Human Annexins A1, A2, and A8 as Potential Molecular Targets for Ni(II)

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Human Annexins A1, A2, and A8 as Potential Molecular Targets for Ni(II) Ions Nina E. Wezynfeld, Karolina Bossak, Wojciech Goch, Arkadiusz Bonna, Wojciech Bal, and Tomasz Frączyk* Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Pawińskiego 5a, 02-106 Warsaw, Poland S Supporting Information *

ABSTRACT: Nickel is harmful for humans, but molecular mechanisms of its toxicity are far from being fully elucidated. One of such mechanisms may be associated with the Ni(II)dependent peptide bond hydrolysis, which occurs before Ser/Thr in Ser/Thr-Xaa-His sequences. Human annexins A1, A2, and A8, proteins modulating the immune system, contain several such sequences. To test if these proteins are potential molecular targets for nickel toxicity we characterized the binding of Ni(II) ions and hydrolysis of peptides Ac-KALTGHLEE-am (A1−1), AcTKYSKHDMN-am (A1−2), and Ac-GVGTRHKAL-am (A1−3), from annexin A1, Ac-KMSTVHEIL-am (A2−1) and Ac-SALSGHLET-am (A2−2), from annexin A2, and Ac-VKSSSHFNP-am (A8−1), from annexin A8, using UV−vis and circular dichroism (CD) spectroscopies, potentiometry, isothermal titration calorimetry, high-performance liquid chromatography (HPLC), and electrospray ionization mass spectrometry (ESI-MS). We found that at physiological conditions (pH 7.4 and 37 °C) peptides A1−2, A1−3, A8−1, and to some extent A2−2 bind Ni(II) ions sufficiently strongly in 4N complexes and are hydrolyzed at sufficiently high rates to justify the notion that these annexins can undergo nickel hydrolysis in vivo. These results are discussed in the context of specific biochemical interactions of respective proteins. Our results also expand the knowledge about Ni(II) binding to histidine peptides by determination of thermodynamic parameters of this process and spectroscopic characterization of 3N complexes. Altogether, our results indicate that human annexins A1, A2, and A8 are potential molecular targets for nickel toxicity and help design appropriate cellular studies.

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INTRODUCTION Nickel is toxic for humans, causing allergy, cancers of the respiratory system, and other serious health problems.1−5 Despite this, nickel is a component of stainless steel and other alloys, which are found in coins, mobile phones, accessories, jewelry, and many implantable materials. These objects and materials remain in contact with human tissues and release nickel.6−13 Furthermore, significant amounts of this metal are delivered into lungs from standard as well as electronic cigarettes and from airborne dust.14−16 Nickel, as presumably the most frequent contact allergen, is responsible for substantial health problems, as more than 5% of the general population of many developed countries (e.g., Germany and Denmark) is sensitized to this metal.3,6 The prevalence of nickel allergy is especially high (17%) among women.17 Although many hypotheses have been proposed for the mechanisms of development of metal-induced diseases, such as nickel allergy, there is a high possibility that many relevant mechanisms are yet to be unveiled. One of such possibility may be related to hydrolysis of a peptide bond catalyzed by Ni(II) ions. The Ni(II)-dependent hydrolysis of peptide bond occurs in amino acid sequences Yaa-Ser/Thr-Xaa-His-Zaa (where Yaa and Zaa stand for any amino acid residue and Xaa means any © XXXX American Chemical Society

amino acid residue except Pro). The hydrolyzed bond is located between the Yaa and Ser/Thr residues. The binding of Ni(II) occurs through the formation of a four nitrogen (4N) square planar complex (by the imidazole and amide of histidine, and two preceding amides), which causes the bending of the peptide chain, and places the nucleophilic Ser/Thr hydroxyl group in a proximity of the peptide bond preceding this residue. As a result, the acyl shift takes place with the formation of an intermediate ester product, which subsequently undergoes spontaneous hydrolysis in aqueous solution.18−20 Sequences susceptible to Ni(II)-dependent hydrolysis are present in many proteins, but only some of them can undergo hydrolysis under physiological conditions. We demonstrated that the type of amino acid residue close to the potential hydrolysis site has a significant influence on the hydrolysis rate. This effect is especially relevant for those amino acids that neighbor the histidine residue, where bulky and hydrophobic residues are preferred for the fast reaction.18,19,21 The formation of the initial 4N Ni(II) complex is prerequisite for the hydrolysis to occur; thus, the accessibility of the potential Received: August 19, 2014

A

dx.doi.org/10.1021/tx500337w | Chem. Res. Toxicol. XXXX, XXX, XXX−XXX

Chemical Research in Toxicology

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cleavage site for Ni2+ ions is required. The protein sequences harboring the cleavage sites should therefore be exposed on the protein surface and sufficiently flexible to adopt the square planar structure of the complex.22 However, local bends of the protein main chain may predispose it to such conformation, resulting in the hydrolysis reaction faster than expected on the basis of sequence alone.23 Annexins are proteins that bind phospholipid membranes in a calcium-dependent manner. This binding is performed by their conserved C-terminal domain. The N-terminal domain interacts with other proteins, depending on the annexin type. The specificity of these interactions has an impact on roles played by different annexins.24 Functions of annexin A1 include (1) regulation of the innate and adaptive immune systems,25 (2) participation in the plasma membrane repair system,26 (3) helicase activity,27 and more. Annexin A2 participates in immunological processes.28 It is worth mentioning that the expression of annexin A2 was found to be induced by nickel in human HaCaT keratinocyte line.29 Annexin A8 is a poorly characterized member of the annexin family. It is known to bind F-actin (similarly as annexin A1 and A230) and is associated with late endosomes.31,32 To find out whether human annexins A1, A2, and A8 are potential targets for the toxicity of Ni(II) ions we synthesized peptides that represent fragments of these proteins potentially susceptible to Ni(II)-dependent hydrolysis. We also synthesized a reference peptide Ac-GGASRHWKF-am, with an amino acid sequence corresponding to that of the positive hydrolysis control sequence established in our previous studies. Molecular modeling, UV−vis and circular dichroism (CD) spectroscopies, potentiometry, isothermal titration calorimetry, high-performance liquid chromatography (HPLC), and electrospray ionization mass spectrometry (ESI-MS) were used to characterize Ni(II) binding to these peptides and Ni(II)-dependent peptide bond hydrolysis.

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nm. The CD spectra were recorded on the J-815 CD spectrometer (JASCO) over the spectral range 300−650 nm. For both methods, path length was 1 cm, and samples containing 0.95 mM peptide and 0.9 mM Ni(NO3)2 were titrated with small portions of concentrated NaOH in the pH range 3.0−11.5, at 25 °C. Potentiometry. Potentiometric titrations were performed on a 907 Titrando Automatic Titrator (Metrohm), using a Biotrode combined glass electrode (Metrohm), calibrated daily by nitric acid titrations.35 One hundred millimolar NaOH (carbon dioxide free) was used as a titrant. Samples (1.5 mL) were prepared by dissolving peptides in 4 mM HNO3/96 mM KNO3 to obtain 0.8−1.5 mM peptide concentrations. The Ni(II) complex formation was studied using samples in which the molar ratios of peptide to Ni(II) were 1:0.9, 1:0.45, and 1:0.3. The pH range for all potentiometric titrations was 2.7−11.6. All experiments were performed under argon at 25 °C. Three titrations were included simultaneously into calculations for protonation, and five for Ni(II) complexation. The data were analyzed using the SUPERQUAD and HYPERQUAD programs.36,37 Standard deviations provided by these software and reported here have statistical nature and do not include potential systematic errors. Isothermal Titration Calorimetry. Calorimetric titrations were carried out on the Nano ITC Standard Volume calorimeter (TA Instruments). The sample cell (950 μL) was filled with a peptide solution, and the reference cell was filled with Milli-Q water. The syringe (250 μL) was loaded with a Ni(II) solution. Milli-Q water was degassed under vacuum for 15 min before sample preparation. Furthermore, peptide and Ni(II) solutions were degassed for 5 min before loading into the cell and the syringe. The peptide solutions contained 2 mM peptide, 100 mM H3BO3, and 64 mM KNO3 at pH 9.0. The ionic strength was 0.1 M. The Ni(II) solutions contained 20 mM Ni(NO3)2 and 64 mM KNO3. Typically, 2 μL of the Ni(II) solution was added to the peptide solution at 1200−1500 s intervals using a stirring speed of 250 rpm. To prevent the metal-dependent hydrolysis of peptide bond during the experiments, the measurements were performed at 5 °C. The blank linear functions were calculated on the basis of the last measurement points, where the observed heat flow resulted almost exclusively from the dilution of Ni(II) solution. The blank corrections were made by subtracting values of the blank linear function from the raw data. The data were analyzed by the NanoAnalyze Software (TA Instruments). HPLC Measurements of Hydrolysis Rates. Samples containing 0.8 mM peptide, 1 mM Ni(NO3)2, and 20 mM HEPES at pH 8.2 or 7.4 were incubated at 25, 37, 45, or 60 °C. Twenty microliter aliquots were periodically collected from the sample and acidified by addition of 20 μL of 2% TFA to break down hydrolytic Ni(II) complexes in order to stop the hydrolysis reaction. The products of hydrolysis were separated by HPLC and their identity checked on ESI-MS. Kinetics Analysis. To determine rate constants for the hydrolysis reaction we used the iterative optimization algorithm based on the Levenberg−Marquardt method, which was adapted to minimize the squared distance between the experimental data and theoretical curves obtained from numerical solutions of the corresponding system of the following equations (eqs 1−3):

