplitude, 5G; modulation frequency, 100 kHz; receiver gain, 30 dB).

are assumed to arise from changes in the protonation states subsequent to freezing.39 Vibrational Spectroscopies. Further characterization of the two forms of Cu-AHH were obtained by measuring Fourier transform infrared (FTIR) and Raman spectra at pH 5.2 and 7.1 (Figure 4). The assignment of the infrared modes discussed below was made according to previous studies, including individual amino acids, Cu-peptide model compounds, and theoretical calculations.40−46 The vibrational spectra of AH and AAH and their Cu(II) complexes are reported in Figures S3− S6. The FTIR spectra of free and Cu-bound AHH as dried films on a diamond ATR crystal at pH 5.2 and 7.1 are shown in Figure 4, left. At pH 5.2, the amide I band is observed at 1663 cm−1, while the stretching (C4C5) vibration of the C


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Figure 4. FTIR (left) and Raman (right) spectra of free (black trace) and Cu-bound (red trace) AHH at pH 5.2 (top) and 7.2 (bottom).

protonated His appears at 1626 cm−1. This signal was not sensitive to H/D exchange (data not shown), which leads to excluding a contribution from the bending (NH3+)as vibration of the N-terminus, which is known to downshift to around 1200 cm−1 upon N-deuteration. Upon Cu(II) binding at pH 5.2, the amide I band at 1626 cm−1 lost intensity. This leads to the conclusion that at least one amide group is deprotonated and binds Cu(II). The band observed at 1256 cm −1 corresponds to the bending (C4H)/stretching (C2Nπ) vibrations of Nπ-protonated form of His. This His band at 1256 cm−1 upshifts to 1272 cm−1 upon Cu(II) binding, a signal characteristic of the stretching (CN) ring vibration of imidazole of either Nπ or Nτ Cu-bound form while Nτ or Nπ is protonated, respectively. At pH 7.2, the amide I band can be found at 1653 cm−1 in the spectrum of the free peptide, and it is lost upon Cu(II) binding. This behavior is typical of the deprotonation of the amide function (CN becomes N−) and subsequent binding to Cu(II). The carboxylate of the C-terminus binds Cu(II) because the stretching (COO−)as vibration downshifts from 1582 cm−1 in the free form to 1568 cm−1 in the Cu-bound form. Note that the His stretching (C4C5) vibration also occurs in this spectral range. Yet, it is known that the frequency of the stretching (C4C5) vibration upshifts upon metal binding. Accordingly, the intense signal observed here is dominated by the stretching (COO−)as vibration of the Cterminus. To complete the assignments performed for FTIR, especially regarding the Cu-binding mode to the His residues, Raman spectra have been recorded on the same samples (see Figure 4, right). The main goal was to determine which N atom of the imidazole ring binds Cu(II). At both pH values, the stretching (NπC5) vibration is seen at 1264 and 1266 cm−1 for pH 5.2 and 7.2, respectively. Upon Cu(II) binding, the signal upshifts toward 1271 at pH 5.2 and 1278 cm−1 at pH 7.2, which is an indicator of His binding to Cu(II) via Nπ while Nτ is protonated.

The IR and Raman spectra confirm the pH dependent binding of Cu(II) at pH 5.2 in the 3N form and at pH 7.2 in the 4N form. For both forms, Raman shows the binding of the Nπ of the His, in line with the favored 6-membered chelate ring (compared to a 7 chelate ring in the case of the Nτ coordination). The only difference is the proposed binding of the COO− to the Cu(II). It is noteworthy that our vibrational spectroscopy measurements were done at very high concentration compared to the other measurements (from a 40 mM solution to drying). This indicates that COO− binds intermolecularly to Cu(II) only weakly and hence is observed only at very high concentration or in a dried sample. Precedence from GHK shows that COO− can bind in the crystalline state but does not in more diluted solution.47,48 Potentiometry of AHH. Potentiometry was applied to obtain the thermodynamic constants and fully quantitative pH distribution of complexes. The pKa values of free AHH are given in Table 1. Four values were obtained for the C-terminal carboxylate, both His imidazoles, and the N-terminal amine. The distribution of Cu(AHH) species, their stability constants, and corresponding pKa values are given in Figure 5 and Table 2, respectively. The agreement between the potentiometry and Table 1. Protonation Constants and Their Corresponding pKa Values of AHH and AHH-NH2 Determined with the Use of Potentiometry at 25 °C and I = 0.1 Ma

