Chem. Res. Toxicol. 1996,8,683-692
683
Interactions of Nickel(I1) with Histones. Stability and Solution Structure of Complexes with CH3CO-Cys-Ala-Ile-His-NH2, a Putative Metal Binding Sequence of Histone H3 Wojciech Bal,*?$?§ Jan Lukszo,II Malgorzata Jezowska-Bojczuk,Land Kazimierz S. Kasprzak$ Laboratory of Comparative Carcinogenesis, National Cancer Institute, FCRDC, Frederick, Maryland 21 702,Laboratory of Molecular Structure, National Institute of Allergy and Infectious Diseases, Rockville, Maryland 20852, and Institute of Chemistry, University of Wroclaw, Wroclaw, Poland Received January 30, 1995@
Nickel(I1) compounds are established human carcinogens, but the molecular mechanisms underlying their activity are only partially known. One mechanism may include mediation by nickel of promutagenic oxidative DNA damage that depends on Ni(I1) binding to chromatin. To characterize such binding at the histone moiety of chromatin, we synthesized the peptide CH3CO-Cys-Ala-Ile-His-NH2(L), a model of the evolutionarily conserved motif in histone H3 with expected affinity for transition metals, and evaluated its reactivity toward Ni(I1). Combined spectroscopic (W/vis, CD, NMR) and potentiometric measurements showed that, a t physiological pH, mixtures of Ni(I1) and L yielded unusual macrochelate complexes, NiL and NiL2, in which the metal cation was bound through Cys and His side chains in a squareplanar arrangement. Above pH 9, a NiH-3L complex was formed, structurally analogous to typical square-planar nickel complexes. These complexes are expected to catalyze oxidation reactions, and therefore, coordination of Ni(I1) by the L motif in core histone H3 may be a key event in oxidative DNA base damage observed in the process of Ni(I1)-induced carcinogenesis.
Introduction Ni(II), especially in the form of complexes with certain natural ligands, enhances oxidative damage of DNA bases both in vivo and in vitro, 8-oxoguanine (8-oxo-Gua)l being one of the major products of this process (1-3). 8-Oxo-Gua was shown to be promutagenic (4-7),and its formation may be responsible for nickel carcinogenicity. Reactivity of the Gua residue toward peroxides in the presence of Ni(I1) complexes was studied in vitro in model systems containing 2’- deoxyguanosine (8-11). Although our understanding of the reactions involved is still limited, it is clear that the mechanism of oxidation of guanine is dependent on the stereochemistry of the nickel complex. Square-planar (low spin) complexes, including those containing thiol groups coordinated to Ni(II), generate oxygen-based free radicals, whereas octahedral complexes seem to catalyze the oxidation via nonradical mechanisms (10-14). Exposure of chromatin, unlike pure DNA, to ambient oxygen in the presence of Ni(I1) added as a noncomplexed salt also results in increased DNA base oxidation (15). Modulation of the damage by the protein component of
* Address correspondence to this author at NCI-FCRDC, Building 538, Room 205, Frederick, MD 21702-1201. Tel: 301-846-5738; FAX: 301-846-5946. National Cancer Institute. On leave from the Institute of Chemistry, University of Wroclaw. II National Institute of Allergy and Infectious Diseases. Institute of Chemistry, University of Wroclaw. Abstract published in Advance ACS Abstracts, May 15, 1995. Abbreviations: 8-oxo-Gua, 7,8-dihydro-8-oxoguanine; CAIH, CH3CO-Cys-Ala-Ile-His-NHz; L, CAIH anion; DMF, dimethylformamide; TFA, trifluoroacetic acid; DSS, 3-(trimethylsily1)propionicacid sodium salt; MALDI-TOF MS, matrix-assisted laser desorption ionization timeof-flight mass spectroscopy.