EXPERIMENTAL SECTION

Materials. N-α-9-Fluorenylmethyloxycarbonyl (Fmoc) amino acids were purchased from Novabiochem (Merck). Trifluoroacetic acid (TFA), piperidine, O-(benzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium hexafluorophosphate (HBTU), triisopropylsilane (TIS), and N,N-diisopropylethylamine (DIEA) were purchased from Merck. Acetic anhydride was purchased from Sigma-Aldrich. TentaGel S RAM resin was obtained from Rapp Polymer GmbH. Molecular Modeling. Crystallographic structures of annexins A1, A2, and A8 (PDB codes: 1MCX, 2HYU, and 1W45, respectively) were taken to simulations performed in Discovery Studio 4.0 Visualizer. Ni(II) ion with a square planar geometry was incorporated to respective potential sites. Conformations of complexes were obtained by geometry optimization utilizing Dreiding-like force field.33 Peptide Synthesis. All peptides were synthesized in the solid phase according to the Fmoc protocol34 using an automatic peptide synthesizer (Prelude, Protein Technology). The syntheses were accomplished on a TentaGel S RAM resin (RAPP Polymere GmbH), using HBTU as a coupling reagent, in the presence of DIEA. Both acetylation of the N-terminus and cleavage were done manually. The acetylation was carried out in 10% acetic anhydride in DCM. The cleavage was done by the cleavage mixture composed of 95% TFA, 2.5% TIS, and 2.5% water. Crude peptides were isolated from cleavage mixtures by precipitation by the addition of cold diethyl ether. Following precipitation, peptides were dissolved in water and lyophilized. Finally, they were purified by HPLC, and their identities were checked by ESI-MS, as described before.19 UV−vis and Circular Dichroism Spectroscopies. The UV−vis spectra were recorded on the LAMBDA 950 UV/vis/NIR spectrophotometer (PerkinElmer) over the spectral range 330−850

dS = − R 4Nk1S(t )(S(t ) + M 0 − S0) dt

(1)

dIP = − R 4Nk1S(t )(S(t ) + M 0 − S0) − k 2IP(t ) dt

(2)

dP = k 2IP(t ) dt

(3)

They were used to calculate the first (k1) and the second rate constant (k2) on the basis of changes in time of substrate (S), intermediate product (IP), and products (P) concentrations depending on the ratio of 4N hydrolytic complexes to the rest of substrate species (R4N) at a given pH, and the initial concentration of the substrate (S0) and metal ions (M0). All calculations were performed in the Wolfram Mathematica 8 environment. B



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The full description of the model used to describe the kinetics of the metal dependent hydrolysis, including approximations that could be used when the concentration of 4N complexes are in excess or in deficiency to the rest of the substrate species throughout the experiment, is available in the Supporting Information.

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RESULTS Analysis of 3D Structures of Annexins A1, A2, and A8. Structures of human annexins A1, A2, and A8 are available in the RCSB PDB database. This allowed us to analyze geometries of potential sites of Ni(II) binding and Ni(II)-dependent peptide bond hydrolysis. However, for human annexin A1, the crystallographic data are available only for Cα atoms of the main chain of the protein (PDB code: 1AIN). The amino acid sequence alignment of porcine and human annexin A1 shows high similarity (89% of identical amino acid residues), with full identity in the regions with potential sites of Ni(II) binding and hydrolysis. Structural alignment of the human (PDB code: 1AIN) and porcine (PDB code: 1MCX) proteins confirms high similarity. Therefore, we used the structure of porcine annexin A1 (PDB code: 1MCX), which includes bound calcium ions. For annexin A2 we chose the crystal structure of human protein complexed with calcium ions and heparin fragments (PDB code: 2HYU). These heparin fragments do not affect the conformation of potential Ni(II) binding and hydrolysis sites (as compared to the respective structure without heparin; PDB code: 2HYW). For annexin A8 we chose the structure of the human protein without calcium ions (PDB code: 1W45), as these ions do not influence the site of potential Ni(II) binding and hydrolysis (as compared to the respective structure with calcium ions; PDB code: 1W3W). Moreover, the 1W45 structure contains a longer fragment of the N-terminus of the protein where the potential Ni(II) binding and hydrolysis site is located. To test the possibility of formation of 4N square planar Ni(II) complexes in the above-mentioned annexins we simulated the Ni(II) binding with the use of geometry optimization utilizing Dreiding-like force field33 in Discovery Studio 4.0 Visualizer. The fragments of proteins containing sites for Ni(II) binding and hydrolysis, together with corresponding simulated complexes are shown in Figure 1. All these sites are located in coils, assuring the flexibility of the polypeptide chain. Accordingly, all simulated complexes were acquired without disturbing large parts of proteins, which means that the process of Ni(II) binding would not need to overcome excessive energy barriers. This analysis shows that the sites are accessible for solutes such as metal ions and can adopt conformations conforming to square planar complexes. These features can promote the binding of Ni(II) ions to potential hydrolytic sites in annexins. In the amino acid sequence of human annexin A1 there are three Yaa-Ser/Thr-Xaa-His-Zaa sites that may potentially bind Ni(II) and undergo Ni(II)-dependent peptide bond hydrolysis. They are 101Thr-Gly-His103, 244Ser-Lys-His246, and 291Thr-ArgHis293 (Figure 1A−F). They are located on the convex side of the molecule. This side binds Ca2+ ions and interacts with phospholipids. The following Ca2+ binding residues are located in the proximity of potential Ni(II) sites: Lys97, Leu100, Glu105, Asp253, Leu256, Gly288, Met286, and Gly290. Although it is not observed in simulated structures, this proximity suggests that the Ni(II) binding can affect Ca2+ binding and vice versa. Interestingly, imidazole groups of His103

Figure 1. Fragments of human annexins A1, A2, and A8 with potential Ni(II) binding sites. Shown are native proteins (A, C, E, G, I) and with Ni(II) ion bound (B, D, F, H, J) as modeled with the use of geometry optimization utilizing Dreiding-like force field33 in Discovery Studio 4.0 Visualizer. Shown are fragments with 101TGH103 (A,B), 244SKH246 (C,D), and 291TRH293 (E,F) from annexin A1, 92SGH94 (G,H) from annexin A2, and 18SSH20 (I,J) from annexin A8. Amino acid residues expected to compose the Ni(II) complex are shown in a stick representation. The sequences studied in this article are violet; Ca, green; Ni2+, yellow; heparin, white sticks.