AHH−CONH2

AHH−COOH

a

D

protonation form

log β ± SD

HL H2L H3L HL H2L H3L H4L

8.032 ± 0.007 14.622 ± 0.008 20.41 ± 0.01 8.176 ± 0.003 15.222 ± 0.003 21.317 ± 0.003 23.572 ± 0.006

pK (deprotonated function) 8.03 6.59 5.79 8.18 7.05 6.10 2.20

(N-term) (His) (His) (N-term) (His) (His) (C-term)

Standard deviations (SD) are provided by Hyperquad software. DOI: 10.1021/acs.inorgchem.7b01996 Inorg. Chem. XXXX, XXX, XXX−XXX

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Inorganic Chemistry

decrease in the complex vs the peptide and the change of the peptide conformation. Cu Binding to AHH-NH2. The motifs of the XH and XZH types are usually not present just as respective di- and tripeptides but rather as N-termini of longer peptides or proteins. To mimic that, we investigated the C-terminal amide form, i.e. AHH-NH2, which reproduces better the downstream peptidic bond in comparison to the carboxylate form. We applied UV−vis, CD, EPR, and potentiometry (Figures 6, S2 and 7) to investigate the Cu(II) binding to this peptide. The

Figure 5. Species distribution diagram for AHH/Cu(II) complexes at 25 °C based on potentiometry determined stability constants that were modeled for 2 mM peptide and 1.8 mM CuCl2. The left-side axis denotes molar fractions of Cu(II) complexes. Calculated species are marked in solid lines as follows: Cu(II) (black), CuHL (purple), CuL (red), CuH−1L (green), CuH−2L (blue) and CuH−1L2 (dark red). The overlap of potentiometric and spectroscopic data is given in Figure S7.

Table 2. Stability Constants and Corresponding pKa Values of Measured Binary Complexes of AHH/Cu(II) Determined by Potentiometry at 25 °C and I = 0.1 Ma species AHH

a

log β ± SD

pK

CuHL

16.538 ± 0.009

CuL

12.660 ± 0.004

3.88

CuH−1L

6.229 ± 0.006

6.43

CuH−2L

−1.868 ± 0.008

8.09

CuH−1L2

9.81 ± 0.02

coordination mode

Figure 7. Species distribution diagram for Cu(II)-AHH-NH 2 complexes at 25 °C based on potentiometry determined stability constants that were modeled for 2 mM peptide and 1.8 mM CuCl2. The left-side axis denotes molar fractions of Cu(II) complexes. Calculated species are marked in solid lines as follows: Cu(II) (black), CuL (red), CuH−1L (green), CuH−2L (blue), and CuH−1L2 (purple).

3N: NNH2, N−, Nπ, protonated COOH 3N: NNH2, N−, Nπ, deprotonated COO− 4N: NNH2, N−, Nπ, protonated His 4N: NNH2, N−, Nπ, deprotonated His 3N+1N: NNH2, N−, Nπ, and Nπ

pKa values for free AHH-NH2 obtained by potentiometry are given in Table 1. Three protonation steps for AHH-NH2 were detected, corresponding to deprotonations of two imidazole rings of His residues and the N-terminal amine. All three values are slightly lower (0.15−0.46 pH units) compared to those of AHH. Potentiometry of AHH-NH2. Based on UV−vis and CD spectra we were able to assign the Cu(II) species distribution according to the pH changes (Figure 7). Already below pH 4 we observed a decrease of Cu(II) aqua ion and the appearance of the 3N complex (CuL), where Cu(II) is bound to NIm of His2, Ala1-His2 amide bond nitrogen and the N-terminal amine.

