DOI: 10.1002/chem.201404546
Full Paper
& Bioinorganic Chemistry
Engineering Short Peptide Sequences for Uranyl Binding Colette Lebrun,[a, b] Matthieu Starck,[a, b] Vicky Gathu,[a, b] Yves Chenavier,[a, b] and Pascale Delangle*[a, b]
Abstract: Peptides are interesting tools to rationalize uranyl–protein interactions, which are relevant to uranium toxicity in vivo. Structured cyclic peptide scaffolds were chosen as promising candidates to coordinate uranyl thanks to four amino acid side chains pre-oriented towards the dioxo cation equatorial plane. The binding of uranyl by a series of decapeptides has been investigated with complementary analytical and spectroscopic methods to determine
Introduction Uranium is a natural element widely found in the environment, due to both natural occurrence in mineral ores or in sea water and industrial applications. In particular, it is a key element for energy production through nuclear fission. This heavy metal belongs to the actinides series and has no essential role in living organisms from plants to man. Uranium displays radiological and chemical toxicity to living organisms.[1, 2] Whatever the route of entry, uranium reaches the blood and then targets mainly the kidneys and the bones. However, despite significant recent advances in the field, there is still a serious lack of knowledge about the molecular interactions responsible for uranium toxicity. The underlying mechanisms need to be unraveled to predict the effect of uranium on living organisms and also to help in designing efficient detoxification agents. Indeed, such chelating agents are essential tools in case of dirty bombs or the accidental release of uranium in the environment.[3] Uranium exhibits several redox states, but the most stable and relevant form in aqueous solutions and in physiological conditions is the hexavalent uranyl cation, UO22 + , which is a rare form of linear trans-dioxo cation. The presence of the two oxo groups favors the coordination of four to six extra ligands in the equatorial plane, which is perpendicular to the OUO bonds. The uranyl ion is classified as a hard Lewis acid [a] C. Lebrun, Dr. M. Starck, V. Gathu, Y. Chenavier, Dr. P. Delangle Univ. Grenoble Alpes, INAC, SCIB 38000 Grenoble (France) [b] C. Lebrun, Dr. M. Starck, V. Gathu, Y. Chenavier, Dr. P. Delangle CEA, INAC, SCIB 38054 Grenoble (France) E-mail: [email protected] Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/chem.201404546. Chem. Eur. J. 2014, 20, 16566 – 16573
the key parameters for the formation of stable uranyl–peptide complexes. The molar ellipticity of the uranyl complex at 195 nm is directly correlated to its stability, which demonstrates that the b-sheet structure is optimal for high stability in the peptide series. Cyclodecapeptides with four glutamate residues exhibit the highest affinities for uranyl with log KC = 8.0–8.4 and, therefore, appear as good starting points for the design of high-affinity uranyl-chelating peptides.
and is highly oxophilic with high affinity for hard oxygen donors. Therefore, the preferred binding sites of uranyl in proteins involve carboxylate donors from aspartates, glutamates, or the carboxyl terminus of the peptide chain.[4] Two abundant serum proteins, namely albumin and transferrin, have been extensively studied as uranium targets in vivo.[2, 5, 6] However, other significant uranyl target proteins have been identified over the past decade by screening human serum proteins for uranium binding,[7] thanks to a combination of in silico approaches[8] and analytical tools.[9] Two proteins demonstrated a particularly large affinity for uranyl. Fetuin, despite its low concentration, could be one of the major target proteins of uranyl in the serum,[10] and the highly phosphorylated protein osteopontin, involved in the organo-mineral homeostasis of the bone, also binds tightly with uranyl.[11] Peptides are frequently used as simple and efficient tools to study metal–protein binding sites and mimic biological molecules such as metal transporters[12, 13] or metal-responsive transcriptional factors.[14] In our laboratory, pseudopeptides based on chemical scaffolds functionalized with amino acids have also proven to be excellent models of metal detoxification proteins.[15] Peptide derivatives may also be powerful metal chelators[16] with potential applications in detoxification or excellent models of enzyme activity.[17] Some peptides have been applied to uranyl complexation with sequences found in albumin,[18] osteopontin[19] or in calcium-binding proteins such as calmodulin[20, 21] to identify the chelation sites and their affinity for uranyl. Interestingly, protein engineering either from nickelbinding sites or from de novo sequences provided very interesting tools for uranyl chelation with large affinities.[22] In this study, we propose to take advantage of structured cyclodecapeptides that form well-defined uranyl complexes to rationalize peptide–uranyl interactions. Indeed, de novo cyclodecapeptides, which incorporate two prolylglycine sequences as b-turn inducers, demonstrate a backbone conformation in
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Full Paper antiparallel b-sheet.[23, 24] This controlled conformation defines two topologically independent faces, which have been exploited successfully to induce biomolecules’ assemblies[25] and drug targeting.[26] Moreover, such scaffolds have been proven to be relevant metal-chelating agents for metal ions such as Cu + , using two cysteine side chains from the “lower face” of the peptide scaffold[27] , and Cu2 + , using four ligands from histidines and aspartates from the “upper face”.[28] A prototype peptide, A, adapted to the coordination of hard cations (Scheme 1, Table 1) has been fully studied a few years ago as a lanthanide-complexing agent.[24] This peptide incorpo-
(Scheme 1) in the same direction. The four carboxylate amino acid side chains have been demonstrated to be involved in the trivalent lanthanide cation coordination.[24, 29] This prototype peptide is very attractive for uranyl complexation because it is expected to provide four predisposed carboxylate side chains of amino acids in the equatorial plane of the trans-dioxo uranyl cation. Also, the tryptophan (Trp) residue is an interesting fluorescent probe, which can be used to follow the complexation reaction with uranyl. The main objective of the present study is to determine the key parameters involved in uranyl chelation with these de novo-designed cyclodecapeptides, which form a well-defined and structured uranyl complex. The effect of the peptide backbone structure and the nature of the coordinating amino acids have been analysed to draw structure–affinity correlations. To do so, a reliable titration method has first been developed to measure the uranyl complexes’ stability constants in the presence of a competing agent.
Results and Discussion Design and synthesis of the peptides
Scheme 1. Design of cyclic peptides to bind uranyl. Arrows represent the preferential orientation of the amino acid’s side chains in the b-sheet structures when X9 is a proline residue.
Table 1. Amino acid sequences of peptides together with conditional KC and KD values[a] for UO22 + at pH 6.0. Potentially coordinating amino acids are indicated in bold. Peptides
Group A 4 acidic residues
Group B Non b-sheet
Group C 3 acidic residues
A A1 A2 A3 A4 A5 B1 B2 C1 C2 C3 C4 C5 C6
Sequence
logKC
KD 109
c(DREPGEWDPG) c(EREPGEWEPG) c(ESEPGEWEPG) c(EREPGEYEPG) c(DRDPGDWDPG) c(DSDPGDWDPG) c(DREPGEWDSG) Ac-EREPGEWEPG-NH2 c(DREPGEWSPG) c(ERSPGEWEPG) c(EREPGSWEPG) c(QREPGEWEPG) c(HREPGEWEPG) c(HREPGEWESG)
8.1(1) 8.2(1) 8.4(1) 8.0(1) 7.2(1) 7.3(1) 7.5(1) 100
[a] Conditional KC and KD values for UO22 + were measured at pH 6.0, NaCl (0.1 m), MES (20 mm) in the presence of IDA as a competitor of known affinity.
rates four amino acids, which carry carboxylate hard donors for hard metal-ion coordination, namely two aspartates and two glutamates. The solution NMR structure of peptide A confirms that the constrained cyclic-peptide backbone orients the side chains of the acidic amino acids in positions 1, 3, 6, and 8 Chem. Eur. J. 2014, 20, 16566 – 16573
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Three series of peptides were synthesized and studied in order to determine the influence of the peptide structure on the ability to bind uranyl. Peptides of group A (Table 1) all display two b-turns that promote the b-sheet structure with different acidic residues in positions 1, 3, 6, and 8 (Scheme 1), namely aspartic acids (Asp) or glutamic acids (Glu). The prototype peptide A, with known solution structure,[24] has one turn flanked with two aspartic acids and a second one with two glutamic acids. The lengths of the coordinating amino acid side chains are expected to influence the metal-binding ability of these constrained peptides. Therefore, A1 and A2, which have four glutamic acid residues with longer side chains, were compared to A4 and A5, which have four aspartic acid residues with shorter side chains. Finally, the influence of the fluorescent group was investigated by replacing tryptophan in A1 by tyrosine (Tyr) in A3. The two peptides belonging to group B were synthesized in order to analyze the effect of the flexibility of the peptide scaffold. B1 is similar to A, with one b-turn (Pro–Gly) replaced by a more flexible loop (Ser–Gly). A similar peptide, mimicking a copper chaperone binding loop, was dedicated to soft metal-ion coordination thanks to two cysteine residues. As expected, the solution structure showed that the xPGx region was well-structured in a b-turn, whereas the xSGx region was very flexible.[12] The peptide B2 is even more flexible because this is a linear decapeptide equivalent to A1. The C–ter and N– ter of B2 are protected with amide functions in order to have the same number of coordinating groups as A1. Peptides of the third group, group C, bear only three acidic residues to check whether the four acidic residues of group A compounds are involved in uranyl coordination and eventually to explore the potential coordination of other amino acid side chains, such as primary amide in glutamine (Gln) or imidazole in histidine (His). Serine in peptides C1, C2 and C3 is not ex-
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Full Paper pected to significantly contribute to the complexes’ stability. Gln and His residues have been introduced in position 1 in peptides C4 and C5. Finally, C6, with the histidine in a loop, was synthesized to release some constraint in the coordination of the cyclic His. The cyclic peptides were synthesized as previously described.[24, 27] The linear protected decapeptides were assembled by automated solid-phase peptide synthesis using Fmoc strategy on 2-chlorotrityl chloride resin. After cleavage, they were cyclized in dichloromethane and deprotected to give the expected peptides after reverse-phase HPLC.
