Probing the Structural Flexibility of the Human Copper

Article pubs.acs.org/JPCB

Probing the Structural Flexibility of the Human Copper Metallochaperone Atox1 Dimer and Its Interaction with the CTR1 C‑Terminal Domain Ariel R. Levy, Valeria Yarmiayev, Yoni Moskovitz, and Sharon Ruthstein* The Department of Chemistry, Faculty of Exact Science, Bar Ilan University, Ramat-Gan, Israel, 5290002 S Supporting Information *

ABSTRACT: Both the essentiality and the toxicity of copper in human, yeast, and bacteria cells require precise mechanisms for acquisition, intimately linked to controlled distribution, which have yet to be fully understood. This work explores one aspect in the copper cycle, by probing the interaction between the human copper chaperone Atox1 and the c-terminal domain of the copper transporter, CTR1, using electron paramagnetic resonance (EPR) spectroscopy and circular dichroism (CD). The data collected here shows that the Atox1 keeps its dimer nature also in the presence of the CTR1 c-terminal domain; however, two geometrical states are assumed by the Atox1. One is similar to the geometrical state reported by the crystal structure, while the latter has not yet been constructed. In the presence of the CTR1 c-terminal domain, both states are assumed; however, the structure of Atox1 is more restricted in the presence of the CTR1 c-terminal domain. This study also shows that the last three amino acids of the CTR1 cterminal domain, HCH, are important for maintaining the crystal structure of the Atox1, allowing less structural flexibility and improved thermal stability of Atox1.

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an overall βαββαβ fold structure. It coordinates one copper atom by the cysteine residues in a conserved MxCxxC motif (where x represents other amino acids).15,21,22 The ATP7A/B copper pumps contain between two and six MxCxxC metalbinding domains (MBDs) at their amino terminal, each with an overall tertiary structure similar to that of Atox1, where each MBD binds a single copper atom in vitro. The Atox1 and ATP7A/B proteins directly interact in a copper-stimulated manner. According to structural predictions, the metal is transferred via a series of ligand-exchange reactions involving two- and three-coordinate intermediates between cysteine ligands in the CxxC motifs on Atox1 and the recipient copper-transporting ATPases.8,12,15,16,21 While the mechanism by which copper is transferred from the Atox1 to the ATP7A/B is better known, other Cu pathways in the cell, such as the Cu import and export mechanisms used by the CTR1 to the Cu chaperones, are less known. Unger et al.23,24 recently reported the 6 Å resolution electron crystallography structure of the human CTR1, indicating that the CTR1 is a trimer oligomerization, where each monomer is 23 kDa, 190 amino acids and contains: (1) a 60 amino acid extracellular N-terminal domain, (2) an intracellular loop of 46 amino acids, connecting the first and second putative membrane-spanning helices, (3) three transmembrane helices, and (4) a short intracellular c-terminal domain with 15 amino acids. Despite this significant progress in resolving the CTR1 3D structure, the Cu(I) translocation mechanism through the

INTRODUCTION Copper is an essential trace metal for living organisms. Many proteins that participate in cellular respiration, antioxidant defense, neurotransmitter biosynthesis, connective-tissue biosynthesis, and pigment formation use copper as their prosthetic, active group. However, because free copper is toxic, cells have developed highly regulated copper transmitting pathways. Today, it is known that Cu(II) is consumed and accumulated in our body through diet, reduced to Cu(I), and delivered to the cell by the copper transporter (CTR1).1,2 When Cu(I) is transferred into the cell, specific Cu chaperones are responsible for delivering it to specific cellular pathways.3−6 These chaperones include Atox1, which delivers copper to the copper transporting ATPases in the Golgi, CCS, required for copper incorporation into cytoplasmic Cu/Zn superoxide dismutase, and Cox17, Sco1, and Sco2, which deliver copper to mitochondrial cytochrome c oxidase.4−9 The mechanism of copper translocation across the cell membrane and within the cell remains largely unknown. This is mostly due to the difficulties associated with determining the structures of integral proteins, and the lack of an in vitro system designed to systematically study copper transport. As a result, the majority of what is known about copper transport−cellular acquisition is based on genetic complementation strategies and measurements of copper transport in vivo.1,10,11 To date, most of the focus has been on the Atox1 copper chaperone and the copper transport ATP7A/B.12−16 Mutations in ATP7A and ATP7B have been found to be the leading cause for Menkes and Wilson’s diseases.8,16−18 The crystal structures of Atox119 and its analogue yeast Atx1 were reported.7,20 Atox1 was shown to be a small soluble protein (68 amino acids) with © 2014 American Chemical Society

