DOI: 10.1002/chem.201404909

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& Disulfide Bonds

Extraordinary Modulation of Disulfide Redox-Responsiveness by Cooperativity of Twin-Disulfide Bonds Linxiang Zhai, Jingjing Liang, Xiangqun Guo, Yibing Zhao, and Chuanliu Wu*[a]

Abstract: Disulfide bonds have frequently been incorporated into synthetic materials to promote sensitivity of the systems towards different redox environments. Although many strategies have been developed to rationally tune the stability of disulfide linkers, methods to tune their responsiveness towards different redox environments remain elusive. In this work we have developed and explored a disulfide linker bearing two independent disulfide bonds, referred to as a twin-disulfide linker. We have demonstrated that the twindisulfide linker displays an ultrahigh stability at lower con-

centrations of reducing agent or in weakly reducing environments without a significant compromise in the sensitivity of its response to highly reducing environments such as cytoplasm, a feature that is in remarkable contrast to the traditional single disulfide bonds. Such an extraordinary responsiveness arises from the cooperativity of the twin-disulfide bonds, which should be of particular interest for applications such as controlled drug delivery and sensing, because relatively large differences in disulfide stability in different redox environments is desired in these applications.

Introduction Disulfide bonds are the most prevalent dynamic covalent bonds in proteins; they not only play an important role in regulating the folding of proteins, but make them responsive to the different redox environments in the body.[1] The dynamic feature of a disulfide bond (R* S S R*) lies in its exchange reaction with thiolates (R S ) through an SN2 mechanism to produce a different disulfide and new thiolates (Figure 1a).[2] Although the reaction mechanism seems straightforward, in proteins, the dynamic feature of disulfides is complicatedly and precisely regulated by the primary sequence and protein structure so that proteins can fold correctly and/or the disulfide bonds can make response to a wide range of different redox potentials.[1e, 2, 3] In particular, disulfides in proteins, upon reduction by thiol–disulfide exchange, have a strong propensity towards the reverse reaction (Figure 1a), a process that could markedly modulate their stability (or redox potential, responsiveness).[3c, d, 4] However, although a disulfide bond formed from two isolated thiolates can be reduced in biological environments, the reverse thiol–disulfide exchange process seldom takes place because of the physical separation between the newly formed thiolate and disulfide, which leads to permanent cleavage.[5]

[a] L. Zhai, J. Liang, Prof. Dr. X. Guo, Prof. Dr. Y. Zhao, Prof. Dr. C. Wu The MOE Key Laboratory of Spectrochemical Analysis and Instrumentation State Key Laboratory of Physical Chemistry of Solid Surfaces Department of Chemistry, College of Chemistry and Chemical Engineering Xiamen University, Xiamen, 361005 (P. R. China) E-mail: [email protected] Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/chem.201404909. Chem. Eur. J. 2014, 20, 17507 – 17514

Figure 1. a) Generic thiol–disulfide exchange reactions taking place during the reduction of disulfide bonds formed from two isolated thiolates and in proteins. b) Thiol–disulfide exchange reactions taking place in twin-disulfide linkers bearing two independent disulfide bonds.

Disulfides have also been frequently incorporated into synthetic materials to promote the sensitivity of the systems towards different redox environments in extra- and intracellular spaces, for example, drug delivery systems and imaging probes.[6] In many instances, the disulfide bond has been exploited as a linker to directly link a payload with a carrier, the responsiveness of these systems relying primarily on the cleavage kinetics of the disulfide linkers.[5b, 6d, 7] In general, to improve the performance of these systems in vivo, the stability of disulfide linkers has to be rationally tuned to prevent prema-

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Full Paper ture cleavage during circulation or in extracellular spaces.[6d, 7b, 8] Thus far, two strategies have been explored for this purpose, namely the introduction of steric hindrance and microenvironment modulation, respectively.[7b, 8, 9] For example, it has been reported that the stability of disulfide bonds can be predictably tuned over around three orders of magnitude by synergistic manipulation of steric hindrance and microenvironment effects.[7a] However, despite the ease of achieving tunable disulfide stability, the degree of difference in disulfide stability under different reducing conditions cannot be tuned, that is, the manner of response of the synthetic disulfide linkers is simply based on the linear relationship between the rate of disulfide cleavage and thiolate concentration (i.e., pseudo-firstorder reaction).[7a, 9a] This is in remarkable contrast to the disulfides in proteins, the stability of which is additionally controlled by the reverse thiol–disulfide exchange process and, as a consequence, these disulfides are capable of making diverse and more complicated responses to different redox environments. Inspired by this, we envisioned the possibility of manipulating the manner of response of disulfide linkers through the control of the reverse disulfide exchange reaction. This exploration would provide an unconventional route to modulating the responsiveness of synthetic disulfide linkers. To accomplish this goal, in this work we developed and explored a disulfide linker bearing two independent disulfide bonds, referred to as a twin-disulfide linker (Figure 1b). The twin-disulfide linker, upon partial reduction at lower concentrations of thiolate (i.e., cleavage of one of the two disulfide bonds), can subsequently undergo a rapid reverse thiol–disulfide exchange due to the effect of cooperativity between the twin disulfide bonds, which leads to the re-formation of the original twin-disulfide linker. However, in the case of higher concentrations of thiolate, concomitant reduction of the twin disulfides was observed (Figure 1b), which led to permanent cleavage of the twin-disulfide linker. We have therefore succeeded in developing a new type of disulfide linker with an extraordinary responsiveness towards different redox environments. Moreover, we explored how the twin-disulfide linker responds to redox buffers that mimic redox environments in the body, such as extracellular spaces, endocytic organelles, and cytoplasm. Finally, we explored the feasibility of modifying the twin-disulfide linker with functional molecules by click reactions and how asymmetric twin-disulfide bonds/linkers can be prepared, which would greatly benefit the development of various redox-responsive synthetic systems through the incorporation of twin-disulfide linkers.