and His293 residues are 4 Å apart and interact via π−π stacking. Human annexin A2 exists in two isoforms resulting from alternative splicing. The second isoform differs from the canonical one by the longer N-terminus (18 residues). Here, we will use the numbering for the canonical sequence. It contains two Ni(II) hydrolysis sites. The N-terminal 3Thr-Val-His5 site is not resolved in crystal structures, indicating that it is flexible or assumes multiple conformations. Indeed, NMR and CD spectroscopic characterization of the N-terminal fragment of annexin A2 showed that, although it adopts an α-helical conformation in a membrane-mimetic environment, it has random structure in aqueous solution.38 The 92Ser-Gly-His94 site is located on the surface of the convex side of molecule (Figure 1G,H). Adjacent amino acid residues Lys88, Leu91, and Glu96 bind Ca2+ ions. Furthermore, His94 interacts with a fragment of heparin. Thus, the Ni(II) binding is likely to affect interactions of annexin A2 with Ca2+ ions and heparin. The potential Ni(II) binding sequence in human annexin A8, Ser-Ser-His 18 20 (Figure 1I,J) is located on the concave side of the protein, opposite to the Ca2+ binding side. The fragment is on the surface of the protein in a coiled, flexible structure. The above analysis shows that all Yaa-Ser/Thr-Xaa-His-Zaa sites in human annexins A1, A2, and A8 are easily accessible for C



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described by particular models were compared to the pH dependence of absorbance at the maximum of d−d bands of low-spin complexes. We used root-mean-square deviation (rmsd) as a parameter to measure differences between UV− vis spectroscopic parameters and species distributions based on these models. Ni(II) can be coordinated by imidazole ring of histidine residue (1N complex), by the imidazole and two amide nitrogens (3N complex), or by the imidazole and three amide nitrogens (4N complex). Previous research showed that 2N complexes are not observed in hydrolytic peptides containing Yaa-Ser/Thr-Xaa-His-Zaa sequences and that 3N complexes are formed from 1N species through cooperative deprotonation of two amide nitrogens.18 We examined at least two models for each peptide. In the first model we assumed the formation of 1N, 3N, and 4N complexes. In the second model we disregarded 3N complexes. The simulated amounts of 3N complexes, whenever considered, were always low compared to corresponding 4N complexes, and never exceeded 25% of all Ni(II) species. We were able to confirm the existence of 3N complexes in UV−vis spectra only for A1−3, A2−1, A2−2, and A8−1 peptides. The comparisons of Ni(II) binding models considered are presented in Figure S1 of the Supporting Information. The verified Ni(II) binding constants are presented in Table 3. In Figure 2 we present examples of Ni(II) species distributions compared to the absorbance at the maximum of respective d−d bands. At pH above 4, Ni(II) binds to peptides via a His imidazole ring nitrogen (1N complex). Its presence could not be confirmed for A2−1, probably as a result of a lower concentration of this peptide in potentiometric experiments, due to its low solubility. In general, 1N complexes did not exceed 20% of all Ni(II) species, which could also significantly hinder the detection of this complex for the A2−1 peptide. As expected, the UV−vis spectra at pH of the maximum of molar fraction for 1N complexes did not differ significantly from those at pH below 4, which confirms that 1N complexes are high-spin (octahedral), similarly to the Ni(II) aqua ion. The good fit of the absorbance at the d−d band maximum with the sum of 3N and 4N complexes helped us conclude that 3N complexes of A1−3 and A8−1 peptides contain low-spin Ni(II). On the contrary, for A2−1 and A2−2 peptides, the absorbance of Ni(II) complexes at the d−d band maxima corresponds to the sum of 4N complexes rather than the sum of 3N and 4N complexes. Thus, Ni(II) in 3N complexes of A2−1 and A2−2 is high-spin. Low-spin 4N complexes are hydrolytically active species. Their Ni(II) complex formation constants are highlighted in bold in Table 3. It is worth noting that almost every studied peptide can form more than one 4N complex, due to deprotonations of Lys or Tyr residues. The only exception,

Ni(II) ions. Therefore, we decided to characterize the Ni(II) binding and susceptibility to Ni(II)-dependent peptide bond hydrolysis of peptides with sequences taken from the annexins. Sequences of these peptides in the protein context and labels used for them in the text are presented in Table 1. Table 1. Sequences of the Studied Peptides human protein annexin A1 annexin A2 annexin A8

label

amino acid sequence of peptide

position of the histidine residue in the proteina

A1−1 A1−2 A1−3 A2−1 A2−2 A8−1

Ac-KALTGHLEE-am Ac-TKYSKHDMN-am Ac-GVGTRHKAL-am Ac-KMSTVHEIL-am Ac-SALSGHLET-am Ac-VKSSSHFNP-am

103 246 293 5b/23c 94b/112c 20

a

The numbering from protein sequences including the initiator methionine is shown. bThe positions are valid for isoform 1, chosen as canonical sequence from the two produced by alternative splicing. c The positions are from isoform 2.

Ni(II) Complexation. We used UV−vis and circular dichroism spectroscopies, potentiometry, and isothermal titration calorimetry to characterize the Ni(II) binding to annexin-derived peptides. Table 2 presents cumulative Table 2. Logarithmic Protonation Constants for Annexin Peptides Determined at 25 °C and I = 0.1 M (KNO3)a peptide

HL

H2L

H3L

H4L

H5L

A1−1 A1−2 A1−3 A2−1 A2−2 A8−1

10.18(1) 10.61(1) 10.05(1) 9.85(1) 6.66(1) 10.19(1)

16.97(1) 20.99(1) 16.13(1) 16.19(1) 10.98(1) 16.56(1)

21.78(1) 30.34(1)

25.68(1) 36.70(1)

40.31(1)

20.51(1)

a Standard deviations on the least significant digits, provided by HYPERQUAD,37 are given in parentheses.

logarithmic protonation constants of all peptides calculated on the basis of potentiometric titrations. All peptides contain a single His residue, with pK values in the range of 6.1 to 6.8. Most of the peptides also contain a Lys or a Tyr residue, with pK 9.3−10.6, and acidic residues, with pK 3.6−4.8. As potentiometry records only H+ concentration changes upon the metal ion binding, rather than metal specific properties, we utilized UV−vis results to select the correct stoichiometric model of Ni(II) binding and to verify Ni(II) complex formation constants calculated on the basis of potentiometric titrations. The distributions of Ni(II) species

Table 3. Logarithmic Ni(II) Complex Formation Constants for Annexin Peptides Determined at 25 °C and I = 0.1 M (KNO3)a peptide

NiH3L

NiH2L

NiHL

NiL

NiH−1L

NiH−2L

NiH−3L

A1−1 A1−2 A1−3 A2−1 A2−2 A8−1

n.d.b 32.43(3) n.d. n.d. n.d n.d.

n.d. n.d. n.d. n.d. n.d. n.d.

12.61(2) n.d. 12.53(2) n.d. n.d. 12.55(3)

n.d. 9.53(1) n.d. n.d. 2.90(1) n.d.

n.d. −0.08(1) −3.13(1) −4.31(3) n.d. −2.86(1)

−11.50(1) −10.67(1) −11.21(1) −12.11(1) −13.33(2) −10.86(1)

−21.59(1) −21.44(1) −21.61(1) −21.80(1) −21.24(1) −21.10(1)

a

Standard deviations on the least significant digits, provided by HYPERQUAD,37 are given in parentheses. bn.d.; not detected. Values for 4N complexes are bold. D



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Figure 2. Species distributions of Ni(II) complexes of selected peptides: Ac-KALTGHLEE-am (A1−1; left), Ac-KMSTVHEIL-am (A2−1; in the middle), and Ac-VKSSSHFNP-am (A8−1; right), at 25 °C, calculated for concentrations used in UV−vis and CD titrations (0.95 mM peptides and 0.9 mM Ni(II)) using stability constants from Tables 2 and 3. The common scale left-side axes represent Ni(II) molar fractions. Ni(II) species are color-marked as follows: Ni(II) aqua ion, black; 1N complex, red; 3N complex, blue; 4N complexes, green and orange. Pink dotted lines show the sum of 4N complexes, while navy dashed lines show the sum of 3N and 4N complexes. The variable scale right-side axes provides values of absorbance and ellipticity at the d−d band maximum of low-spin complexes: UV−vis absorbance, black circles; CD ellipticity, blue, pink, and violet triangles. Absorbance and ellipticity values are omitted for clarity. Species distributions for other peptides are available in Figure S1 of the Supporting Information.