Received: December 24, 2013 Revised: April 20, 2014 Published: May 16, 2014 5832

dx.doi.org/10.1021/jp412589b | J. Phys. Chem. B 2014, 118, 5832−5842

The Journal of Physical Chemistry B

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CTR1 protein and to the various Cu chaperones is still unknown. It was previously suggested that metallochaperones can dock to CTR1, since significant structural similarity exists between two Cu chaperones in the cell, CCS and Atox1.25,26 In addition, the copper exchange assay between the c-terminal segment of the yeast CTR1 and the yeast Atx1, analogue of the human CTR1 and Atox1, showed a controlled Cu transfer between the c-terminal and the chaperone, suggesting a specific interaction between them.27 Resolving the copper transfer mechanism in a cell is essential for understanding the copper cycle in the human cell, and for exposing interferences with copper homeostasis. To comprehend such processes, it is necessary to be sensitive to the structural changes that occur in the metallochaperone upon metal binding, and upon binding to the intracellular segment of CTR1. The conventional methods, NMR and X-ray crystallography, are limited by protein size, and difficulties in protein and protein complex crystallization. Thus, probing a biological process which involves weak interaction between two transient proteins is a challenge with NMR and X-ray crystallography. Electron paramagnetic resonance (EPR) spectroscopy has emerged as an excellent tool for resolving such systems, since it does not require crystallization and does not depend on protein size. EPR measurement can be performed in buffer solution, and even weak interaction between proteins can be detected.28 EPR’s strength lies in its sensitivity to both atomic level changes and nanoscale fluctuations. EPR can characterize properties such as redox state and ligand geometry for different functional states of the protein.29−33 In addition, EPR can measure distances between paramagnetic probes within the protein, and between proteins, up to 80 Å.34−40 In some cases, the measurement of one distance (or a few) is sufficient to establish the plausibility of a mechanism or to corroborate a proposed structure. The most common experiment for obtaining nanosacle structure information is the pulsed electron double resonance (PELDOR) also commonly referred to as the double electron electron resonance experiment (DEER). Pulsed EPR experiments can measure nanometer distances between paramagnetic probes, and continuous wave (CW) EPR can derive the dynamics of protein chains. The combination of CW and pulsed EPR with site-directed spin labeling (SDSL) has become widely used in biophysical research,41−45 where an electron spin introduced into diamagnetic proteins provides information on their local environment and on the mobility of the protein domain. When multiple spin labels are attached, distance distributions between them can be derived.43−50 In this study, CW and pulsed EPR will be combined with SDSL and CD to exploit one copper cycle in the human cell related to the mechanism for transmitting copper from the intracellular CTR1 domain to the Atox1 chaperone. Residue Cys41 in Atox1 will be labeled with a spin label, and its conformational state while interacting with the CTR1 cterminal domain will be probed (see Figure 1). The effect of a mutated CTR1 c-terminal domain on this interaction will also be explored, in order to shed light on the role of the Cys189 residue of the CTR1. Finally, kinetic CD measurements will be performed to obtain better insight into the interaction between Atox1 and the CTR1 c-terminal domain. This study is important to the understanding of copper homeostasis in the human body.

Figure 1. Schematic view presenting the interaction between the copper chaperone, Atox1, and the copper transporter, CTR1, cterminal domain which is explored in this study. The red dots on the Atox1 mark the spin labeling sites. The inset shows the site-directed spin labeling (SDSL) method using a methanesulfonthioate (MTSSL) spin label.

