Revisiting the significance of kinetic inertia in complex formation/decomplexation of metal–ATCUN peptide complexes

Abstract

Research on copper and nickel complexes formed by an amino terminal Cu(II) and Ni(II) binding (ATCUN) motif has greatly progressed in recent decades. These compounds are of considerable interest in bioinorganic chemistry, both as potential metallodrug candidates and as artificial metalloenzymes. Although the high stability of the Cu(II)– and Ni(II)–ATCUN complexes under physiologically relevant conditions is well established, the kinetic inertia associated with their formation has often been underestimated, particularly in the context of their catalytic applications. Here, we prepared ATCUN peptides (GGHWGKRG–Am; GGH–Pep) and investigated the stability of their Cu(II) and Ni(II)–ATCUN complexes in aqueous solutions under conditions enabling 1 : 1 metal-to-peptide complex formation at micromolar concentrations. Systematic pH titration revealed that the low basicity of the N-terminal amine of the peptide contributes to stabilizing the metal–ATCUN complex in aqueous solution. These findings highlight the need to account for kinetic inertia when evaluating ATCUN-like complexes under catalytically relevant conditions.

---

Introduction

The amino terminal Cu(II) and Ni(II) binding (ATCUN) motif is a metal binding site recurrent in proteins and peptides, and it has been studied for more than 40 years.^1–5 The canonical ATCUN sequence is H₂N–X₁–X₂–His, where X₁ and X₂ are arbitrary amino acids. The wide range of applications in chemistry and biology fields envisioned for ATCUN derivatives⁶ has strongly stimulated research on designed metal–ATCUN complexes.^3,7

Peptides containing an ATCUN motif are known to coordinate copper(II) with remarkably high affinity (ca. 10¹⁴ M⁻¹),^8,9 adopting a 4N square planar geometry.^5,10 Nickel(II) also binds to ATCUN, though with considerably weaker affinity (dissociation constant ≈150 nM, about 10⁵ times higher than that of Cu(II)).^11–13 At physiological pH, Cu(II) and Ni(II) coordination involves the N-terminal amine, the proximal δ-nitrogen of the histidine side chain, and both deprotonated amides of the peptide backbone between the N-terminal amine and histidine.^8,9 Therefore, the second residue in the ATCUN sequence cannot be Pro, since its secondary amine lacks a dissociable proton that would otherwise be replaced by the metal ion when it participates in the peptide bond.⁷

The ATCUN coordination can significantly influence the properties of the bound metal ion, particularly in terms of the redox activity and stability of the metal/peptide adduct.¹⁴ Strong metal coordination generally suppresses redox cycling,^7,14,15 while naturally occurring ATCUN complexes with redox-active metals can promote the generation of reactive oxygen species (ROS).¹⁴ This activity has been linked to oxidative damage to biomolecules, including DNA cleavage^2,14 and protein aggregation processes associated with neurodegenerative diseases such as Alzheimer's and Parkinson's.^16–18

ATCUN complexes also proved that they can act as catalysts in several redox and non-redox reactions, as recently reviewed by Faller⁷ and Moura.⁶ Just as an example, in recent decades, ATCUN motifs have been increasingly explored as catalysts for hydrogen evolution in photochemical systems.^19,20 For these reasons, ATCUN has been used as the metal binding site for the design of artificial catalytic metallopeptides and metalloproteins. In this context, we recently reported the design of a catalytically active Cu(II)/ATCUN artificial protein based on the SpyCatcher/SpyTag technology.

In the context of developing new Ni(II)-containing artificial metalloproteins based on the Spy construct, we decided to study in detail the thermodynamics and kinetics of the formation of Ni(II) complexes with a designed heptapeptide (GGHWAKR–NH₂, Table 1). The advantage of using such a designed oligopeptide lies in the possibility to tailor its sequence: the GGHW peptide contains, downstream the ATCUN triad, a tryptophan residue as a spectroscopic (absorption and emission) tag, while arginine and lysine were included to increase its solubility in aqueous media (Fig. 1). Together with this peptide we also studied the complexation of Ni(II) using a peptide recently reported by us which acts as a paradigmatic ATCUN ligand (AAHAWG–NH₂, Table 1).

Fig. 1 Representation of the single-chain GGH–Pep peptide studied in this work. Peptide is reported in its fully protonated form (panel A) and fully protonated and coordinated to the metal ion (M²⁺ = Cu(II) or Ni(II)).

Table 1 Peptide sequence

Short name	Sequence N → C
GGH–Pep	H2N–GGHWGKRG–Am
AAH–Pep	H2N–AAHAWG–Am

---

It is worth highlighting that, different from Cu(II), the kinetics of complexation of Ni(II) were examined in much less detail. Actually, the interest in Cu(II) regarding the onset of neurodegenerative diseases and the clarification of reaction events in the synaptic cleft has prompted several authors to study equilibria and intermediate formation in different Cu(II)/peptide systems, including ATCUN.^6,7,15,18,21 In this paper, we present potentiometric and spectrophotometric studies that demonstrate that the kinetic aspects in metal–ATCUN complexation play a crucial role, as metal complexation occurs on a timescale of minutes for both Cu(II) and Ni(II). More specifically, we shed light on the kinetic inertia of the ATCUN site towards Cu(II) and in particular Ni(II), an aspect that has not been clearly reported in previous studies on these systems that are available in the literature.