amide nitrogen involved in the Ni(II) binding in 4N complexes should cause a larger difference in energy between d orbitals than three nitrogen ligands as in 3N complexes. In other words, bands of 4N complexes should be shifted toward shorter wavelengths relative to the 3N complexes, as we indeed observed for A1−3 and A8−1. This confirms that A1−3 and A8−1 peptides form low-spin 3N complexes. Calorimetric titrations were carried out at pH 9.0 to ensure the high percentage of 4N complexes (>98%) and at 5 °C to inhibit the hydrolysis reaction. Thermodynamic parameters of the analyzed process are presented in Table 5. The representative titration is shown in Figure 4. We could not obtain satisfactory results for Ni(II) binding to A2−1 due to insufficient solubility of this peptide. Conditional Ni(II) complex formation constant values for studied peptides are in the range 1.1−6.8 × 104 M−1. To compare binding constants obtained by ITC and potentiometry, we calculated conditional Ni(II) binding constants at pH 9.0 on the basis of potentiometric results. They are 20−40 times higher than those obtained from ITC. This discrepancy could be a result of an interaction between Ni(II) and the buffer. Although we did not detect such interactions in control titrations of borate with Ni(II) ions, there is a literature report by Tilak et al.39 indicating that borate and nickel form a weak Ni(H2BO3)2 complex. Defining the association constant of the Ni(II) borate (B) complex (Kbuff) by eq 4,

the A2−2 peptide contains neither Lys nor Tyr residue and forms only one 4N complex in the studied pH range. The low-spin 3N and 4N complexes are detectable not only in UV−vis but also in CD spectra in the region specific for d−d transitions (350−600 nm). While UV−vis spectra of Ni(II) complexes are very similar to each other for all studied peptides, we observed a variety of CD spectral patterns, especially in terms of the number of extrema (Figure 3). In the range of 300−650 nm, we observed just one minimum and one maximum for peptides A1−1 and A1−3. The CD spectra of Ni(II) complexes of A2−1 and A8−1 peptides have an additional maximum at about 570 nm. A2−2 has one maximum at 456 nm but also two much smaller minima at 385 and 562 nm. For the A1−2 complex we noticed three extrema of comparable intensities. On the basis of potentiometric data, we used the leastsquares calculations to deconvolute UV−vis and CD spectra for each low-spin complex species. Table 4 presents their molar absorption coefficients. For the A1−2 peptide overlapping deprotonations of two Lys and one Tyr residues yielded four 4N species, which could not be deconvoluted reliably. In the course of preliminary calculations we detected that two intermediate species, NiH−1L and NiH−2L, may have very similar spectra and consequently merged them in the calculations. Generally, the spectra of different Ni(II) complexes for the same peptide have similar patterns (see Figures S2 and S3 of the Supporting Information). The exception is the NiL complex of A1−2 for which we distinguished an additional minimum of a very low intensity at short wavelengths. The wavelengths of the maximum absorbance (λmax) of d−d bands for a given peptide differ by less than 4 nm for A1−1, A1−2, and A2−1 peptides, where, according to stoichiometry of complexes, only 4N species are low-spin. For A1−3 and A8−1, where potentiometric calculations indicated that both 3N and 4N complexes should have low-spin character, the shift was 29 and 24 nm, respectively. According to the ligand field theory, an additional

K buff =

[MB2 ] [M ][B]2

(4)

one can easily obtain a formula converting the apparent binding constant of the Ni(II) complexation to the studied peptide obtained from ITC (Kapp) into the conditional binding constant at pH 9 (Kc, eq 5) as K c = K app(1 + K buff × [B]2 ) E

(5)



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Figure 3. CD spectra of low-spin Ni(II) complexes of peptides: Ac-KALTGHLEE-am (A1−1), Ac-TKYSKHDMN-am (A1−2), Ac-GVGTRHKALam (A1−3), Ac-KMSTVHEIL-am (A2−1), Ac-SALSGHLET-am (A2−2), and Ac-VKSSSHFNP-am (A8−1), at different pH values, coded with rainbow colors from red (the lowest pH ∼3) to dark blue (the highest pH 11.5). The spectra of apopeptides, exhibiting no extrema in this spectral range, are marked by dotted gray lines (mostly not visible as they have zero value throughout all the spectra). The titrations were carried out at 25 °C.

We followed the assumption of Tilak et al. that only H2BO3− ions bind Ni(II).39 Taking that pK of H3BO3 at 5 °C is 9.4,40 the log Kbuff explaining the discrepancy between ITC and potentiometric results should be in the range of 4.4−4.7. This is in a good agreement with results of Tilak et al. (log K = 3.8− 4.9).39 Therefore, we can safely use ITC results to compare Ni(II) affinities of the studied peptides. In agreement with potentiometric results, A1−2 and A8−1 peptides have the highest affinity to Ni(II) ions, followed by A2−2 and A1−3 peptides. A1−1 has the lowest affinity for Ni(II) ions, with the value of binding constant more than six times lower than for A1−2 peptide. The observed enthalpy changes (ΔHobs) range between 27.6 and 47.7 kJ × mol−1 (Table 5). They have positive values, indicating that Ni(II) coordination is an endothermic reaction. The highest ΔH was noticed for A8−1, the lowest for A1−1. The value of the heat absorbed after addition of Ni(II) ions to peptide solution is a result of Ni(II) binding to peptides and the sum of energy of all other processes occurring after injection, including dilution and buffer protonation. The heat of dilution was subtracted. The buffer protonation was the result of proton release from peptides upon Ni(II) binding. At pH 9.0 almost all histidine residues are deprotonated spontaneously, so Ni(II) binding only requires deprotonation of three amide

nitrogens. According to literature data,40 the enthalpy of borate protonation at 5 °C is about −18.6 kJ × mol−1. Assuming that released protons react only with the buffer, the contribution of this process is about −55.8 kJ × mol−1. These considerations yield estimated values of Ni(II) binding enthalpy changes (ΔHNi(II) bind) ranging between 83.4 and 103.5 kJ × mol−1 (Table 5). We did not include the possible protonation of lysine or tyrosine residues, as we calculated that it has a minor buffering effect in experimental conditions because of a much higher concentration of buffer (100 mM) from that of the peptide (2 mM). The observed entropy changes (ΔSobs) range between 176.4 J × mol−1 × K−1 for A1−1 and 260.2 J × mol−1 × K−1 for A8−1 (Table 5). The correction of enthalpy change values by the buffer protonation effect led to a corresponding increase in the entropy changes associated with Ni(II) binding (ΔSNi(II) bind) by about 200 J × mol−1 × K−1. It suggests that Ni(II) binding is the entropy-favored reaction under ITC experiment conditions. It is mainly the result of water release from nickel aqua ions after Ni(II) binding to peptide. Hydrolysis of Peptides. All studied peptides undergo Ni(II)-dependent peptide bond hydrolysis. Ni(II) ions selectively cleave a peptide bond preceding serine or threonine residue in Yaa-Ser/Thr-Xaa-His-Zaa sequences. Separation of F



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Table 4. Parameters of d−d Bands of Low-Spin Square-Planar Ni(II) Complexes of Annexin Peptides UV−vis peptide A1−1

Ni(II) species

CD

λmax (nm)

ε (dm3 × mol−1 × cm−1)

λext (nm)

Δε (dm3 × mol−1 × cm−1)

449 519 448 513 391 443 497 559 438 496 559 436 495 562 442 501 422 493 423 494 433 493 568 430 492 568 385 456 562 448 506 569 428 498 574 419 492 573

1.70(4) −0.69(1) 1.79(6) −0.69(3) −0.11(3) 0.57(3) −1.23(8) 0.74(5) 0.54(7) −1.2(2) 0.5(1) 0.60(7) −1.4(2) 0.6(1) 0.72(4) −1.2(3) 1.15(3) −1.49(3) 1.11(4) −1.56(5) 0.7(1) −1.0(2) 0.06(3) 1.1(1) −1.6(2) 0.15(3) −0.05(5) 1.6(2) −0.07(6) 0.9(1) −1.0(1) 0.18(5) 0.74(2) −1.22(3) 0.10(2) 0.84(2) −1.32(3) 0.08(5)

NiH−2L

4N

455

135(1)

NiH−3L

4N

454

140(2)

NiL

4N

457

100(1)

NiH−1L/NiH−2La

4N

457

102(1)

NiH−3L

4N

455

100(2)

NiH−1L

3N

457

129(7)

NiH−2L

4N

432

119(2)

NiH−3L

4N

428

120(2)

NiH−2L

4N

452

93(3)

NiH−3L

4N

448

103(3)

A2−2

NiH−3L

4N

454

121(1)

A8−1

NiH−1L

3N

464

103(5)

NiH−2L

4N

448

105(2)

NiH−3L

4N

440

104(2)

A1−2

A1−3

A2−1

a Since in the result of preliminary calculations we detected that, for A1−2, two intermediate species, NiH−1L and NiH−2L, may have very similar spectra, we merged them in the calculations.