Standard deviations (SD) are provided by Hyperquad software.

the spectroscopic data (UV−vis and CD) is satisfactory. Four major forms can be observed from low to high pH: free Cu(II) and 3N, 4N, and 4N′ forms. The 3N complex includes CuLH and CuL species, differing by protonation of the carboxylate. The elevation of its pK from 2.2 in the free peptide to 3.9 in the complex can be attributed to a combined effect of charge

Figure 6. UV−vis and CD spectra of Cu-AHH-NH2 at different pH values. Experimental conditions for the UV−vis and CD: Final concentrations: 2 mM peptide, 1.8 mM CuCl2. Titrations were made by adding 1 μL aliquots of 5 M NaOH. The reaction was left to equilibrate for ∼10 min at each NaOH addition. The final volume of the solution was 1.5 mL. E


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absorption and CD spectra in solution. However, at a ratio of 0.9 equiv, a broad feature was observed superimposed to the classical isolated 4N fingerprint. The loss of the hyperfine structure is assigned to the presence of a dimeric or oligomeric forms of copper complexes. Such aggregation pattern is already observable at 0.9 equiv of copper with respect to 2 mM of peptide and becomes largely predominant at overstoichiometry. Similarly, the AHH-NH2 copper complex exhibits comparable behavior at the same copper threshold. It is important to note that such loss of the hyperfine structure was not obtained for AH and AAH copper complexes even for Cu(II) overstoichiometry (Figure 8, lower panel). This indicates that such interaction is specific to AHH (Figure 8, upper panel).

The directness of this reaction was confirmed by the presence of the isosbestic point at 750 nm in the UV−vis spectra. This complex becomes the major species up to pH ∼ 5.4. The CuH−1L complex, assigned as the 4N complex, appears about pH 6.2 (Table 3). With an increase of pH a further Table 3. Stability Constants and Corresponding pKa Values of Measured Binary and Ternary Cu(II) Complexes of AHHAmide Determined by Potentiometry at 25 °C and I = 0.1 M species

log β ± SD

CuL CuH−1L CuH−2L

12.449 ± 0.007 6.94 ± 0.01 −0.85 ± 0.02

CuH−1L2

10.97 ± 0.03

pK

coordination

5.51 7.69

3N: NNH2, N−, Nπ 4N: NNH2, N−, Nπ, protonated His 4N: NNH2, N−, Nπ, deprotonated His 3 + 1 N: NNH2, N−, Nπ, and Nπ

deprotonation occurs and the 4N′complex with the same equatorial coordination sphere as 4N is formed (CuH−2L) (see also above). We could also observe small amounts of a biscomplex CuH−1L2 around pH 6−10. Overall, the behavior of Cu-AHH-NH2 was similar to that of Cu-AHH. These results were expected as COO− was not involved in the equatorial coordination of Cu(II) in solution. However, a surprisingly large shift of the pH distribution between the 3N (CuL) and 4N (CuH−1L) forms was observed. For Cu-AHH, the pH of equal distribution between 3N and 4N was 6.4. The latter drops to pH 5.5 for Cu-AHH-NH2. The shift was corroborated by the UV−vis and CD data (see above). This large pH shift could be due to H-bonding between the COO− and imidazole of the His at position 3 or between the COO− and the H−N of the amide (between the two His). These H-bonds involve an entropically favored six-membered ring and can occur in the 3N form, but they would have to be broken to make the 4N form; therefore, the 3N form is more stable in the Cu-AHH complex compared to that in the CuAHH-NH2 (no COO−) complex. The impact of the COO− versus CONH2 can also been found in the literature from a comparison of Cu-AAH (Cterminus COO−) and AAH-NH2 (C-terminus CONH2). The AAH had a higher pKa of the His ring and a lower Cu affinity at pH 7.4 compared to AAH-NH2, in line with the secondary sphere effect of the H-bonds by COO−.32 A similar impact on Cu(II) coordination by a substitution of a carboxylate by an amide has been described recently for another peptide with a different Cu coordination.49 Indeed, compared to that of AHH-NH2, AHH has also a higher pKa of the His imidazole (Table 1), and the Cu-affinity is higher for AHH-NH2 (compared to AHH) from pH 4−11 (Figure S9), in line with the proposed H-bond. EPR and Dimerization or Oligomerization. Potentiometric studies proposed the presence of a minor bis-complex (CuH−1L2). A closer inspection of the EPR spectra obtained for Cu-AHH in Figure 3 indicated that there might be a partial interaction between some of the Cu(II) centers due to the presence of a broad featureless band underneath the classical spectrum for the 4N form. To investigate this effect, we performed a Cu titration to AHH and compared it to those of AH and AAH. Titrations using 0.3, 0.6, 0.9, 1.2, and 1.5 equiv of copper with respect to a 2 mM peptide solution at pH 7.2 were performed. At 0.3 equiv of Cu, the main species observed had the classical EPR of the 4N form, correlating with the UV−vis