We believe that a deep understanding of metal–ATCUN complexes is essential for both elucidating their biological roles and harnessing their potential in biomedical²² and catalytic applications.^23,24 Stability and kinetics of formation of these complexes are therefore aspects of predominant importance in view of designing new catalytic artificial ATCUN proteins.

Results and discussion

Peptide protonation equilibria

The protonation constants of the peptide GGH–Pep (sequence in Table 1) were determined in aqueous solution by potentiometric titrations. Data are presented in Table 2 as protonation constants (log β) and acid dissociation constants (pK_a) and the corresponding species distribution diagrams are shown in the SI (Fig. S3A). GGH–Pep in its fully protonated form is a tetraprotic acid ([H₄L]⁴⁺). The pK_a values determined by potentiometric titrations are in agreement with the primary sequences of the peptides (Table 2). These dissociation processes correspond to the sequential deprotonation of the imidazole groups of His, the amino terminal NH₂, the ε-amino group of Lys and the guanidine group of Arg (Table 2). The pK_a values and the distribution diagram of GGH–Pep show that the deprotonation equilibria of the acidic groups are quite well separated. Surprisingly, the pK_a associated with the deprotonation of the guanidinium group of arginine is lower than the values widely reported in the literature for that group (11.3 vs. ca. 13.8, respectively).^25–27 Overall, with the exception of that of the arginine guanidinium group, the pK_a values of GGH–Pep are in full agreement with pK_a values reported for ATCUN motifs in the literature,^{4,5,7,28–30} indicating that the ATCUN binding site is not affected by the tail of the peptide.

Table 2 Proton dissociation constants (pK_a) of the fully protonated GGH–Pep peptide (H₄L⁴⁺), and logarithms of overall formation constants (log β) and acid dissociation constants (pK_a) of the copper(II) and nickel(II) complexes of GGH–Pep in aqueous solutions (T = 298.2 K, I = 0.1 M in KCl). Standard deviations are given in parentheses. L represents the completely deprotonated form of the peptide

Species	Log β	pKa	Dissociation site
[LH]^+	11.3(1)	11.3(1)	NH3^+ Arg
[LH2]^2+	20.39(9)	9.11(9)	NH3^+ Lys
[LH3]^3+	27.67(8)	7.28(8)	NH3^+ N-terminal
[LH4]^4+	33.58(7)	5.91(7)	NimH^+ His
[CuL]^2+	20.04(4)	8.8(2)
[CuLH−1]^+	11.2(2)
[NiL]^2+	13.14(2)	9.4(1)
[NiLH−1]^+	3.7(1)	10.0(1)	NH3^+ Lys
[NiLH−2]	−6.25(7)

---

Given the pK_a values, at physiological pH, GGH–Pep is deprotonated at the imidazole group and partially deprotonated at the amino terminal NH₂ group. As a consequence, at pH 7.4, GGH–Pep exists as [H₃L]³⁺ and [H₂L]²⁺ (50% each) and the peptide is positively charged under these conditions.

Protonation constants of the peptide AAH–Pep (sequence in Table 1) were previously determined by Perinelli et al.²⁹ Data are presented in Table S2 and the corresponding species distribution diagrams are shown in the SI (Fig. S3B). Given the pK_a values, at physiological pH, AAH–Pep exists as [H₂L]²⁺ and [HL]⁺ (50% each), with a deprotonated imidazole group and a partially deprotonated amino terminal NH₂ group.

Copper(II)/peptide complex formation

The formation of copper(II) complexes of GGH–Pep was studied by potentiometric titrations in the presence of slight excess of the ligand (Cu : L 1 : 1.2). A careful analysis of the potentiometric titration curves of Cu(II)/GGH–Pep showed the systematic presence of discontinuity around pH 4 in the form of a slight jump to higher e.m.f. values (lower pH, Fig. S4). In order to exclude the possibility that this discontinuity was due to experimental or instrumental biases, potentiometric curves were recorded using different glass electrodes, and freshly prepared solutions of peptides. Moreover, different peptide stock solutions were prepared using solid batches of the compound originating from different syntheses. As a result, the discontinuity was present in all potentiometric curves recorded.

We put forward the hypothesis that this discontinuity could be due to kinetic inertia in the formation of copper(II) species in that pH region. Actually, until the appearance of this discontinuity the e.m.f. values of the titration curves in the presence of copper(II) were lower than those in its absence. Therefore expectedly, the displacement of protons on the peptide upon coordination of copper(II) occurs in that pH range, however, the species formed may not be the most thermodynamically stable under those conditions. Under this perspective, the discontinuity may arise from a fast rearrangement of the coordination environment of copper(II) to form different species.