*
@
chromatin clearly indicates complexation of Ni(II), most likely by the histones. The potential role of histone H1 in nickel carcinogenesis has recently been studied (16). The question of specific binding of nickel ions to H1 and other histones has, however, not been addressed. Hydroxyl radical-dependent formation of cross-links between core histones (H2A, H2B, H3, and H4) was found to be a major destructive process in y-irradiated chromatin (17),but participation of particular histones and their functional groups in cross-linking has not been studied so far. Two general modes of interaction of transition metal ions, including Ni(II), with peptides and proteins, exist. The “peptide mode” involves the binding to the terminal groups and deprotonated amide bonds of the peptide backbone (18). The “protein mode” employs only the reactive side chain groups, histidine imidazole and cysteine thiol being most effective in Ni(I1) binding (19). Exceptions to this rule for proteins are very few and always involve a histidine residue in position 2 or 3, e.g., in albumin (20). Therefore, the search for potential nickel binding sites in the histones should be focused on terminal sequences, histidines, and cysteines. Inspection of the available histone sequences (21) revealed several histidine residues; none of them, however, clustered in a possible binding motif, resembling, e.g., a “His zipper” (22). On the other hand, the positions 110-113 of the histone H3 sequence contain the sequence -Cys-Ala-IleHis-, which is evolutionarily strictly conserved among animal species (21). This sequence was suggested as a potential binding site for Zn(I1) as a result of identification of the zinc finger binding patterns (23),but no study seems to have followed. Cys-110 is the only free thiol in H3, and as such has often been employed as a chemical
This article not subject to U.S.Copyright. Published 1995 by the American Chemical Society
Bal et al.
684 Chem. Res. Toxicol., Vol. 8, No. 5, 1995
labeling site (24). Also, Hg(1I) bound to Cys-110 was detected as the sole derivative in the process of heavy atom labeling for the X-ray determination of the structure of histone octamer (25). Having all this information in mind, we decided to synthesize this peptide (CAIH throughout the text for brevity) as a minimal model for the protein sequence, and to characterize its complex formation with Ni(I1) in order to seek possible implications for the process of oxidative damage of DNA and chromatin. This paper presents our first set of experiments, focused on chemical description of complexation. The potential of nickel complexes with CAIH to mediate oxidative DNA damage will be discussed in our next report.
Materials and Methods Materials2 NaOH and HC1 used in spectroscopic measurements were purchased from Fisher Scientific (Pittsburgh, PA). D20, NaOD, DC1, and 3-(trimethylsily1)propionicacid sodium salt (DSS) were purchased from Aldrich Chemical Co. Inc. (Milwaukee, WI). Peptide Synthesis. CH3CO-Cys-Ala-Ile-His-NH2 peptide sequence was assembled by Fmoc strategy (26) on a solid support, using a n Applied Biosystems Inc. (ABI) Model 430A automated peptide synthesizer. The substrates, Na-Fmocprotected amino acids, were obtained from AB1 (Foster City, CA), and amide resin was from Bachem Bioscience Inc. (Philadelphia, PA). Acetylation of the N-terminal was performed using N-acetylimidazole (Sigma Chemical Co., St. Louis, MO) in dimethylformamide (DMF). Cleavage was effected on a 500 pmol scale using a mixture of trifluoroacetic acid (TFA), thioanisole, anisole, and ethanedithiol (90/5/2/3 v/v) over a period of 2 h. The crude peptide (80 mg) was purified by preparative HPLC on the Waters Delta-Pak (Milford, MA) CIS column, 300 A pore size, 15 m m particle size, 19 mm x 300 mm, eluting with 0.1% TFNwater (solvent A) and 0.1% TFN70% acetonitrile/ water (solvent B) using a gradient of 100% NO% B to 80% N20% B over 40 min a t a flow rate of 12.5 m u m i n . Fractions containing product were pooled and lyophilized to yield 29 mg of fluffy, white solid of purity '98% a s assessed by HPLC analysis (detection at 215 nm, Rainin Dynamax (Ridgefield, N J ) CIS, 300 A, 5 mm, 250 x 4.6 mm column, gradient 100% NO% B to 65% N35% B over 20 min, flow rate 1 m u m i n , elution time 12.43 min); MALDI-TOF MS ( m i z = 484.2, calcd L H = 484.6). Potentiometry. The stability constants of H+ and Ni2+ complexes of CAIH in the presence of 0.1 M KNo3 were determined a t 25 "C using pH-metric titrations over the pH range 3-11.5 (Molspin automatic titrator, Molspin Ltd., Newcastle-upon-Tyne, U.K.) with NaOH as titrant. pH changes were monitored with a combined glass-calomel electrode calibrated in hydrogen concentrations by HN03 titrations (27). Sample volumes of 1.5 mL, 1mM CAIH, and CAIH:Ni2+ molar ratios 1.1:l to 2.5:l were used. The data were analyzed using the SUPERQUAD program (28). Standard deviations computed by SUPERQUAD refer to random errors. CD. Spectra were recorded at 25 "C on a Jasco J-500A spectropolarimeter equipped with an IF-500 analog-to-digital interface and controlled by Jasco software operating on a n IBMPC compatible. All spectra were recorded over the range of 190-750 nm, using 1, 0.1, and 0.02 cm cuvettes. Peptide concentrations were 1.0 and 0.1 mM. For complexation experiments, samples with 1 : l and 2:l peptide-to-metal ratios were
+
Certain commercial equipment and materials are indentified in this paper in order to specify adequately the experimental procedure. Such identification does not imply recommendation or endorsement by the National Cancer Institute or the National Institute of Allergy and Infectious Diseases (National Institutes of Health, Department of Health and Human Services, U S . Government) nor does it imply that the materials or equipment identified are necessarily the best available for the purpose.