Results and Discussion Molecular design and synthesis Previous work on twin disulfides formed by the oxidation of peptides containing the CXC motif (cysteine-any-cysteine) unambiguously revealed the mechanism of thiol–disulfide exchange in which the equilibrium between the twin-disulfide CXC dimer and its close-loop monomer is directed towards the dimer.[10] However, owing to the structural flexibility of the CXC Chem. Eur. J. 2014, 20, 17507 – 17514

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motif, once one of the twin-disulfide bonds has been cleaved, the reduced cysteine has a propensity to attack the other disulfide bond and, as a result, both disulfide bonds will be cleaved, a process that can reduce, to a great extent, the difference in the responsiveness of twin-disulfide bonds in redox media as compared with single-disulfide bonds.[10] In addition, the CPPC motif (cysteine-proline-proline-cysteine) has been found to show a high propensity towards the formation of a parallel dimer upon oxidation.[11] However, the conformation of the CPPC motif is still not rigid enough to prevent intramolecular disulfide formation. Thus, in this study we aimed to design a twin-cysteine-containing building block bearing a structurally rigid backbone for the development of twin disulfides. To this end, 1,4,5,8-naphthalenediimide (NDI) bearing two terminal cysteines (C-NDI-C) was designed and synthesized (Figure 2a). The synthetic procedures have been reported in the literature[12] and its characterization is reported in Figure S1 in the Supporting Information. C-NDI-C was originally synthesized for the construction of dynamic combinatorial libraries.[12c, d] The single-cysteine analogue was also synthesized by blocking one of the two thiols with 2-iodoacetamide as a capping agent (Figures 2a and S2 in the Supporting Information). It is worth mentioning that the terminal cysteines also provide additional carboxylate anions, which is not only important for water solubility, but they can also be exploited as functional groups for further modification or functionalization. Formation of a twin-disulfide dimer/linker We then investigated whether C-NDI-C can be oxidized to form twin disulfides. C-NDI-C was first incubated in phosphate buffer and aqueous DMSO solution. Oxidation was monitored by HPLC and mass spectrometry. We observed that the major product formed after the oxidation was the C-NDI-C dimer (Figure 2b, as identified by mass spectrometry, see Figure S3 in the Supporting Information). In particular, the closed-loop monomer was not observed due to the structural rigidity of the NDI moiety, which makes the formation of an intramolecular disulfide bond impossible. This would significantly push the equilibrium of disulfide formation towards the twin-disulfide dimer. The trimer of C-NDI-C was also observed during the oxidation, but the yield was significantly less than that of the dimer (Figures 2b and S4). In addition, the addition of DMSO can dramatically accelerate the rate of oxidation, as expected,[9a, 13] and it has no effect on the equilibrium of disulfide formation. C-NDI-C was then incubated in phosphate buffer containing oxidized glutathione (GSSG; glutathione, GSH). As a result of thiol–disulfide exchange, C-NDI-C can form mixed GSH disulfides, C-NDI-CGSH or CGSH-NDI-CGSH (Figure 2c, as identified by mass spectrometry, see Figures S5 and S6 in the Supporting Information), which is in marked contrast to what was observed with molecules containing two thiols isolated with a flexible linker, for example, the CXC motif, from which the formation of mixed disulfides is negligible.[10] In addition, it was found that C-NDI-C and the mixed disulfides are subject to intermolecular disulfide isomerization and, as a result, the twin-disul-

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Full Paper that is, no thiol–disulfide exchange was observed between them.

Redox-responsiveness of the twin-disulfide dimer/linker To explore the responsiveness of the twin-disulfide dimer in redox buffers, the C-NDI-C dimer was purified by preparative HPLC and then its thiol–disulfide exchange reaction with GSH was studied. The HPLC chromatograms shown in Figure 3a

Figure 2. a) Molecular structures of C-NDI-C and its single-cysteine analogue. b) HPLC chromatograms of C-NDI-C and the C-NDI-C dimer formed by oxidation in water/DMSO mixed buffer (concentration of C-NDI-C: 0.5 mm). c) HPLC chromatograms showing the products formed after the reaction of C-NDI-C (5 mm) with different concentrations of GSSG (line 1: 0.5 mm; line 2: 5 mm). d) Schematic drawing illustrating the formation of the C-NDI-C dimer in oxidizing conditions, in which the disulfide RSSR is the oxidant. Note that dimeric intermediates were not observed during the experiment.

fide dimer of C-NDI-C formed (Figures 2c,d). This process was more clearly observed when the concentration of GSSG was relatively low, because an excess of GSSG can direct the thiol– disulfide exchange equilibrium towards the formation of CGSHNDI-CGSH. Therefore these results demonstrate that C-NDI-C has a strong propensity to form twin disulfides under oxidizing conditions (the mechanism for the formation of the C-NDI-C dimer is given in Figure 2d). Moreover, the two thiols in C-NDIC have been demonstrated to be independent of each other, Chem. Eur. J. 2014, 20, 17507 – 17514

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Figure 3. a) Representative chromatograms for the thiol–disulfide exchange cleavage of the C-NDI-C dimer (5 mM) in 10 mm GSH at pH 7.4 (phosphate buffer, 100 mm) after different times. b) Plots of the disappearance of the CNDI-C dimer (top panel) and its single-disulfide analogue (bottom panel) as a function of time in GSH buffers of different concentrations. The data are presented as the mean  SD (n = 3).