To prove that kinetic aspects are relevant to the appearance of this discontinuity and that equilibration time after each addition of titrant aliquot is a significant variable, we collected the potentiometric curves while changing the equilibration time after each aliquot addition. Fig. S4 (GGH–Pep) and Fig. S5 (AAH–Pep) report the curve using 60 s to 700 s as the equilibration time. Using this experimental setup in data collection, the discontinuity was systematically absent in the collected curves.

We have used potentiometric curves collected with long equilibration times to study the speciation of the Cu(II)–GGH–Pep system, which is reported in Table 2. The formation of solely mononuclear complexes was observed, and a representative distribution diagram for the Cu(II)–GGH–Pep complex is reported in Fig. 2A.

Fig. 2 Representative species distribution diagram for complex-formation of GGH–Pep with Cu(II), at T = 298.15 K and I = 0.1 M (KCl). (Cu(II) : GGH–Pep = 1 : 1.2, [GGH–Pep] = 0.3 mM, [Cu(II)] = 0.25 mM).

The peptide forms two complex species that differ in their protonation states, but all corresponding to the 1 : 1 Cu(II) : peptide stoichiometry as stated above.

Between a pH range of 4.5–9.0, the predominant species is [CuL]²⁺, and it reaches almost 100% total copper between pH 5 and pH 7.4 (Fig. 2). This species is consistent with Cu(II) bound in the 4N equatorial coordination (NH₂, 2× N⁻, N_im) at the ATCUN site.²⁸ The observed maximum absorption wavelength at physiological pH (Fig. 3A) at λ_max = 527 nm is in agreement with the expected λ_max value of 536 nm taking into account the Billo's parameters.^31,32 Moreover, the d–d band at λ_max = 527 nm (80 M⁻¹ cm⁻¹) is characteristic of the 4N complex.^7,33–37 Analogously, the CD spectra of the same UV-Vis samples confirm the coordination number and geometry (Fig. S6). A positive band is observed at 504 nm and a negative one at 602 nm (Fig. S6).²⁹ These spectroscopic features confirm that Cu(II) binds to GGH–Pep at the ATCUN site with a 1 : 1 metal : peptide stoichiometry. It is worth noting that the significantly lower intensity of the negative peak of Cu–GGH–Pep compared to other Cu(II)–ATCUN peptides reported in the literature⁵ is due to the fact that GGH–Pep bears two non-chiral residues that are involved in Cu(II) coordination,^38–40 However, the distinctive CD spectrum profile of a Cu(II)–ATCUN complex upon the addition of 1 eq. of Cu(II) has been observed.

Fig. 3 UV-vis absorption spectra for the titration of GGH–Pep solution with up to 1.6 eq. of Cu(II) (A). Absorbance at 527 plotted as a function of Cu(II) eq. (B). (GGH–Pep 0.3 mM, 50 mM HEPES solution at pH 7.4.) Fluorescence normalized (C) and fluorescence plotted at 460 nm registered after 1 min of equilibration time (D). Apo peptide is depicted in black.

At physiological pH, copper(II)–peptide complex formation is almost immediate (less than 30 s after metal addition) (Fig. S7). This is in complete agreement with the previous literature where the binding reaction between Cu(II) and GGH tripeptide was determined to take almost one second to be complete.^40,41

However, this experimental pH condition is widely beyond the value of the observed kinetic inertia for Cu(II) complexation (around pH 4). We tried to further investigate the kinetics of complex equilibria by UV-Vis, working just before the spike (pH 3.75), where the spike was observed (pH 4.0) and just after (pH 4.25). Nevertheless, under our experimental conditions we were unable to observe complex formation. As reported in Fig. S8, the characteristic band at 527 nm starts to be observed above pH 4.2; making it nearly impossible to monitor the Cu(II)–ATCUN complex formation by UV-Vis in proximity to the observed spike.

Complexation at pH 7.4 is well beyond the pH range affected by the Cu(II) complex kinetic inertia. In particular, the speciation close to physiological pH (pH 7.4) foresees that [CuL]²⁺ is the almost unique species present in solution. Both the UV-visible and CD spectra collected at this pH are in full agreement with the formation of a 4N square planar geometry complex of Cu(II), in which the donor set consist of (NH₂, 2× N_am, N_im) groups typically reported for the ATCUN motif.^4,5,42

Copper(II) coordination by GGH–Pep at pH 7.4 has also been investigated by spectrofluorimetric titration, taking advantage of the presence of a Trp residue in proximity of the binding site. The Trp emission at 350 nm is completely quenched upon the addition of 1 eq. of copper(II) (Fig. 3C and D).