Table 1. Protonation and Stability Constants of CAIH-Ni(I1) System at Z = 0.1 M (KNo3)and 25 "CU species
1% P
pKa (macro)
pKa (microIb
H2L HL NiL NiL2 NiH-IL NiH-2L NiH-3L
8.59(1) 15.04(1) 4.03(5) 7.98(5) -4.89(9) -13.87(9) -22.85(3)
8.59 6.45
8.4(1) 6.7(1)
8.92 8.98 8.98
a P(Ni,H,Lk) = [Ni,H,LkY[Ni]l[HY[LIk.Standard deviations on the last digit are given in parentheses. Protonation microconstants obtained from NMR in D2O at 27 "C are given for comparison. Not corrected for isotopic effects, see text.
used. The concentration of Ni2+ was kept constant at 1 mM. All samples were prepared and their pHs measured under argon. Spectra are expressed in terms of A6 = €1 - cr. W N i s . Spectra were recorded on a Hewlett-Packard H P 8452A diode array spectrophotometer over the spectral range of 190-820 nm in 1and 0.1 cm cuvettes, using the same samples as i n CD measurements. NMR. lH NMR spectra were recorded at 27 "C with the Varian VXR500S NMR spectrometer (Palo Alto, CA) at 499.84 MHz. DSS was used as internal standard. Concentration of the peptide was 5 mM for the acid-base titration and 1.5 mM for Ni(I1) complexation. In the latter experiment, 2.7:1,2:1, and 1.3:l peptide-to-metal ratios were employed. pH* (pH reading of the electrode not corrected for isotope effects) of the samples was adjusted by adding small volumes of concentrated DC1 or NaOD and monitored with an Aldrich extra long stem electrode inside the NMR tubes. All samples were prepared under argon.
Results Protonation of the Peptide. CAIH contains two titratable groups: the pyrrolic nitrogen atom of the imidazole ring (N-3) on His and the thiol group of Cys. Protonation macroconstants obtained from pH-metric titrations are presented in Table 1. In order to correlate these with microconstants corresponding to molecular events, a series of lH NMR spectra at various pH* values was recorded. A double quantum filtered COSY (DQ COSY) experiment was performed at pH* 12.5 to assign fully the spectra (high pH* was chosen for the optimal separation of lines for the assignment procedure). Figure 1 presents selected 1D spectra of CAIH, compared to spectra of its nickel complexes at the same pH*. Assignments of these spectra are given in Table 2. Figure 2 presents the pH* dependence of selected resonances. These data were used for calculation of protonation microconstants (see Table 1).They show excellent agreement with macroconstants obtained potentiometrically. No attempt was made to correct these values for deuterium isotope effect. Mutual canceling of the isotope effect of the electrode reading and of the effect of deuterium substitution on intrinsic pKa is, however, a well-known phenomenon, and thus the uncorrected results give a reasonable approximation of the situation in the HzO solution (29). Protonation equilibria at His and Cys are independent and well separated (cf. the difference of behavior between His signals and Cys and Ile signals, Figure 2). Interestingly, chemical shifts of a and methyl protons of Ala and Ile are sensitive only to the thiol deprotonation, whereas His resonances show little change above pH* 8. CD spectra of the peptide (Table 3) were recorded over the pH range of 2.0-12.0. The spectra recorded between pH 2.0 and 7.5, i.e., covering the fully protonated as well as His-deprotonated form, are essentially unchanged.
