Thermodynamic and spectroscopic study of Cu(II) and Zn(II) complexes with the (148–156) peptide fragment of C4YJH2, a putative metal transporter of Candida albicans

Denise Bellotti ab, Cinzia Tocchio a, Remo Guerrini a, Magdalena Rowińska-Żyrek b and Maurizio Remelli *a
aDepartment of Chemical and Pharmaceutical Sciences, University of Ferrara, Via L. Borsari 46, 44121, Ferrara, Italy. E-mail: rmm@unife.it
bFaculty of Chemistry, University of Wrocław, F. Joliot-Curie 14, 50-383, Wrocław, Poland

Received 9th October 2019 , Accepted 8th November 2019

First published on 8th November 2019


Candida albicans is a widespread human pathogen which can infect humans at different levels. Like the majority of microorganisms, it needs transition metals as micronutrients for its subsistence. In order to acquire these nutrients from the host, C. albicans employs various strategies, also involving chelating proteins specifically expressed to sequester metals from the environment. A histidine-rich protein sequence identified in the C. albicans genome, named C4YJH2, has been recently studied for its putative role in Zn(II) transport. Two outer membrane major histidine-rich clusters of C4YJH2, namely the domains 131–148 (FHEHGHSHSHGSGGGGGG) and 157–165 (SHSHSHSHS), have been confirmed as strong binding sites for the Cu(II) and Zn(II) ions. Nevertheless, the 9-residue “linker” sequence 148–156 (GSDHSGDSK) between the two His-rich fragments of C4YJH2, containing an additional His residue, can also contribute to metal binding. In the present work, the protected peptide Ac-GSDHSGDSK-NH2 and some analogues (Ac-GSDHSGASK-NH2, Ac-GADHAGDAK-NH2, Ac-GSDH-NH2, and Ac-HSGD-NH2) have been synthesized and their metal binding properties have been studied in detail. The thermodynamics of complex-formation equilibria of the above reported ligands with Cu(II) and Zn(II) ions have been studied by potentiometry in a wide pH range and the stoichiometry of the formed species has been confirmed by mass spectrometry; the most likely solution structures of the metal complexes are also discussed on the basis of NMR, UV-vis, circular dichroism (CD) and EPR data. The results show the importance of Asp7 in the stabilization of Zn(II) complexes and suggest a significant role of the (quite abundant) Ser residues in the task of metal uptake and regulation.



Significance to metallomics

The mechanism of metal acquisition at the host/pathogen interface can be a fertile ground for the design of new antibiotics. The present paper deals with the characterization of Cu(II) and Zn(II) binding to the putative metal transporter C4JYH2, found in the genome of Candida albicans. The thermodynamic investigation of the formed metal complexes and their spectroscopic characterization in solution opens the way to the discovery of possible new pharmacological targets based on metal ions.

Introduction

The constant increase of the drug-resistance phenomenon against antibiotics and antifungal agents, with a high incidence especially in hospitals, represents a heavy burden for the healthcare systems: a deep reconsideration of commonly used therapies and the availability of innovative drugs are definitely required. Candida albicans is one of the most diffused opportunistic pathogenic yeasts that can affect the human organism. The pathological conditions associated to C. albicans frequently involve skin, oral cavity, oesophageal, vaginal and gastrointestinal tract infections ranging from superficial disorders to invasive systemic diseases that can eventually involve the vascular system and various organs anywhere in the body. Infections caused by C. albicans represent a serious threat for the subsistence of immunocompromised or debilitated individuals and it is by far the major cause of severe mucosal infections (invasive candidiasis) with an elevated mortality rate.1–5 Furthermore, the phenomenon of drug resistant mycoses has become a serious clinical and financial burden on the world healthcare systems: in the long run it may become a critical and dangerous threat comparable to the increase of “superbug” infections.6,7 As a consequence, the need of innovative antifungal treatments with high selectivity, specificity and effectiveness is undeniable.

Since the eukaryotic cells of human and fungal pathogens share several biological processes, in order to identify new pharmacological targets, one possible strategy is to focus on the differences in the metabolic pathways of these species. One difference concerns the mechanism of metal uptake and transport into the fungal cell. In fact, several lines of evidence confirm that one critical aspect of fungal infection and survival is the ability of the pathogenic microorganism to assimilate metal nutrients from the host environment.8 Since metals are essential for many vital cellular functions, to avoid infection, the host restricts the access to the essential nutrients by means of a process known as “nutritional immunity”.9 The first step towards the design of a highly specific metal transport targeting therapeutic is therefore to obtain relevant information about thermodynamics and coordination chemistry of the interacting systems (metal–metal transporter).9–12

Zn(II), as well as several other transition metals, including Cu(II), is crucial for life and it is frequently involved in many cellular processes, where it can play the role of coenzyme or cofactor. Since Zn(II) concentration in a free, nonprotein-bound form is estimated to be sub-nanomolar,13 its uptake becomes extremely challenging for the fungal pathogen. C. albicans has evolved several mechanisms to overcome host nutritional immunity by expressing zinc transporters (e.g. Zrt1/Zrt2/Pra1 for Zn(II))14,15 or redundant enzymes that withhold the host stocks. Also Cu(II) is an endogenous metal and a necessary nutrient for C. albicans; it can compete with Zn(II) for the same binding sites and actively participate in the host–pathogen strife.16,17

In this context, we recently started a study on Zn(II) and Cu(II) binding behaviour towards C4YJH2 (UniProt Knowledgebase),18 a protein sequence of 199 amino acid residues, found in the genome of Candida albicans (strain WO-1), which is supposed to be involved in metal transport processes. In fact, it shares a high percentage of identity with putative Zn(II) transporters and proteins involved in metal homeostasis.19 Its amino acid sequence contains a remarkably high number of alternating histidine and serine residues, mostly located in the domains 131–148 (FHEHGHSHSHGSGGGGGG) and 157–165 (SHSHSHSHS) and confirmed to be involved in Zn(II) and Cu(II) coordination.20 In the native protein, these two His-rich domains are linked by the 9-residue sequence GSDHSGDSK (148–156), also containing a histidine and thus possibly contributing to the metal binding. Therefore, we decided to extend our investigation focusing on this “linker”. The protected peptide WT = Ac-GSDHSGDSK-NH2 has been considered, along with its analogues D7A = Ac-GSDHSGASK-NH2, S2A/S5A/S8A = Ac-GADHAGDAK-NH2, GSDH = Ac-GSDH-NH2, and HSGD = Ac-HSGD-NH2. The peptide sequence of D7A lacks the aspartic acid in position 7, which is substituted by an alanine in order to investigate the role of Asp-7 in the wild-type peptide (WT) protonation (possible hydrogen bond with Lys-9) and metal coordination. The nonapeptide S2A/S5A/S8A is an analogue of WT where serine residues have been replaced by alanines. Serine residues are rather abundant in C4YJH2 protein sequence. Although the serine side hydroxymethyl group has no acidic properties in the explored pH range and it is generally not expected to significantly interact with Cu(II) or Zn(II) ions, an electronic effect on the histidine residues and/or on the amides of the peptide backbone has previously been suggested.20 Finally, the study of the two short peptides, Ac-GSDH-NH2 and Ac-HSGD-NH2, corresponding to the left-hand and the right-hand side fragments around histidine, provides information on the Cu(II) attitude to coordinate amidic nitrogen atoms in the amino- or carboxyl-terminus direction.