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EXPERIMENTAL SECTION Peptide Synthesis, Purification, and Labeling. Table 1 lists the sequences of the peptides used in this study. All Table 1. CTR1 c-Terminal Domain Segments Used in This Work peptide

sequence

pep1 pep2 pep3 pep4 pep5

SWKKAVVVDITEHCH SWKKAVVVDITEHC-(MTSSL)H SWKKAVVVDITE SWKKAVVVDITEHGH SWKKAVVVDITEHMH

peptides were synthesized on a rink amide resin (Applied Biosystems). Couplings of standard Fmoc (9-fluorenylmethoxy-carbonyl)-protected amino acids were achieved with (Obenzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium (HBTU, Dchem) in N,N-dimethylformamide (DMF, Bio lab) in combination with N,N-diisopropylethylamine (DIPEA, Bio lab) for a 1 h cycle. Fmoc deprotection was achieved with piperidine (Bio lab). Side-chain deprotection and peptide cleavage from the resin were achieved by treating the resin bound peptides with 5 mL of 95% (90% trifluoroacetic acid (TFA, Bio lab), 5% ethane dithiol (EDT, Alfa Aesar), 2.5% triisopropylsilane (TIS, Alfa Aesar), and 2.5% thioanisole (Alfa Aesar) for 2.5 h under N2. An additional 65 μL of bromotrimethylsilane (TMSBr, Alfa Aesar) was added during the final 30 min to minimize methionine oxidation. The peptides were washed four times with cold diethyl ether, vortexed, and then centrifuged for 5 min at 3500 rpm. After evaporation of TFA under N2, 10 mM DTT (dithiothreitol, Sigma) was added to the peptide and was dissolved in HPLC water. The peptide was then purified by preparative reversedphase HPLC (vydac, C18, 5 cm). The mass of the peptide was confirmed either by MALDI-TOF MS-Autoflex III-TOF/TOF mass spectrometer (Bruker, Bermen, Germany) equipped with a 337 nm nitrogen laser or by ESI (electron spray ionization) mass spectrometry on a Q-TOF (quadruple time-of-flight) lowresolution micromass spectrometer (Micromass-Waters, Corp.) 5833



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observed that a similar CW-EPR spectrum is obtained at any sequence addition. Moreover, after adding pep1 to the Atox1 solution, no spin label was transferred from the Atox1 to the pep1, as was verified by HPLC and mass spec. CD. Circular dichroism (CD) measurements were carried out using a Chirascan spectrometer (Applied Photophysics, U.K.). Measurements were carried out over a range of 20−90 °C in a 1 mm optical path length cell. The protein concentration was 0.2 mM. Buffer scans were subtracted from the sample scans. The data was recorded from 190 to 260 nm with a step size and a bandwidth of 0.5 nm. The CD signal was averaged for three scans for each sample. The temperature measurements were performed with a rate of 1 °C/min. EPR. CW-EPR (continuous wave EPR) spectra were recorded using an E500 Elexsys Bruker spectrometer operating at 9.0−9.5 GHz. The spectra were recorded at room temperature (295 ± 2 K) at a microwave power of 20.0 mW, modulation amplitude of 1.0 G, time constant of 60 ms, and receiver gain of 60.0 dB. The samples were measured in 0.8 mm capillary quartz tubes (vitrocom). CW-EPR simulations were carried out on MATLAB, using the easyspin toolbox.52 The constant time four-pulse DEER experiment π/2(νobs)τ1-π(νobs)-t′-π(νpump)-(τ1+τ2-t′)-π(νobs)-τ2(νobs)-τ2-echo was carried out at 80 ± 1.0 K on a Q-band Elexsys E580 (equipped with a 2 mm probehead, bandwidth = 220 MHz). A two-step phase cycle was employed on the first pulse. The echo was measured as a function of t′, while τ2 was kept constant to eliminate relaxation effects. The observer pulse was set 60 MHz higher than the pump pulse (see Figure S1, Supporting Information). The observer π/2 and π pulses had a length of 40 ns, and the π pump pulse had a length of 40 ns as well. The dwell time was 20 ns. The observer frequency was 33.82 GHz. The power of the 40 ns π pulse was 20.0 mW. The samples were measured in 1.6 mm capillary quartz tubes (Wilmand). The data was analyzed using the DeerAnalysis 2011 program, using Tikhonov regularization.53 The regularization parameter in the L curve was optimized by examining the fit of the time domain data. The data presented in this manuscript is after 3D homogeneous background subtraction.