Ni(II) Binding to Histone H3 Motif
Chem. Res. Toxicol., Vol. 8, No. 5, 1995 686
A
C
L D
8
7
6
5
A
4
3
2
1
II 0
PPm
7
Figure 1. The 500-MHz l H NMR spectra of CAIH and CAIH-Ni(I1) complexes in DzO at 27 "C: ( A ) , C a !pH* 8.35. (€3) CAIH: Ni(I1) = 2.3:1, pH* 8.35. (C) CAIH, pH* 12.5. (D) CAIH:Ni(II) = 2.3:1, pH* 12.5. For detailed description c . Table 2. Large increases of the signal intensities as well as bandshifts occur in high pH, thus qualitatively supporting the pattern of changes seen in the NMR spectra. Unfortunately, a very high far-UV absorption prevented recording the CD spectra of the fully deprotonated CAIH below ca. 200 nm. Therefore, only a qualitative indication of an apparent intense positive band beyond 202 nm is given in Table 3. Complex Formation. Coordination of Ni(1I) to CAIH was studied with the use of potentiometry and W/vis, CD, and NMR spectroscopies. Preliminary tests revealed that a red-brown precipitate forms at higher concentrations above pH 7.5, and seems to redissolve partially above pH 10. Therefore, concentrations of Ni(1I) not higher than 1 mM were used in all subsequent experiments. Solutions containing nickel chloride and the peptide were turning yellow above pH 6.5, a phenomenon indicative of the formation of low spin, square-planar complexes in Ni(I1)-peptide interactions. Unexpectedly, after adding nickel ions to peptide solutions or adjusting
pH of Ni(I1)-CAIH mixtures, systems equilibrated rapidly (typically within a minute, as monitored by pH change and spectroscopicmeasurements), whereas squareplanar nickel complexes with other peptides are known to equilibrate more slowly (18,30). Table 1 presents stoichiometry of the complexes and their stability constants. Figure 3 presents speciation curves computed for conditions of typical spectroscopic experiments. Figure 4 presents selected electronic absorption and CD spectra of CAIH complexes with Ni(I1) over the visible and n e a r - W range. Inspection of the visible range of CD spectra clearly reveals the existence of two major types of complex species: one present at neutral to weakly alkaline pH, having very weak Cotton effects in the d-d range, and another in strongly alkaline pH, with intense d-d bands. Spectral parameters of particular complex species, calculated on the basis of species distribution plots, together with spectral band assignments, are presented in Table 4. A sample containing 1 mM CAIH and Ni(I1) at pH 8.2 was diluted
Bal et al.
686 Chem. Res. Toxicol., Vol. 8, No. 5, 1995 Table 2. NMR of CAIH and Its Ni(I1) Complexes in DzO at 27 "C (A) NiL2 Complex Compared to Free CAIH a t pH* 8.35 6 (ppm), 6 (ppm), complex, 8 (ppm), complex, set A (sharp) set B (broad) peptide
amino acid
assignment
CYS
Ca H CP H2 N-ACH3
4.338 2.874 2.039
1.3'~~ 0.8'~~ 2.056
Ala
Ca H CP H3 Ca H CP H CYHa cy Hb Cd H3 Cy' H3
4.319 1.372
4.362 1.350
4.019 1.78 1.39 1.12 0.809 0.694
C2 H C4 H Ca H CP Ha CP Hb
7.687 6.997 4.593 3.137 3.043
Ile
His
Ad Aa (ppm)
Ad Bb (ppm)
-3.0 -2.0 +0.017
f0.101
+0.043 -0.022
0.0 $0.003
4.104 1.80 1.40 1.12 0.834 0.821
e e 2.14d 4.31gd 1.375d 4.180 1.80 1.40 1.12 0.732 0.678
+0.095 +0.02 fO.O1 0.0 +0.025 +0.127
f0.161 f0.02 $0.01 0.0 -0.077 -0.016
7.683 6.967 4.662 3.182 2.937
8.3EId 7.12d 4.617 3.121 3.018
-0.004 -0.03 +0.069 +0.045 -0.106
+0.667 +0.153 f0.024 -0.016 -0.025
(B) NiH-sL Complex Compared to Free CAIH a t pH* 12.5 peptide
complex
amino acid
assignment
6 (ppm)
CYS
Ca H CP Ha CP Hb N-ACH3
4.216 2.863 2.848 2.037
7.0 (CP Ha), 7.5 (CP Hb) 14.8 (CP Hb), 7.0 (Ca H) 14.8 (CP Hb), 7.5 (Ca H)
3.597 1.082d 1.082d 2.154
g g g
-0.619 -1.781 -1.766 +0.117
Ala
Ca H CP H3 Ca H CP H Cy Ha cy Hb C6 H3 CY' H3
4.316 1.418
7.3 (CP H3) 7.3 (Ca H)
3.800 1.350
6.6 (CP H3) 6.6 (Ca H)
-0.516 -0.068
3.962 1.78 1.41 1.13 0.810 0.587
9.3 (CP H) 9.3 ( C a H), 6.7 (Cy' H3Y 7.4 (C6 H3)h 7.4 (Cd H3Ih 7.4 (Cy H) 6.7 (CP H3)
4.105 1.82 1.52 1.15 0.901 0.82'
7.6 (CP H) 7.6 (Ca H)* 7.3 (C6 H3p 7.3 (C6 H3Ih 7.3 (Cy H) g
+0.143 +0.04 +0.11 $0.02 +0.091 f0.23
C2 H C4 H Ca H CP Ha CP Hb
7.665 7.030 4.589 3.172 3.086
1.0 (C4 H) 1.0 (Cz HI, 0.8 (CP Ha) 5.1 (CP Ha), 10.2 (CP Hb) 5.1 (Ca H), 14.8 (CP Hb), 0.8 (cq H) 10.2 (Ca H), 14.8 (CP Ha)
7.770 7.139 4.661 3.116 1.128
6.1 (CP Ha), 8.0 (CP Hb) 6.0 (Cb Hb)d 6.0 (Cb H a y
+0.105 +0.109 +0.07 -0.056 -1.958
Ile
His
J (Hz)
6 (ppm)
J (Hz)
A@
G(comp1ex set A) - &peptide). d(comp1ex set B) - &peptide). May be interchanged. Very broad peak (vu2 '10 Hz). e Signals not detected (broadened out). f d(comp1ex) - &peptide). g Coupling not resolved. Coupling partially resolved. Assignment tentative, based on signal integration. J Partially obscured by HDO signal, coupling lost due to water presaturation.