Results and discussion

Ligand protonation

All the investigated peptides, WT, D7A, S2A/S5A/S8A, GSDH and HSGD, were protected at their N-terminus by acetylation and at their C-terminus by amidation; therefore, their acid–base behaviour depends on the amino acid side-chain properties, i.e. on the imidazole ring of His, the carboxylic group of Asp and the amino group of Lys. The amidic protons of the peptide backbone cannot be spontaneously released in the pH range explored by potentiometry, since they are very weak acids (pKa ≈ 15),21 but they can be displaced by Cu(II) at a mildly acidic pH value, to form a coordination bond. The protonation constants of WT, D7A, S2A/S5A/S8A, GSDH and HSGD are reported in Table 1 while the corresponding distribution diagrams are shown in Fig. S1–S5 (ESI).
Table 1 Overall (log[thin space (1/6-em)]β) and step (log[thin space (1/6-em)]K) protonation constants for WT, D7A, S2A/S5A/S8A, HSGD and GSDH at T = 298 K and I = 0.1 mol dm−3 (KCl). Values in parentheses are standard deviations on the last significant figure
Species WT S2A/S5A/S8A Species D7A HSGD GSDH
log[thin space (1/6-em)]β log[thin space (1/6-em)]Kstep log[thin space (1/6-em)]β log[thin space (1/6-em)]Kstep log[thin space (1/6-em)]β log[thin space (1/6-em)]Kstep log[thin space (1/6-em)]β log[thin space (1/6-em)]Kstep log[thin space (1/6-em)]β log[thin space (1/6-em)]Kstep
HL 10.84 (2) 10.84 10.39 (1) 10.39 HL 10.21 (2) 10.21 6.58 (2) 6.58 6.83 (2) 6.83
H2L 17.72 (3) 6.88 17.19 (2) 6.80 H2L+ 16.70 (3) 6.49 10.44 (2) 3.86 10.60 (2) 3.77
H3L+ 21.82 (3) 4.10 21.29 (2) 4.10 H3L2+ 20.46 (3) 3.76
H4L2+ 25.01 (3) 3.19 24.63 (2) 3.34


The obtained protonation constants are consistent with the literature values for similar systems22 and show reasonable similarities among the considered peptides; the variability can be mainly ascribed to the charge of the different species. The histidine protonation constant ranges from 6.49 to 6.88; this interval is rather small, suggesting the absence of specific behaviours depending on the ligand sequence. Furthermore, the presence or absence of the serine residues nearby the histidine does not seem to significantly affect its protonation constant. The lysine log[thin space (1/6-em)]K value is rather high (10.84) for the “wild-type” peptide (Ac-GSDHSGDSK-NH2), although still measurable under the employed experimental conditions. Finally, the Asp residues have the lowest log[thin space (1/6-em)]K values, due to their acidic moiety, which vary between 3.19 and 4.10, depending on the primary structure of the peptide and its charge.

The comparison between WT and its mutants does not point out any significant change in the protonation behaviour attributable to Ala substitutions. The difference in the log[thin space (1/6-em)]K values of Lys residues between WT and D7A does not suggest the formation of a hydrogen bond between the protonated amino group of Lys-9 and the carboxylate group of Asp-7, but it can be simply due to the higher charge of the former ligand, which contains an additional carboxylate group. Analogously, the Ala-Ser substitutions do not significantly affect the protonation constants of peptide side chains.

Cu(II) complexes

All the investigated peptides proved to be able to form stable 1[thin space (1/6-em)]:[thin space (1/6-em)]1 complexes with the Cu(II) ion; no poly-nuclear or bis-complexes have been detected either by potentiometry, mass spectrometry or EPR. No precipitation has been observed over the explored pH range. Overall complex-formation constants (log[thin space (1/6-em)]β) and corresponding acidity constants (pKa) are reported in Table 2 and the corresponding distribution diagrams are plotted in Fig. 1 and Fig. S6–S9 (ESI); ESI-MS results are shown in Table S1 (ESI). UV-vis, CD and EPR results are reported in Tables S2–S6 (ESI) and Fig. 2, 3 and Fig. S10–S17 (ESI).
Table 2 Overall stability constants (log[thin space (1/6-em)]β) and acid dissociation constants (pKa) of Cu(II) complexes with WT, S2A/S5A/S8A, D7A, HSGD and GSDH at T = 298 K and I = 0.1 mol dm−3 (KCl). Values in parentheses are standard deviations on the last significant figure
Species WT S2A/S5A/S8A Species D7A HSGD GSDH
log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa
[CuHL]+ 15.66 (4) 14.63 (3) 6.58 [CuHL]2+ 14.04 (6) 5.57
[CuL] 8.05 (3) 6.63 [CuL]+ 8.48 (3) 5.84 4.21 (2) 6.79 4.24 (5) 5.87
[CuH−1L] 3.54 (3) 8.15 1.42 (3) 9.59 [CuH−1L] 2.64 (2) 8.29 −2.58 (2) 6.91 −1.63 (3) 5.92
[CuH−2L]2− −4.61 (5) 10.46 −8.27 (4) 10.65 [CuH−2L] −5.65 (4) 10.38 −9.50 (2) 8.78 −7.56 (2) 8.53
[CuH−3L]3− −15.07 (7) −18.81 (5) [CuH−3L]2− −16.03 (5) −18.27 (2) −16.09 (4)



image file: c9mt00251k-f1.tif
Fig. 1 Representative species distribution diagram relative to Cu(II) complexes with WT; Cu(II)[thin space (1/6-em)]:[thin space (1/6-em)]L molar ratio = 0.8[thin space (1/6-em)]:[thin space (1/6-em)]1.

image file: c9mt00251k-f2.tif
Fig. 2 Vis absorption spectra [350–900 nm; optical path 1 cm] for Cu(II) complexes with WT; Cu(II)[thin space (1/6-em)]:[thin space (1/6-em)]L molar ratio = 0.8[thin space (1/6-em)]:[thin space (1/6-em)]1.

image file: c9mt00251k-f3.tif
Fig. 3 Circular dichroism spectra [200–800 nm; optical path 1 cm] for Cu(II) complexes with WT; Cu(II)[thin space (1/6-em)]:[thin space (1/6-em)]L molar ratio = 0.8[thin space (1/6-em)]:[thin space (1/6-em)]1.