Uranyl binding to the prototype peptide A Electrospray-ionization mass spectrometry demonstrates the formation of the 1:1 uranyl complex with the prototype peptide A, either in the positive or negative detection mode. The spectrum recorded in negative mode (Figure 1) shows that the
Figure 1. () ESI-MS spectrum of A (100 mm) with equimolar UO22 + . Inset: isotopic patterns, experimental and calculated. * Acetic acid and sodium acetate adducts. ** Sodium adduct.
major species is the complex UO2–A, which is detected as a mono- or di-anion. Smaller signals characteristic of the free peptide are also present in this spectrum. The experimental peak envelopes of the metal complex are consistent with the theoretical isotopic patterns as shown in the insets of Figure 1. The complexation reaction was also followed by titrating peptide A with uranyl and recording circular dichroism or tryptophan fluorescence spectra. A precipitate is observed in excess of uranyl when the titration is run in HEPES buffer at pH 7, whereas no precipitation was observed when working in MES buffer at pH 6, even in significant excess of UO22 + (ca. 4 equiv). Therefore, we chose to study uranyl complexation at pH 6 to get reliable data and disfavor the formation of uranyl hydroxo complexes of low solubility. Moreover, the pKa values Chem. Eur. J. 2014, 20, 16566 – 16573
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of the peptide A indicate that the four carboxylic acid functions are deprotonated at pH 6 and therefore the complexation behaviour of the peptide at this pH should not be signicantly different from physiological pH. The acidity constants of peptide A were measured by potentiometric titrations in KCl (0.1 m) solution at 298 K. As expected, five basic sites are detected. The highest one, pKa5 = 11.7(2), corresponds to the protonation of the guanidine group of the arginine side chains. The pKa of the four acidic side chains are fitted to the following values: pKa1 = 3.23(2), pKa2 = 4.16(6); pKa3 = 4.69(2) and pKa4 = 5.41(8). The side chain of Asp is more acidic than that of Glu because of the larger withdrawing effect of the peptide backbone through shorter side chains. Incidentally, this effect is seen on the pKa values of the carboxylic acid side-chain of aspartic acid and glutamic acid, which are 3.71 and 4.15, respectively.[30] Therefore, pKa1 and pKa2 are assigned to aspartic acid residues, whereas the two slightly higher values, pKa3 and pKa4, may be assigned to glutamic acid side chains. The titration of A with UO22 + , followed by CD, shows characteristic features of the evolution of the peptide-scaffold structure upon metal binding (Figure 2 a). A unique complex, UO2– A, is formed as indicated by two isodichroic points and no significant evolution of the spectrum in an excess of metal ion. The spectra of the free peptide A and the UO2–A complex are dramatically different at pH 6. Indeed, the CD spectrum of the free compound is reminiscent of a random coil conformation with a negative band near 200 nm.[31] This is consistent with the four carboxylic acids being deprotonated, which leads to major electrostatic repulsions between the four negatively charged amino acid side chains in position 1, 3, 6, and 8, and to destabilization of the b-sheet structure of the peptide scaffold at this pH. In addition, the CD spectrum of the 1:1 complex UO2–A at the same pH shows a weak negative band at 224 nm and a large positive band at 195 nm, characteristic of a class B spectrum in Woody’s theoretical approach[32] and consistent with b-turn conformations.[33] These CD signatures demonstrate the formation of a well-defined uranyl complex, UO2– A, in which the prototype peptide A adopts a b-sheet structure to promote the coordination of the carboxylate amino acid side chains in the equatorial plane of the uranyl cation. Uranyl complexation was also efficiently detected through a large quenching of tryptophan fluorescence at 350 nm upon excitation at 280 nm. A similar effect was reported with tyrosine, with which quenching was used to study uranyl complexation with calmodulin derivatives[21] or transferrin.[6] The evolution of Trp fluorescence spectra during the titration of peptide A with uranyl nitrate is shown in Figure 3. An endpoint was detected for one UO22 + equivalent and confirms the formation of the complex UO2–A. The conditional stability constant of UO2–A at pH 6 was measured by fitting florescence titrations. Fluorescence spectra were recorded in dilute solutions ([A] 10 mm, Abs (280 nm) < 0.05) to avoid inner-filter effects and have a linear relationship between the fluorescence intensity and the concentrations. Moreover, uranyl is known to form many hydroxo complexes (more than 6 are reported in the NIST database),[30] which are
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Full Paper UO2 þ P ¼ UO2 P
K C ¼ 1=K D ð1Þ
Figure 2. CD titration of representative peptides (ca. 10 mm) with UO22 + (0 to 1.5 equiv, aliquots of 0.25 equiv) at pH 6.0, i indicates isodichroic points.
Six different conditions were used and these point consistently to a logKC value of 8.1 0.2 (Table 2). The pKa values of A account for the slightly higher KC value measured at pH 6.5 (entry 3, Table 2). Indeed, the difference calculated in logKC value from pH 6 to 6.5 is only + 0.1. In the following sections, the conditions of entry 5 were used because uranyl complexes of IDA display a simpler speciation. In conclusion, the prototype cyclodecapeptide A forms a well-defined 1:1 uranyl complex UO2–A, which exhibits a bsheet structure consistent with the coordination of the four carboxylate side chains to uranyl. This de novo-designed peptide exhibits a higher affinity for
Table 2. Conditional stability constant values (logKC) and dissociation constants KD for the complex UO2A in MES 20 mm obtained in various experimental conditions. [A] 10 mm. Entry pH Salt [0.1 M] Competitor [Competitor] [mM] logKC KD 109 1 2 3 4 5 6[a]
6.0 6.0 6.5 6.0 6.0 6.0
KNO3 KNO3 KNO3 KNO3 NaCl NaCl
CO32 CO32 CO32 IDA IDA IDA
0.5 0.1 0.1 0.1 0.1 0.02
8.0(1) 7.8(1) 8.3(1) 7.9(1) 8.1(1) 8.1(1)
10 2 16 4 51 13 3 82 82
[a] [A] 2 mm
Figure 3. Fluorescence titration of A (ca. 10 mm) with UO22 + (0 to 4 equiv) at pH 6.0, MES buffer (20 mm), KNO3 (0.1 m) with excitation at 280 nm. Inset: variation of the intensity at the peak maximum (350 nm) with UO22 + .
significantly present even at pH 6. To control the nature of the uranyl complexes formed during the titrations, two different competing ligands with well-defined speciation with UO22 + were added to the solution: carbonate (CO32) or iminodiacetic acid (IDA). The competing ligand concentration was chosen, 1) to minimize the formation of hydroxo species (< 2 %), and 2) to get reliable binding curves. The conditional stability constants of the uranyl complexes formed with the competing ligands were calculated from published values[30, 34] and inserted in the fitting procedure according to models described in the experimental section (see the Supporting Information). The KC constants reported in Table 2 were obtained for the formation of a unique UO2–A complex [Eq. (1)] as demonstrated by MS and CD experiments in similar conditions. Chem. Eur. J. 2014, 20, 16566 – 16573
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uranyl than calcium binding sites directly derived from calmodulin (logKC = 7.5), which organize similar coordinating atoms in a structured loop.[21] The series of decapeptides listed in Table 1 with various sequences and structures are studied in the following section to determine the key parameters that induce a large affinity for uranyl. Influence of the nature of the four-coordinating acidic residues (Asp or Glu): peptides of group A The nature of the acidic residues (Asp or Glu) introduced in positions 1, 3, 6, and 8 of the cyclic peptide scaffold for metal coordination is varied in the first series of peptides (Group A in Table 1). A1 and A2 are variants of the prototype peptide A with 4 glutamic acids as potential coordinating amino acids, whereas A4 and A5 bear 4 aspartic acids. ESI-MS spectra show the formation of 1:1 uranyl complexes for all these peptides.
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Full Paper Examples of MS spectra recorded in the positive mode are shown in Figure 4. Even though MS is not a quantitative method, the response coefficients are expected to be very similar for the series of peptides that have the same charge, such as A, A1, and A4. The relative peak intensities of the uranyl complex and the corresponding free peptide (in MS spectra acquired in the same conditions), clearly show that UO2–A1 is more abundant than UO2–A4.
Figure 5. Titration of 10 mm peptide solutions (MES buffer 20 mm, pH 6.0, 298 K) with uranyl in the presence of 100 mm IDA. Top: Fluorescence intensity of tryptophan at 350 nm upon excitation at 280 nm as a function of uranyl equivalents. Bottom: Corresponding Stern–Volmer plots.
Figure 4. (+) ESI-MS spectra of the UO22 + complexes (ca. 100 mm) with representative peptides in ammonium acetate buffer (20 mm, pH 6.9). Adducts with sodium are detected at higher m/z than the main signals indicated on the figure.
The higher stability of the former complex is confirmed by the analyses of the fluorescence titrations. Striking differences are detected in the tryptophan fluorescence quenching during titration with uranyl. Figure 5 displays the evolution of the Trp emission at 350 nm as a function of uranyl content for three peptides of group A. The quenching is a lot more efficient for A1 containing four glutamate residues than for A4 containing four aspartate residues. The Stern–Volmer plots shown in the bottom half of Figure 5 also reflect this tendency. Indeed, the slope of the plot, which gives an evaluation of the conditional stability constant with respect to IDA, is one order of magnitude larger for A1 (logKSV = 5.6) than for A4 (logKSV = 4.6). The conditional stability constants obtained by fitting the titrations with IDA, taking into account the constants of uranyl complexes of IDA, are reported in Table 1 and also demonstrate one order of magnitude difference between A1 and A4. Whereas A1 has a very similar affinity to the prototype peptide A, the incorporation of 4 aspartic acids in A4 significantly decreases the uranyl complex stability. One explanation for the greater affinity of glutamate-based peptides compared to aspartate-based peptides is the larger basicity of glutamate with respect to aspartate. Because carboxylate–uranyl interactions are mainly electrostatic due to the hard nature of both the metal cation and the donor, the stability should correlate with the basicity of simple donors, as deChem. Eur. J. 2014, 20, 16566 – 16573
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scribed for lanthanide complexes with aminocarboxylate ligands.[35] The carboxylate side chain of Asp is therefore expected to have a lower affinity for UO22 + than that of Glu. A significant structural effect is also detected on the circular dichroism spectra. Whereas A and A1 show characteristic CD features of b-sheet formation upon uranyl coordination (Figures 2 a and 2 b, respectively), peptide A4 demonstrates a poor evolution in the CD spectra and, more importantly, the absence of characteristic features of a b-sheet structure for the uranyl complex (Figure 2 c). This indicates that the cyclic peptide backbone is too constrained to benefit from the stable b-sheet conformation and the tight coordination of the four short side chains of the aspartate residues in A4. In these cyclic peptides, both steric constraints and basicity favor the glutamate–uranyl interaction. For amino acids in positions 2 and 7, the side-chain oriented on the opposite face to the metal ion coordination was also varied. The positively charged guanidinium function of arginine in position 2 in peptides A1 and A4 was replaced by the polar alcohol group of serine in peptides A2 and A5, without any significant effect on the affinity for uranyl. Similarly, peptide A3 with a fluorescent tyrosine (Tyr) shows similar uranyl coordination properties to A1, with a tryptophan. This confirms that the two faces defined by the cyclic peptide scaffold are topologically independent.