stepwise to the final 0.1 mM with water, and the 335 nm band was followed in order to investigate the possibility of the existence of oligomeric complexes. The dependence of intensity of this band on concentration is compared to the theoretical curve computed from stability constants in Figure 5. The pH* values for the NMR spectra were chosen to approach maximum formation of the major species evidenced by other methods. All spectra recorded at lower pH* show line broadening, but this effect is absent from the higher pH* spectra. Composite spectral patterns including up to four sets of peaks were observed at high pH* and ascribed to partial oxidation and decomposition of CAIH in the course of measurements (Table 2). In order to identify signals of decomposition products, NMR spectra of samples exposed to air and alkali were recorded (data not shown). Examples of complex spectra are given in Figure 2.
Discussion Values of protonation constants obtained from pHmetric titrations of CAIH are well within the range typical for peptides. As noted in the Results section, the
deprotonations of His and Cys residues are clearly independent, and pH ranges for these processes practically do not overlap. The NMR titration curves did not reveal any systematic deviation from the behavior described by simple pKa formula. Therefore, an interaction between these groups can be excluded, and macroconstants are equivalent to microconstants: the lower for the imidazole nitrogen and the higher for the thiol. On the other hand, the analysis of relations between the pH* and positions of signals in the NMR spectra of the peptide suggests the existence of two moieties of the CAIH molecule: only His protons appear sensitive to the imidazole deprotonation, whereas Cys, Ala, and some of Ile protons cotitrate with the thiol. This behavior indicates a conformational relationship in the molecule involving the CAI moiety. A support for this observation comes from CD spectra. Those recorded in acidic or neutral solutions are typical for short, unordered peptides, although a negative shoulder at 221 nm may indicate some ordering (31). Deprotonation of the Cys thiol results in a major change of the spectral pattern, probably due to the sulfur-centered transitions. Such a short peptide would not be expected to possess a rigid structure in aqueous solution. An increase of intensity
Ni(IZ) Binding to Histone H3 Motif
Chem. Res. Toxicol., Vol. 8, No. 5, 1995 687 A
1
1
His C2H 76 7.6
4 84.6 4.44.2DH
4.0-
His p-1 3.1 3.0
\
.-
E
LL
"'I
6
1.3
0.9-
-
"
(3
0.5
l
i
1
1
I Y
1
1
1
1
l
1
l
peptide form
A
A6
HzL
221 (sh)b 198 221 (shIb 198 213 (shIb 202
-5.1 -27.0 -5.1 -32.3 -13.3
HL L
1
+
+c
(sh) denotes a a A€ units are dm3 cm-l (mol of peptide)-'. denotes a very strong positive band shoulder on the spectrum. c (see text).
++
of the band at 213 nm may, however, indicate a bent conformation (31, 32). A possibility of a conformational effect of thiol deprotonation on the peptide backbone is also suggested by changes in chemical shifts of the a protons. They are accompanied by specificincrease of shielding on terminal methyl groups (6, y? of the Ile residue (Figure 21,whereas other Ile signals show little effect. The upfield shifts of Ile-6 and Cys-NAc protons are small and similar (0.04 and 0.03 ppm, respectively), as opposed to a large effect on Ile-y' protons (0.2 ppm). Such a result strongly suggests a change of hydrophobic packing of the Ile side chain, rather than an overall increase of electronic deqsity due to the increased electronic charge on Cys. This conformational potential of a fairly short peptide
7
I
NIL.