Cu(II) starts to interact with WT and its mutants around pH 3.5 and the first detected complex is the mono-protonated species ([CuHL]+ for WT and S2A/S5A/S8A; [CuHL]2+ in the case of D7A). The stoichiometry of these species indicates that the ligand is mono-protonated, most likely at its Lys residue. Doubtless, Cu(II) is anchored to the imidazole nitrogen of histidine, while the possible participation in coordination by the carboxylic group(s) of the Asp residue(s) is arguable. The wavelength of maximum absorption expected for a coordination (NIm, COO) is 731 nm,21 in good agreement with the VIS absorption data in the pH range 5–6 for WT and S2A/S5A/S8A (λmax = 726 nm and 735 nm, respectively) (see Tables S2 and S3, ESI). Otherwise, in the case of D7A, we found λmax = 750 nm, a value closer to that expected for a Cu(II) complex with a (1NIm) configuration (760 nm). Increasing the pH value, the species [CuL] (for S2A/S5A/S8A) and [CuL]+ (for D7A) are formed with pKa values of 6.58 and 5.57, respectively. This complex was not observed in the case of WT. It is well known23 that, in this pH range, the cupric ion already bound to the ligand can displace one peptide hydrogen of the backbone to bind the amide nitrogen. In that case, the coordination geometry should be (1NIm, 1N) or (1NIm, 1N, COO). The presence of two nitrogens in the first coordination sphere of Cu(II) is confirmed by EPR spectra (see Tables S3 and S4, ESI). At higher pH, the ionization/binding of a second amide occurs with values of pKa = 6.63 (for S2A/S5A/S8A) and 5.84 (for D7A) and leads to the formation of the species [CuH−1L] and [CuH−1L], respectively. The complex [CuH−1L] was also detected in the system containing WT, where, however, a cooperative binding effect leads to the simultaneous coordination of the first and second amide nitrogens. These complexes reach about 90% of formation at physiological pH. The most plausible coordination hypothesis for these species is (1NIm, 2N) (see Fig. 4). In fact, the wavelengths of maximum absorption recorded at neutral pH are consistent with the expected value for a Cu(II) complex with this donor-atom set (λmax = 583 nm) and the EPR results at physiological pH confirm a 3N coordination.


image file: c9mt00251k-f4.tif
Fig. 4 Proposed coordination sphere of Cu(II) complexes with WT ligand at physiological pH. Explicit hydrogen atoms and water molecules are omitted for clarity.

In the alkaline pH range, the deprotonation of the third amide likely occurs (pKa values = 8.15, 9.59 and 8.29 for WT, S2A/S5A/S8A and D7A, respectively). The visible absorption bands always shift to shorter wavelengths, suggesting the increase of the number of coordinated nitrogen atoms. Based on spectroscopic results it is possible to suggest two possible configurations. In the case of ligands WT and D7A, the values of λmax (≈560 nm) suggest the substitution of the imidazole nitrogen by the N-amide in the Cu(II) equatorial plane, to obtain a (3N) coordination mode (expected λmax value: 563 nm) with a water molecule in the fourth equatorial position. On the other hand, in the case of ligand S2A/S5A/S8A, the obtained wavelength of maximum absorption at pH 10.3 (529 nm) strongly suggests a (1NIm, 3N) configuration (expected λmax value: 522 nm). The experimental values of A and g (Tables S2–S4, ESI) for all three systems agree with the hypothesis of both a 3N coordination and a 4N coordination. It is worth noting that the acidity constants of the three amide groups of S2A/S5A/S8A involved in complexation are approximately one order of magnitude lower (pKa one unit higher) than the corresponding constants of WT and D7A. Since no steric effect comes into play, an electronic effect due to the hydroxyl side chain of Ser can be the source of this difference and of the consequent lower stability of Cu(II) complexes with S2A/S5A/S8A with respect to the analogous complexes with the other mutants. A similar hypothesis was already put forward in the previous study on the main metal binding domains of C4YJH2.20 In addition, among the three Ser residues of S2A/S5A/S8A, a significant role should be played by Ser-2, as suggested by the constant values obtained for the systems Cu(II)/GSDH and Cu(II)/HSGD (see below).

Finally, under the most alkaline conditions, a further deprotonation step is observed and [CuH−3L]2− (for WT and S2A/S5A/S8A) and [CuH−3L]2− (for D7A) species are formed, without any significant variation in the spectroscopic parameters. This suggests that the last deprotonation step does not affect the Cu(II) coordination and likely corresponds to the deprotonation of a not-coordinated lysine. This hypothesis is also in good agreement with the thermodynamics results, since the corresponding pKa values (10.46, 10.65 and 10.38) are practically identical to those obtained from ligand protonation measurements.

Cu(II) interaction with HSGD and GSDH begins at pH lower than 4. The speciation model is the same for the two ligands, with four mononuclear 1[thin space (1/6-em)]:[thin space (1/6-em)]1 complexes, variously protonated. However, the distribution diagrams (Fig. S8 and S9, ESI) show that the behaviour is not exactly the same, as explained below.

The first detected complex is [CuL]+; the stoichiometry of this species suggests that both the histidine and the aspartic acid are not protonated. The imidazole nitrogen of the histidine can be deprotonated at such an acidic pH value only if it is bound to the metal ion; instead, the side carboxylate of the Asp residue can be involved or not in the metal coordination. In the case of HSGD, the obtained wavelength of maximum absorption at pH 6 – where the [CuL]+ complex is the most abundant species in solution – is 715 nm, suggesting the formation of a macrocycle with a (1NIm, COO) coordination mode (expected λmax value: 731 nm). On the other hand, in the case of GSDH, the wavelength of maximum absorption at pH 5.5 is significantly higher (763 nm), suggesting a (1NIm) coordination (expected λmax value: 760 nm). Most likely, in the latter case, the carboxylate group of the Asp residue (close to histidine), being at the opposite side of the backbone with respect to the imidazole ring, cannot bind the same Cu(II) ion due to steric hindrance. On increasing the pH value, three successive deprotonation steps are observed, giving rise to the species [CuH−1L], [CuH−2L] and [CuH−3L].2 The corresponding pKa values (6.79, 6.91 and 8.78 for HSGD and 5.87, 5.92 and 8.53 for GSDH, respectively) suggest the progressive coordination of amide nitrogens of the peptide backbone. Considering the complex [CuH−1L] formed by HSGD, a (1NIm, 1N, and COO) coordination mode can be suggested. In fact, the wavelength of maximum absorption at pH 7 – where this species reaches its higher percentage – (λmax = 636 nm) is very close to the expected λmax value (638 nm).21 It is not possible, from the present experimental data, to state which one of the backbone amides is bound to copper; as a matter of fact, the coordination of the amide nitrogen most close to His will lead to the formation of a 7-membered chelate ring. On the other hand, in the case of GSDH, the amide nitrogen which binds the metal ion should be that of His, the first one in the N-terminal direction, forming a stable 6-membered ring. The lower pKa value of this step for GSDH and, consequently, the higher value of the overall stability constant of [CuH−1L] (about one order of magnitude with respect to that of HSGD) support the above hypothesis.