reveal the cleavage process of the C-NDI-C dimer in 10 mm GSH at pH 7.4. A gradual decrease in C-NDI-C dimer is associated with an increase in reduced C-NDI-C and mixed disulfides (C-NDI-CGSH or CGSH-NDI-CGSH). Figure 3b shows the kinetics of the cleavage of the C-NDI-C dimer in buffers containing different concentrations of GSH through plots of the percentage of residual dimer as a function of time. In addition, the kinetics of cleavage of the single-disulfide bond analogue under the same conditions is shown for comparison. It can be seen that the twin-disulfide displays different degrees of enhancement in stability compared to single-disulfide bond at different concen-

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Full Paper trations of GSH. Interestingly, the cleavage of both the C-NDI-C dimer and its single-disulfide analogue is very rapid at higher GSH concentrations (5 or 10 mm), with the half-lives of dimer disappearance (t = ) less than 15 min. However, as the concentration of GSH decreases, the difference in stability becomes more and more pronounced. In particular, at low GSH concentrations (0.1 or 0.2 mm), the C-NDI-C dimer is extremely stable, that is, only a negligible amount has been cleaved after 30 h incubation, whereas its single-disulfide analogue is steadily cleaved with half-lives of less than 2 h. To further compare the response of the twin-disulfide dimer to different concentrations of GSH with that of the single-disulfide analogue, Figure 4a shows the plots of t = as a function of 1

2

1

2

Figure 4. a) Correlation between t = and 1/[GSH] for the C-NDI-C dimer (*) and its single-disulfide analogue (&). The data is also presented on a logarithmic scale in the bottom panel for comparison. Predicted values based on the pseudo-first-order rate equation are also shown (*). *See the Supporting Information for t = calculations. b) Equilibria of the C-NDI-C dimer in redox buffers (RSH/RSSR). 1

2

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2

the inverse concentration of GSH ([GSH] 1). The half-lives of the single-disulfide analogue can be extracted easily from the curves in Figure 3b by using a pseudo-first-order rate equation (see Figure S7 and Table S1 in the Supporting Information).[9a] For the single-disulfide analogue, a linear correlation between t = and [GSH] 1 should be observed (pseudo-first-order rate equation),[7a, 9a] and is verified by the data shown in Figure 4a (filled squares). However, because the cleavage kinetics of the C-NDI-C dimer cannot be simply fitted by an existing reaction 1

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model, their half-lives were extracted by either nonlinear- or linear-fitting, depending on the shape of response curve (see Figures S8 and S9, and Table S1). In view of the similarity in responsiveness for both types of dimer at higher concentrations of GSH (e.g., 10 mm), we also predicted the half-lives of the CNDI-C dimer at different GSH concentrations through the 10 mm data point according to the pseudo-first-order rate equation (Figure 4a, open circles, and Table S2). As shown in Figure 4a, there is a clear deviation from the predicted linear trend by the experimental data (filled circles), that is, the twindisulfide dimer displays an extraordinary nonlinear responsiveness towards different concentrations of GSH, which should be impossible to be achieved by the single-disulfide bonds. In addition, the nonlinear responsiveness is made up of an ultrahigh stability in weakly reducing media and a sensitivity not significantly compromised at high concentrations of reducing agents, a property that would be of particular interest for drug delivery applications. It was speculated that the reverse thiol–disulfide exchange reaction in the C-NDI-C dimer should be responsible for the observed difference in responsiveness (Figure 4b). To verify this, we monitored the cleavage kinetics of the C-NDI-C dimer in phosphate buffers containing GSH and different concentrations of GSSG. If the reverse thiol–disulfide exchange took place while the twin-disulfide was partially reduced, then the presence of GSSG in solution would compete with the intramolecular disulfide exchange reaction by reacting with the newly formed thiolate in the intermediate (Figure 4b), a process that should accelerate the rate of cleavage of the C-NDI-C dimer. Interestingly, this is what we observed experimentally, that is, the addition of GSSG can markedly increase the rate of cleavage (see Figure S10 in the Supporting Information), particularly when the concentration of GSH is relatively low. However, when the concentration of GSH is high (e.g., 10 mm), the effect of GSSG on the cleavage kinetics is no longer so clear (see Figure S10), which implies that both of the twin-disulfide bonds could be cleaved rapidly through thiol–disulfide exchange with GSH, which leads to permanent cleavage of the twin-disulfide dimer/linker (i.e., the reverse thiol–disulfide exchange reaction is no longer the rate-determining step for the cleavage of the C-NDI-C dimer). In contrast, the addition of GSSG has no remarkable effect on the cleavage kinetics of the single-disulfide analogue (see Figure S11), which demonstrates the absence of a reverse thiol–disulfide exchange process in a single-disulfide-based system. Therefore our results have clearly demonstrated the extraordinary responsiveness of the twin-disulfide linker/dimer towards different redox environments, a feature that is in remarkable contrast to single-disulfide bonds. In addition, this nonlinear responsiveness has been demonstrated to stem from the reverse thiol–disulfide exchange reaction that occurs while the twin-disulfide dimer is partially reduced. Responsiveness in mimetic redox environments In light of its extraordinary responsiveness, we further evaluated how the twin-disulfide linker (C-NDI-C dimer) makes re-

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Full Paper sponse to redox buffers mimicking different redox environments in the body. First, the buffers with redox potentials of 260, 220, 170, and 140 mV at pH 7.4 were explored, which mimic the steady-state redox potential of the cytoplasm of proliferating, nondividing, and apoptotic cells, and the extracellular redox potential, respectively.[9a, 14] Figure 5 shows the

drug conjugates for cancer therapy.[6d] In addition, in light of the large differences in the kinetics of cleavage of the twin-disulfide linker observed in mimetic cytoplasm of different redox potentials, the twin-disulfide linker has the potential to be exploited for the development of new redox-sensitive synthetic systems that might be responsive to differing cellular states.