\
0.54
8
9
10
11
12
PH Figure 3. Species distribution diagrams for mixtures of CAIH and Ni2+. (A) 1 mM CAIH and 1mM Ni2+.(B) 2 mM CAIH and 1 mM NP+.
sequence, apart from being interesting on its own, may contribute to coordination and reactivity and thus warrants further research. Electronic absorption and CD spectra help to identify the coordination geometry and binding mode of nickel complexes with thiol-containing peptides through the analysis of d-d and charge transfer (CT) transitions (3335). At the 1:l ratio, complexation manifests itself in the electronic spectra with the appearance of an absorption band around 335 nm, associated with a sulfur to nickel CT (34, 36). This band is discernible at pH as low as 6.5, and its intensity rises up to pH around 8, accompanied by the appearance of relatively intense d-d bands at 430 and 500 nm (Figure 4,Table 3) that are typical for low spin Ni(I1). The same charge transfer band is seen in the CD spectra. However, instead of corresponding d-d bands of a magnitude k2-3 AEunits that would be expected for square-planar, sulfur-coordinated nickel peptide species (33-35), weak (A€ values of the range of 2~0.2)and poorly resolved features are present. Although they qualitatively resemble patterns observed for high spin octahedral nickel complexes with peptides (37, 38), they are unlikely t o be of such origin. Cotton effect magnitudes require that such an octahedral species be a major complex form in solution. This is contrary to the absorption spectra, and also N M R results (see below). Rather, the spectral pattern should be assigned to a distorted diamagnetic complex (39). At the 2:l ratio, the complex formed a t neutral pH easily precipitates from the solution as red-brown powder. This strongly indicates the lack of charge on the complex species and, therefore, supports the binding of
Bal et al.
688 Chem. Res. Toxicol., Vol. 8, No. 5, 1995 A
b
30001
0.5
,
"
\\'\
\
10ooj
I
0 ) , I 400 450 500 550 600 650 700
,p\,-\
\
,
-. 0 200
B
I
250
300
350
400
450
500
550
600
700
650
57
I
0.1
I
I
q \\--$+(,
-0.1 /'
-0.2 400 450 500 550 600 650 700
200
250
300
350
400
450
500
550
600
650
700
h [nml
- - - - - - pH = 6.55 -.-.-
pH = 8.07
----
pH = 9.23
PH = 7.50 -..-..-
-pH = 11.50
Figure 4. Spectra of solutions containing 1 mM CAIH and 1 mM Ni2+a t various pH. (A) UV/vis. (B) CD.
two deprotonated thiol donors. The pH dependence of the CD pattern closely resembles the one for 1:l ratio, the only difference being a slight hindrance of formation of the high pH CD pattern. All these observations fit very well with potentiometric results. Species distribution diagrams (Figure 3) allowed assignment of the optical activity in neutral and weakly alkaline solutions to NiL and N i L complexes, both present in all solutions used for spectroscopic analysis. Examination of their distribution vs pH allowed calculation of the spectral parameters presented in Table 4. Metal ion centered (dd) bands were not assigned due to the effective symmetry of the system lower than D 4 h . The ligand-to-metal CT bands of low spin Ni(I1) complexes can be qualitatively analyzed with the use of the model developed for Cu(I1) thiolate complexes (36) due to similarity of their electronic structures. The main difference is that the O!,Z-~Z orbital of Ni(I1) is much more destabilized, and so all bands are blue-shifted. According to this model, further supported for Ni(I1)thiol complexes by Kozlowski et al. (34, 35), three S Ni(I1) CT bands in addition to N Ni(I1) bands can be observed in the CD spectra. Spectroscopic features presented in Table
-
-
4 can be explained by the following model: the NiL and