At higher pH, the deprotonation/coordination of a second amide occurs in a quick sequence in both the systems, giving rise to the [CuH−2L] complex, the predominant species at physiological pH. In the case of HSGD, the positions of the d–d band in the UV-vis spectra both at pH 7.60 and 8.12 (615 nm and 612 nm, respectively, see Fig. S12 and Table S5 ESI) suggest a coordination mode where the two amides and the carboxyl group occupy the equatorial positions of the metal coordination sphere (2N, COO; expected λmax = 614 nm). EPR data at pH 8.7 agree with a [2N,O] coordination mode; in the CD spectra, the typical24 charge transfer band at 301 nm, due to the formation of the amide-copper bond, has been observed (Table S5, ESI). As for GSDH, the wavelength of maximum absorption at neutral pH (λmax = 585 nm) is very close to the one expected for a coordination (1NIm, 2N) (583 nm), where Cu(II) coordinates one imidazole and two amide nitrogens in the equatorial plane of the complex. The EPR data at pH 8.6 (A = 191.6, and g = 2.215, Table S6, ESI) clearly indicate a 3N coordination mode. The last deprotonation step is associated with the binding of a third amide in the main coordination plane; the corresponding species is [CuH−3L]2−. In the case of HSGD, a further blue-shift is observed in the vis absorption spectra with a wavelength of maximum absorption at pH 11 of 575 nm, only slightly higher than the predicted value for a (3N, COO) complex (547 nm). This difference can be ascribed to the presence of an axial binding group, most likely the His imidazole.21 EPR spectra at pH 10–11 are in agreement with both 3N and 4N coordination modes.25 Also in the case of the ligand GSDH, at very alkaline pH, the deprotonation of a further amide occurs, with a pKa 8.53. The visible absorption band shifts to a lower wavelength (560 nm), suggesting that the third backbone amide substitutes the imidazole nitrogen in the Cu(II) main coordination plane; in fact, the expected value of λmax for a (3N) coordination mode is 563 nm, very close to the experimental one. As for the (1NIm, 3N) coordination mode, the expected λmax value would be 522 nm, rather far from the experimental one. A similar behaviour was already previously observed for the protected tetrapeptide Boc-Ala-Gly-Gly-His.26 Actually, EPR spectra at pH 10–11 indicate a 4N coordination mode, thus suggesting that the imidazole ring can be still bound in an axial position.

Zn(II) complexes

The characterization of the Zn(II) complexes has been achieved by means of mass spectrometry, potentiometry and nuclear magnetic resonance. Mass spectrometric measurements provided information on the stoichiometry of the formed species, potentiometry allowed the partial and overall stability constants of the detected metal complexes to be determined and NMR spectra recorded both in the presence and in the absence of Zn(II) ion pointed out the precise metal binding sites. All the peptides studied here are able to form 1[thin space (1/6-em)]:[thin space (1/6-em)]1 complexes with the Zn(II) ion; no poly-nuclear or bis-complexes have been detected either by potentiometry or by mass spectrometry. No precipitation has been observed in the explored pH range. The formation constant values are shown in Table 3 and the corresponding distribution diagrams are plotted in Fig. 5 and Fig. S18–S21 (ESI); ESI-MS results are reported in Table S1, ESI.
Table 3 Overall stability constants (log[thin space (1/6-em)]β) and acid dissociation constants (pKa) of Zn(II) complexes with WT, S2A/S5A/S8A, D7A, HSGD and GSDH at T = 298 K and I = 0.1 mol dm−3 (KCl). Values in parentheses are standard deviations on the last significant figure
Species WT S2A/S5A/S8A Species D7A HSGD GSDH
log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa log[thin space (1/6-em)]β pKa
[ZnHL]+ 14.11 (6) 13.37 (9)
[ZnL] [ZnL]+ 2.9 (1) 2.85 (9)
[ZnH−1L] −1.86 (4) 10.39 −2.15 (4) 10.35 [ZnH−1L] −2.57 (3) 10.36
[ZnH−2L]2− −12.25 (5) −12.50 (6) [ZnH−2L] −12.93 (6) −12.43 (4) −12.41 (3)



image file: c9mt00251k-f5.tif
Fig. 5 Representative species distribution diagram relative to Zn(II) complexes with WT; Zn(II)[thin space (1/6-em)]:[thin space (1/6-em)]L molar ratio = 0.8[thin space (1/6-em)]:[thin space (1/6-em)]1.

WT and S2A/S5A/S8A peptides behave in a very similar manner in coordination to the Zn(II) ion. In both systems, the metal/ligand interaction starts at pH 4 and the first detected species is [ZnHL]+, which is also the prevailing complex at physiological pH. The stoichiometry of this complex suggests that only the side chain of Lys is protonated. The metal ion should be anchored at the His residue and one or both of the carboxylate moieties of the Asp residues can be involved in complexation; the corresponding coordination mode can either be (1NIm, COO) or (1NIm, 2COO). The suggested geometry can be either tetrahedral or trigonal bipyramid, as most frequently found in proteins,27 where the additional coordination positions are occupied by water molecules (Fig. 6). On increasing the pH value, two simultaneous deprotonation steps occur, giving rise to the [ZnH−1L] complex, the most abundant species in the pH range 8–10. It is reasonable to suggest the hydrolysis of two coordinated water molecules. The [ZnL] complex has not been detected by potentiometry, thus suggesting it is only a transient species. 1H–1H TOCSY spectra recorded at physiological pH (Fig. S22 and S23, ESI) confirm the suggested coordination hypotheses, since the major Zn(II)-induced shifts are those of histidine protons (Hε1–Hδ2), (Hα–Hβ1), and (Hα–Hβ2) and aspartic acid protons (Hα–Hβ1), and (Hα–Hβ2). Furthermore, in the case of WT ligand, the pronounced deshielding effect exhibited by His Hε1 protons (Δδ = 0.125 ppm) compared to the shift of His Hδ2 (Δδ = 0.050 ppm) (Fig. S22, ESI), in the presence of Zn(II) ions, suggests a coordination through the imidazole-Nδ.28–31 This difference is less pronounced in S2A/S5A/S8A, thus suggesting that the presence of the Ser residues may influence the binding behaviour of histidine. It is also worth noting that in the Zn(II)/S2A/S5A/S8A system, the protons of lysine display a moderate downfield shift (Fig. S23, ESI): since Lys does not directly participate in coordination, these perturbations can suggest an interaction of Zn(II) with the aspartic acid in position 7, the nearest the Lys residue. In the case of the WT ligand, only Lys-Hα exhibits a small perturbation after Zn(II) addition, possibly because the presence of serine residues reduces the deshielding effect on Lys. Finally, starting from pH 8.5, the complex [ZnH−1L] releases a further proton, giving rise to the species [ZnH−2L]2−. The corresponding pKa values for the two systems (10.39 and 10.35 for WT and S2A/S5A/S8A, respectively) are very similar to those obtained in the absence of Zn(II), suggesting the simple deprotonation of the Lys residue without any involvement in complexation.


image file: c9mt00251k-f6.tif
Fig. 6 Proposed molecular structure of Zn(II) complexes with WT ligand at physiological pH. Explicit hydrogen atoms are omitted for clarity.