Synthetic modification of C-NDI-C We also explored the feasibility of modifying the C-NDI-C with functional groups so that novel redox-responsive synthetic systems can be easily developed. The C-NDI-C molecule bears two carboxy groups, which enable functional molecules with amino groups to be linked directly through amide bond formation. However, amide-forming reactions are subject to poor site-specificity, which might give rise to heterogeneous populations of conjugates, especially when multiple amino or carboxy groups are present in the functional molecules of interest. Thus, it will be useful to introduce artificial and bio-orthogonal groups into the C-NDI-C molecule for the convenience of postfunctional modification. To this end, azide-derivatized C-NDI-C (Azide-C-NDI-C) was synthesized through the conjugation of CNDI-C with 2-azidoethylamine (Figures 6a and S16 in the Sup-

Figure 5. Correlation between t1=2 and the redox potential of GSH/GSSG redox buffers (pH 7.4): & represents the twin-disulfide dimer and * the single-disulfide analogue The total concentration of GSH and GSSG was 10 mm.

half-lives of the twin-disulfide in the relevant buffers along with those of the single-disulfide analogue presented for comparison. It was found that both the twin- and single-disulfide linkers are cleaved rapidly in buffers of 260 and 220 mV (t = < 15 min). However, in the buffers of 170 and 140 mV, the cleavage of the twin-disulfide linker was significantly slower than that of the single-disulfide linker (t = : 9- and 18fold slower, respectively), which indicates the effect of reverse thiol–disulfide exchange on the stability of the twin-disulfide linker in weakly reducing environments. In the next step, a buffer of pH 4.9 and 10 mm concentration of GSH was explored, which mimics the acidic conditions in late endosomal or lysosomal compartments. At this pH, thiol–disulfide exchange was significantly slower than in neutral conditions (pH 7.4) for both disulfide linkers because of the decreased concentration of deprotonated GSH (i.e., the active form of GSH participating in thiol–disulfide exchange; see Figure S14 in the Supporting Information). Finally, we explored the cleavage of the twin-disulfide linker in buffer (pH 7.4) containing 100 mm cysteine, which is the major low-molecular-weight thiol in blood.[15] The twin-disulfide linker was vastly more stable than the single-disulfide bond in these conditions (see Figure S15), as in other weakly reducing environments. Overall, the promising results presented in this section indicate that the twin-disulfide linker shows not only a rapid responsiveness to highly reducing conditions of cytoplasm, but an ultrahigh stability in conditions that mimic extracellular spaces, blood, and endocytic organelles. This might be particularly appealing for drug delivery applications for which relatively large differences in the stability of disulfide linkers in different environments in the body is desired, for example, antibody 1

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Figure 6. a) Synthesis of azide-derivatized C-NDI-C: 1) conjugation of CTrtNDI-CTrt with 2-azidoethylamine; 2) deprotection of thiol groups. b) Protocol for the synthesis of an asymmetric twin-disulfide dimer/linker.

porting Information), which has the potential to conjugate specifically with alkyne-derivatized functional molecules through click reactions. In general, asymmetric disulfides can be prepared by the thiol–disulfide exchange reaction of a free thiol molecule with another thiol that was activated by 2,2’-dithiopyridine. Therefore we examined whether this protocol could be applied to the preparation of an asymmetric twin-disulfide dimer/linker. First, our results show that both thiol groups in C-NDI-C can

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Full Paper be activated by incubation with a 25-fold excess of 2,2’-dithiopyridine (Figures 6b and S17 in the Supporting Information). It is worth emphasizing that an excess of 2,2’-dithiopyridine is a prerequisite for the formation of thiol-activated C-NDI-C, otherwise the twin-disulfide dimer predominantly formed during the activation step. In the next step, we demonstrated that free C-NDI-C can effectively react with the thiol-activated CNDI-C to form the desired twin disulfides in high yield (> 90 %, Figures 6b and S17). The preparation of asymmetric twin-disulfide bonds/linkers is thus as straightforward as the synthesis of traditional single-disulfides. This study is particularly important when considering that in many applications, including drug delivery, bioimaging, and sensing, two different functional entities have to be linked together through a disulfide linker. General discussion Twin-disulfides exploit the cooperativity between two independent disulfide bonds to modulate redox-responsiveness. The cooperative behavior is analogous to chelate cooperativity, which arises from the presence of multiple intermolecular binding interactions (i.e., multivalency).[16] In the presence of (chelate) cooperativity, the partially bound intermediate is usually present in minor amounts or even completely absent, a feature that is characteristic of cooperative systems. This is exactly what we observed in this work with the C-NDI-C system, in which no partially reduced intermediate was detected, which indicates the ease of the reverse/intramolecular thiol–disulfide exchange process. In cooperative systems, the effective molarity is usually used to evaluate the ease of intramolecular processes,[16] which, in the case of our twin-disulfide system, might denote the threshold concentration of thiol reducing agent below which the intermolecular thiol–disulfide exchange responsible for the cleavage of twin-disulfide linker no longer competes with the reverse/intramolecular process. Assuming the absence of the reverse thiol–disulfide exchange process and a sufficient excess of GSH, the cleavage of the twin-disulfide should be as rapid as that of its single-disulfide analogue according to the pseudo-first-order reaction model. However, our results show that, although the twin-disulfide linker can be cleaved rapidly under high concentrations of GSH, the kinetics of cleavage is still slower than that of the single-disulfide analogue. This implies that the effective molarity for the C-NDI-C dimerization should exceed the maximum concentration of GSH (10 mm) explored in this study. It is expected that tunable redox-responsiveness might be further achieved by tuning the backbone structure (e.g., length and flexibility) of twin-disulfides and the intermolecular noncovalent interactions between two backbone building blocks, which determines the effective molarity. In addition, the factors that influence intermolecular thiol–disulfide exchange, including steric hindrance and the microenvironment, may also be rationally tuned to further modulate the responsiveness of twin-disulfide-based linkers.[7a, 9a] As the nonlinear responsiveness of twin disulfides strongly relies on the reverse thiol–disulfide exchange reaction, this process might be susceptible to nonspecific interactions beChem. Eur. J. 2014, 20, 17507 – 17514