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Full Paper Influence of the structure: peptides of group B The two decapeptides B1 and B2 were studied to determine the influence of the scaffold on the uranyl complexation properties. In peptide B1, one Pro–Gly motif, which induces the formation of a b-turn, was replaced by a less constrained sequence Ser–Gly, which promotes the formation of a flexible loop. The 1:1 complex UO2–B1 is detected by MS and its CD spectrum is reminiscent of a b-sheet structure (Figure 2 d) even though the molar ellipticity ([V]195nm 153 000 deg cm2 dmol1) is significantly lower than the one measured for UO2–A ([V]195nm 189 000 deg cm2 dmol1). Moreover, the stability constant of UO2–B1 is 4 times lower than for the prototype peptide A (Table 1), as shown by the binding isotherms and on the Stern–Volmer plots in Figure 5. The comparison between the b-sheet prototype peptide A and its more flexible analogue B1 is clear evidence that the b-sheet structure is optimal when constraints associated with short side chains are not too large. The behaviour of the linear equivalent of A1, peptide B2, is even more striking. Indeed, it is totally different from all the cyclic compounds. The ES–MS spectrum in Figure 4 shows almost no formation of the uranyl complex and also displays signals due to polymetallic adducts. Furthermore, the slight fluorescence quenching upon uranyl addition to B2 is indicative of a dramatically lower metal-binding affinity. The comparison of peptides of group A with the two analogues B1 and B2 exemplifies the perfect adjustment of the bsheet cyclic peptide scaffold in A or A1 for uranyl complexation.
Influence of nonacidic-coordinating amino acids: peptides of group C All the peptides of group C have only three acidic residues. All of them demonstrate a significantly lower affinity for uranyl. Indeed, in contrast to peptides of group A and B1, their uranyl complexes are barely detectable in the MS spectra (see Figure S1 in the Supporting Information). Moreover, the fluorescence quenching of tryptophan is a lot smaller and no real binding isotherms could be observed (see Figure S2 in the Supporting Information). The Stern–Volmer constants are lower than 4.5 (in log units) and only an upper limit of the affinity constant could be determined (logKC < 7). In this group, only peptide C2 demonstrates a larger fluorescence quenching corresponding to a logKC = 7.4. C2 is equivalent to A1 with a serine instead of a glutamate in position 3. The slightly larger affinity of C2 for uranyl, in comparison with other peptides of group C, suggests that the thermodynamic contribution to the affinity constant of the glutamate in position 3 in A1 is lower. Finally, the substitution of glutamate by either glutamine or histidine in peptides C4 and C5, respectively, leads to a decrease of more than one order of magnitude in uranyl affinity. Histidine is a cyclic nitrogen donor that is more constrained than the oxygen donors studied in this work. Therefore, peptide C6 was also considered with the histidine in a loop to release some constraint in its coordination. Chem. Eur. J. 2014, 20, 16566 – 16573
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Peptides of group C, lacking one carboxylate side chain compared with peptides of group A, show a significant loss in affinity for uranyl. This proves the involvement of the four amino acid side chains of Asp and Glu in the dioxo-cation coordination with peptides of group A. Moreover, neutral donors, such as glutamine in C4, or histidine in C5 and C6, do not play a significant role in the uranyl complex stability.
Conclusion The ability of a series of decapeptides to bind uranyl has been investigated with complementary analytical and spectroscopic methods to determine the key parameters for the formation of stable uranyl–peptide complexes. Structured cyclic peptide scaffolds were chosen as promising candidates to coordinate uranyl thanks to four amino acid side chains pre-oriented towards the dioxo-cation equatorial plane. The b-sheet structure of the prototype peptide A with two aspartates and two glutamates, and of its equivalents A1 and A2 incorporating four glutamates, is perfectly adapted to the coordination of UO22 + . Indeed, the more flexible cyclic compound B1, having only one turn, displays a lower affinity for uranyl. The behaviour of the linear decapeptide B2 is even more striking, because it forms nondefined polymetallic uranyl complexes of low stability. Compounds with four aspartate residues, A4 and A5, with short carboxylate side chains, show the role of the conformational constraints in the metal complex formation. These two cyclic peptides also complex uranyl in a 1:1 complex, but their circular dichroism signature reveals that the scaffold cannot adopt a b-sheet structure. As a result, the stability constants are one order of magnitude lower than those obtained with peptides A and A1, which have glutamate residues. Actually, the molar ellipticity at 195 nm of the uranyl complex is directly correlated to its stability constant, which confirms that the bsheet structure is a key parameter for high stability in the peptide series. Sequences with only three acidic amino acids display significantly lower affinities for uranyl, which proves the involvement of the four amino acid side chains of Asp and Glu in the dioxo-cation coordination with peptides of group A. Importantly, the de novo-designed peptides A, A1 and A2 exhibit a higher affinity for uranyl (logKC = 8.1–8.4) than calcium binding sites derived from calmodulin, which had their affinity for uranyl measured in exactly the same conditions (logKC = 7.5).[21] This suggests that the presentation of four carboxylate side chains in the b-sheet structure is particularly adapted to uranyl coordination even in comparison with the structured EF-hand loops of calmodulin. Moreover the cyclodecapeptide A is selective for uranyl with respect to calcium by more than four orders of magnitude. Indeed, the affinity of A for calcium is only logKC = 3.7.[24] This study shows that simple uranyl-binding sequences in vivo, built with aspartates and glutamates, should display conditional affinity constants in the range logKC = 7–9, but not significantly higher. Furthermore, the peptide scaffold structure is a key parameter for the design of efficient uranyl-binding peptides that control metal–complex speciation. Within the stud-
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Full Paper ied sequences, A1 and A2 appear as good starting points for the design of high-affinity uranyl-chelating peptides. Phosphorous groups are known to strongly bind hard ions and, in particular, uranyl.[36] For instance, the insertion of a phosphoserine residue in the binding loop of a calmodulin-derived peptide was successful in enhancing the uranyl complex stability.[21] Therefore, phosphorylated amino acids are currently inserted in the cyclic peptide sequence in order to test the complexation properties of peptides bearing both carboxylate and phosphate side chains, which may also be relevant to some uranylbinding sites in vivo.[11, 19] Another strategy involves developing unnatural amino acids bearing metal-chelating groups on their side chains. Peptides incorporating such unnatural residues have been shown to significantly improve the coordination properties of peptides for lanthanides[37] and are currently being developed for uranyl chelation.
cell in centimeters, and c is the peptide concentration in moles per liter.
Fluorescence titrations Spectra were recorded on a LS50B spectrofluorimeter connected to a computer equipped with FLWILAB 2.0. The measurements were performed at 298 K. Trp fluorescence titrations were performed with 280 nm excitation (excitation slit: 3.0 nm). The emission slit was adjusted (3.5–4.5 nm) to avoid signal saturation. Experiments were repeated to ensure reproducibility.
Acknowledgements
Experimental Section
We thank Olivier Snque and Didier Boturyn for the initial synthesis of the prototype peptide A. This research was supported by the “Programme Transversal Toxicologie du CEA”, the NRBC-E program and the Labex ARCANE (Grant ANR-11LABX-0003-01).
The peptide syntheses and additional experimental procedures for physico-chemical experiments are reported in the Supporting Information.
Keywords: bioinorganic chemistry · carboxylate ligands · fluorescence · peptides · uranium
Preparation of aqueous solutions A stock uranyl solution (ca. 10 mm) was prepared from uranyl dinitrate hexahydrate in nitric acid (0.01 m). The precise uranyl concentration was obtained by measuring the absorbance of an aliquot compared to an ICP Uranium standard (1000 mg U per mL in 1 % HNO3) at l = 415 nm. Peptide solutions were prepared freshly before use and the precise peptide concentration was determined by recording a UV spectrum (e280 = 5690 L mol1 cm1 owing to the presence of a Trp residue). MES buffer was prepared by dissolving solid 2-(N-morpholino)ethanesulfonic acid in H2O and by adjusting the pH to 6.0 or 6.5 with KOH. Solutions of IDA or carbonate were prepared in the buffer from solid iminodiacetic acid or sodium carbonate.
ESI-MS Peptide solutions (100 mm) were prepared in an ammonium acetate buffer (20 mm, pH 7). The stock uranyl solution was added to prepare aliquots containing 0, 0.5, 1, and 2 equiv of UO22 + per peptide. Mass spectra were recorded on a LXQ type THERMO SCIENTIFIC spectrometer, equipped with an electrospray ionization source and a linear-trap detector. Solutions were injected in the spectrometer at 10 mL min1 flow rate. Ionization voltage and capillary temperature were about 2 kV and 250 8C, respectively. The source settings were the same (sheath gas, auxiliary gas, capillary voltage, and tube lens) for all series of peptides with comparable data.
CD titrations CD spectra were recorded at 25 8C on an Applied Photophysics Chirascan Spectrometer in a 1 cm path cell. The peptide concentration was 10 mm in water and the pH was adjusted to 6.0 with KOH. All spectra were obtained from 320 to 190 nm with a 1 nm data interval, a time constant of 2 s and a band width of 1 nm, with 3 scans. CD spectra are reported in molar ellipticity ([V] in units of deg cm2 dmol1). [V] = qobs/(10lc), where qobs is the observed ellipticity in millidegrees, l is the optical path length of the Chem. Eur. J. 2014, 20, 16566 – 16573
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[1] a) P. W. Durbin, Actinides in Animals and Man in The Chemistry of the Actinide and Transactinide Elements (Eds.: L. R. E. Morss, N. Fuger, J. Katz), Springer, Dordrecht (The Netherlands), 2006, pp. 3339 – 3440; b) D. M. Taylor, S. K. Taylor, Rev. Environ. Health 1997, 12, 147 – 158. [2] E. Ansoborlo, O. Prat, P. Moisy, C. Den Auwer, P. Guilbaud, M. Carriere, B. Gouget, J. Duffield, D. Doizi, T. Vercouter, C. Moulin, V. Moulin, Biochimie 2006, 88, 1605 – 1618. [3] a) E. Ansoborlo, B. Amekraz, C. Moulin, V. Moulin, F. Taran, T. Bailly, R. Burgada, M. H. Henge-Napoli, A. Jeanson, C. Den Auwer, L. Bonin, P. Moisy, C. R. Chim. 2007, 10, 1010 – 1019; b) A. E. V. Gorden, J. D. Xu, K. N. Raymond, P. Durbin, Chem. Rev. 2003, 103, 4207 – 4282; c) R. J. Abergel, P. W. Durbin, B. Kullgren, S. N. Ebbe, J. D. Xu, P. Y. Chang, D. I. Bunin, E. A. Blakely, K. A. Bjornstad, C. J. Rosen, D. K. Shuh, K. N. Raymond, Health Phys. 2010, 99, 401 – 407; d) M. Sturzbecher-Hoehne, G. J.-P. Deblonde, R. J. Abergel, Radiochim. Acta 2013, 101, 359 – 366. [4] J. D. Van Horn, H. Huang, Coord. Chem. Rev. 2006, 250, 765 – 775. [5] a) S. Scapolan, E. Ansoborlo, C. Moulin, C. Madic, Radiat. Prot. Dosim. 1998, 79, 505 – 508; C. Vidaud, S. Gourion-Arsiquaud, F. Rollin-Genetet, C. Torne-Celer, S. Plantevin, O. Pible, C. Berthomieu, E. Quemeneur, Biochemistry 2007, 46, 2215 – 2226; b) G. Montavon, C. Apostolidis, F. Bruchertseifer, U. Repinc, A. Morgenstern, J. Inorg. Biochem. 2009, 103, 1609 – 1616. [6] J. Michon, S. Frelon, C. Garnier, F. Coppin, J. Fluoresc. 2010, 20, 581 – 590. [7] C. Vidaud, A. Dedieu, C. Basset, S. Plantevin, I. Dany, O. Pible, E. Quemeneur, Chem. Res. Toxicol. 2005, 18, 946 – 953. [8] a) O. Pible, C. Vidaud, S. Plantevin, J. L. Pellequer, E. Quemeneur, Protein Sci. 2010, 19, 2219 – 2230; b) O. Pible, P. Guilbaud, J. L. Pellequer, C. Vidaud, E. Quemeneur, Biochimie 2006, 88, 1631 – 1638. [9] a) A. Dedieu, F. Berenguer, C. Basset, O. Prat, E. Quemeneur, O. Pible, C. Vidaud, J. Chromatogr. A 2009, 1216, 5365 – 5376; b) C. Basset, A. Dedieu, P. Guerin, E. Quemeneur, D. Meyer, C. Vidaud, J. Chromatogr. A 2008, 1185, 233 – 240; c) L. Reisser-Rubrecht, C. Torne-Celer, W. Renier, O. Averseng, S. Plantevin, E. Quemeneur, L. Bellanger, C. Vidaud, Chem. Res. Toxicol. 2008, 21, 349 – 357; d) O. Averseng, A. Hagege, F. Taran, C. Vidaud, Anal. Chem. 2010, 82, 9797 – 9802. [10] C. Basset, O. Averseng, P. J. Ferron, N. Richaud, A. Hagege, O. Pible, C. Vidaud, Chem. Res. Toxicol. 2013, 26, 645 – 653. [11] L. Qi, C. Basset, O. Averseng, E. Quemeneur, A. Hagege, C. Vidaud, Metallomics 2014, 6, 166 – 176. [12] a) P. Rousselot-Pailley, O. Snque, C. Lebrun, S. Crouzy, D. Boturyn, P. Dumy, M. Ferrand, P. Delangle, Inorg. Chem. 2006, 45, 5510 – 5520; b) O.