As for peptide D7A, Zn(II) complexes begin to form only at pH 6.5, i.e. when imidazole nitrogen is deprotonated and available for complexation. Evidently, the presence, at lower pH, of only one Asp carboxylate is not sufficient to stably bind the metal ion. Thus, the first detected species is [ZnH−1L], most likely derived from the binding of D7A to the hydrolytic species [ZnOH]+ already present in solution in small amount (see distribution diagram of Fig. S19, ESI). In the complex [ZnH−1L], the dominant species throughout all the explored pH range, the Zn(II) ion is certainly bound to the histidine residue. Rather unexpectedly, the 1H–1H TOCSY spectra recorded at pH 7.2 (Fig. S24, ESI) suggest that the aspartic acid in position 3 should not participate in complex formation. Indeed, the metal addition causes only a selective shift of histidine (Hε1–Hδ2) imidazole protons (more pronounced for His Hε1 protons, as already observed above for WT and possibly due to the presence of Ser residues) leaving the signals of Hα–Hβ1/Hβ2 unchanged, therefore suggesting that no other residues are involved in the complexation. Signals corresponding to the aspartic acid (Hα–Hβ1), and (Hα–Hβ2) protons undergo only a slight shift, supposedly due to the proximity to the histidine anchor site. Starting form pH 8.5, the complex [ZnH−2L] begins to form with pKa = 10.36, likely corresponding to the deprotonation of a not-coordinated lysine.

For the sake of completeness, Zn(II) complexes with ligands HSGD and GSDH have been also investigated. These peptides only possess two possible donor groups: one histidine and one aspartic acid. No precipitation has been observed in the explored pH range. In both the systems Zn(II) complexes begin to form at pH 4 and only two major species were detected by potentiometry: [ZnL]+, where both the Asp and His residues should be bound to the metal ion, and [ZnH−2L], where the Zn(II) coordination sphere possibly involves also two ionized water molecules, as already hypothesised for the systems WT, D7A and S2A/S5A/S8A. The participation of His and Asp residues in the Zn(II) complex formation is also confirmed by the general changes of their proton correlations in 1H–1H TOCSY spectra (Fig. S25 and S26, ESI). It is also reasonable to assume that both the low availability of donor groups and the relatively small dimensions of these peptides can encourage the formation of bis-/(poly)-complexes, although they should represent only a minor species under the experimental conditions employed here (ligand and metal ion in almost equimolar concentration) and therefore not detectable. The MS spectra recorded at pH 6 confirm the presence of mononuclear complexes as major species (see Table S1, ESI).

Comparison of complex stability

In an attempt to better understand the binding ability of the investigated ligands towards Zn(II) and Cu(II) ions, competition plots have been built up at equimolar concentrations of ligands and metal ions. The diagrams plotted in Fig. 7 compare the thermodynamic stability of WT, D7A and S2A/S5A/S8A peptide complexes with Cu(II) and Zn(II). The replacement of the Asp residue in position 7 seems to affect the affinity with both the metal ions. In the case of Cu(II), the absence of Asp7 makes the metal complexation only slightly less efficient under acidic pH values (Fig. 7a). It is noteworthy that at pH 6.5 the competition curves of Cu(II)-WT and Cu(II)-D7A coincide; this is in agreement with the proposed speciation models, since at that pH value, the species [CuH−1L] dominate in solution and have the same coordination geometry in both systems. The importance of the Asp residue in position 7 is much more evident in the case of Zn(II) (Fig. 7b), since WT and S2A/S5A/S8A show a much better binding ability throughout all the explored pH range, thus suggesting that the carboxyl group of Asp7 is of fundamental importance for the complex stability. As for the possible role of serine residues, in the case of Cu(II) (Fig. 7a), the presence of non-coordinating serines seems to have an impact on amide deprotonation, making complexes with WT and D7A more stable than the species formed by S2A/S5A/S8A, starting from pH 5 on, when amides come into play. On the contrary, this trend does not occur for Zn(II) complexes where amides are not involved in the coordination. This is in agreement with the above hypothesis that serines are responsible for an electronic effect that makes amide protons much more acidic.
image file: c9mt00251k-f7.tif
Fig. 7 Competition plots for solutions containing equimolar concentrations of M(II), WT, D7A and S2A/S5A/S8A. M(II) = (a) Cu(II) and (b) Zn(II).

The calculated competition diagram for the Cu(II)/WT/HSGD/GSDH system is shown in Fig. S27 (ESI). The HSGD and GSDH tetra-peptides coincide with the 151–154 and 148–151 amino acid sequence of the C4YJH2 protein and are coincidentally composed by the same amino acid residues with a different order. They also correspond to the minimal coordination site for a Cu(II) ion when anchored to the His residue of the linker sequence WT (C4YJH2148–156). After the histidine binding, the N-amide coordination can only proceed towards the N-terminus in the GSDH ligand – forming a 6-membered ring between the imidazole nitrogen and the first amide – and towards the C-terminus in the case of HSGD – with instead the formation of an initial 7-membered ring. These two different coordination modes are certainly in competition also in the WT peptide (Ac-GSDHSGDSK-NH2). The comparison among these systems reveals that under acidic conditions the two short peptides have similar behaviour, while starting from pH 5.5 (when the amide coordination comes into play), GSDH complexes are by far more stable than HSGD species, suggesting that in WT the amide coordination is unbalanced in favour of the N-terminal direction. In the case of the Zn(II) ion, which is not able to deprotonate and bind amide nitrogens, the two tetra-peptides HSGD and GSDH have exactly the same behaviour all over the explored pH range (Fig. S28, ESI). On the other hand, WT ligand shows a slightly higher affinity for Zn(II) under acidic and physiological pH, probably due to the presence of two Asp residues, which can both contribute to the metal coordination. At alkaline pH, the stability of WT complexes becomes a little lower than that of the species formed by the two short fragments, possibly only because the lower negative charge of the latter species stabilizes the hydroxo-complexes.

Conclusions

The present work represents an in-depth analysis of the thermodynamic and spectroscopic properties of Ac-GSDHSGDSK-NH2 (WT) solution complex-formation equilibria. Four analogues were also taken into account, for the sake of comparison. WT, corresponding to the 148–156 amino acid sequence of C4YJH2, acts as a linker between the two main poly-histidine domains of the mentioned protein and proved able to coordinate both the Cu(II) and Zn(II) ions.

For all the investigated peptides (Ac-GSDHSGDSK-NH2, Ac-GSDHSGASK-NH2, Ac-GADHAGDAK-NH2, Ac-GSDH-NH2 and Ac-HSGD-NH2) the Cu(II) coordination begins at about pH 3.5. After a first anchoring step to the histidine residue, up to three deprotonated amidic groups of the peptide backbone can bind the Cu(II) ion, occupying the equatorial position of its coordination sphere. At physiological pH, the species (1NIm, 2N) is always the most abundant in solution. This complex is rather stable and reaches about 90% of formation with all the investigated peptides. The study of the two tetra-peptides, Ac-GSDH-NH2 and Ac-HSGD-NH2 allowed us to qualitatively describe the Cu(II) N-amide-coordination attitude: it can be assumed that the nona-peptide WT is able to form a mixture of isomeric species in which the amide coordination towards the N-terminal direction is favoured. Furthermore, the substitution of serine with alanine residues significantly lowers the peptide affinity for the Cu(II) ion, suggesting a possible role of Ser residues (quite abundant in C4YHJ2 protein sequence) in the biological task of metal uptake and regulation.

In Zn(II) complexes, the metal coordination at physiological pH occurs mainly by means of (at least) one aspartic acid moiety and the histidine residue. The Asp in position 7 is likely a crucial residue for the formation and stabilization of Zn(II) complexes, as indicated by the decrease of Zn(II) affinity for ligand Ac-GSDHSGASK-NH2 (Fig. 7b), where the proposed metal coordination does not involve the aspartic acid.