The Cu(II) binding affinity of the ATCUN site can be determined based on the potentiometric model using Hyss2009,⁴³ which yielded a log K_app value of 13.8 in Hepes 50 mM pH 7.4. This log K value is fully consistent with those expected for the binding of Cu(II) at an ATCUN site in oligopeptides.^4,5,7,29 On this ground, the range of log K_app values for ATCUN sites is in the 12–14 range, the lowest value measured for the GGH–OH³⁵ (ATCUN tripeptide, log K = 12.2).

The other species observed for the Cu(II)–GGH–Pep complex is [Cu(LH₋₁)]⁺; which starts to form above pH 8.0, becoming the most abundant species above pH 9.0, and reaching its maximum at pH 10 (almost 100% total copper). The deprotonation step that leads to [Cu(LH₋₁)]⁺ from [CuL]²⁺ likely involves only the deprotonation of the lysine residue, which does not impact copper coordination. The arginine side chain would appear at higher pH and it would lead to [Cu(LH₋₂)]; however, under the conditions reported here it was not observed. Copper(II) coordination does not change with pH, as confirmed also by UV-Vis spectra collected up to pH 11 (Fig. S8).

Having a reliable speciation of the Cu(II)–GGH–Pep system at hand, we tried to analyse the potentiometric curves collected at shorter equilibration times (i.e. those with discontinuity). As shown in Fig. S9, the small discontinuity seems in some way negligible, however its effect on the pK_a determination is not so negligible. As reported in Table S3, treating the potentiometric curves showing the spike induced by the competition between the deprotonation process and the Cu(II) complexation equilibrium would lead to overestimation of approximately 0.3 logarithmic units in the Cu(II)–ATCUN dissociation constants.

Overall, our results are in partial agreement with those reported in the seminal paper by Bal and coworkers,⁴¹ who have examined the microscopic processes of recruitment by Cu(II) on a short time scale. In that work, a 2N coordination occurred first, followed by the formation of the expected 4N ATCUN complex. Our results suggest the possibility that a second structural rearrangement, from 4N to a different 4N conformation may occur on longer time scale, in particular minutes (corresponding to that of potentiometric titrations).

Nickel(II)/peptide complex formation

The mild kinetic inertia observed during Cu(II)–ATCUN complex formation becomes far more pronounced with Ni(II).

This effect manifests as a distinct spike in the potentiometric titration curves around neutral pH, which cannot be neglected during data analysis.

As previously reported for Cu(II), the formation of Ni(II) complexes with GGH–Pep was investigated by potentiometric titrations in the presence of a slight excess of the ligand (Cu : L 1 : 1.2). The same experimental approach to assess the reasons behind the titration spike in Cu(II) studies were widely applied here for Ni(II), including testing different electrodes and peptide batches. However, these measures did not resolve the problem. The kinetic inertia associated with ATCUN–metal complexation was only overcome by modifying the titration protocol (Fig. S10). The corresponding parameters are summarized in Table S1. The most effective protocol involved increasing the minimum equilibration time after each addition from 10 s (Method #1) to 120 s (Method #3), ensuring that each new measurement was not biased by incomplete equilibration from the preceding complexation step.

This effect is clearly illustrated in the potentiometric curve obtained by Method #2 (Fig. S10, black trace). When insufficient equilibration time was allowed, the successive titration point is registered at a lower potential than its true value. Such deviations complicate, or even prevent, assignment of the observed event to either a genuine deprotonation process or a carryover from the previous process of deprotonation.

Herein, the apparent “negative spike” arises because the pH increase induces peptide deprotonation and hence a proton release, temporarily altering the measured f.e.m. Under normal circumstances, this equilibrium is rapidly established. However, Ni(II) binding to the ATCUN motif requires significantly longer to achieve full deprotonation and coordination compared with Cu(II). The optimized protocol (Method #3), employing equilibration times of 300–900 s after each KOH addition, provided the best compromise between measurement reliability, experiment duration, and minimization of solvent evaporation. This method yielded stable potentials and reproducible complexation data.

A similar kinetic inertia was observed for Ni(II) binding to AAH–Pep (Fig. S11). Once again, only the implementation of Method #3 produced reliable potentiometric measurements. For completeness, the formation constants of Ni(II)–AAH–Pep complexes are reported in the SI (Table S2).

Having established a reliable potentiometric method to determine Ni(II) speciation in solution, we were able to characterize its complexation with GGH–Pep. This peptide forms three distinct Ni(II) complex species, all with a 1 : 1 Ni(II) : peptide stoichiometry (Fig. 4). The detailed speciation is reported in Table 2, with the species reported differing only in their protonation states. Among them, [NiL]²⁺ is the predominant species in the physiological pH range (7.0–8.5), accounting for ca. 99% of the total nickel at pH 7.1 and ca. 98% at pH 7.4.

Fig. 4 Representative species distribution diagram for complex-formation of GGH–Pep with Ni(II), at T = 298.15 K and I = 0.1 M (KCl) (Ni(II) : GGH–Pep = 1 : 1.2, [GGH–Pep] = 0.3 mM, [Ni(II)] = 0.25 mM).