Figure 8. Frozen-solution EPR of (a) AHH−COO, (b) AHH-NH2, (c) AAH, and (d) AH. Conditions: 2 mM peptide, pH 7.2 in MES buffer, 30% glycerol (EPR experimental parameters: microwave power, 0.1 mW; modulation amplitude, 5G; modulation frequency, 100 kHz; receiver gain, 30 dB).

Cyclic Voltammetry. The cyclic voltammograms (CV) of Cu bound to AH, AHH, and AAH are plotted in Figure 9. The Cu(II)-AAH complex (Figure 9d) shows an anodic process at E = 0.73 V vs Ag/AgCl (0.929 V vs NHE). Unlike for the parent Cu(II)-DAHK complex, the process is irreversible.29 This might be linked to the oxidation ability of the electrogenerated Cu(III) complex in the present case. The Cu(II)-AH complex (Figure 9a) presents a more complex electrochemical signature, although strongly reminiscent of what was previously described for the Cu(II)-GHK species.29 In this case a chemical− electrochemical−chemical (CEC) process is observed: the Cu(II)-AH complex is reduced at E = −0.64 V vs Ag/AgCl (−0.441 V vs NHE), leading to a Cu(I)-AH species that chemically evolves toward a Cu(I)-AH′ species that is oxidized at E = 0.0 V vs SCE (0.244 V vs NHE). The exact nature of the Cu(I)-AH′ species is not known, but a linearly bound Cu(I) might be proposed with the N-terminal amine and the imidazole group of the His as ligands. The CV trace of the Cu-AHH complex at pH 6.2 corresponds almost perfectly to the superimposition of the two formerly described CVs (AAH, AH), correlating with the ability of the Cu-AHH to be either in F


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a 3N (like AH) or 4N (like AAH) environment. When scanning toward the anodic potential (Figure 9c)), an irreversible oxidation peak is detected at E = 0.76 V vs Ag/ AgCl (0.959 V vs NHE) and when scanning toward the cathodic potential, an electrochemical−chemical−electrochemical (ECE) process is observed with the first cathodic peak at E = −0.57 V vs Ag/AgCl (−0.371 V vs NHE) and an anodic peak on the reverse scan at E = 0.26 V vs Ag/AgCl (0.459 V vs NHE). Note that this latter peak is not present when scanning toward the anodic potential from −0.1 V vs Ag/AgCl (0.299 V vs NHE). This proves that the redox process detected at E = 0.26 V vs Ag/AgCl mirrors the oxidation of the species resulting from the chemical evolution of the Cu(I)-AHH formed at the electrode. The main difference between the CuAH and Cu-AHH is the values of the reoxidation peaks detected at E = 0.0 V vs Ag/AgCl (AH-Cu) and E = 0.26 V vs Ag/AgCl (Cu-AHH). This may be linked to the formation of a linearly bound Cu(I) with imidazole groups of the two His in the latter case that is expected to be more stable than Cu(I) bonding by the N-terminal amine and the imidazole group of the His proposed in the former case due to a softer character of Nim vs NH2. Cu-Exchange Reaction Investigated by NMR. It has been reported by NMR that Cu(II) is in fast exchange between peptides in GHK (3N form) and very slow for DAHK (4N form), i.e. Cu(II) bound in the 3N form exchanges faster than the NMR time scale (ms or faster) and the 4N forms slower, i.e. seconds or slower at pH 7.4.29 This raised the question of whether the 3N and 4N species in AHH behave the same way. We first set up to verify if the pure 3N and 4N, i.e. AH and AAH, behave similarly to the canonical GHK and DAHK peptides. The experiments showed that the addition of 5% Cu induced a broadening of all signals in AH. Only the Ala CH3-