An overall comparison of the stabilities of the studied domains of C4YJH2 is shown in Fig. 8. The calculated competition plots take into account the Cu(II) and Zn(II) binding abilities of the studied “linker” Ac-GSDHSGDSK-NH2 (148–156) and of the two poly-His sequences Ac-FHEHGHSHSHGSGGGGGG-NH2 (131–148) and Ac-SHSHSHSHS-NH2 (157–165).20 The obtained results emphasize that the ligand metal binding efficiency comes mainly from the high number of histidine residues, however highlighting a minor, but non-negligible contribution of the linker sequence in the complex formation, which, in the acidic pH range, is most likely due to the impact of the further histidine and the additional serine residues.


image file: c9mt00251k-f8.tif
Fig. 8 Competition plots for solutions containing equimolar concentrations of M(II), Ac-FHEHGHSHSHGSGGGGGG-NH2 (131–148), Ac-GSDHSGDSK-NH2 (148–156) and Ac-SHSHSHSHS-NH2 (157–165). M(II) = (a) Cu(II) and (b) Zn(II).

As for the histidine rich fragments of C4YJH2, it is possible to summarize that Zn(II) coordination mostly occurs by means of three histidine residues (3NIm) and an oxygen atom belonging to the aspartic acid side chain (in the case of Ac-FHEHGHSHSHGSGGGGGG-NH2) or to a water molecule (in the case of Ac-SHSHSHSHS-NH2). Similarly, in the case of Cu(II) complexation, histidine residues promote the metal binding at acidic pH, while, moving to alkaline conditions, the amide nitrogens of the peptide backbone gradually bind copper, eventually substituting the imidazole nitrogens in the equatorial positions. No direct evidence for the preferred metal binding site of the C4YJH2 sequence is available, although the higher affinity of Ac-FHEHGHSHSHGSGGGGGG-NH2 suggests that the distribution of mono-nuclear species is in favour of this fragment, most likely due to the availability of both a higher number of His imidazoles and the Asp carboxylic side-chain.

As a matter of fact, the presence of specific, highly conserved, histidine-rich motifs (e.g. HXHXH or HXXHXH) determines the metal-binding ability of the protein and then its capability to affect the host/pathogen Zn(II) homeostasis.32–34 However, it is also clear that different coordination modes, like those found at different pH values, can definitely affect the efficacy of the protein activity in the task of metal recruitment, as previously shown in the case of calcitermin, a human antimicrobial peptide.32 The rational understanding of such correlation is still ongoing, especially in the case of C4YJH2, the biological activity of which is still completely unknown, and the door is open to further investigations.

Experimental

Materials

ZnCl2 and CuCl2 were extra pure products (Sigma-Aldrich); the concentrations of their stock solutions were standardised by EDTA titration and periodically checked via ICP-MS. The carbonate-free stock solutions of 0.1 mol dm−3 KOH were prepared by diluting concentrated KOH (Sigma-Aldrich) and then potentiometrically standardized with the primary standard potassium hydrogen phthalate (99.9% purity). All sample solutions were prepared with freshly prepared Milli-Q® water. The HCl and HNO3 stock solutions were prepared by diluting concentrated HCl and HNO3 (Sigma-Aldrich) and then standardized with KOH. The ionic strength was adjusted to 0.1 mol dm−3 by adding KCl (Sigma-Aldrich). Grade A glassware was employed throughout.

Peptide synthesis and purification

Peptides: Ac-GSDHSGDSK-NH2; Ac-GSDHSGASK-NH2; Ac-GADHAGDAK-NH2; Ac-GSDH-NH2; and Ac-HSGD-NH2 were synthesized according to published methods35 using Fmoc/t-butyl chemistry with a Syro XP multiple peptide synthesizer (MultiSynTech GmbH, Witten, Germany). Rink amide MBHA resin was used as a solid support for the synthesis of all derivatives. Fmoc-amino acids (4-fold excess) were sequentially coupled to the growing peptide chain using DIPCDI/HOBt (N,N′-diisopropylcarbodiimide/1-hydroxybenzotriazole) (4-fold excess) as an activating mixture for 1 h at room temperature. Cycles of deprotection of Fmoc (40% piperidine/N,N-dimethylformamide) and coupling with the subsequent amino acids were repeated until the desired peptide-bound resin was completed. N-Terminal acetylation was performed with acetic anhydride (0.5 M) in the presence of N-methylmorpholine (0.25 M) (3[thin space (1/6-em)]:[thin space (1/6-em)]1 v/v; 2 mL/0.2 g of resin) as the last synthetic step. The protected peptide-resin was treated with reagent B36 (trifluoroacetic acid (TFA)/H2O/phenol/triisopropylsilane 88[thin space (1/6-em)]:[thin space (1/6-em)]5[thin space (1/6-em)]:[thin space (1/6-em)]5[thin space (1/6-em)]:[thin space (1/6-em)]2; v/v; 10 mL/0.2 g of resin) for 1.5 h at room temperature. After filtration of the resin, the solvent was concentrated in vacuo and the residue triturated with ethyl ether. Crude peptides were purified by preparative reversed-phase HPLC using a Water Delta Prep 3000 system with a Jupiter column C18 (250 × 30 mm, 300 A, and 15 μm spherical particle size). The column was perfused at a flow rate of 20 mL min−1 with a mobile phase containing solvent A (5%, v/v, acetonitrile in 0.1% TFA), and a linear gradient from 0 to 30% of solvent B (60%, v/v, acetonitrile in 0.1% TFA) over 25 min for the elution of peptides. Analytical HPLC analyses were performed on a Beckman 116 liquid chromatograph equipped with a Beckman 166 diode array detector. Analytical purity of the peptides has been assessed using a Zorbax C18 column (4.6 × 150 mm, 3 μm particle size) with the above solvent system (solvents A and B) programmed at a flow rate of 0.5 mL min−1 using a linear gradient from 0% to 50% B over 25 min. All analogues showed ≥95% purity when monitored at 220 nm. The molecular weight of the final compounds was determined by an ESI Micromass ZMD-2000 mass spectrometer.