Table 5. Thermodynamic Parameters for Ni(II) Binding to Annexin Peptides Obtained from ITC Experiments in 100 mM H3BO3, I = 0.1 M, at pH 9.0 and 5 °C peptide

Kaobs (M−1 × 104)a

Katheor (M−1 × 104)b

ΔGobs (kJ × mol−1)a

ΔHobs (kJ × mol−1)a

ΔHNi(II) bind (kJ × mol−1)c

ΔSobs (J × mol−1 × K−1)a

ΔSNi(II) bind (J × mol−1 × K−1)c

A1−1 A1−2 A1−3 A2−2 A8−1

1.1(1) 6.8(8) 1.8(2) 1.6(2) 4.3(3)

21.4 131.8 58.9 63.1 100.0

−21.5 −25.7 −22.5 −22.4 −24.7

27.6(6) 32.1(3) 39.7(5) 33.7(7) 47.7(5)

83.4 87.9 95.5 89.5 103.5

176.4 208.1 223.5 202.0 260.2

377.1 408.4 424.2 402.3 460.9

a

The values obtained directly from ITC experiments. bThe values calculated on the basis of potentiometric results. cThe values calculated with consideration of the protonation of buffer.

reactants by HPLC allowed us to detect four species in all cases: a substrate, an intermediate product (IP), and N- and Cterminal final products (see Figures S4 and S5 of the Supporting Information). The identity of these species was confirmed by ESI-MS, but because of the same mass of the

substrate and the IP, a distinction between them required an additional comparison of their differing retention times. We performed the peptide hydrolysis reactions at pH 7.4 and 8.2. The former value was chosen to refer to physiological conditions; the latter, to easily compare our results with those G



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corresponding to the first pH-dependent constant, which we used and presented previously, can be easily obtained by multiplying the first pH independent constant (k1Ind) by the molar fraction of the hydrolytic 4N complex at the beginning of the reaction. From such calculation one can obtain the values of rate constants observed at specific pH, e.g., 7.4 (k17.4) or 8.2 (k18.2). The second step of the reaction, which is the hydrolysis of the intermediate product described by the k2 value, is independent from the Ni(II) species distribution, although it is also pH-dependent. The pH dependences of the first and the second step of the metal-dependent hydrolysis are different. For the first step of reaction, it is associated with the formation of 4N complexes; for the second one, with the concentration of hydroxide ions, which can catalyze hydrolysis of an ester bond in the intermediate product.41 Thus, in the case of the second rate constant (k2) we calculated values valid for specific pH. Because of the low amount of 4N complexes at pH 7.4 (e.g., for A1−1 about 0.16%), we decided to present rate constants calculated on the basis of experiments performed at pH 8.2 because they provide smaller errors of determination of the amount of 4N complexes. Rate constants, k1Ind, k18.2, and k28.2, for the hydrolysis at pH 8.2 are presented in Table 6. At 37 °C, the highest k1Ind and k18.2 values were for A1−3, followed by A8−1. In the group of peptides with the highest k28.2 values, we found A8−1 and A1−2. The lowest k1Ind, k18.2, and k28.2 constants were noted for A2−1 and A1−1. We used the linear Arrhenius plot to describe how temperature influences the rate constants k18.2 and k28.2 (Figure 5). The values of k18.2 and k28.2 (Figure 5), as well as k1Ind, k17.4, and k27.4 (Figure S6 of the Supporting Information), fulfill the Arrhenius equation for all studied peptides. Analyzing the temperature dependence of individual rate constants, we observe no significant differences in slopes between the peptides. However, the values of k1 are more dependent on temperature than k2. The hydrolysis of the ester-containing IP is the rate limiting step of the peptide cleavage for all studied peptides at the experimental conditions. The amount of IP accumulating in the initial phase of all hydrolysis experiments is a combined effect of the rates of the first and the second step of the reaction. Specifically, the higher the k1 in relation to k2, the higher the amount of IP observed. The highest amount of IP was detected for A1−3 (e.g., 85% of the total peptide at pH 8.2 and 37 °C; Figure S7 of the Supporting Information), as its k18.2 was 17.5 times higher than k28.2. The lowest amount of IP was observed

Figure 4. ITC titration of 2 mM Ac-TKYSKHDMN-am peptide (A1− 2), 100 mM H3BO3, 64 mM KNO3, pH 9.0 with 20 mM Ni(NO3)2 and 64 mM KNO3 at 5 °C. The upper plot shows raw data from the experiment; the lower plot shows the absorbed heat (per mol of injectant) in each injection (black dots), with the fitting of the model with assumed 1:1 interaction stoichiometry (green line).

published earlier by us for other amino acid sequences. The amount of 4N complexes was low at pH 7.4, as calculated using potentiometric and spectroscopic results. Under conditions of hydrolysis experiments, the concentration of 4N complexes did not exceed 2% of the total peptide concentration. Furthermore, the product of hydrolysis binds Ni(II) ions much more tightly than the substrate,18 scavenging Ni(II) from the system, so the concentrations of Ni(II) aqua ions and 4N Ni(II) substrate complexes decreased systematically during the experiment. These conditions have a major impact on equations describing hydrolysis used for the analysis of results of our experiments. The rate constant equations used previously are valid for higher pH values, where the degree of complexation is much higher.18,19 In our experimental conditions the hydrolytic 4N complex is not only consumed by the reaction, but its concentration is additionally decreased by a shift of complexation equilibrium accompanying the effective dilution of the reacting system in the course of reaction. In order to accommodate this effect, we developed new equations (given in the Supporting Information), which use potentiometryderived species distribution data and directly yield the maximum at the specific temperature, pH-independent rate constant for the first step of the reaction (k1Ind), corresponding to a 100% formation of the hydrolytic 4N complex. The value

Table 6. First Order Rate Constants Determined for Ni(II)-Dependent Hydrolysis of the Studied Peptides Calculated on the Basis of Experiments Performed at pH 8.2a k (s−1 × 10−5) 25 °C

37 °C

45 °C

peptide

k1Ind

k18.2

k28.2

k1Ind

k18.2

k28.2

k1Ind

k18.2

k28.2

A1−1 A1−2 A1−3 A2−1 A2−2 A8−1

0.321(1) 1.148(2) 21.18(9) 0.944(2) 1.014(5) 11.38(5)

0.081(1) 0.648(2) 5.910(3) 0.169(1) 0.289(1) 3.891(2)

0.031(1) 2.47(2) 0.353(1) 0.145(1) 0.547(3) 2.523(9)

1.941(6) 6.75(2) 100.0(6) 6.04(2) 6.06(2) 56.6(2)

0.492(2) 3.811(8) 27.9(2) 1.082(4) 1.878(6) 19.37(7)

0.126(1) 8.67(4) 1.581(6) 0.455(2) 1.770(6) 10.33(3)

6.27(2) 19.86(2) 480(5) 15.6(1) 16.79(5) 139.3(7)

1.589(5) 11.22(2) 134(2) 2.789(2) 4.781(2) 47.65(3)

0.294(1) 20.32(4) 3.86(2) 0.968(4) 3.467(9) 22.08(8)

a

k1Ind, maximum at specific temperature and pH-independent rate constant for the 1st step of the Ni(II)-dependent hydrolysis; k18.2, the rate constant for the 1st step of the Ni(II)-dependent hydrolysis at pH 8.2; k28.2, the rate constant for the 2nd step of the Ni(II)-dependent hydrolysis at pH 8.2. H



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A1−1 to about 82 times for A2−2. The t1/2P was shortened from about 6 times for A1−1 up to 24 times for A2−2 peptide. At pH 8.2, differences between t1/2S and t1/2P were bigger when compared to the values observed at pH 7.4. The hydrolysis of the reference peptide (Ac-GGASRHWKFam) was the fastest among all of the studied peptides. At pH 7.4, t1/2S and t1/2P were 2.2 times shorter as compared to annexin peptides characterized by the shortest t1/2S and t1/2P. At pH 8.2, these differences were not so big, and t1/2S of the reference peptide was similar to t1/2S of A1−3 or A8−1. The t1/2P for the reference peptide was comparable to t1/2P for A8−1 (Table 7).