Figure 9. Cyclic voltammograms of (a) Cu-AH, (b and c) Cu-AHH, and (d) Cu-AAH complexes. [peptide] = 1.0 mM, [Cu] = 0.9 mM, [NaClO4] = 0.1 M, pH = 6.2 ± 0.1. Working electrode = glassy carbon, counter electrode = Pt. Scan rate 100 mV/sec. The arrow indicates the scanning direction.

Figure 10. 1H NMR of AH (top left), AHH (top right), AHH (bottom left), and AHH-NH2 (bottom right) without Cu (upper) and with 5% Cu (bottom). Conditions: 2.5 mM peptide, 0.125 mM Cu (when added) water suppression using excitation sculpting with gradients, pH 8.5, 100 mM phosphate buffer. G


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resonance was clearly detected. Whereas in AAH, the signals generally were not broadened (Figure 10). The intensity is slightly diminished, which is in line with a slow exchange, resulting in a 95% of the 1H NMR identical to the apo-AAH and 5% very strongly broadened due to the Cu(II) binding. Addition of 5% to AHH at pH 8.5 showed a strong broadening of the signals, nearly as much as in the AH. This indicates that the Cu(II) exchange between two AHH was rapid within the NMR time scale. pH 8.5 was chosen because at this pH, Cu(II) is bound predominantly in the 4N form in AHH (Figure 5). Likewise, for AHH-NH2, there was >99% of the 4N form at pH 8.5 is (see Figure 7). Even so, AHH-NH2 showed a fast Cu(II)-exchange, resembling that of AH rather than AAH. The broadening was less pronounced for AHH-NH2 compared to that for AHH but still closer to AH than AAH (Figure 10). These results indicate that despite Cu(II) being bound mainly to the 4N form in AHH or AHH-NH2 (>99%), the Cu exchange is still relatively fast. It is conceivable that the low populated but accessible 3N form is responsible for the fast exchange of Cu(II) observed in AHH and AHH-NH2.