Potentiometric measurements

Stability constants for proton and metal complexes were calculated from pH-metric titration curves registered at T = 298 K and ionic strength 0.1 mol dm−3 (KCl). The potentiometric apparatus consisted of an Orion EA 940 pH-meter system provided with a Metrohm 6.0234.100, glass-body, micro combination pH electrode and a Hamilton MICROLAB 500 dosing system, equipped with a 0.5 mL micro burette. The thermostated glass-cell was equipped with a magnetic stirring system, a microburet delivery tube and an inlet–outlet tube for nitrogen. High purity grade nitrogen gas was gently blown over the test solution in order to maintain an inert atmosphere. Constant-speed magnetic stirring was applied throughout. Solutions were titrated with 0.1 mol dm−3 carbonate-free KOH. The electrode was daily calibrated for hydrogen ion concentration by titrating HNO3 with alkaline solution under the same experimental conditions as above. The standard potential and the slope of the electrode couple were computed by means of the SUPERQUAD37 and Glee38 programs. The purities and the exact concentrations of the ligand solutions were determined by the Gran method.39 The HYPERQUAD40 program was employed for the overall formation constant (β) calculations, referring to the following equilibrium:
pM + qL + rH ⇆ MpLqHr
(charges omitted; p is 0 in the case of ligand protonation; r can be negative). Step formation constants (Kstep) and/or acid dissociation constants (Ka) are also reported. The computed standard deviations (referring to random errors only) were given by the program itself and are shown in parentheses as uncertainties on the last significant figure. Hydrolysis constants for Zn(II) and Cu(II) ions were taken from the literature (Table S7, ESI).22,41 The distribution and the competition diagrams were computed using the HYSS program.42 In fact, the overall metal binding ability of the different ligands can be compared in a wide pH range by computing the competition diagrams, starting from the binary speciation models. A solution containing the metal and the two (or more) ligands (or vice versa) is simulated, admitting that all the components compete with each other to form the respective binary complexes, without mixed species formation. This is a reasonable approximation in the case of peptides, which most often form only 1[thin space (1/6-em)]:[thin space (1/6-em)]1 complexes in which the peptide completely wraps the metal ion.

Mass spectrometric measurements

High-resolution mass spectra were obtained on a BrukerQ-FTMS spectrometer (Bruker Daltonik, Bremen, Germany), equipped with an Apollo II electrospray ionization source with an ion funnel. The mass spectrometer was operated in the positive and negative ion modes. The instrumental parameters were as follows: scan range m/z 300–4000, dry gas-nitrogen, temperature 453 K and ion energy 5 eV. The capillary voltage was optimized to the highest S/N ratio and it was 4500 V. The small changes in voltage (±500 V) did not significantly affect the optimized spectra. The samples were prepared in a 1[thin space (1/6-em)]:[thin space (1/6-em)]1 methanol–water mixture at different pH values. The samples (Zn(II)[thin space (1/6-em)]:[thin space (1/6-em)]ligand and Cu(II)[thin space (1/6-em)]:[thin space (1/6-em)]ligand in a 0.8[thin space (1/6-em)]:[thin space (1/6-em)]1 stoichiometry, [ligand]tot = 5 × 10−4 mol dm−3) were infused at a flow rate of 3 μL min−1. The instrument was calibrated externally with a Tunemix™ mixture (Bruker Daltonik, Germany) in a quadratic regression mode. Data were processed using the Bruker Compass DataAnalysis 4.2 program. The mass accuracy for the calibration was better than 5 ppm, enabling, together with the true isotopic pattern (using SigmaFit), an unambiguous confirmation of the elemental composition of the obtained complex.

Spectroscopic measurements

The absorption spectra were recorded on a Varian Cary50 Probe spectrophotometer, in the range 350–900 nm, using a quartz cuvette with an optical path of 1 cm. Circular dichroism (CD) spectra were recorded on a Jasco J-1500 CD spectrometer in the 200–800 nm range, using a quartz cuvette with an optical path of 1 cm in the visible and near-UV range. Electron paramagnetic resonance (EPR) spectra were recorded in liquid nitrogen on a Bruker ELEXSYS E500 CW-EPR spectrometer at X-band frequency (9.5 GHz) and equipped with an ER 036TM NMR teslameter and an E41 FC frequency counter. Ethylene glycol (30%) was used as a cryoprotectant for EPR measurements. The EPR parameters were analysed by computer simulation of the experimental spectra using the WIN-EPR SIMFONIA software, version 1.2 (Bruker). The concentrations of sample solutions used for spectroscopic studies were similar to those employed in the potentiometric experiment. The UV-vis, CD and EPR spectroscopic parameters were calculated from the spectra obtained at the pH values corresponding to the maximum concentration of each particular species, based on distribution diagrams.

NMR measurements

NMR spectra were recorded at 14.1 T on a Bruker Avance III 600 MHz system equipped with a Silicon Graphics workstation. The temperatures were controlled with an accuracy of ±0.1 K. Suppression of the residual water signal was achieved by excitation sculpting, using a selective square pulse 2 ms long on water. All the samples were prepared in D2O (99.9% from Merck) solution. Proton resonance assignment was accomplished by 2D 1H–1H total correlation spectroscopy (TOCSY) experiments carried out with standard pulse sequences. Samples of analysed complexes (Zn(II)[thin space (1/6-em)]:[thin space (1/6-em)]ligand in a 1[thin space (1/6-em)]:[thin space (1/6-em)]1 stoichiometry, [ligand]tot = 0.003 mol dm−3) were prepared by adding metal ions to the acidic solution of a ligand (pH 3), and the pH was then increased to a higher value (pH 7.4). Spectral processing and analysis was performed using Bruker TOPSPIN 2.1 and Sparky.43

Complex structure drawings

The pictures of the complex structural hypotheses have been drawn with ACD/ChemSketch and the 3D structure visualisation was obtained with Mercury (The Cambridge Crystallographic Data Centre, CCDC).

Author contributions

D. Bellotti carried out potentiometric, CD, EPR, ESI-MS and NMR measurements on Zn(II) and Cu(II) systems. C. Tocchio carried out potentiometric and UV-Vis measurements on Cu(II) systems. R. Guerrini performed peptide synthesis and purification. M. Remelli and M. Rowińska-Żyrek designed and coordinated the experiments. D. B., M. R. and M. R.-Z. contributed to the data interpretation. All the authors contributed equally to the writing of the paper.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

The present research was financially supported by the National Science Centre (UMO-2017/26/A/ST5/00364, SONATA BIS grant to MR-Z), University of Ferrara (FAR 2018), CIRCMSB (Consorzio Interuniversitario di Ricerca in Chimica dei Metalli nei Sistemi Biologici, Bari, Italy), MIUR (Ministero dell’Istruzione, dell’Università e della Ricerca) projects (PRIN2015-2015MP34H3 and PRIN2015-2015T778JW) and Erasmus+ programme of the European Union. The CD measurements were carried out with the equipment purchased thanks to the financial support of the Polish National Science Centre (Grant UMO 2015/19/B/ST5/00413).