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tween the linker and the conjugated functional unities. Thus, for synthetic systems in which twin-disulfide linkers are incorporated, responsiveness of the disulfide linkers should be verified. In addition, in contrast to the single-disulfide bond, the twin-disulfide linker is extremely bulky. Thus, a new route to explore is the miniaturization of the twin-disulfide linker (e.g., exploring and exploiting a single benzene ring as a rigid scaffold for twin-disulfide construction). In doing this, functional groups that can be used for the conjugation of functional unities have to be wisely placed, and the twin-thiol groups need to be mutually independent, unless rationally designed to avoid this. A variety of NDI derivatives containing two thiol groups have been exploited for the construction of dynamic combinatorial libraries.[12b–d, 17] In these studies, intra- and intermolecular thiol–disulfide exchange reactions play an important role in determining the distribution of synthetic libraries. However, the interplay between the nature of the dithiol building blocks and the redox environments in the media has thus far not been discussed. The present contribution might be a first step towards understanding the contribution of cooperativity between disulfide bonds to the stability of individual dynamic complexes (i.e., library members) in redox media.

Conclusion We have developed and explored an extraordinary twin-disulfide linker that shows responsiveness to different redox environments in a manner that is in remarkable contrast to the traditional single-disulfide bond. The twin-disulfide linker displays an ultrahigh stability at low concentrations of reducing agent or in weakly reducing environments without significant compromise of the sensitivity of the response to highly reducing environments such as cytoplasm. This feature is of particular interest for applications such as drug delivery and redox-sensing, because in these applications relatively large differences in the stability of disulfide linkers in different environments in the body are desired. To the best of our knowledge, this is the first example of a redox-responsive linker to show a nonlinear responsiveness to the concentration of the reducing agent. We have also demonstrated that this extraordinary nonlinear responsiveness arises from the cooperativity of the twin-disulfide bonds, on the basis of which a reverse/intramolecular thiol–disulfide exchange reaction can take place while the twin-disulfide linker is partially reduced. In addition, our study has shown the feasibility of modifying C-NDI-C with azide groups, which can be subsequently used for postmodification by click reactions. Finally, we have demonstrated that asymmetric twindisulfide linkers can be easily prepared by activation of the twin-thiol groups with 2,2’-dithiopyridine followed by an intermolecular thiol–disulfide exchange reaction, a procedure frequently used for the preparation of asymmetric single-disulfide bonds. In light of the interesting findings reported herein, the twin-disulfide linker has great potential for being incorporated into synthetic systems.[6c, d, j] In addition, a comparison of the responsiveness of twin-disulfide linkers with that of single-disulfide bonds in biological environments (in vitro or in vivo) is of