The coordination mode at pH 7.4 was investigated by UV–Vis spectroscopy, which showed an absorption maximum at 425 nm (Fig. 5A and B). This is consistent with the low-spin Ni(II)–ATCUN complex adopting a square-planar 4N coordination geometry.⁴⁴ CD spectroscopy further supported this assignment, displaying a positive d–d band at λ_max = 417 nm and a negative band at λ_max = 490 nm (Fig. S13). These spectroscopic features confirm that Ni(II) binds to GGH–Pep at the ATCUN motif in a 1 : 1 stoichiometry. As in the case of Cu(II) ([CuL]²⁺), [NiL]²⁺ is coordinated through the imidazole nitrogen of histidine and the deprotonated amino-terminal nitrogen (Fig. 6).

Fig. 5 UV-vis absorption spectra for the titration of GGH–Pep solution with up to 1.6 eq. of Ni(II) (A). Absorbance at 425 plotted as a function of Ni(II) equiv. (B). (GGH–Pep 0.3 mM, 50 mM HEPES solution at pH 7.4.) Fluorescence normalized overnight (C) and fluorescence plotted at 630 nm after 1 min (light blue), 5 min (magenta), 10 min (blue), 15 min (teal) of equilibration time and after overnight incubation (purple) (D). Apo peptide is depicted in black. Dashed lines have been added in panels B and D to determine the titration endpoint for the longest equilibration times used.

Fig. 6 Representation of deprotonation and complexation equilibrium which leads to [ML]²⁺ coordination mode to form the [ML]²⁺ complex with GGH–Pep. The sphere in pale green represents the metal ion. Representation of proposed nickel(II) coordination modes has been obtained using HyperChem.

The pH-dependent formation of Ni(II)–GGH–Pep complexes was also monitored by UV–Vis spectroscopy. The onset of complex formation was observed starting at ca. pH 4.5, with the overall complexation pK_a determined as 6.15(6) (Fig. S13).

Notably, the kinetic inertia in potentiometric titrations occurs in the same pH range (around pH 6.5), underscoring the importance of complementary spectroscopic analysis.

Herein, to further probe this effect, UV–Vis kinetic experiments were performed just before the spike (at pH 6.25), at pH 6.5 (spike region), and just post spike (at pH 6.7) (Fig. S14). At pH 6.25, Ni(II) remained essentially unbound, while beginning from pH 6.5 the complexation begins, as evidenced by a gradual shift of the absorption band from 393 nm (free Ni(II)) to 420 nm (Ni(II) in the low spin, 4N square planar geometry). In contrast, outside this narrow pH region, conversion from free Ni(II) to the Ni(II)–ATCUN complex occurred rapidly, with a single band at 420 nm observed throughout the titration (Fig. S14).

Finally, the kinetics of the Ni(II)–GGH–Pep complex formation were examined at pH 7.4 using both UV–Vis and circular dichroism spectroscopy, at a metal–peptide molar ratio of 1 : 1.2 (Fig. S15). These experiments confirm that under near-physiological conditions, Ni(II) rapidly achieves the square-planar 4N coordination characteristic of the ATCUN motif.

In contrast to Cu(II), the nickel(II)–peptide complex formation is not immediate even working in the millimolar range concentration, as highlighted by potentiometric studies. In fact, the kinetic inertia takes place at around pH 6.5 affecting the ATCUN coordination. We observe that complex formation required at least 10 minutes. Furthermore, complexation time increases as the concentration decreases, indicating that the process is governed by the reagent concentration. Herein, fluorescence experiments will require longer equilibration time, even overnight incubation if working in a micro- or nano-molar concentration range.

A longer equilibration time (overnight) was allowed for the Ni(II)–ATCUN complex (Fig. 5C and D). If, on the one hand, working at millimolar concentrations as in the UV-Vis experiments, the incubation time does not seem to significantly impact the complexation kinetics, on the other hand, the equilibration time becomes crucial in the spectrofluorimetric experiments.

Fluorimetric titrations are performed 100-times more dilution than UV-Vis titration. The titration at pH 7.4 was carried out by allowing 0, 5, 10 or 15 minutes of equilibration after each addition, or working in batch mode and performing the measures after overnight incubation (Fig. 5C and D). The fluorescence Trp emission at 350 nm was completely quenched upon the addition of 1 equivalent of nickel(II) after overnight incubation (Fig. 5D). Working in the micromolar concentration range, metal complex formation is incomplete if not enough equilibration time is allowed. In fact, when waiting 15 minutes between titration additions, complete Trp quenching was observed at 1.5 equivalents, suggesting an incorrect metal : peptide stoichiometry.