■

DISCUSSION Characteristics of Complexes. To characterize Ni(II) complexes, we used methods applied successfully in our previous research on Ni(II) complexes of peptides susceptible to Ni(II)-dependent hydrolysis: potentiometry and UV−vis and CD spectroscopies. We supplemented our research methodology with ITC as an auxiliary method for determination of binding constants and thermodynamic parameters. According to our best knowledge, systematic studies of kinetics of formation of various Ni(II) complexes of His-n (n > 3, where n denotes position in peptide) and His-3 peptides were not performed. Nevertheless, it seems clear that His-n complexes are formed much faster than His-3 complexes. For example, slow reactivity of the order of minutes to hours is observed frequently for the latter, often precluding potentiometric determination of their stabilities. Such behavior was not seen in His-n complexes studied by us, as well as other research groups.18,41−44 In the case of annexin peptides we observed the equilibration of complex formation reactions essentially within the mixing time (few seconds) of the samples. The 1N complexes are octahedral species anchored at the imidazole nitrogen, the only nitrogen donor accessible, due to its low pKa value. We detected 1N, 3N, and 4N complexes. Among 3N complexes we found high-spin species for A2−1 and A2−2, and low-spin (square planar) species for A1−3 and A8−1, judged by the absence or presence of a relatively intense d−d band close to 450 nm. The low-spin 3N complexes have spectra similar to low-spin 4N; however, they are systematically red-shifted by 24 to 29 nm. It is, according to our best knowledge, the first clear-cut account of d−d parameters of low-spin 3N Ni(II) complexes of peptides. We demonstrated that 3N complexes of annexin model peptides are either low- or high-spin. In previous studies on Ni(II) complexes of peptides susceptible to Ni(II)-dependent hydrolysis, also a mixture of high- and low-spin 3N complexes was observed among Ni(II) complexes for the same peptide, e.g., for Ac-GASRHAKFLam.18 The 3N complexes in His peptides are a little mysterious. In many cases they were found to be inexistent or very minor species. When observed in amounts sufficient to characterize their spectroscopic properties, they were found to be high-spin, low-spin, or a mixture of these two cases. In general, the spin state of a Ni(II) complex depends on the relationship between the ligand field strength of the equatorial vs potential axial ligands (e.g., thermochromism and solvatochromism).45−48 The gradual substitutions of water oxygens with peptide nitrogens bring the initially octahedral complexes closer to the spin crossover limit. In complexes involving main chain peptide nitrogen coordination equatorial 1N and 2N complexes are always high-spin and 4N complexes are always low-spin. The

Figure 5. Arrhenius plot of the first rate constants, k18.2 (A), and the second rate constants, k28.2 (B), describing Ni(II)-dependent peptide bond hydrolysis at pH 8.2 for peptides: Ac-KALTGHLEE-am (A1−1), black squares; Ac-TKYSKHDMN-am (A1−2), red circles; AcGVGTRHKAL-am (A1−3), blue triangles; Ac-KMSTVHEIL-am (A2−1), pink triangles; Ac-SALSGHLET-am (A2−2), green squares; and Ac-VKSSSHFNP-am (A8−1), navy triangles.

for A1−2 (e.g., 25% of the total peptide at pH 8.2 and 37 °C), as its k18.2 was even lower than k28.2. As k1 decreased more rapidly than k2 at pH 7.4, when compared to pH 8.2, the amounts of observed IP were smaller, ranging from 7% of the total A1−2 to 42% of the total A1−3, at 37 °C. We also detected a linear temperature dependence of the maximum amount of IP (except A1−3 at pH 8.2; Figure S7 of the Supporting Information), which results from the abovementioned dependences of inverse temperature (T−1) on the ln(k1) and ln(k2). We also determined the half-times of substrate decay (t1/2S) and the half-times of formation of final products (t1/2P) (Table 7) because both of these processes could be important from the Table 7. Half-Times of Substrate Decay (t1/2S) and Formation of Hydrolysis Products (t1/2P) of the Studied Peptides at 37 °C and pH 7.4 and 8.2 t1/2S (h)

t1/2P (h)

peptide

pH 7.4

pH 8.2

pH 7.4

pH 8.2

A1−1 A1−2 A1−3 A2−1 A2−2 A8−1 referencea

1251 168 29 1076 610 45 13

31 4.1 0.6 16 7.5 0.9 0.5

1321 182 109 1295 702 60 28

231 7.7 14 79 24 3.0 2.2

a

The peptide Ac-GGASRHWKF-am, with an amino acid sequence corresponding to that of the positive nickel hydrolysis control sequence established in our previous research.18,19 Values for the fastest processes for annexin peptides in physiological conditions are bold.

toxicological point of view. At pH 7.4 and 37 °C t1/2S and t1/2P were the shortest for A1−3 and A8−1 and the longest for A2− 1 and A1−1. Increasing pH to 8.2 does not change the order of peptides in terms of susceptibility to Ni(II)-dependent hydrolysis. Nevertheless, we observed a significant reduction of t1/2S at pH 8.2 compared to t1/2S at pH 7.4, from about 40 times for I



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chain lowers the basicity of its amine nitrogen down to pKa 7.11−7.5349,51−54 from a typical value of ∼7.8. This effect may also have an influence on proximate amide nitrogens coordinating the metal ion. Mlynarz et al. showed that the lower is the basicity of an amide nitrogen, the higher is the stability of the Ni(II) complexes.55 Altogether, this principle explains the highest stabilities of 3N complexes with the Arg residue. While we observed heterogeneity of spectroscopic properties of 3N complexes, d−d band parameters of 4N complexes for all studied peptides are very similar to each other and consistent with the results for two species of 4N complex (NiH−2L and NiH−3L) of the peptide derived from histone H2A and its analogues42,43 and the peptide derived from the C-terminus of histone H4.44 The protonation-corrected stability constants of the first (at the lowest pH) 4N complexes, *K4N, for annexin peptides, calculated according to eq 6, revealed the highest stability of Ni(II) complexes with A1−2, A1−3, and A8−1 peptides, with log *K4N values −27.17, −27.34, and −27.41, respectively. The remaining log *K4N values are −28.03 for A2−1, −28.30 for A2−2, and −28.46 for A1−1. The range of log *K4N values observed for annexin peptides is similar to those obtained in our previous study on model hydrolytic peptides (from −28.12 to −27.05)18 as well as to the Ni(II) complexes of the peptide derived from histone H2A and its analogues (from −28.58 to −27.26),42,43 and model peptide of C-terminus of histone H4 (−27.92).44 Therefore, the 4N Ni(II) complexes containing imidazole nitrogen and three amide nitrogen donors have rather uniform stabilities (within 1.6 log units). When comparing patterns of CD spectra of Ni(II) complexes of model annexin peptides, we noted that A1−1 and A1−3 peptides have the same pattern as observed previously for AcGASRHWKFL-am, a model hydrolytic peptide,56 and for peptide Ac-GTHS-am.57,58 The patterns of CD spectra of A2− 1 and A8−1 peptides are more similar to that observed for AcTESAHK-am,42 with an additional small positive band at longer wavelengths. The A1−2 peptide has the same CD spectra pattern as that for Ac-TYTEHA-am, the model peptide from Cterminus of histone H4,44 with more pronounced positive band at 550 nm. CD spectra of A2−2 peptide are unusual, as they generally have only a positive band in the measured spectral range. Complex Stabilities and Hydrolysis in the Context of Biology. The nickel-dependent peptide bond hydrolysis was observed in viable cultures of CHO (Chinese hamster ovary), NRK-52 (rat renal tubular epithelium), and HPL1D (human lung epithelium) cells treated with Ni(II) ions for one to seven days.59 The cleaved bond was found to be in histone H2A between Glu and Ser residues in sequence ESHHK. It should be noted that the analytical method used in the cited study allowed us to observe only the histones. The above-mentioned value of −28.58 for the protonation-corrected logarithmic stability constant (log *K4N) for the Ni(II) complex with AcTESHHK-am (a model sequence from histone H2A) was reported.42 The comparison of this value with the respective values for peptides studied here shows that annexin peptides form more stable Ni(II) complexes, compared to the peptide derived from H2A. Furthermore, we calculated that the amounts of 4N hydrolytic complexes for Ac-TESHHK-am peptide at pH 7.4 and for concentrations used in our hydrolysis experiments are significantly (from 5.5 to 65 times, Figure S8 of the Supporting Information) lower than those observed for