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Experimental Section Materials and instruments: All chemicals were purchased commercially and used without further purification. Millipore ultrapure water was used throughout the experiments. Analytical HPLC was performed by using a Shimadzu instrument equipped with a Prominence LC-20AD solvent delivery unit, a Prominence DGU-20A5 degassing unit, and a Prominence SPD-20A UV/Vis detector. Chromatograms were recorded by using an Inertsil ODS-SP C18 (5 mm, 4.6  250 mm) column by using gradients of water + 0.1 % trifluoroacetic acid (TFA) and acetonitrile (ACN) + 0.1 % TFA at a flow rate of 1 mL min 1. Preparative HPLC was performed by using a Shimadzu system equipped with two LC-6AD solvent delivery units, a Prominence communications bus module, a fraction collector, and a Prominence SPD-20A UV/Vis detector. An Hitachi U-3900H UV/Vis spectrometer was used to record the UV absorption spectra. A Bruker Esquire 3000plus ion-trap ESI mass spectrometer was used to identify the synthesized compound and the products formed during the thiol–disulfide exchange reactions. Synthesis of C-NDI-C: 1,4,5,8-Naphthalenediimide (NDI, 100 mg, 0.373 mmol) and S-trityl-l-cysteine (2 equiv, 271 mg, 0.746 mmol) were suspended in DMF (5 mL) in a pressure-tight 10 mL microwave tube. Triethylamine (52 mL) was added to this suspension and the suspension was sonicated until the mixture became homogenous. The mixture was then heated under microwave irradiation for 5 min at 100 8C. After that, the solution was added slowly to a vigorously stirred aqueous solution of 1 m HCl. The precipitate was S-trityl-protected C-NDI-C, which was then filtered and washed with water and dried in vacuo to yield the product as a yellow solid (350 mg, 98 %). 1H NMR (400 MHz, [D6]DMSO): d = 13.16 (br, 2 H; COOH), 8.74 (s, 4 H; NDI), 7.30–7.10 (m, 30 H; Trt), 5.55 (dd, J = 10.3, 4.6 Hz, 2 H; a-Cys), 3.11(dd, J = 12.9, 4.5 Hz, 2 H; b-Cys), 2.96 ppm (dd, J = 12.8, 10.4 Hz, 2 H; b-Cys); MS (ESI): calcd for C58H42N2O8S2 [M + Na] + 981.2; found: 981.4. Then, TFA (2 mL), dichloromethane (DCM, 2 mL), and triethylsilane (0.2 mL) were added to a 25 mL round-bottomed flask containing S-trityl-protected C-NDI-C (100 mg, 0.104 mmol), and the reaction mixture was stirred vigorously for 1 h. All the volatiles were then removed under reduced pressure and the residue dissolved in ACN and purified by C18 reversed-phase preparative chromatography to yield the product (C-NDI-C) as a yellow solid after lyophilization (22.4 mg, 45 %). 1H NMR (400 MHz, [D6]DMSO): d = 13.21 (br, 2 H; COOH), 8.77 (s, 4 H; NDI), 5.72 (dd, J = 9.3, 5.4 Hz, 2 H; a-Cys), 3.21 (dd, J = 9.2, 4.9 Hz, 2 H; b-Cys), 2.71 ppm (dd, J = 8.9, 4.0 Hz, 2 H; bCys); MS (ESI): calcd for C20H14N2O8S2 [M + H] + 475.02; found: 475.03. Synthesis of a single-cysteine analogue: 2-Iodoacetamide (3.5 mg, 0.019 mmol) and C-NDI-C (10 mg, 0.021 mmol) were dissolved in phosphate buffer (2 mL, 100 mm, pH 7.4), and then the mixture was stirred at room temperature for 1 h in the dark. The solution was purified by preparative HPLC to yield the product as a light-yellow solid after lyophilization (4.9 mg, 49 %). MS (ESI): calcd for C22H17N3O9S2 [M + H] + 532.04; found: 532.05. Synthesis of azide-derivatized C-NDI-C: C-NDI-C (356 mg, 0.372 mmol) was dissolved in anhydrous DMF (5 mL), and then 2azidoethylamine (63.9 mL, 0.743 mmol) and N,N-diisopropylethylamine (DIEA, 246 mL, 1.544 mmol) were added. After stirring at room temperature for 15 min, N-[(dimethylamino)-1H-1,2,3-triazolo[4,5-b]pyridin-1-ylmethylene]-N-methylmethanaminium hexafluoroChem. Eur. J. 2014, 20, 17507 – 17514

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phosphate N-oxide (HATU, 311 mg, 0.818 mmol) was added, and the mixture was stirred at room temperature for 2 h. The product was precipitated and washed three times with water. The crude product (Azide-CTrt-NDI-CTrt) was then purified by preparative HPLC to afford the product as a yellow solid (160 mg, 39.3 %). 1H NMR (400 MHz, [D6]DMSO): d = 8.73 (s, 4 H; NDI), 8.24 (t, J = 5.4 Hz, 2 H; NH), 7.21–7.15 (m, 30 H; Trt), 5.46 (dd, J = 10.4, 4.5 Hz, 2 H; a-Cys), 3.27 (dd, J = 11.9, 6.0 Hz, 2 H; b-Cys), 3.20 (dd, J = 12.5, 4.8 Hz, 4 H; N3-CH2), 3.14–3.11 (m, 4 H; CH2-NH), 2.84 ppm (dd, J = 12.6, 10.6 Hz, 2 H; b-Cys); MS (ESI): calcd for C62H50N10O6S2 [M + Na] + 1117.33; found: 1117.32. The obtained Azide-CTrt-NDI-CTrt was then deprotected as described above to yield Azide-C-NDI-C. MS (ESI): calcd for C24H22N10O6S2 [M + H] + 611.12; found: 611.12. Formation of the twin/single-disulfide dimer: The twin-disulfide dimer was prepared by oxidation in 20 % aqueous DMSO and purified by preparative HPLC. In a typical experiment, C-NDI-C (7 mg, 0.015 mmol) was first dissolved in phosphate buffer (14 mL, 100 mm, pH 7.4) and then DMSO was added. The solution was incubated at room temperature for 24 h. HPLC (flow rate 1 mL min 1, isocratic with 5 % v/v ACN (+ 0.1 % TFA) for 5 min followed by a linear gradient of ACN (+ 0.1 % TFA) over 40 min (5–60 % v/v)) was performed to isolate the C-NDI-C dimer. The single-disulfide dimer was prepared by oxidation in phosphate buffer (100 mm, pH 7.4) at room temperature for 30 h and then isolated by HPLC. The isolated dimers were identified by mass spectrometry. Quantification of the twin/single-disulfide dimers: Stock solutions of the twin/single-disulfide dimers were prepared by dissolving the lyophilized solid in phosphate buffer (pH 7.4). The concentrations of the twin/single-disulfide dimers were determined by UV/Vis spectroscopy. To do this, the dimer was first completely reduced with dithiothreitol (DTT) to form the C-NDI-C monomer and then diluted with phosphate buffer. Absorption spectra in the range 200–500 nm were recorded. The concentration of the solution was calculated by using Beer–Lambert’s law with the molar extinction coefficient of C-NDI-C monomer (2.216  104 cm 1 m 1 at 363 nm, determined from the standard absorption curve). Determination of thiol–disulfide exchange kinetics: All solutions used in the kinetics studies were deoxygenated by bubbling with oxygen-scrubbed nitrogen through a round-bottomed flask sealed with a rubber stopper. Furthermore, to exclude oxygen, experiments were carried out in an anaerobic incubator. In a typical experiment, phosphate buffer (0.2 mL, 100 mm, pH 7.4) was added to a 1.5 mL Eppendorf tube and then the C-NDI-C dimer (or single-disulfide analogue) stock solution (50 mL, 50 mM, dissolved in 100 mm phosphate buffer, pH 7.4) and GSH (0.25 mL, 20 mm or other concentrations) in 100 mm phosphate buffer were added. At predefined times, aliquots (40 mL) were extracted into an empty tube and the reaction was quenched by using liquid nitrogen. The samples were then analyzed by analytical HPLC (30 mL injection volume, 1 mL min 1 flow rate, isocratic with 1 % v/v ACN (+ 0.1 % TFA) for 5 min followed by a linear gradient of ACN (+ 0.1 % TFA) over 40 min (1–70 % v/v)). The HPLC peaks were monitored at 363 nm. The peak area of the dimer was used to calculate the residual percentage of dimer in the reaction mixture. To monitor the thiol–disulfide exchange kinetics of the dimer at pH 7.4 in GSH/ GSSG redox buffers with four different redox potentials ( 260, 220, 170, and 140 mV), a calculated concentration ratio of GSH/GSSG in phosphate buffer solution (100 mm, pH 7.4) was used. A mixture of GSH (0.247 mL, 20 mm) and GSSG (67 mL, 1 mm), GSH (0.2 mL, 20 mm) and GSSG (99 mL, 10 mm), GSH (0.113 mL, 10 mm) and GSSG (0.194 mL, 20 mm), or GSH (0.077 mL, 5 mm) and GSSG (0.231 mL, 20 mm) in phosphate buffer (100 mm, pH 7.4) was added to the solution of the C-NDI-C dimer to gener-