The Ni(II)–ATCUN coordination is not affected by increasing the pH above pH 8.0, at which point the formation of [NiLH₋₁]⁺ begins. This species becomes the most abundant species above pH 9.6, reaching 58.2% of the total Ni content. This deprotonation step involves solely the deprotonation of the lysine residue. At higher pH, a successive deprotonation of the Arg residue forms [NiLH₋₂] which becomes the prevalent species above pH 10.5 (Fig. 4). These two last deprotonations do not impact nickel coordination, as confirmed also by UV-Vis (Fig. S13) where the maximum absorbance does not change.

Having a reliable speciation of the Ni(II)–GGH–Pep system at hand, we tried to analyse the potentiometric curves recorded at shorter equilibration times (i.e. those with discontinuity). It is clear that, if the kinetic inertia of Ni(II)–ATCUN complex formation around pH 6.5 is overlooked, an underestimation (up to 0.5 logarithmic units) of the pK_a associated of metal ion coordination occurs. This is the most impactful with Ni(II) (Fig. S16 and Table S3), where the spike observed is important and Ni(II) coordination has already started.

Finally, data analysis based on the potentiometric model with Hyss2009⁴³ yielded a log K_app value of 7.18 in Hepes 50 mM pH 7.4. This log K value is absolutely consistent with those expected for the binding of Ni(II) at an ATCUN site.^2,45

Conclusions

In this work, we examined the kinetic inertia of Ni(II) and Cu(II) complexation in ATCUN peptides. Metal ion coordination was studied using a variety of analytical techniques. Potentiometric titration revealed that, although complexation is thermodynamically favored by the high affinity constants, it is kinetically hindered by proton release and by reorganization of the metal-binding site. Given sufficient time, however, the system undergoes a structural rearrangement, and the potentiometric data demonstrate how the ATCUN motif modulates the acidity of neighboring amino acid residues.

The potentiometric studies highlight that the coordination of Ni(II) by ATCUN peptides requires rearrangements of nitrogen ligands in order to form the [ML]²⁺ complex with GGH–Pep (Fig. 6) or the [MLH₋₂] complex with AAH–Pep. In the case of Ni(II), the initial binding involves a transient 2N coordination mode (NH₂, N_im, 2 × O_water), which subsequently evolves into the final 4N coordination mode (NH₂, 2 × N⁻, N_im).⁴¹ Furthermore, as shown in Fig. S14, the Ni(II) coordination into an ATCUN motif is triggered by a transient form ascribable to the 2N coordination mode around pH 6.5–6.75, which evolves to 4N. In contrast, at a higher pH value (i.e. pH 7.4), Ni(II) complexation to the ATCUN site is quicker than the time scale used in our experiments. Taking together, these data suggest a change in the coordination mode from 2N to 4N for the Ni(II)–ATCUN complex, on a timescale much longer than that previously observed by Bal and co-workers⁴¹ for Cu(II). Herein, this is an aspect that should be considered in the future when designing new ATCUN peptides.

In contrast, evaluating the lower potentiometric inertia observed in the presence of Cu(II), together with the absence of UV-Vis spectral changes near the potentiometric spike, we cannot propose a change in coordination mode (2N → 4N). Rather, ATCUN peptides GGH–Pep and AAH–Pep likely undergo a structural rearrangement that does not affect the 4N ATCUN coordination motif.

Therefore, the kinetic inertia in metal(II)–ATCUN complex formation with GGH–Pep and AAH–Pep possibly could be observed and taken into account with other peptides, on a non-negligible time scale even at millimolar concentrations.

Analysis of stability constants allowed us to fully describe the speciation and complex formation at different pH values, revealing that the identity of the transition metal plays a non-negligible role in peptide reorganization, particularly in the kinetics of the rearrangement process. The N-terminal amine and the histidine imidazole group serve as anchoring sites for Cu(II) and Ni(II), initiating coordination.^46,47 This anchoring induces deprotonation of the additional donor groups, thereby modulating their acidity.

Particular attention was paid to peptide behavior at pH 7.4, since this is the physiological pH at which catalytic assays involving ATCUN peptides are typically performed. The kinetics of Ni(II) complex formation has shed light on the importance of equilibration time, especially at catalytic concentrations, suggesting that incomplete equilibration may influence the evaluation of new artificial metalloenzymes. This is especially critical at lower pH values, where the equilibrium between transient 2N and final 4N coordination modes is more sensitive to proton concentration, even at relatively high peptide levels.

Although the ATCUN site is well known for its high-affinity coordination of Cu(II) and Ni(II), the intrinsic differences between these metal ions play a fundamental role in their kinetic complexation. Ultimately, the strong chelating ability of the ATCUN motif is primarily dictated by the acidity of the amino groups in the two N-terminal residues. However, one must be cautious not to overinterpret the high affinity constants without considering the kinetic and structural reorganizations required for complex formation.

Author contributions

All authors have given approval to the final version of the manuscript.

Conflicts of interest

There are no conflicts to declare.