16572
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Full Paper
[13] [14]
[15]
[16]
[17] [18] [19] [20] [21] [22]
[23]
[24] [25]
Snque, S. Crouzy, D. Boturyn, P. Dumy, M. Ferrand, P. Delangle, Chem. Commun. 2004, 770 – 771. E. Rossy, O. Snque, D. Lascoux, D. Lemaire, S. Crouzy, P. Delangle, J. Coves, Febs Lett. 2004, 575, 86 – 90. a) A. Jancs, D. Szunyogh, F. H. Larsen, P. W. Thulstrup, N. J. Christensen, B. Gyurcsik, L. Hemmingsen, Metallomics 2011, 3, 1331 – 1339; b) V. L. Pecoraro, A. F. A. Peacock, O. Iranzo, M. Luczkowski, Understanding the Biological Chemistry of Mercury Using a de novo Protein Design Strategy in Bioinorganic Chemistry: Cellular Systems and Synthetic Models (Eds.: E. C. Long, M. J. Baldwin), ACS, Washington D.C., 2009, pp. 183 – 197. a) A.-S. Jullien, C. Gateau, C. Lebrun, I. Kieffer, D. Testemale, P. Delangle, Inorg. Chem. 2014, 53, 5229 – 5239; b) A.-S. Jullien, C. Gateau, I. Kieffer, D. Testemale, P. Delangle, Inorg. Chem. 2013, 52, 9954 – 9961; c) A. M. Pujol, C. Lebrun, C. Gateau, A. Manceau, P. Delangle, Eur. J. Inorg. Chem. 2012, 3835 – 3843; d) A. M. Pujol, C. Gateau, C. Lebrun, P. Delangle, Chem. Eur. J. 2011, 17, 4418 – 4428; e) A. M. Pujol, C. Gateau, C. Lebrun, P. Delangle, J. Am. Chem. Soc. 2009, 131, 6928 – 6929. a) S. Pires, J. Habjanic, M. Sezer, C. M. Soares, L. Hemmingsen, O. Iranzo, Inorg. Chem. 2012, 51, 11339 – 11348; b) C. Gateau, P. Delangle, Ann. N. Y. Acad. Sci. 2014, 1315, 30 – 36; c) A. M. Pujol, M. Cuillel, A.-S. Jullien, C. Lebrun, D. Cassio, E. Mintz, C. Gateau, P. Delangle, Angew. Chem. Int. Ed. 2012, 51, 7445 – 7448; Angew. Chem. 2012, 124, 7563 – 7566. M. Tegoni, F. T. Yu, M. Bersellini, J. E. Penner-Hahn, V. L. Pecoraro, Proc. Natl. Acad. Sci. USA 2012, 109, 21234 – 21239. H. Huang, S. Chaudhary, J. D. Van Horn, Inorg. Chem. 2005, 44, 813 – 815. S. Safi, G. Creff, A. Jeanson, L. Qi, C. Basset, J. Roques, P. L. Solari, E. Simoni, C. Vidaud, C. Den Auwer, Chem. Eur. J. 2013, 19, 11261 – 11269. L. Le Clainche, C. Vita, Environ. Chem. Lett. 2006, 4, 45 – 49. R. Pardoux, S. Sauge-Merle, D. Lemaire, P. Delangle, L. Guilloreau, J.-M. Adriano, C. Berthomieu, PLOS ONE 2012, 7, e41922. a) L. Zhou, M. Bosscher, C. Zhang, S. Ozcubukcu, L. Zhang, W. Zhang, C. J. Li, J. Liu, M. P. Jensen, L. Lai, C. He, Nat. Chem. 2014, 6, 236 – 241; b) S. V. Wegner, H. Boyaci, H. Chen, M. P. Jensen, C. He, Angew. Chem. Int. Ed. 2009, 48, 2339 – 2341; Angew. Chem. 2009, 121, 2375 – 2377. a) P. Dumy, I. M. Eggleston, G. Esposito, S. Nicula, M. Mutter, Biopolymers 1996, 39, 297 – 308; b) S. Peluso, T. Ruckle, C. Lehmann, M. Mutter, C. Peggion, M. Crisma, ChemBioChem 2001, 2, 432 – 437. C. S. Bonnet, P. H. Fries, S. Crouzy, O. Snque, F. Cisnetti, D. Boturyn, P. Dumy, P. Delangle, Chem. Eur. J. 2009, 15, 7083 – 7093. a) G. T. Dolphin, P. Dumy, J. Garcia, Angew. Chem. Int. Ed. 2006, 45, 2699 – 2702; Angew. Chem. 2006, 118, 2765 – 2768; b) P. Murat, R. Bonnet, A. Van der Heyden, N. Spinelli, P. Labbe, D. Monchaud, M. P. Teulade-Fichou, P. Dumy, E. Defrancq, Chem. Eur. J. 2010, 16, 6106 –
Chem. Eur. J. 2014, 20, 16566 – 16573
www.chemeurj.org
[26]
[27]
[28]
[29] [30]
[31]
[32] [33] [34] [35] [36]
[37]
6114; c) M. Galibert, O. Renaudet, P. Dumy, D. Boturyn, Angew. Chem. Int. Ed. 2011, 50, 1901 – 1904; Angew. Chem. 2011, 123, 1941 – 1944. a) D. Boturyn, J. L. Coll, E. Garanger, M. C. Favrot, P. Dumy, J. Am. Chem. Soc. 2004, 126, 5730 – 5739; b) S. Grigalevicius, S. Chierici, O. Renaudet, R. Lo-Man, E. Deriaud, C. Leclerc, P. Dumy, Bioconjugate Chem. 2005, 16, 1149 – 1159. A. M. Pujol, M. Cuillel, O. Renaudet, C. Lebrun, P. Charbonnier, D. Cassio, C. Gateau, P. Dumy, E. Mintz, P. Delangle, J. Am. Chem. Soc. 2011, 133, 286 – 296. a) A. Fragoso, P. Lamosa, R. Delgado, O. Iranzo, Chem. Eur. J. 2013, 19, 2076 – 2088; b) A. Fragoso, R. Delgado, O. Iranzo, Dalton Trans. 2013, 42, 6182 – 6192. C. S. Bonnet, P. H. Fries, S. Crouzy, P. Delangle, J. Phys. Chem. B 2010, 114, 8770 – 8781. R. M. Smith, A. E. Martell, R. J. Motekaitis, NIST Critically Selected Stability Constants of Metal Complexes Database, NIST Standard Reference Database 46, 2001. N. Sreerama, R. W. Woody in Circular Dichroism of peptides and proteins, in Circular Dichroism: Principles and Applications, 2ednd ed(Eds.: N. Berova, K. Nakanishi, R. W. Woody), Wiley, New York, 2000, pp. 601 – 620. J. A. Smith, L. G. Pease, CRC Crit. Rev. Biochem. 1980, 8, 315 – 399. B. Imperiali, S. L. Fisher, R. A. Moats, T. J. Prins, J. Am. Chem. Soc. 1992, 114, 3182 – 3188. J. Jiang, M. J. Sarsfield, J. C. Renshaw, F. R. Livens, D. Collison, J. M. Charnock, M. Helliwell, H. Eccles, Inorg. Chem. 2002, 41, 2799 – 2806. T. F. Gritmon, M. P. Goedken, G. R. Choppin, J. Inorg. Nucl. Chem. 1977, 39, 2021 – 2023. a) W. A. Reed, L. F. Rao, P. Zanonato, A. Y. Garnov, B. A. Powell, K. L. Nash, Inorg. Chem. 2007, 46, 2870 – 2876; b) M. Sawicki, J. M. Siaugue, C. Jacopin, C. Moulin, T. Bailly, R. Burgada, S. Meunier, P. Baret, J. L. Pierre, F. Taran, Chem. Eur. J. 2005, 11, 3689 – 3697. a) L. Ancel, A. Niedz´wiecka, C. Lebrun, C. Gateau, P. Delangle, C. R. Chim. 2013, 16, 515 – 523; b) A. Niedz´wiecka, F. Cisnetti, C. Lebrun, C. Gateau, P. Delangle, Dalton Trans. 2012, 41, 3239 – 3247; c) A. Niedz´wiecka, F. Cisnetti, C. Lebrun, P. Delangle, Inorg. Chem. 2012, 51, 5458 – 5464; d) F. Cisnetti, C. Lebrun, P. Delangle, Dalton Trans. 2010, 39, 3560 – 3562; e) F. Cisnetti, C. Gateau, C. Lebrun, P. Delangle, Chem. Eur. J. 2009, 15, 7456 – 7469.