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Full Paper ate buffers (total volume: 0.5 mL) with redox potentials of 260, 220, 170, and 140 mV, respectively. To monitor the thiol–disulfide exchange kinetics between the dimer and GSH at pH 4.9, acetate buffer was used instead of phosphate buffer.

Acknowledgements We would like to acknowledge the financial support of the National Natural Science Foundation of China (Grant 21305114 and 21375110), the National Basic Research 973 Program of China (Grant 2014CB932004), and the Fundamental Research Funds for the Central Universities (Grant ZK1047).

[7]

[8]

Keywords: conjugation · cooperative effects · disulfide bonds · redox chemistry · thiol–disulfide exchange [1] a) J. Riemer, N. Bulleid, J. M. Herrmann, Science 2009, 324, 1284 – 1287; b) D. Fass, Annu. Rev. Biophys. Bioeng. 2012, 41, 63 – 79; c) A. W. Zhou, R. W. Carrell, M. P. Murphy, Z. Q. Wei, Y. H. Yan, P. L. D. Stanley, P. E. Stein, F. B. Pipkin, R. J. Read, Nature 2010, 468, 108 – 111; d) B. Schmidt, L. Ho, P. J. Hogg, Biochemistry 2006, 45, 7429 – 7433; e) B. S. Mamathambika, J. C. Bardwell, Annu. Rev. Cell Dev. Biol. 2008, 24, 211 – 235. [2] J. Alegre-Cebollada, P. Kosuri, J. A. Rivas-Pardo, J. M. Fernandez, Nat. Chem. 2011, 3, 882 – 887. [3] a) P. Kosuri, J. Alegre-Cebollada, J. Feng, A. Kaplan, A. Ingles-Prieto, C. L. Badilla, B. R. Stockwell, J. M. Sanchez-Ruiz, A. Holmgren, J. M. Fernandez, Cell 2012, 151, 794 – 806; b) W. J. Wedemeyer, E. Welker, M. Narayan, H. A. Scheraga, Biochemistry 2000, 39, 4207 – 4216; c) P. J. Hogg, Nat. Rev. Cancer 2013, 13, 425 – 431; d) P. J. Hogg, Trends Biochem. Sci. 2003, 28, 210 – 214. [4] a) K. S. Jensen, R. E. Hansen, J. R. Winther, Antioxid. Redox Signaling 2009, 11, 1047 – 1058; b) K. M. Cook, P. J. Hogg, Antioxid. Redox Signaling 2013, 18, 1987 – 2015; c) Z. Y. Cheng, J. F. Zhang, D. P. Ballou, C. H. Williams, Chem. Rev. 2011, 111, 5768 – 5783; d) U. Grauschopf, J. R. Winther, P. Korber, T. Zander, P. Dallinger, J. C. A. Bardwell, Cell 1995, 83, 947 – 955. [5] a) J. Yang, H. Chen, I. R. Vlahov, J. X. Cheng, P. S. Low, Proc. Natl. Acad. Sci. USA 2006, 103, 13872 – 13877; b) G. D. Lewis Phillips, G. M. Li, D. L. Dugger, L. M. Crocker, K. L. Parsons, E. Mai, W. A. Blattler, J. M. Lambert, R. V. J. Chari, R. J. Lutz, W. L. T. Wong, F. S. Jacobson, H. Koeppen, R. H. Schwall, S. R. Kenkare-Mitra, S. D. Spencer, M. X. Sliwkowski, Cancer Res. 2008, 68, 9280 – 9290. [6] a) G. Saito, J. A. Swanson, K. D. Lee, Adv. Drug Delivery Rev. 2003, 55, 199 – 215; b) S. Bauhuber, C. Hozsa, M. Breunig, A. Gopferich, Adv. Mater. 2009, 21, 3286 – 3306; c) F. H. Meng, W. E. Hennink, Z. Zhong, Biomaterials 2009, 30, 2180 – 2198; d) A. M. Wu, P. D. Senter, Nat. Biotechnol. 2005, 23, 1137 – 1146; e) G. Gasparini, E. K. Bang, G. Molinard, D. V. Tulu-