Data availability

Supplementary information (SI) is available: experimental procedures, LC-MS chromatograms of peptides, protonation diagram of PepGGH and PepAAH peptides, potentiometric titration curves and dissociation constant for their Cu(II) and Ni(II) complexes, UV-Vis, fluorescence and circular dichroism spectra of PepGGH and PepAAH with Cu(II) or Ni(II). See DOI: https://doi.org/10.1039/d5dt02014j.

Acknowledgements

Dr Sabrina Capodaglio is gratefully acknowledged for her support in the setup of the instrumental methodologies for peptide synthesis and characterization. This work has been supported by project ART-2-HYDROGEN “Artificial enzymes for the photocatalytic production of hydrogen in photosynthetic bacteria” funded under the National Recovery and Resilience Plan (NRRP), Mission 2 Component 2 Investment 3.5 - Call for tender No. 4 of March 23, 2022 of the Italian Ministry of Ecologic Transition funded by the European Union – NextGenerationEU. Project code RSH2A_000009, Concession Decree 445 of December 29, 2022 adopted by the Italian Ministry of Environment and Energy Security, CUP F97G22000270006. M. T. also thanks the project of national interest (PRIN) 2022 “Bioinspired systems for ROS regulation: metalloporphyrinoids in neurodegeneration and artificial biocatalysis” prot. 2022RCRWE5 – Italian Ministry of University and Research (MUR), funded by the European Union – NextGenerationEU. This work has benefited from the equipment and framework of the COMP-HUB and COMP-R Initiatives, funded by the ‘Departments of Excellence’ program of the Italian Ministry for University and Research (MIUR, 2018–2022 and MUR, 2023–2027).