3N complexes of peptides studied in this work appear to be near or at the boundary, and so their high-spin (octahedral) or low-spin (square planar) character is determined by minor alterations of electronic density of the third nitrogen in the structure of the complex, that is the amide nitrogen of the residue preceding the His residue. The difficulty of studying this phenomenon stems from the fact that these 3N complexes are usually minor species, present only at low concentrations, due to high cooperativity in the formation of three-ring systems of final 4N complexes, as Hill coefficient for this process was always higher than 2 for all studied peptides. In many of the peptides studied they were not even detected. In order to find the rules for the formation of 3N complexes we calculated protonation-corrected stability constants *K3N according to eq 6:18,49,50 log *KxN = log βNiH

n − xL

− log βH L n

(6)

The βHnL corresponds to the protonation of the histidine residue and the βNiHn−xL is a Ni(II) cumulative stability constant of complexes with x nitrogen atoms coordinating metal ion. The protonation-corrected stability constants enable comparisons of complexes having different protonation stoichiometries. In particular, they allowed us to compensate for the numeric effects of deprotonations of nonbonding Tyr and Lys residues in values of β constants. In Figure 6 we compared log

Figure 6. Protonation-corrected logarithmic stability constants of Ni(II) 3N complexes, log *K3N, and spin states of these complexes. The considered peptides are annexin peptides marked by green squares, Ac-GVGTRHKAL-am (TRHK); Ac-KMSTVHEIL-am (TVHE); Ac-SALSGHLET-am (SGHL); Ac-VKSSSHFNP-am (SSHF); and other, previously studied peptides18,42 marked by black squares, Ac-GASGHAKFL-am (SGHA); Ac-GASAHWKFL-am (SAHW); Ac-GASKHWKFL-am (SKHW); Ac-GASRHAKFL-am (SRHA); Ac-GASRHWKFL-am (SRHW); Ac-GATRHWKFL-am (TRHW); Ac-TESAHK-am (SAHK); Ac-TESHAK-am (ESHA).

*K3N values of high-spin, low-spin, and mixed 3N complexes of annexin peptides studied here, of hydrolytic peptides used in our previous research,18 and of analogues of the model peptide of histone H2A.42 Generally, among 3N complexes, low-spin species are the most, and high-spin are the least stable ones. Complexes with intermediate log *K3N values are a mixture of high- and low-spin species. The exception is AcGASKHWKFL-am, which forms high-spin 3N complex despite a high value of log *K3N.18 The residue preceding His seems to be the most important for the 3N complex spin state, and Arg residue seems to be preferred in this position for low-spin complexes. Noteworthy, it was demonstrated that Arg side J



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(2) influence the stability of the protein, and (3) create epitopes recognized by the immune system as non-native. As mentioned above, annexins regulate the immune system. Thus, malfunctioning of the immune system observed, e.g., in allergy can be induced by any of these three modes of action. Annexin A1 can inhibit phospholipase A2, cyclooxygenase-2, and inducible nitric oxide synthase, enzymes involved in inflammatory processes.66 Furthermore, one of the potential Ni(II) binding sites (244SKH246) in annexin A1 is part of 246HDMNKVLDL254, a sequence that was found to inhibit antigen-induced proliferation of T cells.67 Interestingly, the same site is close to 254LELKGD259 sequence, which is known as a nuclear export signal.68 Finally, all three potential Ni(II) binding sites are in the proximity of Ca2+ binding residues. Thus, one can assume that binding of Ni(II) and subsequent change of the conformation, being a result either of a binding or an acyl shift, as well as hydrolysis of peptide bond in annexin A1 may have an influence on important physiological processes. The N-terminal part of annexin A2, containing potential Ni(II) binding residues 3TVH5, consists of a nuclear export sequence: 4VHEILCKLSL13.69 Furthermore, residues Val4, Ile7, Leu8, and Leu11 participate in the interaction with the S100A10 protein.69 There is a redox active Cys9 residue in the same region. This cysteine residue is reversibly oxidized by H2O2 and reduced by the thioredoxin system; thus, annexin A2 is a probable antioxidant molecule.70 Annexin A2 accumulates in the nucleus in response to oxidative stress, and this is probably one of the means of DNA protection. In the absence of oxidative stress, annexin A2 is exported from the nucleus.70 In this context, it is important to mention that Ni(II) complexes with peptides were found to induce DNA breakage in the presence of H2O2.53,71−73 Thus, binding of Ni(II) to 3TVH5 may have an influence on these processes. The Ni(II)dependent cleavage of peptide bond between Ser2 and Thr3 could also transform isoform 2 of annexin A2 with the longer N-terminal part to the analogue of isoform 1. As mentioned above, another potential Ni(II) binding site, 92SGH94, is close to Ca2+ binding residues. In the result, Ni(II) binding may also impair calcium-dependent functions of annexin A2. The binding of Ni(II) and peptide bond hydrolysis in annexin A8 is probably most relevant for smokers and other humans exposed to high nickel level in the air, as the highest levels of this protein was observed in human lung endothelium.74 The results presented above permit us to consider the toxicological impact of three chemical processes differing in their velocity: (1) Ni(II) binding, (2) formation of the intermediate product, and (3) formation of the final products. Chelation of nickel by amino acid sequences including the His residue in position further than 3 is fast (in the range of seconds, as mentioned above), and its duration is correlated with a short contact of human skin with, e.g., coins. It has to be emphasized that even such short contact is sufficient to elicit allergic reaction.11 However, formation of IP and final products is slower. It seems to correlate with the fact that the nickel allergy is usually linked with type IV hypersensitivity, which is known to develop days after the contact with allergen.75 Perspective. We demonstrated that all peptides from human annexins studied here undergo Ni(II)-dependent hydrolysis. Analysis of 3D structures of annexins A1, A2, and A8 showed that potential hydrolytic sites are accessible for Ni(II) ions and can adopt conformations required for the square planar complex formation in native proteins. The next

model peptides of annexins presented here. Since Ni(II) was shown to bind and cleave H2A in cultured cells, it seems to be likely that the same processes can also occur for annexin A1, A2, or A8. Peptides from histone H2B were thoroughly tested considering binding of Ni2+ ions.60−63 It was suggested that formation of such complexes may change conformation of the whole protein inducing epigenetic and promutagenic effects. However, taking into account relatively weak binding at physiological pH, the significance of these findings remains to be established. Toll-like receptor 4 (TLR4) was proposed by Schmidt et al.64 as a target molecule for Ni2+ ions in nickel allergy. They stated that a Ni2+ ion bridges two monomers of TLR4 leading to its direct activation. However, the relevance of that finding in the context of nickel allergy was called into question in the recent paper of Vennegaard et al.,65 who found that epicutaneous exposure (resembling common challenge of the human skin by nickel from, e.g., jewelry) of mice to nickel can induce allergy independently of TLR4, in contrast to intradermal injections of nickel performed by Schmidt et al.64 This example shows that the mechanisms of nickel allergy are far from being fully elucidated. There is a need for further research to reveal the complete spectrum of pathophysiology of Ni(II). Rates of Hydrolysis. From the physiological point of view, it is more intuitive to compare the amounts of reaction products at pH 7.4 and 37 °C, after, e.g., 24 h (Figure 7),