NiLz are low spin planar complexes containing nickel ion bound through the deprotonated Cys thiol and a nitrogen of the imidazole ring. The geometry of these complexes is distorted from regular square due to sterical crowding between the thiol and the imidazole ring, a phenomenon similar to those observed previously for other complexes with His-containing peptides (37,40,41).The d-d bands of this species would be expected to have a very weak CD due t o the existence of only the large macrochelate loop and the resulting conformational nonrigidity (42). In such a situation, the asymmetric effects of the electronic density localized on chirality centers of the peptide molecule effectively cancel out. The ground states of charge transfer transitions are, however, located primarily on the peptide molecule, and thus the chirality transfer is expected to be much more effective, allowing the moderate rotational strength in those transitions. The NMR spectra in neutral and weakly alkaline solutions at CAIH:Ni(II) ratios between 1.3:l and 2.7:l reveal the presence of two distinct sets of spectral lines, as presented in Table 2A (sets A and B). None of these sets correspond to the free peptide. Analysis of species
NiUI) Binding to Histone H3 Motif
Chem.Res. Toxicol., Vol. 8, No. 5, 1995 689
Table 4. W N i s and CD Parameters of Ni(II)-CAIH Complexeg uvlvis CD species NiL
NiL2
NiH-3L
i
E
500 (sh)b 415 (sh)b 336
280 740 5200
500 (shIb 410 (sh)b 328
510 (shIb 445 285
300 720 5700
70 120 3000
i
A€
'700 550 480 430 (sh)b 348 276 250 (shIb
-0.28 -0.16 f0.20 -0.20 -1.95 +LOO -0.40
'700 560 480 430 (shIb 343 278 250 (sh)b
-0.21 -0.20 +0.15 -0.15 -2.48 $1.80 -0.40
S, S, S,
541 421 307 273 238
-1.32 -1.30 -0.44 +2.65 +4.2
d-d (A) d-d (E) S, Ni(I1) S, Ni(I1) S, Ni(I1)
assignment d-d d-d
S
-
Ni(I1)
d-d (A) d-d (E) S Ni(I1)
-
d-d (A) d-d (E) S Ni(I1)
-
assignment d-d d-d d-d d-d
S, S, S-,
Ni(I1) Ni(I1) Ni(I1)
+ N,,
-
Ni(1I)
+ N,
-
Ni(I1)
d-d d-d d-d d-d
--Ni(I1) Ni(I1)
--
Ni(I1)
+ N- - Ni(I1)
units are dm3 cm-l (mol of nickel)-l. N,,, N- denote imidazole and deprotonated peptide nitrogens, respectively. S,, a E and A6 = €1 S, denote u and n orbitals of thiolate sulfur, respectively. (sh) denotes a shoulder on the spectrum. Experlmental and simulated values of A,
0
1
0.5
mM Ni(li)
250
300
350
400
450
500
550
600
tnml Figure 5. Effect of dilution of the equimolar mixture of CAIH and Ni2+ on UV/vis spectra. Comparison of experimental values of with those calculated from a simulation based on potentiometric results.
A335
distributions indicates that free peptide, Ni2+,NiL, and N i b are present in all solutions studied. The existence of only two sets of lines thus indicates the fast (in terms of the NMR experiment time frame) exchange case. Observed positions of lines result from a dynamic equilibrium between several species and cannot be ascribed to particular complexes. It is probable that the broader set B corresponds to the exchange of the peptide between L and NiL (and thus undergoes more dipolar broadening from Ni2+1, and set A to the exchange between NiL and NiL, but they cannot be assigned unambiguously from available data. Nevertheless, information on the binding modes can be obtained. Both sets indicate the involvement of the Cys thiol in Ni(I1) binding. Extensive broadening and very large shifts of signals of Cys a and
/3 protons are observed for set A, and these signals are completely lost in set B. These effects result from the electronic interaction with the Ni(I1) ion bound to sulfur. Positions of His CZand Cq resonances are hardly affected in set A, as opposed to set B. This might indicate a monodentate binding (Cys only) of the second CAIH molecule in NiL2. However, analysis of stability constants suggests that the second molecule of peptide binds as efficiently as the first one. Statistical considerations of binding of bidentate ligands to a square-planar metal ion yield the expression: 2 log /3101 - log 8102 = log 4, i.e., 0.6, for noninteracting ligands with no steric hindrance. For CMH-Ni(I1) complexes the value of this expression is only 0.08, thus indicating an actual increase of affinity of the second molecule by ca. 0.5 log unit. This would
Bal et al.
690 Chem. Res. Toxicol., Vol. 8, No. 5, 1995
I
li
\ I
V
I
Mtf
0.25
6
B
H
! ? i
0 S-Ni-N
Figure 6. Proposed structures of different complexes formed by Ni(I1) with CAIH. (A) NiLz complex. (B) NiH-3L complex.