References

1. T. Peters, Biochim. Biophys. Acta, 1960, 39, 546–547.
2. C. Harford and B. Sarkar, Acc. Chem. Res., 1997, 30, 123–130.
3. K. P. Neupane, A. R. Aldous and J. A. Kritzer, Inorg. Chem., 2013, 52, 2729–2735.
4. T. Miyamoto, S. Kamino, A. Odani, M. Hiromura and S. Enomoto, Chem. Lett., 2013, 42, 1099–1101.
5. T. Miyamoto, Y. Fukino, S. Kamino, M. Ueda and S. Enomoto, Dalton Trans., 2016, 45, 9436–9445.
6. B. K. Maiti, N. Govil, T. Kundu and J. J. G. Moura, iScience, 2020, 23, 101792.
7. P. Gonzalez, K. Bossak, E. Stefaniak, C. Hureau, L. Raibaut, W. Bal and P. Faller, Chem. – Eur. J., 2018, 24, 8029–8041.
8. M. Mital, N. E. Wezynfeld, T. Frączyk, M. Z. Wiloch, U. E. Wawrzyniak, A. Bonna, C. Tumpach, K. J. Barnham, C. L. Haigh, W. Bal and S. C. Drew, Angew. Chem., Int. Ed., 2015, 54, 10460–10464.
9. J. D. Barritt and J. H. Viles, J. Biol. Chem., 2015, 290, 27791–27802.
10. T. Peters and F. A. Blumenstock, J. Biol. Chem., 1967, 242, 1574–1578.
11. J. D. Glennon and B. Sarkar, Biochem. J., 1982, 203, 15–23.
12. W. Bal, J. Christodoulou, P. J. Sadler and A. Tucker, J. Inorg. Biochem., 1998, 70, 33–39.
13. M. Rózga, M. Sokołowska, A. M. Protas, W. Bal and J. B. I. Chem, JBIC, J. Biol. Inorg. Chem., 2007, 12, 913–917.
14. J. Heinrich, K. Bossak-Ahmad, M. Riisom, H. H. Haeri, T. R. Steel, V. Hergl, A. Langhans, C. Schattschneider, J. Barrera, S. M. F. Jamieson, M. Stein, D. Hinderberger, C. G. Hartinger, W. Bal and N. Kulak, Chem. – Eur. J., 2021, 27, 18093–18102.
15. V. Borghesani, B. Alies and C. Hureau, Eur. J. Inorg. Chem., 2018, 2018, 7–15.
16. B. Jafari, M. Muthuvel and G. G. Botte, J. Electroanal. Chem., 2021, 895, 115547.
17. A. Sedjahtera, L. Gunawan, L. Bray, L. W. Hung, J. Parsons, N. Okamura, V. L. Villemagne, K. Yanai, X. M. Liu, J. Chan, A. I. Bush, D. I. Finkelstein, K. J. Barnham, R. A. Cherny and P. A. Adlard, Metallomics, 2018, 10, 1339–1347.
18. M. Lefèvre, K. P. Malikidogo, C. Esmieu and C. Hureau, Molecules, 2022, 27, 7903.
19. S. Chakraborty, E. H. Edwards, B. Kandemir and K. L. Bren, Inorg. Chem., 2019, 58, 16402–16410.
20. B. Kandemir, L. Kubie, Y. Guo, B. Sheldon and K. L. Bren, Inorg. Chem., 2016, 55, 1355–1357.
21. P. H. Nguyen, A. Ramamoorthy, B. R. Sahoo, J. Zheng, P. Faller, J. E. Straub, L. Dominguez, J.-E. Shea, N. V. Dokholyan, A. De Simone, B. Ma, R. Nussinov, S. Najafi, S. T. Ngo, A. Loquet, M. Chiricotto, P. Ganguly, J. McCarty, M. S. Li, C. Hall, Y. Wang, Y. Miller, S. Melchionna, B. Habenstein, S. Timr, J. Chen, B. Hnath, B. Strodel, R. Kayed, S. Lesné, G. Wei, F. Sterpone, A. J. Doig and P. Derreumaux, Chem. Rev., 2021, 121, 2545–2647.
22. E. Kimoto, H. Tanaka, J. Gyotoku, F. Morishige and L. Pauling, Cancer Res., 1983, 43, 824–828.
23. S. Chakraborty, E. H. Edwards, B. Kandemir and K. L. Bren, Inorg. Chem., 2019, 58, 16402–16410.
24. B. Kandemir, L. Kubie, Y. Guo, B. Sheldon and K. L. Bren, Inorg. Chem., 2016, 55, 1355–1357.
25. C. A. Fitch, G. Platzer, M. Okon, B. E. Garcia-Moreno and L. P. McIntosh, Protein Sci., 2015, 24, 752–761.
26. N. F. Hall and M. R. Sprinkle, J. Am. Chem. Soc., 1932, 54, 3469–3485.
27. S. J. Angyal and W. K. Warburton, J. Chem. Soc., 1951, 2492–2494.
28. K. Szarszoń, A. Mikołajczyk, M. Grelich-Mucha, R. Wieczorek, A. Matera-Witkiewicz, J. Olesiak-Bańska, M. Rowińska-Żyrek and J. Wątły, J. Inorg. Biochem., 2024, 253, 112476.
29. M. Perinelli, R. Guerrini, V. Albanese, N. Marchetti, D. Bellotti, S. Gentili, M. Tegoni and M. Remelli, J. Inorg. Biochem., 2020, 205, 110980.
30. J. Watly, E. Simonovsky, R. Wieczorek, N. Barbosa, Y. Miller and H. Kozlowski, Inorg. Chem., 2014, 53, 6675–6683.
31. E. J. Billo, Inorg. Nucl. Chem. Lett., 1974, 10, 613–617.
32. E. Prenesti, P. G. Daniele, M. Prencipe and G. Ostacoli, Polyhedron, 1999, 18, 3233–3241.
33. E. Farkas and T. Kiss, Polyhedron, 1989, 8, 2463–2467.
34. R. W. Hay, M. M. Hassan and C. You-Quan, J. Inorg. Biochem., 1993, 52, 17–25.
35. K. Bossak-Ahmad, T. Frączyk, W. Bal and S. C. Drew, ChemBioChem, 2020, 21, 331–334.
36. H. Sigel and R. B. Martin, Chem. Rev., 1982, 82, 385–426.
37. H. Kozłowski, W. Bal, M. Dyba and T. Kowalik-Jankowska, Coord. Chem. Rev., 1999, 184, 319–346.
38. B. Gyurcsik, I. Vosekalna and E. Larsen, J. Inorg. Biochem., 2001, 85, 89–98.
39. N. I. Jakab, B. Gyurcsik, T. Körtvélyesi, I. Vosekalna, J. Jensen and E. Larsen, J. Inorg. Biochem., 2007, 101, 1376–1385.
40. R. Kotuniak, P. Szczerba, D. Sudzik, M. J. F. Strampraad, P.-L. Hagedoorn and W. Bal, Dalton Trans., 2022, 51, 17553–17557.
41. R. Kotuniak, M. J. F. Strampraad, K. Bossak-Ahmad, U. E. Wawrzyniak, I. Ufnalska, P.-L. Hagedoorn and W. Bal, Angew. Chem., Int. Ed., 2020, 59, 11234–11239.
42. J. Wątły, K. Szarszoń, A. Mikołajczyk, M. Grelich-Mucha, A. Matera-Witkiewicz, J. Olesiak-Bańska and M. Rowińska-Żyrek, Inorg. Chem., 2023, 62, 19786–19794.
43. L. Alderighi, P. Gans, A. Ienco, D. Peters, A. Sabatini and A. Vacca, Coord. Chem. Rev., 1999, 184, 311–318.
44. J. Dolovich, S. L. Evans and E. Nieboer, Br. J. Ind. Med., 1984, 41, 51.
45. Y. Jin, M. A. Lewis, N. H. Gokhale, E. C. Long and J. A. Cowan, J. Am. Chem. Soc., 2007, 129, 8353–8361.
46. I. Sóvágó, C. Kállay and K. Várnagy, Coord. Chem. Rev., 2012, 256, 2225–2233.
47. I. Sóvágó, K. Várnagy, N. Lihi and Á. Grenács, Coord. Chem. Rev., 2016, 327–328, 43–54.

---