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EXPERIMENTAL SECTION

All solutions were prepared with Milli-Q (18 MW) water. Stock solutions of peptides were prepared by dissolving the peptides in MiliQ, and concentrations were determined by titration followed by UV/ vis absorption spectroscopy as indicated in the Supporting Information. This experimental determination led to concentration values that are 10−20% off compared to those obtained using the molecular mass of the peptide and its counterions, thereby suggesting that counterion salts coprecipitate during peptide synthesis. Stock solutions were stored at −20 °C. The Cu(II) ion source was hydrated CuCl2 2H2O. The concentration of Cu(II) was determined by dissolving the Cu salt in Milli-Q water to prepare a 100 mM solution considering an extinction coefficient of ϵ = 12 Mol−1cm−1 Synthesis of Peptides. The AHH, AHH-NH2, AAH, and AH peptides were synthesized manually with standard 9-fluorenylmethoxycarbonyl (Fmoc) chemistry on a Fmoc-L-His(Trt)-Wang resin (0.63 mmol/g from Iris Biotech GMBH) through solid phase peptide synthesis protocols. HATU ((1-[bis(dimethylamino)methylene]-1H1,2,3-triazolo[4,5-b]pyridinium 3-oxid hexafluorophosphate) was used as the coupling reagent, DIPEA (N,N-diisopropylethylamine) as the base, and DMF (dimethylformamide) as the solvent. The coupling reactions were performed by using a 4-fold excess of amino acid, 3.8fold excess of HATU, and 6-fold excess of DIPEA in DMF under mixing for 30 min; Fmoc deprotection was carried out using 20% of 4methylpiperidine in DMF for 20 min. Coupling reactions were controlled by the Kaiser test and repeated if the test was positive. Crude peptides were cleaved and the side chain deprotected at the same time by treatment with 95% TFA, 2.5% TIPS, and 2.5% water for 2 h. The peptides were precipitated with cold ether from the cleaving solution, centrifuged, dissolved in a CH3CN/H2O (1/1), 0.1% TFA solution, filtered from the resin, and lyophilized. Purity of the peptides was assessed by NMR. UV−Vis. UV−vis absorption measurements in the pH range of ∼2−10 were performed in the absence of buffer. The peptide:Cu ratio was slightly over 1 to ensure total binding of Cu(II) (2 mM peptide and 1.8 mM Cu(II)). The spectra were recorded in the 200−800 nm spectral range using a dual beam spectrometer (Cary 5000 UV−vis− NIR) in a 1 cm path length quartz cuvette with a final sample volume of 1.5 mL. Titrations were made by adding 1 μL of 5 M NaOH directly to samples. Circular Dichroism. Circular dichroism spectra were recorded on a JASCO J-810 spectropolarimeter coupled to a Peltier Jasco PFD425S system for temperature control (20 °C). Measurements were done in the 800−250 nm range with a 1 cm path quartz cuvette with a sample final volume of 1.5 mL, 2 mM final concentration of peptide, and 1.8 mM of CuCl2. Titrations were made by adding 1 μL of 5 M NaOH (no buffer). EPR Spectroscopy. X-band EPR spectra (9.4 GHz) were obtained on a continuous-wave EMX-plus spectrometer (Bruker Biospin GmbH, Germany) equipped with a high sensitivity resonator (4119HS-W1, Bruker). The g factor was calibrated in the experimental conditions using the Bruker strong pitch (g = 2.0028). All samples were supplemented by 30% v/v glycerol to ensure homogeneous protein distributions and avoid water crystallization-induced phase separation. Then, samples were introduced to 4 mm outer diameter quartz tubes (Wilmad-Labglass) and freeze-quenched into liquid nitrogen prior to their introduction into the precooled cavity (100 K, achieved by continuous flow liquid nitrogen cryostat). The principal experimental parameter values were modulation amplitude 5 G; microwave power ca. 0.1 mW; conversion time and time constant were set up at ca. 310 and 80 ms, respectively; 1500 G were swept in 5 min; and 1−4 scans were accumulated to achieve reasonable signal-to-noise (S/N) ratio. Simulations based on experimental data were performed under Matlab environment using Easyspin Toolbox.50 NMR Spectroscopy. 1H experiments were recorded using a Bruker Avance III 400 MHz spectrometer. All spectra were calibrated with respect to the D2O signal (4.79 ppm). NMR spectra were collected at 298 K in 10% D2O/H2O, 100 mM phosphate buffer at pH 8.5 using the Watergate suppression technique.

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CONCLUSIONS In the work presented here, several spectroscopic techniques were used to provide structural details of the peptide AHH and the Cu(II)-AHH complex. The two main complex species 3N and 4N are pH dependent, with the 3N form dominating at around pH 4.5−6.5 and the 4N at pH 6.5−10. This pH dependence is explained by the number of amides involved in the Cu(II) coordination, one for the 3N form (N− of Ala1-His2 bond) and two for the 4N form (N− of Ala1-His2 and His2-His3 amide bonds). The equilibrium between the 3N and the 4N forms can be easily affected, as in the case of the replacement of the N-terminal carboxylate by an amide, in which the pKa of the 3N/4N equilibrium is shifted by one pH unit. This we attribute to the second sphere effect via a H-bond of COO− to H−N in the imidazole of His3 or in the amide bond. Also, the freezing of the sample induced a significant shift of the population of the forms. This indicates that the equilibrium can be modified by second (or further) sphere interactions, opening a possibility to control the coordination mode by external stimuli. AHH represents the general motif of XHH; thus, an equilibrium between the two forms is expected for the motif and would apply for all peptides and proteins with a nonburied N-terminal XHH sequence, considering that several proteins and peptides with such sequence are known. It is also expected that the identity of the first amino acid could impact the pKa value of the 3N/4N transition via second sphere (or further) interactions. Although the Cu(II)-AHH complex is mainly in the 4N form around the neutral pH and has the same type of coordination and very similar spectroscopic signatures (EPR, FTIR, Raman, UV−vis, and CD) as in the XXH motif (e.g., the canonical DAHK), there is a striking difference in terms of the Cu(II) exchange kinetics, as probed by NMR. It is suggested that the 3N form, although populated to a very low extent via the equilibrium, is nevertheless accessible and able to accelerate the Cu(II) exchange considerably. Thus, the motif AHH or XHH in general has the potential to have the thermodynamic stability of the well-known XZH motif (as found in HSA), but in contrast allows a fast exchange reaction (ms or faster) of Cu(II) with interesting biological consequences. H