Received: July 23, 2014 Published online on October 16, 2014
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& Bioinorganic Chemistry
Engineering Short Peptide Sequences for Uranyl Binding Colette Lebrun,[a, b] Matthieu Starck,[a, b] Vicky Gathu,[a, b] Yves Chenavier,[a, b] and Pascale Delangle*[a, b]
Abstract: Peptides are interesting tools to rationalize uranyl–protein interactions, which are relevant to uranium toxicity in vivo. Structured cyclic peptide scaffolds were chosen as promising candidates to coordinate uranyl thanks to four amino acid side chains pre-oriented towards the dioxo cation equatorial plane. The binding of uranyl by a series of decapeptides has been investigated with complementary analytical and spectroscopic methods to determine
Introduction Uranium is a natural element widely found in the environment, due to both natural occurrence in mineral ores or in sea water and industrial applications. In particular, it is a key element for energy production through nuclear fission. This heavy metal belongs to the actinides series and has no essential role in living organisms from plants to man. Uranium displays radiological and chemical toxicity to living organisms.[1, 2] Whatever the route of entry, uranium reaches the blood and then targets mainly the kidneys and the bones. However, despite significant recent advances in the field, there is still a serious lack of knowledge about the molecular interactions responsible for uranium toxicity. The underlying mechanisms need to be unraveled to predict the effect of uranium on living organisms and also to help in designing efficient detoxification agents. Indeed, such chelating agents are essential tools in case of dirty bombs or the accidental release of uranium in the environment.[3] Uranium exhibits several redox states, but the most stable and relevant form in aqueous solutions and in physiological conditions is the hexavalent uranyl cation, UO22 + , which is a rare form of linear trans-dioxo cation. The presence of the two oxo groups favors the coordination of four to six extra ligands in the equatorial plane, which is perpendicular to the OUO bonds. The uranyl ion is classified as a hard Lewis acid [a] C. Lebrun, Dr. M. Starck, V. Gathu, Y. Chenavier, Dr. P. Delangle Univ. Grenoble Alpes, INAC, SCIB 38000 Grenoble (France) [b] C. Lebrun, Dr. M. Starck, V. Gathu, Y. Chenavier, Dr. P. Delangle CEA, INAC, SCIB 38054 Grenoble (France) E-mail: [email protected] Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/chem.201404546. Chem. Eur. J. 2014, 20, 16566 – 16573
the key parameters for the formation of stable uranyl–peptide complexes. The molar ellipticity of the uranyl complex at 195 nm is directly correlated to its stability, which demonstrates that the b-sheet structure is optimal for high stability in the peptide series. Cyclodecapeptides with four glutamate residues exhibit the highest affinities for uranyl with log KC = 8.0–8.4 and, therefore, appear as good starting points for the design of high-affinity uranyl-chelating peptides.
and is highly oxophilic with high affinity for hard oxygen donors. Therefore, the preferred binding sites of uranyl in proteins involve carboxylate donors from aspartates, glutamates, or the carboxyl terminus of the peptide chain.[4] Two abundant serum proteins, namely albumin and transferrin, have been extensively studied as uranium targets in vivo.[2, 5, 6] However, other significant uranyl target proteins have been identified over the past decade by screening human serum proteins for uranium binding,[7] thanks to a combination of in silico approaches[8] and analytical tools.[9] Two proteins demonstrated a particularly large affinity for uranyl. Fetuin, despite its low concentration, could be one of the major target proteins of uranyl in the serum,[10] and the highly phosphorylated protein osteopontin, involved in the organo-mineral homeostasis of the bone, also binds tightly with uranyl.[11] Peptides are frequently used as simple and efficient tools to study metal–protein binding sites and mimic biological molecules such as metal transporters[12, 13] or metal-responsive transcriptional factors.[14] In our laboratory, pseudopeptides based on chemical scaffolds functionalized with amino acids have also proven to be excellent models of metal detoxification proteins.[15] Peptide derivatives may also be powerful metal chelators[16] with potential applications in detoxification or excellent models of enzyme activity.[17] Some peptides have been applied to uranyl complexation with sequences found in albumin,[18] osteopontin[19] or in calcium-binding proteins such as calmodulin[20, 21] to identify the chelation sites and their affinity for uranyl. Interestingly, protein engineering either from nickelbinding sites or from de novo sequences provided very interesting tools for uranyl chelation with large affinities.[22] In this study, we propose to take advantage of structured cyclodecapeptides that form well-defined uranyl complexes to rationalize peptide–uranyl interactions. Indeed, de novo cyclodecapeptides, which incorporate two prolylglycine sequences as b-turn inducers, demonstrate a backbone conformation in
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Full Paper antiparallel b-sheet.[23, 24] This controlled conformation defines two topologically independent faces, which have been exploited successfully to induce biomolecules’ assemblies[25] and drug targeting.[26] Moreover, such scaffolds have been proven to be relevant metal-chelating agents for metal ions such as Cu + , using two cysteine side chains from the “lower face” of the peptide scaffold[27] , and Cu2 + , using four ligands from histidines and aspartates from the “upper face”.[28] A prototype peptide, A, adapted to the coordination of hard cations (Scheme 1, Table 1) has been fully studied a few years ago as a lanthanide-complexing agent.[24] This peptide incorpo-
(Scheme 1) in the same direction. The four carboxylate amino acid side chains have been demonstrated to be involved in the trivalent lanthanide cation coordination.[24, 29] This prototype peptide is very attractive for uranyl complexation because it is expected to provide four predisposed carboxylate side chains of amino acids in the equatorial plane of the trans-dioxo uranyl cation. Also, the tryptophan (Trp) residue is an interesting fluorescent probe, which can be used to follow the complexation reaction with uranyl. The main objective of the present study is to determine the key parameters involved in uranyl chelation with these de novo-designed cyclodecapeptides, which form a well-defined and structured uranyl complex. The effect of the peptide backbone structure and the nature of the coordinating amino acids have been analysed to draw structure–affinity correlations. To do so, a reliable titration method has first been developed to measure the uranyl complexes’ stability constants in the presence of a competing agent.
Results and Discussion Design and synthesis of the peptides
Scheme 1. Design of cyclic peptides to bind uranyl. Arrows represent the preferential orientation of the amino acid’s side chains in the b-sheet structures when X9 is a proline residue.
Table 1. Amino acid sequences of peptides together with conditional KC and KD values[a] for UO22 + at pH 6.0. Potentially coordinating amino acids are indicated in bold. Peptides
Group A 4 acidic residues
Group B Non b-sheet
Group C 3 acidic residues
A A1 A2 A3 A4 A5 B1 B2 C1 C2 C3 C4 C5 C6
Sequence
logKC
KD 109
c(DREPGEWDPG) c(EREPGEWEPG) c(ESEPGEWEPG) c(EREPGEYEPG) c(DRDPGDWDPG) c(DSDPGDWDPG) c(DREPGEWDSG) Ac-EREPGEWEPG-NH2 c(DREPGEWSPG) c(ERSPGEWEPG) c(EREPGSWEPG) c(QREPGEWEPG) c(HREPGEWEPG) c(HREPGEWESG)
8.1(1) 8.2(1) 8.4(1) 8.0(1) 7.2(1) 7.3(1) 7.5(1) 100
[a] Conditional KC and KD values for UO22 + were measured at pH 6.0, NaCl (0.1 m), MES (20 mm) in the presence of IDA as a competitor of known affinity.
rates four amino acids, which carry carboxylate hard donors for hard metal-ion coordination, namely two aspartates and two glutamates. The solution NMR structure of peptide A confirms that the constrained cyclic-peptide backbone orients the side chains of the acidic amino acids in positions 1, 3, 6, and 8 Chem. Eur. J. 2014, 20, 16566 – 16573
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Three series of peptides were synthesized and studied in order to determine the influence of the peptide structure on the ability to bind uranyl. Peptides of group A (Table 1) all display two b-turns that promote the b-sheet structure with different acidic residues in positions 1, 3, 6, and 8 (Scheme 1), namely aspartic acids (Asp) or glutamic acids (Glu). The prototype peptide A, with known solution structure,[24] has one turn flanked with two aspartic acids and a second one with two glutamic acids. The lengths of the coordinating amino acid side chains are expected to influence the metal-binding ability of these constrained peptides. Therefore, A1 and A2, which have four glutamic acid residues with longer side chains, were compared to A4 and A5, which have four aspartic acid residues with shorter side chains. Finally, the influence of the fluorescent group was investigated by replacing tryptophan in A1 by tyrosine (Tyr) in A3. The two peptides belonging to group B were synthesized in order to analyze the effect of the flexibility of the peptide scaffold. B1 is similar to A, with one b-turn (Pro–Gly) replaced by a more flexible loop (Ser–Gly). A similar peptide, mimicking a copper chaperone binding loop, was dedicated to soft metal-ion coordination thanks to two cysteine residues. As expected, the solution structure showed that the xPGx region was well-structured in a b-turn, whereas the xSGx region was very flexible.[12] The peptide B2 is even more flexible because this is a linear decapeptide equivalent to A1. The C–ter and N– ter of B2 are protected with amide functions in order to have the same number of coordinating groups as A1. Peptides of the third group, group C, bear only three acidic residues to check whether the four acidic residues of group A compounds are involved in uranyl coordination and eventually to explore the potential coordination of other amino acid side chains, such as primary amide in glutamine (Gln) or imidazole in histidine (His). Serine in peptides C1, C2 and C3 is not ex-
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Full Paper pected to significantly contribute to the complexes’ stability. Gln and His residues have been introduced in position 1 in peptides C4 and C5. Finally, C6, with the histidine in a loop, was synthesized to release some constraint in the coordination of the cyclic His. The cyclic peptides were synthesized as previously described.[24, 27] The linear protected decapeptides were assembled by automated solid-phase peptide synthesis using Fmoc strategy on 2-chlorotrityl chloride resin. After cleavage, they were cyclized in dichloromethane and deprotected to give the expected peptides after reverse-phase HPLC.