Peptide samples were typically mixed with two volumes of premade dihydrobezoic acid (DHB) matrix solution, deposited onto the stainless steel target surfaces, and allowed to dry at room temperature. For SDSL: A 1 mg portion of lyophilized peptide was dissolved in 1.0 mL of phosphate buffer (25 mM KPi) (pH 7.4−7.6). A 0.25 mg portion of S-(2,2,5,5-tetramethyl-2,5dihydro-1H-pyrrol-3-yl)methylmethanesulfonothioate (MTSSL, TRC) dissolved in 15 μL of dimethyl sulfoxide (DMSO, Bio lab) was added to the solution (50-fold molar excess of MTSSL). The spin label and peptide solution were then vortexed overnight at 4 °C. The free spin label was removed by HPLC semi preparative (Vydac, C18, 1 cm). The mass of the spin-labeled peptide was confirmed by mass spectrometer. Atox1 Cloning, Expression, Purification, and Labeling. The human Atox1 construct pYTB12-Atox1 was kindly given to us by the lab of Prof. Svetlana Lutsenko (Johns Hopkins University). This construct encodes for the fusion protein composed of Atox1, with an intein and a chitin-binding domain. It was transformed to the Escherichia coli strain BL21 (DE3). The Atox1 construct was expressed in BL21 cells, grown to an optical density of 0.5−0.8 at 600 nm, and induced with 0.5 mM isopropyl-β-D-thiogalactopyranoside (Calbiochem) for 18 h at 18 °C. The cells were then harvested by centrifugation and suspended in lysis buffer (25 mM Na2HPO4, 150 mM NaCl, 20 μM PMSF, pH 7.5). The cells were sonicated by six cycles of 1 min each with a 1 min cooling tense between each cycle (65% amplitude). After sonication, cells were centrifuged and the soluble fraction of the lysate was run through a chitin bead column (New England Biolabs), allowing the Atox1 fusion to bind to the resin via its chitin-binding domain. The resin was then washed with 30-column volumes of lysis buffer (pH 8.9). To induce the intein-mediated cleavage, the beads were incubated in 50 mM DTT, 25 mM Na2HPO4, pH 8.9, 150 mM NaCl, for 40 h at room temperature. Atox1 was then collected in elution fractions and analyzed by SDS-PAGE (Tricine 19%) and MALDI-TOF mass spectrometer (Bruker, Bermen, Germany) or by ESI (electron spray ionization) mass spectrometry on a Q-TOF (quadruple time-of-flight) lowresolution micromass spectrometer (Micromass-Waters, Corp.). Before labeling, 10 mM DTT was added to the protein solution and mixed for 10 h at 4 °C. DTT was dialyzed out using 2 kDa dialysis cassettes (Pierce). A 0.25 mg portion of S(2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-3-yl)methylmethanesulfonothioate (MTSSL, TRC) dissolved in 15 μL of dimethyl sulfoxide (DMSO, Bio lab) was added to 0.75 mL of 0.5 mM protein solution (100-fold molar excess of MTSSL). The protein solution was then vortexed overnight at 4 °C. The free spin label was removed by several dialysis cycles over 4 days. The mass of the spin-labeled protein was confirmed by mass spectrometer, and the concentration was determined by a Lowry assay.51 Addition of the Metal Ion. Cu(I) (Tetrakis (acetonitrile) copper(I) hexafluorophosphate (Aldrich)) was added to a 0.5 mM protein solution under nitrogen gas to preserve anaerobic conditions. Cu(II) EPR signal was not observed at any time. When Atox1 and c-terminal CTR1 peptide were present in the solution, the Atox1 protein concentration, like the peptide concentration, was 0.25 mM. At low temperature measurements, 30% glycerol was added to the protein solution. The sequence of addition of metal ion and CTR1 c-terminal domain to the Atox1 protein solution was tested, and it was