Inorganic Chemistry

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FTIR and Raman. The FTIR spectra were recorded on a Vertex 70 spectrometer (Bruker, Karlsruhe, Germany) equipped with a Globar source and a MCT detector. A 3 μL drop of the sample was used to form a film on a diamond ATR-crystal (Harrick). The spectrum of the dried film was recorded with a resolution of 4 cm−1. At least 5 spectra of 128 scans each were averaged to get the spectrum of each sample. The Raman spectra were recorded on a Renishaw via a Raman microscope equipped with a CCD detector. A 514 nm laser (argon) was used to excite the sample (output power of 16.5 mW). The laser was focused with a 50× objective. Three microliters of the sample were deposited on a Si window. For each sample, several accumulations were averaged with an exposure time of 10 s. The spectral range for all the FTIR and Raman spectra is 1750−1200 cm−1. Potentiometry. Potentiometric titrations were carried out on a 907 Titrando Automatic Titrator (Metrohm) using a Biotrode combined glass electrode (Metrohm) calibrated daily by nitric acid titrations.51 Purified and lyophilized AHH-NH2 and AHH were dissolved in 4.0 mM HNO3/96 mM KNO3 to obtain stock solutions of ∼2.0 mM each. Peptide stock concentration calculation was performed by at least three consecutive measurements of the Cu(II)free peptide. Next, 1.5 mL samples of 1.0 mM peptides were prepared alone for determination of protonation constants and with various Cu(II) concentrations for measurement of metal binding constant. A total of 3 samples of peptide alone and 6 samples of 1:1, 1:2, and 1:3 metal to ligand molar ratios were prepared for each system, in duplicate for each ratio. Samples were titrated with 0.1 M standardized NaOH (carbon-dioxide-free). All experiments were performed under argon at 25 °C. The data were analyzed using the SUPERQUAD (NaOH stock and electrode calibration) and HYPERQUAD (stability constant determination).52,53

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REFERENCES

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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.inorgchem.7b01996. Details of Cu titration to the peptides, table of physicochemical properties, and additional information about EPR and IR spectrometry (PDF)

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Article

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. ORCID

Peter Faller: 0000-0001-8013-0806 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Dr. Vladimir Torbeev and Régis Boehringer (ISIS Strasbourg) for help with the peptide synthesis, and Dr. Marco Cecchini, Florian Blanc, Joel Montalvo-Acosta, and Simone Conti (ISIS Strasbourg) for helpful discussion. Dr. Bruno Vincent (UMR 7177, Strasbourg) and Amandine Conte-Daban (LCC, Toulouse) are acknowledged for help with NMR and electrochemistry measurements, respectively. We acknowledge financial support from the University of Strasbourg Institute for Advanced Study (USIAS), University of Strasbourg (IDEX program), the Frontier Research in Chemistry Foundation (Strasbourg), the REseau NAtional de Rpe interDisciplinaire (RENARD, Fédération RPE CNRS #3443), and the Narodowe Centrum Nauki (Poland), Grant 2016/23/B/ST5/02253. I


Article

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