Uranyl binding to the prototype peptide A Electrospray-ionization mass spectrometry demonstrates the formation of the 1:1 uranyl complex with the prototype peptide A, either in the positive or negative detection mode. The spectrum recorded in negative mode (Figure 1) shows that the
Figure 1. () ESI-MS spectrum of A (100 mm) with equimolar UO22 + . Inset: isotopic patterns, experimental and calculated. * Acetic acid and sodium acetate adducts. ** Sodium adduct.
major species is the complex UO2–A, which is detected as a mono- or di-anion. Smaller signals characteristic of the free peptide are also present in this spectrum. The experimental peak envelopes of the metal complex are consistent with the theoretical isotopic patterns as shown in the insets of Figure 1. The complexation reaction was also followed by titrating peptide A with uranyl and recording circular dichroism or tryptophan fluorescence spectra. A precipitate is observed in excess of uranyl when the titration is run in HEPES buffer at pH 7, whereas no precipitation was observed when working in MES buffer at pH 6, even in significant excess of UO22 + (ca. 4 equiv). Therefore, we chose to study uranyl complexation at pH 6 to get reliable data and disfavor the formation of uranyl hydroxo complexes of low solubility. Moreover, the pKa values Chem. Eur. J. 2014, 20, 16566 – 16573
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of the peptide A indicate that the four carboxylic acid functions are deprotonated at pH 6 and therefore the complexation behaviour of the peptide at this pH should not be signicantly different from physiological pH. The acidity constants of peptide A were measured by potentiometric titrations in KCl (0.1 m) solution at 298 K. As expected, five basic sites are detected. The highest one, pKa5 = 11.7(2), corresponds to the protonation of the guanidine group of the arginine side chains. The pKa of the four acidic side chains are fitted to the following values: pKa1 = 3.23(2), pKa2 = 4.16(6); pKa3 = 4.69(2) and pKa4 = 5.41(8). The side chain of Asp is more acidic than that of Glu because of the larger withdrawing effect of the peptide backbone through shorter side chains. Incidentally, this effect is seen on the pKa values of the carboxylic acid side-chain of aspartic acid and glutamic acid, which are 3.71 and 4.15, respectively.[30] Therefore, pKa1 and pKa2 are assigned to aspartic acid residues, whereas the two slightly higher values, pKa3 and pKa4, may be assigned to glutamic acid side chains. The titration of A with UO22 + , followed by CD, shows characteristic features of the evolution of the peptide-scaffold structure upon metal binding (Figure 2 a). A unique complex, UO2– A, is formed as indicated by two isodichroic points and no significant evolution of the spectrum in an excess of metal ion. The spectra of the free peptide A and the UO2–A complex are dramatically different at pH 6. Indeed, the CD spectrum of the free compound is reminiscent of a random coil conformation with a negative band near 200 nm.[31] This is consistent with the four carboxylic acids being deprotonated, which leads to major electrostatic repulsions between the four negatively charged amino acid side chains in position 1, 3, 6, and 8, and to destabilization of the b-sheet structure of the peptide scaffold at this pH. In addition, the CD spectrum of the 1:1 complex UO2–A at the same pH shows a weak negative band at 224 nm and a large positive band at 195 nm, characteristic of a class B spectrum in Woody’s theoretical approach[32] and consistent with b-turn conformations.[33] These CD signatures demonstrate the formation of a well-defined uranyl complex, UO2– A, in which the prototype peptide A adopts a b-sheet structure to promote the coordination of the carboxylate amino acid side chains in the equatorial plane of the uranyl cation. Uranyl complexation was also efficiently detected through a large quenching of tryptophan fluorescence at 350 nm upon excitation at 280 nm. A similar effect was reported with tyrosine, with which quenching was used to study uranyl complexation with calmodulin derivatives[21] or transferrin.[6] The evolution of Trp fluorescence spectra during the titration of peptide A with uranyl nitrate is shown in Figure 3. An endpoint was detected for one UO22 + equivalent and confirms the formation of the complex UO2–A. The conditional stability constant of UO2–A at pH 6 was measured by fitting florescence titrations. Fluorescence spectra were recorded in dilute solutions ([A] 10 mm, Abs (280 nm) < 0.05) to avoid inner-filter effects and have a linear relationship between the fluorescence intensity and the concentrations. Moreover, uranyl is known to form many hydroxo complexes (more than 6 are reported in the NIST database),[30] which are
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Full Paper UO2 þ P ¼ UO2 P
K C ¼ 1=K D ð1Þ
Figure 2. CD titration of representative peptides (ca. 10 mm) with UO22 + (0 to 1.5 equiv, aliquots of 0.25 equiv) at pH 6.0, i indicates isodichroic points.
Six different conditions were used and these point consistently to a logKC value of 8.1 0.2 (Table 2). The pKa values of A account for the slightly higher KC value measured at pH 6.5 (entry 3, Table 2). Indeed, the difference calculated in logKC value from pH 6 to 6.5 is only + 0.1. In the following sections, the conditions of entry 5 were used because uranyl complexes of IDA display a simpler speciation. In conclusion, the prototype cyclodecapeptide A forms a well-defined 1:1 uranyl complex UO2–A, which exhibits a bsheet structure consistent with the coordination of the four carboxylate side chains to uranyl. This de novo-designed peptide exhibits a higher affinity for
Table 2. Conditional stability constant values (logKC) and dissociation constants KD for the complex UO2A in MES 20 mm obtained in various experimental conditions. [A] 10 mm. Entry pH Salt [0.1 M] Competitor [Competitor] [mM] logKC KD 109 1 2 3 4 5 6[a]
6.0 6.0 6.5 6.0 6.0 6.0
KNO3 KNO3 KNO3 KNO3 NaCl NaCl
CO32 CO32 CO32 IDA IDA IDA
0.5 0.1 0.1 0.1 0.1 0.02
8.0(1) 7.8(1) 8.3(1) 7.9(1) 8.1(1) 8.1(1)
10 2 16 4 51 13 3 82 82
[a] [A] 2 mm
Figure 3. Fluorescence titration of A (ca. 10 mm) with UO22 + (0 to 4 equiv) at pH 6.0, MES buffer (20 mm), KNO3 (0.1 m) with excitation at 280 nm. Inset: variation of the intensity at the peak maximum (350 nm) with UO22 + .
significantly present even at pH 6. To control the nature of the uranyl complexes formed during the titrations, two different competing ligands with well-defined speciation with UO22 + were added to the solution: carbonate (CO32) or iminodiacetic acid (IDA). The competing ligand concentration was chosen, 1) to minimize the formation of hydroxo species (< 2 %), and 2) to get reliable binding curves. The conditional stability constants of the uranyl complexes formed with the competing ligands were calculated from published values[30, 34] and inserted in the fitting procedure according to models described in the experimental section (see the Supporting Information). The KC constants reported in Table 2 were obtained for the formation of a unique UO2–A complex [Eq. (1)] as demonstrated by MS and CD experiments in similar conditions. Chem. Eur. J. 2014, 20, 16566 – 16573
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uranyl than calcium binding sites directly derived from calmodulin (logKC = 7.5), which organize similar coordinating atoms in a structured loop.[21] The series of decapeptides listed in Table 1 with various sequences and structures are studied in the following section to determine the key parameters that induce a large affinity for uranyl. Influence of the nature of the four-coordinating acidic residues (Asp or Glu): peptides of group A The nature of the acidic residues (Asp or Glu) introduced in positions 1, 3, 6, and 8 of the cyclic peptide scaffold for metal coordination is varied in the first series of peptides (Group A in Table 1). A1 and A2 are variants of the prototype peptide A with 4 glutamic acids as potential coordinating amino acids, whereas A4 and A5 bear 4 aspartic acids. ESI-MS spectra show the formation of 1:1 uranyl complexes for all these peptides.
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Full Paper Examples of MS spectra recorded in the positive mode are shown in Figure 4. Even though MS is not a quantitative method, the response coefficients are expected to be very similar for the series of peptides that have the same charge, such as A, A1, and A4. The relative peak intensities of the uranyl complex and the corresponding free peptide (in MS spectra acquired in the same conditions), clearly show that UO2–A1 is more abundant than UO2–A4.
Figure 5. Titration of 10 mm peptide solutions (MES buffer 20 mm, pH 6.0, 298 K) with uranyl in the presence of 100 mm IDA. Top: Fluorescence intensity of tryptophan at 350 nm upon excitation at 280 nm as a function of uranyl equivalents. Bottom: Corresponding Stern–Volmer plots.
Figure 4. (+) ESI-MS spectra of the UO22 + complexes (ca. 100 mm) with representative peptides in ammonium acetate buffer (20 mm, pH 6.9). Adducts with sodium are detected at higher m/z than the main signals indicated on the figure.
The higher stability of the former complex is confirmed by the analyses of the fluorescence titrations. Striking differences are detected in the tryptophan fluorescence quenching during titration with uranyl. Figure 5 displays the evolution of the Trp emission at 350 nm as a function of uranyl content for three peptides of group A. The quenching is a lot more efficient for A1 containing four glutamate residues than for A4 containing four aspartate residues. The Stern–Volmer plots shown in the bottom half of Figure 5 also reflect this tendency. Indeed, the slope of the plot, which gives an evaluation of the conditional stability constant with respect to IDA, is one order of magnitude larger for A1 (logKSV = 5.6) than for A4 (logKSV = 4.6). The conditional stability constants obtained by fitting the titrations with IDA, taking into account the constants of uranyl complexes of IDA, are reported in Table 1 and also demonstrate one order of magnitude difference between A1 and A4. Whereas A1 has a very similar affinity to the prototype peptide A, the incorporation of 4 aspartic acids in A4 significantly decreases the uranyl complex stability. One explanation for the greater affinity of glutamate-based peptides compared to aspartate-based peptides is the larger basicity of glutamate with respect to aspartate. Because carboxylate–uranyl interactions are mainly electrostatic due to the hard nature of both the metal cation and the donor, the stability should correlate with the basicity of simple donors, as deChem. Eur. J. 2014, 20, 16566 – 16573
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scribed for lanthanide complexes with aminocarboxylate ligands.[35] The carboxylate side chain of Asp is therefore expected to have a lower affinity for UO22 + than that of Glu. A significant structural effect is also detected on the circular dichroism spectra. Whereas A and A1 show characteristic CD features of b-sheet formation upon uranyl coordination (Figures 2 a and 2 b, respectively), peptide A4 demonstrates a poor evolution in the CD spectra and, more importantly, the absence of characteristic features of a b-sheet structure for the uranyl complex (Figure 2 c). This indicates that the cyclic peptide backbone is too constrained to benefit from the stable b-sheet conformation and the tight coordination of the four short side chains of the aspartate residues in A4. In these cyclic peptides, both steric constraints and basicity favor the glutamate–uranyl interaction. For amino acids in positions 2 and 7, the side-chain oriented on the opposite face to the metal ion coordination was also varied. The positively charged guanidinium function of arginine in position 2 in peptides A1 and A4 was replaced by the polar alcohol group of serine in peptides A2 and A5, without any significant effect on the affinity for uranyl. Similarly, peptide A3 with a fluorescent tyrosine (Tyr) shows similar uranyl coordination properties to A1, with a tryptophan. This confirms that the two faces defined by the cyclic peptide scaffold are topologically independent.