Figure 7. Molar fraction of an intermediate product (blue) and hydrolysis final products (red) after 24 h at pH 7.4 and 37 °C, for peptides Ac-KALTGHLEE-am (A1−1), Ac-TKYSKHDMN-am (A1− 2), Ac-GVGTRHKAL-am (A1−3), Ac-KMSTVHEIL-am (A2−1), AcSALSGHLET-am (A2−2), and Ac-VKSSSHFNP-am (A8−1).

instead of using values of rate constants. In these conditions more than one-third of the peptide A1−3 with the sequence including His293 of annexin A1 is converted to the estercontaining intermediate product. At the same time, one-fifth of the peptide A8−1 with the sequence from annexin A8 underwent a complete two-step hydrolysis reaction. In the case of peptides from annexin A2, the largest amount of IP (3%) was observed for A2−2. These results suggest that annexins A1 and A8, and to some extent also A2, may be toxicological targets for Ni(II) ions in the context of peptide bond hydrolysis. Potential Impact of Ni(II) Binding and Hydrolysis on Properties of Proteins. The binding of Ni(II) or subsequent hydrolysis can have several different effects; it can (1) change the conformation of the protein and disturb its native function, K



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(6) Schnuch, A., Wolter, J., Geier, J., and Uter, W. (2011) Nickel allergy is still frequent in young German females, probably because of insufficient protection from nickel-releasing objects. Contact Dermatitis 64, 142−150. (7) Darlenski, R., Kazandjieva, J., and Pramatarov, K. (2012) The many faces of nickel allergy. Int. J. Dermatol. 51, 523−530. (8) Hamann, C. R., Hamann, D., Hamann, C., Thyssen, J. P., and Liden, C. (2012) The cost of nickel allergy: a global investigation of coin composition and nickel and cobalt release. Contact Dermatitis 68, 15−22. (9) Thyssen, J. P., Gawkrodger, D. J., White, I. R., Julander, A., Menne, T., and Liden, C. (2012) Coin exposure may cause allergic nickel dermatitis: a review. Contact Dermatitis 68, 3−14. (10) Aquino, M., Mucci, T., Chong, M., Lorton, M. D., and Fonacier, L. (2013) Mobile phones: potential sources of nickel and cobalt exposure for metal allergic patients. Pediatr. Allergy Immunol. Pulmonol. 26, 181−186. (11) Julander, A., Midander, K., Herting, G., Thyssen, J. P., White, I. R., Wallinder, I. O., and Liden, C. (2013) New UK nickel-plated steel coins constitute an increased allergy and eczema risk. Contact Dermatitis 68, 323−330. (12) Svedman, C., Möller, H., Gruvberger, B., Gustavsson, C. G., Dahlin, J., Persson, L., and Bruze, M. (2014) Implants and contact allergy: are sensitizing metals released as haptens from coronary stents? Contact Dermatitis 71, 92−97. (13) Midander, K., Kettelarij, J., Julander, A., and Liden, C. (2014) Nickel release from white gold. Contact Dermatitis 71, 108−128. (14) Mann, E., Ranft, U., Eberwein, G., Gladtke, D., Sugiri, D., Behrendt, H., Ring, J., Schafer, T., Begerow, J., Wittsiepe, J., Kramer, U., and Wilhelm, M. (2010) Does airborne nickel exposure induce nickel sensitization? Contact Dermatitis 62, 355−362. (15) Pappas, R. S., Fresquez, M. R., Martone, N., and Watson, C. H. (2014) Toxic metal concentrations in mainstream smoke from cigarettes available in the USA. J. Anal. Toxicol. 38, 204−211. (16) Williams, M., Villarreal, A., Bozhilov, K., Lin, S., and Talbot, P. (2013) Metal and silicate particles including nanoparticles are present in electronic cigarette cartomizer fluid and aerosol. PLoS One 8, e57987. (17) Thyssen, J. P., Linneberg, A., Menne, T., and Johansen, J. D. (2007) The epidemiology of contact allergy in the general population: prevalence and main findings. Contact Dermatitis 57, 287−299. (18) Kopera, E., Krężel, A., Protas, A. M., Belczyk, A., Bonna, A., Wysłouch-Cieszyńska, A., Poznański, J., and Bal, W. (2010) Sequencespecific Ni(II)-dependent peptide bond hydrolysis for protein engineering: reaction conditions and molecular mechanism. Inorg. Chem. 49, 6636−6645. (19) Krężel, A., Kopera, E., Protas, A. M., Poznański, J., WysłouchCieszyńska, A., and Bal, W. (2010) Sequence-specific Ni(II)dependent peptide bond hydrolysis for protein engineering. Combinatorial library determination of optimal sequences. J. Am. Chem. Soc. 132, 3355−3366. (20) Podobas, E. I., Bonna, A., Polkowska-Nowakowska, A., and Bal, W. (2014) Dual catalytic role of the metal ion in nickel-assisted peptide bond hydrolysis. J. Inorg. Biochem. 136, 107−114. (21) Protas, A. M., Ariani, H. H., Bonna, A., Polkowska-Nowakowska, A., Poznański, J., and Bal, W. (2013) Sequence-specific Ni(II)dependent peptide bond hydrolysis for protein engineering: active sequence optimization. J. Inorg. Biochem. 127, 99−106. (22) Kopera, E., Belczyk-Ciesielska, A., and Bal, W. (2012) Application of Ni(II)-assisted peptide bond hydrolysis to nonenzymatic tag removal. PLoS One 7, e36350. (23) Kurowska, E., Sasin-Kurowska, J., Bonna, A., Grynberg, M., Poznański, J., Knizewski, L., Ginalski, K., and Bal, W. (2011) The C2H2 zinc finger transcription factors are likely targets for Ni(II) toxicity. Metallomics 3, 1227−1231. (24) Gerke, V., Creutz, C. E., and Moss, S. E. (2005) Annexins: linking Ca2+ signaling to membrane dynamics. Nat. Rev. Mol. Cell Biol. 6, 449−461.

step of our research is to react whole proteins with Ni(II) ions, followed by experiments on cell lines. These studies will allow us to verify possible toxicological effects of Ni(II)-dependent hydrolysis on the structure and functions of annexins.

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ASSOCIATED CONTENT

S Supporting Information *

Detailed description of kinetics of the metal-dependent hydrolysis; species distributions of Ni(II) complexes for considered binding models, including rmsd results; deconvoluted UV−vis and CD spectra of low-spin Ni(II) complexes; representative HPLC chromatograms and the time dependence of relative amount of peptide species during hydrolysis; Arrhenius plots for hydrolysis performed at pH 7.4; a temperature dependence of the maximum amount of IP; a comparison of 4N complexes amounts at pH 7.4. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Tel: +48-22-592-5766. Fax: +48-22-659-4636. E-mail: [email protected]. Funding

This study was partially financed by Polish National Science Centre, Grant No. DEC-2011/01/D/NZ1/03501. This work was also supported in part by the project “Metal dependent peptide hydrolysis. Tools and mechanisms for biotechnology, toxicology and supramolecular chemistry”, carried out as part of the Foundation for Polish Science TEAM/2009-4/1 program, cofinanced from European Regional Development Fund resources within the framework of Operational Program Innovative Economy. The equipment used was sponsored in part by the Centre for Preclinical Research and Technology (CePT), a project cosponsored by European Regional Development Fund and Innovative Economy, The National Cohesion Strategy of Poland. Notes

The authors declare no competing financial interest.

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ABBREVIATIONS DCM, dichloromethane; DIEA, N,N-diisopropylethylamine; HBTU, O-(benzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium hexafluorophosphate; ITC, isothermal titration calorimetry; IP, intermediate product; P, products; S, substrate; TIS, triisopropylsilane; TFA, trifluoroacetic acid; TLR4, Toll-like receptor 4; Xaa, any amino acid residue except proline; Yaa, Zaa, any amino acid residue

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REFERENCES

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Human Annexins A1, A2, and A8 as Potential Molecular Targets for Ni(II)

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