Chem. Eur. J. 2014, 20, 17507 – 17514

www.chemeurj.org

[9]

[10] [11] [12]

[13] [14]

[15] [16]

[17]

mello, S. Ward, S. O. Kelley, A. Roux, N. Sakai, S. Matile, J. Am. Chem. Soc. 2014, 136, 6069 – 6074; f) X. M. Wu, X. R. Sun, Z. Q. Guo, J. B. Tang, Y. Q. Shen, T. D. James, H. Tian, W. H. Zhu, J. Am. Chem. Soc. 2014, 136, 3579 – 3588; g) L. Brlisauer, N. Kathriner, M. Prenrecaj, M. A. Gauthier, J. C. Leroux, Angew. Chem. Int. Ed. 2012, 51, 12454 – 12458; Angew. Chem. 2012, 124, 12622 – 12626; h) S. Durocher, A. Rezaee, C. Hamm, C. Rangan, S. Mittler, B. Mutus, J. Am. Chem. Soc. 2009, 131, 2475 – 2477; i) H. Mok, S. H. Lee, J. W. Park, T. G. Park, Nat. Mater. 2010, 9, 272 – 278; j) M. H. Lee, Z. Yang, C. W. Lim, Y. H. Lee, S. Dongbang, C. Kang, J. S. Kim, Chem. Rev. 2013, 113, 5071 – 5109. a) C. L. Wu, S. Wang, L. Brulisauer, J. C. Leroux, M. A. Gauthier, Biomacromolecules 2013, 14, 2383 – 2388; b) B. A. Kellogg, L. Garrett, Y. Kovtun, K. C. Lai, B. Leece, M. Miller, G. Payne, R. Steeves, K. R. Whiteman, W. Widdison, H. S. Xie, R. Singh, R. V. J. Chari, J. M. Lambert, R. J. Lutz, Bioconjugate Chem. 2011, 22, 717 – 727. P. E. Thorpe, P. M. Wallace, P. P. Knowles, M. G. Relf, A. N. F. Brown, G. J. Watson, R. E. Knyba, E. J. Wawrzynczak, D. C. Blakey, Cancer Res. 1987, 47, 5924 – 5931. a) C. L. Wu, C. Belenda, J. C. Leroux, M. A. Gauthier, Chem. Eur. J. 2011, 17, 10064 – 10070; b) S. Arpicco, F. Dosio, P. Brusa, P. Crosasso, L. Cattel, Bioconjugate Chem. 1997, 8, 327 – 337. C. L. Wu, J. C. Leroux, M. A. Gauthier, Nat. Chem. 2012, 4, 1044 – 1049. L. Moroder, D. Besse, H. J. Musiol, S. Rudolph-Bçhner, F. Siedler, Biopolymers 1996, 40, 207 – 234. a) K. Tambara, N. Ponnuswamy, G. Hennrich, G. D. Pantos, J. Org. Chem. 2011, 76, 3338 – 3347; b) H. Y. Au-Yeung, G. D. Pantos, J. K. M. Sanders, J. Org. Chem. 2011, 76, 1257 – 1268; c) H. Y. Au-Yeung, G. D. Pantos, J. K. M. Sanders, Proc. Natl. Acad. Sci. USA 2009, 106, 10466 – 10470; d) H. Y. AuYeung, P. Pengo, G. D. Pantos, S. Otto, J. K. M. Sanders, Chem. Commun. 2009, 419 – 421. J. P. Tam, C. R. Wu, W. Liu, J. W. Zhang, J. Am. Chem. Soc. 1991, 113, 6657 – 6662. a) Y. M. Go, D. P. Jones, Biochim. Biophys. Acta Gen. Subj. 2008, 1780, 1273 – 1290; b) F. Q. Schafer, G. R. Buettner, Free Radical Biol. Med. 2001, 30, 1191 – 1212. D. P. Jones, V. C. Mody, J. L. Carlson, M. J. Lynn, P. Sternberg, Free Radical Biol. Med. 2002, 33, 1290 – 1300. a) G. Ercolani, L. Schiaffino, Angew. Chem. Int. Ed. 2011, 50, 1762 – 1768; Angew. Chem. 2011, 123, 1800 – 1807; b) C. A. Hunter, H. L. Anderson, Angew. Chem. Int. Ed. 2009, 48, 7488 – 7499; Angew. Chem. 2009, 121, 7624 – 7636. a) F. B. L. Cougnon, H. Y. Au-Yeung, G. D. Pantos, J. K. M. Sanders, J. Am. Chem. Soc. 2011, 133, 3198 – 3207; b) F. B. L. Cougnon, N. Ponnuswamy, N. A. Jenkins, G. D. Pantos, J. K. M. Sanders, J. Am. Chem. Soc. 2012, 134, 19129 – 19135; c) J. W. Li, P. Nowak, S. Otto, J. Am. Chem. Soc. 2013, 135, 9222 – 9239; d) N. Ponnuswamy, F. B. L. Cougnon, J. M. Clough, G. D. Pantos, J. K. M. Sanders, Science 2012, 338, 783 – 785.

Received: August 19, 2014 Published online on October 29, 2014

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