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Full Paper Influence of the structure: peptides of group B The two decapeptides B1 and B2 were studied to determine the influence of the scaffold on the uranyl complexation properties. In peptide B1, one Pro–Gly motif, which induces the formation of a b-turn, was replaced by a less constrained sequence Ser–Gly, which promotes the formation of a flexible loop. The 1:1 complex UO2–B1 is detected by MS and its CD spectrum is reminiscent of a b-sheet structure (Figure 2 d) even though the molar ellipticity ([V]195nm 153 000 deg cm2 dmol1) is significantly lower than the one measured for UO2–A ([V]195nm 189 000 deg cm2 dmol1). Moreover, the stability constant of UO2–B1 is 4 times lower than for the prototype peptide A (Table 1), as shown by the binding isotherms and on the Stern–Volmer plots in Figure 5. The comparison between the b-sheet prototype peptide A and its more flexible analogue B1 is clear evidence that the b-sheet structure is optimal when constraints associated with short side chains are not too large. The behaviour of the linear equivalent of A1, peptide B2, is even more striking. Indeed, it is totally different from all the cyclic compounds. The ES–MS spectrum in Figure 4 shows almost no formation of the uranyl complex and also displays signals due to polymetallic adducts. Furthermore, the slight fluorescence quenching upon uranyl addition to B2 is indicative of a dramatically lower metal-binding affinity. The comparison of peptides of group A with the two analogues B1 and B2 exemplifies the perfect adjustment of the bsheet cyclic peptide scaffold in A or A1 for uranyl complexation.
Influence of nonacidic-coordinating amino acids: peptides of group C All the peptides of group C have only three acidic residues. All of them demonstrate a significantly lower affinity for uranyl. Indeed, in contrast to peptides of group A and B1, their uranyl complexes are barely detectable in the MS spectra (see Figure S1 in the Supporting Information). Moreover, the fluorescence quenching of tryptophan is a lot smaller and no real binding isotherms could be observed (see Figure S2 in the Supporting Information). The Stern–Volmer constants are lower than 4.5 (in log units) and only an upper limit of the affinity constant could be determined (logKC < 7). In this group, only peptide C2 demonstrates a larger fluorescence quenching corresponding to a logKC = 7.4. C2 is equivalent to A1 with a serine instead of a glutamate in position 3. The slightly larger affinity of C2 for uranyl, in comparison with other peptides of group C, suggests that the thermodynamic contribution to the affinity constant of the glutamate in position 3 in A1 is lower. Finally, the substitution of glutamate by either glutamine or histidine in peptides C4 and C5, respectively, leads to a decrease of more than one order of magnitude in uranyl affinity. Histidine is a cyclic nitrogen donor that is more constrained than the oxygen donors studied in this work. Therefore, peptide C6 was also considered with the histidine in a loop to release some constraint in its coordination. Chem. Eur. J. 2014, 20, 16566 – 16573
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Peptides of group C, lacking one carboxylate side chain compared with peptides of group A, show a significant loss in affinity for uranyl. This proves the involvement of the four amino acid side chains of Asp and Glu in the dioxo-cation coordination with peptides of group A. Moreover, neutral donors, such as glutamine in C4, or histidine in C5 and C6, do not play a significant role in the uranyl complex stability.
Conclusion The ability of a series of decapeptides to bind uranyl has been investigated with complementary analytical and spectroscopic methods to determine the key parameters for the formation of stable uranyl–peptide complexes. Structured cyclic peptide scaffolds were chosen as promising candidates to coordinate uranyl thanks to four amino acid side chains pre-oriented towards the dioxo-cation equatorial plane. The b-sheet structure of the prototype peptide A with two aspartates and two glutamates, and of its equivalents A1 and A2 incorporating four glutamates, is perfectly adapted to the coordination of UO22 + . Indeed, the more flexible cyclic compound B1, having only one turn, displays a lower affinity for uranyl. The behaviour of the linear decapeptide B2 is even more striking, because it forms nondefined polymetallic uranyl complexes of low stability. Compounds with four aspartate residues, A4 and A5, with short carboxylate side chains, show the role of the conformational constraints in the metal complex formation. These two cyclic peptides also complex uranyl in a 1:1 complex, but their circular dichroism signature reveals that the scaffold cannot adopt a b-sheet structure. As a result, the stability constants are one order of magnitude lower than those obtained with peptides A and A1, which have glutamate residues. Actually, the molar ellipticity at 195 nm of the uranyl complex is directly correlated to its stability constant, which confirms that the bsheet structure is a key parameter for high stability in the peptide series. Sequences with only three acidic amino acids display significantly lower affinities for uranyl, which proves the involvement of the four amino acid side chains of Asp and Glu in the dioxo-cation coordination with peptides of group A. Importantly, the de novo-designed peptides A, A1 and A2 exhibit a higher affinity for uranyl (logKC = 8.1–8.4) than calcium binding sites derived from calmodulin, which had their affinity for uranyl measured in exactly the same conditions (logKC = 7.5).[21] This suggests that the presentation of four carboxylate side chains in the b-sheet structure is particularly adapted to uranyl coordination even in comparison with the structured EF-hand loops of calmodulin. Moreover the cyclodecapeptide A is selective for uranyl with respect to calcium by more than four orders of magnitude. Indeed, the affinity of A for calcium is only logKC = 3.7.[24] This study shows that simple uranyl-binding sequences in vivo, built with aspartates and glutamates, should display conditional affinity constants in the range logKC = 7–9, but not significantly higher. Furthermore, the peptide scaffold structure is a key parameter for the design of efficient uranyl-binding peptides that control metal–complex speciation. Within the stud-
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Full Paper ied sequences, A1 and A2 appear as good starting points for the design of high-affinity uranyl-chelating peptides. Phosphorous groups are known to strongly bind hard ions and, in particular, uranyl.[36] For instance, the insertion of a phosphoserine residue in the binding loop of a calmodulin-derived peptide was successful in enhancing the uranyl complex stability.[21] Therefore, phosphorylated amino acids are currently inserted in the cyclic peptide sequence in order to test the complexation properties of peptides bearing both carboxylate and phosphate side chains, which may also be relevant to some uranylbinding sites in vivo.[11, 19] Another strategy involves developing unnatural amino acids bearing metal-chelating groups on their side chains. Peptides incorporating such unnatural residues have been shown to significantly improve the coordination properties of peptides for lanthanides[37] and are currently being developed for uranyl chelation.
cell in centimeters, and c is the peptide concentration in moles per liter.
Fluorescence titrations Spectra were recorded on a LS50B spectrofluorimeter connected to a computer equipped with FLWILAB 2.0. The measurements were performed at 298 K. Trp fluorescence titrations were performed with 280 nm excitation (excitation slit: 3.0 nm). The emission slit was adjusted (3.5–4.5 nm) to avoid signal saturation. Experiments were repeated to ensure reproducibility.
Acknowledgements
Experimental Section
We thank Olivier Snque and Didier Boturyn for the initial synthesis of the prototype peptide A. This research was supported by the “Programme Transversal Toxicologie du CEA”, the NRBC-E program and the Labex ARCANE (Grant ANR-11LABX-0003-01).
The peptide syntheses and additional experimental procedures for physico-chemical experiments are reported in the Supporting Information.
Keywords: bioinorganic chemistry · carboxylate ligands · fluorescence · peptides · uranium
Preparation of aqueous solutions A stock uranyl solution (ca. 10 mm) was prepared from uranyl dinitrate hexahydrate in nitric acid (0.01 m). The precise uranyl concentration was obtained by measuring the absorbance of an aliquot compared to an ICP Uranium standard (1000 mg U per mL in 1 % HNO3) at l = 415 nm. Peptide solutions were prepared freshly before use and the precise peptide concentration was determined by recording a UV spectrum (e280 = 5690 L mol1 cm1 owing to the presence of a Trp residue). MES buffer was prepared by dissolving solid 2-(N-morpholino)ethanesulfonic acid in H2O and by adjusting the pH to 6.0 or 6.5 with KOH. Solutions of IDA or carbonate were prepared in the buffer from solid iminodiacetic acid or sodium carbonate.
ESI-MS Peptide solutions (100 mm) were prepared in an ammonium acetate buffer (20 mm, pH 7). The stock uranyl solution was added to prepare aliquots containing 0, 0.5, 1, and 2 equiv of UO22 + per peptide. Mass spectra were recorded on a LXQ type THERMO SCIENTIFIC spectrometer, equipped with an electrospray ionization source and a linear-trap detector. Solutions were injected in the spectrometer at 10 mL min1 flow rate. Ionization voltage and capillary temperature were about 2 kV and 250 8C, respectively. The source settings were the same (sheath gas, auxiliary gas, capillary voltage, and tube lens) for all series of peptides with comparable data.
CD titrations CD spectra were recorded at 25 8C on an Applied Photophysics Chirascan Spectrometer in a 1 cm path cell. The peptide concentration was 10 mm in water and the pH was adjusted to 6.0 with KOH. All spectra were obtained from 320 to 190 nm with a 1 nm data interval, a time constant of 2 s and a band width of 1 nm, with 3 scans. CD spectra are reported in molar ellipticity ([V] in units of deg cm2 dmol1). [V] = qobs/(10lc), where qobs is the observed ellipticity in millidegrees, l is the optical path length of the Chem. Eur. J. 2014, 20, 16566 – 16573
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[1] a) P. W. Durbin, Actinides in Animals and Man in The Chemistry of the Actinide and Transactinide Elements (Eds.: L. R. E. Morss, N. Fuger, J. Katz), Springer, Dordrecht (The Netherlands), 2006, pp. 3339 – 3440; b) D. M. Taylor, S. K. Taylor, Rev. Environ. Health 1997, 12, 147 – 158. [2] E. Ansoborlo, O. Prat, P. Moisy, C. Den Auwer, P. Guilbaud, M. Carriere, B. Gouget, J. Duffield, D. Doizi, T. Vercouter, C. Moulin, V. Moulin, Biochimie 2006, 88, 1605 – 1618. [3] a) E. Ansoborlo, B. Amekraz, C. Moulin, V. Moulin, F. Taran, T. Bailly, R. Burgada, M. H. Henge-Napoli, A. Jeanson, C. Den Auwer, L. Bonin, P. Moisy, C. R. Chim. 2007, 10, 1010 – 1019; b) A. E. V. Gorden, J. D. Xu, K. N. Raymond, P. Durbin, Chem. Rev. 2003, 103, 4207 – 4282; c) R. J. Abergel, P. W. Durbin, B. Kullgren, S. N. Ebbe, J. D. Xu, P. Y. Chang, D. I. Bunin, E. A. Blakely, K. A. Bjornstad, C. J. Rosen, D. K. Shuh, K. N. Raymond, Health Phys. 2010, 99, 401 – 407; d) M. Sturzbecher-Hoehne, G. J.-P. Deblonde, R. J. Abergel, Radiochim. Acta 2013, 101, 359 – 366. [4] J. D. Van Horn, H. Huang, Coord. Chem. Rev. 2006, 250, 765 – 775. [5] a) S. Scapolan, E. Ansoborlo, C. Moulin, C. Madic, Radiat. Prot. Dosim. 1998, 79, 505 – 508; C. Vidaud, S. Gourion-Arsiquaud, F. Rollin-Genetet, C. Torne-Celer, S. Plantevin, O. Pible, C. Berthomieu, E. Quemeneur, Biochemistry 2007, 46, 2215 – 2226; b) G. Montavon, C. Apostolidis, F. Bruchertseifer, U. Repinc, A. Morgenstern, J. Inorg. Biochem. 2009, 103, 1609 – 1616. [6] J. Michon, S. Frelon, C. Garnier, F. Coppin, J. Fluoresc. 2010, 20, 581 – 590. [7] C. Vidaud, A. Dedieu, C. Basset, S. Plantevin, I. Dany, O. Pible, E. Quemeneur, Chem. Res. Toxicol. 2005, 18, 946 – 953. [8] a) O. Pible, C. Vidaud, S. Plantevin, J. L. Pellequer, E. Quemeneur, Protein Sci. 2010, 19, 2219 – 2230; b) O. Pible, P. Guilbaud, J. L. Pellequer, C. Vidaud, E. Quemeneur, Biochimie 2006, 88, 1631 – 1638. [9] a) A. Dedieu, F. Berenguer, C. Basset, O. Prat, E. Quemeneur, O. Pible, C. Vidaud, J. Chromatogr. A 2009, 1216, 5365 – 5376; b) C. Basset, A. Dedieu, P. Guerin, E. Quemeneur, D. Meyer, C. Vidaud, J. Chromatogr. A 2008, 1185, 233 – 240; c) L. Reisser-Rubrecht, C. Torne-Celer, W. Renier, O. Averseng, S. Plantevin, E. Quemeneur, L. Bellanger, C. Vidaud, Chem. Res. Toxicol. 2008, 21, 349 – 357; d) O. Averseng, A. Hagege, F. Taran, C. Vidaud, Anal. Chem. 2010, 82, 9797 – 9802. [10] C. Basset, O. Averseng, P. J. Ferron, N. Richaud, A. Hagege, O. Pible, C. Vidaud, Chem. Res. Toxicol. 2013, 26, 645 – 653. [11] L. Qi, C. Basset, O. Averseng, E. Quemeneur, A. Hagege, C. Vidaud, Metallomics 2014, 6, 166 – 176. [12] a) P. Rousselot-Pailley, O. Snque, C. Lebrun, S. Crouzy, D. Boturyn, P. Dumy, M. Ferrand, P. Delangle, Inorg. Chem. 2006, 45, 5510 – 5520; b) O.