not be possible for a monodentately bound ligand. Involvement of a binding mode different from (Cys, His) is also impossible on stoichiometric grounds, as this would require additional deprotonations of the peptide molecule. The apparent lack of shift on His CZ and Cq in set A probably results from the complex dynamics of the exchange process. It may indicate that breaking of the Ni-S bond rather than the Ni-N bond is a rate-limiting step. This should be expected by taking into account that dissociation of the thiol would involve the low spin to high spin transition on Ni(I1). Effects on other parts of the peptide molecule are relatively minor, and most Ad values are within 0.1 ppm, indicating that the conformations at Ala and Ile are not much different between free and bound CAIH molecules. The structure of the NiLz complex consistent with the above features is presented in Figure 6A, and the structure of the NiL complex can be derived from this one by substituting one of the CAIH molecules with coordinated water. Around pH 9, a major rearrangement of the complex occurs, with the vanishing of the 335-350 nm band and appearance of strong CD in the d-d region. In NMR, formation of a new species characterized by sharp resonance lines was observed, and the spectrum of the free peptide reappeared. These phenomena correlate with the cooperative deprotonation of three amide nitrogens and formation of a NiH+L complex. Such cooperative processes (all peptide protons leave with very similar pK, values, see Table 1) are typical for Ni(I1). The formula NiH-3L indicates the presence of the chelate ring system resembling those formed by Ni(I1) with peptides like Gly-Gly-Cys or Ala-Ala-Cys (34, 351, although the Cys residue of CAIH is in the initial rather than terminal position of the peptide chain. The binding through the (S, 3 x N-) donor set is supported by CD spectroscopic results (see Table 4). The N M R spectra at very high pH present a complex pattern most likely because of rapid and complicated degradation processes, including oxidation of the Ni(I1)CAIH molecule by traces of molecular oxygen and possible hydrolysis of peptide bonds. Signals of these
7
6
9
10
11
12
PH
Figure 7. Species distribution diagram calculated for hypothetical “nucleus-like’’concentrations: 10 mM CAIH and 1mM N?+.
products can be seen in the spectrum presented in Figure 2D. Analysis was facilitated by identifying signals of degradation products in samples exposed to ambient oxygen and/or high pH for several hours and by integration of the spectra. In this manner consistent assignments were obtained and are presented in Table 2B. The presence of free peptide lines indicates slow exchange, as would be expected for a tightly coordinated, inert complex. Detailed analysis of chemical shifts also suggests that nickel in NiH-3L is bound to sulfur and to amide nitrogens of Ala, Ile, and His residues, leaving the imidazole unbound. The major piece of N M R evidence for this conclusion is a small Ad on the CZproton of the imidazole: ca. 0.1 ppm instead of ca. 0.4-0.6 observed for imidazole-bound nickel in other histidyl peptide complexes with slow exchange (37, 43). Moreover, no change of chemical shift for those protons was observed upon increasing pH* up to ca. 13. Such a change, indicative of deprotonation of the NB atom, would be expected for a nickel-bound but not for a free imidazole ring (18). Inspection of the effect of coordination on chemical shifts indicates a complicated interplay of electronic and structural effects in the CAIH molecule; e.g., a very strong upfield shift of one of the His$ protons suggests its axial position relative to coordination plane (44). Generally speaking, however, these effects are within normal range for Ni(I1) peptide complexes with fused chelate rings (34,35,37,43-45). The structure of the NiH-3L complex is presented in Figure 6B.
Conclusions The results presented above show that in the physiological pH range CAIH binds Ni(I1) through thiolate and imidazole donors, forming slightly distorted squareplanar NiL and NiLz complexes. This binding mode is likely to occur with the whole histone H3, from consideration of pH range of existence and rapid kinetics of formation. Although these complexes are not very stable at concentrations and molecular ratios suitable for their determination, they may be very important in conditions more similar to those of cell nucleus, i.e., at high abundance of histone and low concentration of Ni(I1). Figure 7 presents such a hypothetical species distribution, in which the NiLz complex is the major species at physiological pH. It should be, however, noted that the formation of the NiL or NiLz complex with the histone
Ni(ZZ) Binding to Histone H3 Motif
odamer remains to be proven. Furthermore, the binding to the intact protein can be expected to be more tight, due to multiple nonbonding interactions available there (e.g., formation of a hydrophobic pocket). On the other hand, particular electrostatic conditions of the highly positively charged protein might enhance the accessibility of the NiH-SL binding mode. Planar complexes of Ni(I1) are known to be very reactive toward DNA in the presence of molecular oxygen or hydrogen peroxide. We are currently investigating this reactivity using CAIH as an introductory model. We are also continuing our study with the use of histone H3 and the nucleohistone complex.
Acknowledgment. The authors would like to thank Mr. John R. Klose for recording the NMR spectra and Dr. Gwendolyn N. Chmurny (both of PRI, NCI-FCRDC) for processing and analyzing the DQ COSY spectrum, Drs. Joseph E. Saavedra and Lucy M. Anderson for helpful comments on the manuscript, and Dr. Anthony Dipple for making his CD spectrometer available to us.
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