Notes and references

  1. T. Kourkoumpetis, D. Manolakaki, G. C. Velmahos, Y. C. Chang, H. B. Alam, M. M. De Moya, E. A. Sailhamer and E. Mylonakis, Virulence, 2010, 1, 359 CrossRef PubMed.
  2. S. Campoy and J. L. Adrio, Biochem. Pharmacol., 2017, 133, 86 CrossRef CAS PubMed.
  3. A. L. Mavor, S. Thewes and B. Hube, Curr. Drug Targets, 2005, 6, 863 CrossRef CAS PubMed.
  4. T. J. Walsh, Expert Rev. Anti-Infect. Ther., 2017, 15, 577 CrossRef PubMed.
  5. A. Pitarch, C. Nombela and C. Gil, Curr. Top. Med. Chem., 2018, 18, 1375 CrossRef CAS PubMed.
  6. V. Srivastava, R. K. Singla and A. K. Dubey, Curr. Top. Med. Chem., 2018, 18, 759 CrossRef CAS PubMed.
  7. M. A. Pfaller and D. J. Diekema, Clin. Microbiol. Rev., 2007, 20, 133 CrossRef CAS PubMed.
  8. P. Chandrangsu, C. Rensing and J. D. Helmann, Nat. Rev. Microbiol., 2017, 15, 338 CrossRef CAS PubMed.
  9. M. I. Hood and E. P. Skaar, Nat. Rev. Microbiol., 2012, 10, 525 CrossRef CAS PubMed.
  10. M. Blatzer and J.-P. Latgé, Curr. Opin. Microbiol., 2017, 40, 152 CrossRef CAS PubMed.
  11. E. R. Ballou and D. Wilson, Curr. Opin. Microbiol., 2016, 32, 128 CrossRef CAS PubMed.
  12. K. W. Paulina, W. Joanna and R.-Z. Magdalena, Curr. Med. Chem., 2016, 23, 3717 CrossRef PubMed.
  13. I. Bremner and P. M. May, in Zinc in Human Biology, ed. C. F. Mills, Springer London, London, 1989, p. 95 Search PubMed.
  14. T. Yi-Hsuan, H. Adela Ya-Ting, C. Po-Yu, C. Hui-Ting and K. Chai-Lin, Curr. Pharm. Des., 2011, 17, 2308 CrossRef PubMed.
  15. F. Citiulo, I. D. Jacobsen, P. Miramón, L. Schild, S. Brunke, P. Zipfel, M. Brock, B. Hube and D. Wilson, PLoS Pathog., 2012, 8, e1002777 CrossRef CAS PubMed.
  16. G. Butler, M. D. Rasmussen, M. F. Lin, M. A. S. Santos, S. Sakthikumar, C. A. Munro, E. Rheinbay, M. Grabherr, A. Forche, J. L. Reedy, I. Agrafioti, M. B. Arnaud, S. Bates, A. J. P. Brown, S. Brunke, M. C. Costanzo, D. A. Fitzpatrick, P. W. J. de Groot, D. Harris, L. L. Hoyer, B. Hube, F. M. Klis, C. Kodira, N. Lennard, M. E. Logue, R. Martin, A. M. Neiman, E. Nikolaou, M. A. Quail, J. Quinn, M. C. Santos, F. F. Schmitzberger, G. Sherlock, P. Shah, K. A. T. Silverstein, M. S. Skrzypek, D. Soll, R. Staggs, I. Stansfield, M. P. H. Stumpf, P. E. Sudbery, T. Srikantha, Q. Zeng, J. Berman, M. Berriman, J. Heitman, N. A. R. Gow, M. C. Lorenz, B. W. Birren, M. Kellis and C. A. Cuomo, Nature, 2009, 459, 657 CrossRef CAS PubMed.
  17. A. N. Besold, B. A. Gilston, J. N. Radin, C. Ramsoomair, E. M. Culbertson, C. X. Li, B. P. Cormack, W. J. Chazin, T. E. Kehl-Fie and V. C. Culotta, Infect. Immun., 2018, 86, e00779 CAS.
  18. T. UniProt Consortium, Nucleic Acids Res., 2018, 46, 2699 CrossRef PubMed.
  19. G. Binkley, J. Binkley, M. S. Skrzypek, M. Simison, S. R. Miyasato and G. Sherlock, Nucleic Acids Res., 2016, 45, D592 Search PubMed.
  20. D. Bellotti, D. Łoboda, M. Rowińska-Żyrek and M. Remelli, New J. Chem., 2018, 42, 8123 RSC.
  21. H. Sigel and R. B. Martin, Chem. Rev., 1982, 82, 385 CrossRef CAS.
  22. L. D. Pettit and H. K. J. Powell, The IUPAC Stability Constants Database, Royal Society of Chemistry, London, 1992–2000 Search PubMed.
  23. R. J. Sundberg and R. B. Martin, Chem. Rev., 1974, 74, 471 CrossRef CAS.
  24. R. Hennig, A. Veser, S. Kirchhof and A. Goepferich, Mol. Pharmaceutics, 2015, 12, 3292 CrossRef CAS PubMed.
  25. J. Peisach and W. E. Blumberg, Arch. Biochem. Biophys., 1974, 165, 691 CrossRef CAS PubMed.
  26. L. D. Pettit, S. Pyburn, W. Bal, H. Kozlowski and M. Bataille, J. Chem. Soc., Dalton Trans., 1990, 3565 RSC.
  27. A. R. Borges and C. L. Schengrund, Curr. Drug Targets: Infect. Disord., 2005, 5, 247 CrossRef CAS PubMed.
  28. M. Remelli, M. Peana, S. Medici, L. G. Delogu and M. A. Zoroddu, Dalton Trans., 2013, 42, 5964 RSC.
  29. S. Medici, M. Peana, L. G. Delogu and M. A. Zoroddu, Dalton Trans., 2012, 41, 4378 RSC.
  30. M. A. Zoroddu, S. Medici, M. Peana and R. Anedda, Dalton Trans., 2010, 39, 1282 RSC.
  31. A. Urbani, R. Bazzo, M. C. Nardi, D. O. Cicero, R. De Francesco, C. Steinkuhler and G. Barbato, J. Biol. Chem., 1998, 273, 18760 CrossRef CAS PubMed.
  32. D. Bellotti, M. Toniolo, D. Dudek, A. Mikołajczyk, R. Guerrini, A. Matera-Witkiewicz, M. Remelli and M. Rowińska-Żyrek, Dalton Trans., 2019, 48, 13740 RSC.
  33. D. Łoboda and M. Rowińska-Żyrek, Dalton Trans., 2017, 46, 13695 RSC.
  34. A. Miller, D. Dudek, S. Potocki, H. Czapor-Irzabek, H. Kozłowski and M. Rowińska-Żyrek, Metallomics, 2018, 10, 1631 RSC.
  35. N. L. Benoiton, Chemistry of Peptide Synthesis, Taylor & Francis, 2005 Search PubMed.
  36. N. A. Sole and G. Barany, J. Org. Chem., 1992, 57, 5399 CrossRef CAS.
  37. P. Gans, A. Sabatini and A. Vacca, J. Chem. Soc., Dalton Trans., 1985, 1195 RSC.
  38. P. Gans and B. O'Sullivan, Talanta, 2000, 51, 33 CrossRef CAS PubMed.
  39. G. Gran, Acta Chem. Scand., 1950, 4, 559 CrossRef CAS.
  40. P. Gans, A. Sabatini and A. Vacca, Talanta, 1996, 43, 1739 CrossRef CAS PubMed.
  41. G. Arena, R. Cali, E. Rizzarelli and S. Sammartano, Thermochim. Acta, 1976, 16, 315 CrossRef CAS.
  42. L. Alderighi, P. Gans, A. Ienco, D. Peters, A. Sabatini and A. Vacca, Coord. Chem. Rev., 1999, 184, 311 CrossRef CAS.
  43. W. Lee, M. Tonelli and J. L. Markley, Bioinformatics, 2015, 31, 1325 CrossRef PubMed.

Footnote

Electronic supplementary information (ESI) available. See DOI: 10.1039/c9mt00251k

This journal is © The Royal Society of Chemistry 2019