16572
2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Full Paper
[13] [14]
[15]
[16]
[17] [18] [19] [20] [21] [22]
[23]
[24] [25]
Snque, S. Crouzy, D. Boturyn, P. Dumy, M. Ferrand, P. Delangle, Chem. Commun. 2004, 770 – 771. E. Rossy, O. Snque, D. Lascoux, D. Lemaire, S. Crouzy, P. Delangle, J. Coves, Febs Lett. 2004, 575, 86 – 90. a) A. Jancs, D. Szunyogh, F. H. Larsen, P. W. Thulstrup, N. J. Christensen, B. Gyurcsik, L. Hemmingsen, Metallomics 2011, 3, 1331 – 1339; b) V. L. Pecoraro, A. F. A. Peacock, O. Iranzo, M. Luczkowski, Understanding the Biological Chemistry of Mercury Using a de novo Protein Design Strategy in Bioinorganic Chemistry: Cellular Systems and Synthetic Models (Eds.: E. C. Long, M. J. Baldwin), ACS, Washington D.C., 2009, pp. 183 – 197. a) A.-S. Jullien, C. Gateau, C. Lebrun, I. Kieffer, D. Testemale, P. Delangle, Inorg. Chem. 2014, 53, 5229 – 5239; b) A.-S. Jullien, C. Gateau, I. Kieffer, D. Testemale, P. Delangle, Inorg. Chem. 2013, 52, 9954 – 9961; c) A. M. Pujol, C. Lebrun, C. Gateau, A. Manceau, P. Delangle, Eur. J. Inorg. Chem. 2012, 3835 – 3843; d) A. M. Pujol, C. Gateau, C. Lebrun, P. Delangle, Chem. Eur. J. 2011, 17, 4418 – 4428; e) A. M. Pujol, C. Gateau, C. Lebrun, P. Delangle, J. Am. Chem. Soc. 2009, 131, 6928 – 6929. a) S. Pires, J. Habjanic, M. Sezer, C. M. Soares, L. Hemmingsen, O. Iranzo, Inorg. Chem. 2012, 51, 11339 – 11348; b) C. Gateau, P. Delangle, Ann. N. Y. Acad. Sci. 2014, 1315, 30 – 36; c) A. M. Pujol, M. Cuillel, A.-S. Jullien, C. Lebrun, D. Cassio, E. Mintz, C. Gateau, P. Delangle, Angew. Chem. Int. Ed. 2012, 51, 7445 – 7448; Angew. Chem. 2012, 124, 7563 – 7566. M. Tegoni, F. T. Yu, M. Bersellini, J. E. Penner-Hahn, V. L. Pecoraro, Proc. Natl. Acad. Sci. USA 2012, 109, 21234 – 21239. H. Huang, S. Chaudhary, J. D. Van Horn, Inorg. Chem. 2005, 44, 813 – 815. S. Safi, G. Creff, A. Jeanson, L. Qi, C. Basset, J. Roques, P. L. Solari, E. Simoni, C. Vidaud, C. Den Auwer, Chem. Eur. J. 2013, 19, 11261 – 11269. L. Le Clainche, C. Vita, Environ. Chem. Lett. 2006, 4, 45 – 49. R. Pardoux, S. Sauge-Merle, D. Lemaire, P. Delangle, L. Guilloreau, J.-M. Adriano, C. Berthomieu, PLOS ONE 2012, 7, e41922. a) L. Zhou, M. Bosscher, C. Zhang, S. Ozcubukcu, L. Zhang, W. Zhang, C. J. Li, J. Liu, M. P. Jensen, L. Lai, C. He, Nat. Chem. 2014, 6, 236 – 241; b) S. V. Wegner, H. Boyaci, H. Chen, M. P. Jensen, C. He, Angew. Chem. Int. Ed. 2009, 48, 2339 – 2341; Angew. Chem. 2009, 121, 2375 – 2377. a) P. Dumy, I. M. Eggleston, G. Esposito, S. Nicula, M. Mutter, Biopolymers 1996, 39, 297 – 308; b) S. Peluso, T. Ruckle, C. Lehmann, M. Mutter, C. Peggion, M. Crisma, ChemBioChem 2001, 2, 432 – 437. C. S. Bonnet, P. H. Fries, S. Crouzy, O. Snque, F. Cisnetti, D. Boturyn, P. Dumy, P. Delangle, Chem. Eur. J. 2009, 15, 7083 – 7093. a) G. T. Dolphin, P. Dumy, J. Garcia, Angew. Chem. Int. Ed. 2006, 45, 2699 – 2702; Angew. Chem. 2006, 118, 2765 – 2768; b) P. Murat, R. Bonnet, A. Van der Heyden, N. Spinelli, P. Labbe, D. Monchaud, M. P. Teulade-Fichou, P. Dumy, E. Defrancq, Chem. Eur. J. 2010, 16, 6106 –
Chem. Eur. J. 2014, 20, 16566 – 16573
www.chemeurj.org
[26]
[27]
[28]
[29] [30]
[31]
[32] [33] [34] [35] [36]
[37]
6114; c) M. Galibert, O. Renaudet, P. Dumy, D. Boturyn, Angew. Chem. Int. Ed. 2011, 50, 1901 – 1904; Angew. Chem. 2011, 123, 1941 – 1944. a) D. Boturyn, J. L. Coll, E. Garanger, M. C. Favrot, P. Dumy, J. Am. Chem. Soc. 2004, 126, 5730 – 5739; b) S. Grigalevicius, S. Chierici, O. Renaudet, R. Lo-Man, E. Deriaud, C. Leclerc, P. Dumy, Bioconjugate Chem. 2005, 16, 1149 – 1159. A. M. Pujol, M. Cuillel, O. Renaudet, C. Lebrun, P. Charbonnier, D. Cassio, C. Gateau, P. Dumy, E. Mintz, P. Delangle, J. Am. Chem. Soc. 2011, 133, 286 – 296. a) A. Fragoso, P. Lamosa, R. Delgado, O. Iranzo, Chem. Eur. J. 2013, 19, 2076 – 2088; b) A. Fragoso, R. Delgado, O. Iranzo, Dalton Trans. 2013, 42, 6182 – 6192. C. S. Bonnet, P. H. Fries, S. Crouzy, P. Delangle, J. Phys. Chem. B 2010, 114, 8770 – 8781. R. M. Smith, A. E. Martell, R. J. Motekaitis, NIST Critically Selected Stability Constants of Metal Complexes Database, NIST Standard Reference Database 46, 2001. N. Sreerama, R. W. Woody in Circular Dichroism of peptides and proteins, in Circular Dichroism: Principles and Applications, 2ednd ed(Eds.: N. Berova, K. Nakanishi, R. W. Woody), Wiley, New York, 2000, pp. 601 – 620. J. A. Smith, L. G. Pease, CRC Crit. Rev. Biochem. 1980, 8, 315 – 399. B. Imperiali, S. L. Fisher, R. A. Moats, T. J. Prins, J. Am. Chem. Soc. 1992, 114, 3182 – 3188. J. Jiang, M. J. Sarsfield, J. C. Renshaw, F. R. Livens, D. Collison, J. M. Charnock, M. Helliwell, H. Eccles, Inorg. Chem. 2002, 41, 2799 – 2806. T. F. Gritmon, M. P. Goedken, G. R. Choppin, J. Inorg. Nucl. Chem. 1977, 39, 2021 – 2023. a) W. A. Reed, L. F. Rao, P. Zanonato, A. Y. Garnov, B. A. Powell, K. L. Nash, Inorg. Chem. 2007, 46, 2870 – 2876; b) M. Sawicki, J. M. Siaugue, C. Jacopin, C. Moulin, T. Bailly, R. Burgada, S. Meunier, P. Baret, J. L. Pierre, F. Taran, Chem. Eur. J. 2005, 11, 3689 – 3697. a) L. Ancel, A. Niedz´wiecka, C. Lebrun, C. Gateau, P. Delangle, C. R. Chim. 2013, 16, 515 – 523; b) A. Niedz´wiecka, F. Cisnetti, C. Lebrun, C. Gateau, P. Delangle, Dalton Trans. 2012, 41, 3239 – 3247; c) A. Niedz´wiecka, F. Cisnetti, C. Lebrun, P. Delangle, Inorg. Chem. 2012, 51, 5458 – 5464; d) F. Cisnetti, C. Lebrun, P. Delangle, Dalton Trans. 2010, 39, 3560 – 3562; e) F. Cisnetti, C. Gateau, C. Lebrun, P. Delangle, Chem. Eur. J. 2009, 15, 7456 – 7469.
Received: July 23, 2014 Published online on October 16, 2014
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