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RESULTS The CTR1 c-terminal domain contains 15 amino acids: SWKKAVVVDITEHCH. Studies have shown that mutation of the single cysteine residue in the human CTR1 c-terminal domain does not affect the Cu(I) in cell concentration;1,2 however, it might affect the copper regulation.11 Other studies suggest that the cysteine residue is important for cisplatin coordination,54 and for the interaction with a general methionine segment.28 Herein, we probed the structural changes that occurred in Atox1 in the presence of the CTR1 c-terminal domain with the cysteine residue and without it. Three additional peptides were synthesized, one without the last three amino residues, HCH (pep3), one that replaced the cysteine with a glycine (pep4), and another one which replaced the cysteine with a methionine (pep5). The CTR1 c-terminal peptides that were synthesized are listed in Table 1. In this study, we used the site-directed spin labeling method, as shown in Figure 1. There are three cysteine residues in Atox1 protein: Cys12 and Cys15 are involved in the cysteine bridge and cannot be labeled, whereas Cys41 is accessible for labeling. All EPR data presented here was performed on a labeled Atox1. Figure 2A presents the mass spectra of labeled pep2, confirming 100% of labeled peptide (CTR1 c-terminal domain). Figure 2B 5834



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Probing the Interaction between the CTR1 c-Terminal Domain and the Metallochaperone Atox1 by CW-EPR at Room Temperature. The CW-EPR spectra of Atox1 at room temperature, with and without the presence of Cu(I) and pep1, are presented in Figure 3A. The CW-EPR spectra were simulated using slow-motion theory derived by Freed and coworkers55 as implemented in the easyspin toolbox,52 and are

Figure 2. (A) ESI-mass spectrum of spin-labeled pep2 (expected mass 1935 Da). (B) MALDI-mass spectrum of Atox1 (expected mass 7672 Da). (C) ESI-mass spectrum of labeled Atox1, confirming addition of one MTSSL for an Atox1 monomer (expected mass 7847 Da). (D) SDS-PAGE gel of Atox1, confirming purified protein. Figure 3. CW-EPR spectra of (A) Atox1, Atox1+Cu(I), Atox1+pep1, and Atox1+pep1+Cu(I). The dotted lines are the simulated spectra using the slow motion theory implemented in the easyspin toolbox. (B) Atox1+pep1 at various Cu(I) concentrations. (C) Atox1 in the presence of pep1, pep3, pep4, and pep5; the arrows mark the characteristic signals corresponding to exchange interaction.

presents the mass spectra of Atox1, and labeled Atox1 is presented in Figure 2C, confirming the addition of a single spin label to an Atox1 monomer. Figure 2D shows the SDS-PAGE gel of Atox1. 5835



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Figure 4. Q-band DEER signals and corresponding distance distributions for Atox1, Atox1+pep1, and Atox1+Cu(I). The gray distance distribution overlaid on the black distance distributions for Atox1 and Atox1+pep1 corresponds to the MMM simulations, described in the text. The inset shows the structure of Atox1 dimer with the distribution of spin label conformers attached to cys41, obtained from the MMM simulations.

Atox1 structure differently than with pep1. The addition of Cu(I) to these solutions did not affect the CW-EPR spectra. These experiments suggest that the last three amino residues of the c-terminal domain are important to the correct folding mechanism of the metallochaperone Atox1, and to its interaction with the CTR1 c-terminal domain. Probing the Interaction between Various Mutated CTR1 c-Terminal Domains and the Metallochaperone Atox1 by DEER at 80 K. To further explore the conformational changes that the c-terminal domain and the Atox1 experience upon interaction, double electron electron resonance (DEER) experiments were carried out. We performed DEER experiments on labeled Atox1 in the presence of Cu(I) and the CTR1 c-terminal domain (pep1). The DEER signals are presented in Figure 4. The presence of a dipolar interaction between spin labels confirms that the Atox1 is indeed a dimer. The DEER experiments on Atox1 revealed a bimodal distance distribution of 2.8 ± 0.7 and 4.5 ± 0.4 nm, after a 3D homogeneous background subtraction (see Figure S2, Supporting Information). The distribution larger than 4.0 nm might have been a ghost distribution within the 2 μs time domain signal.56,57 Hence, we checked the Tikhonov validation of this distribution (noise, background start, and dimensionality (between 2.0 and 3.0)) and the effect of the L parameter on this distribution within the DeerAnalysis program (Figure S3, Supporting Information). The validation confirmed that the distribution around 4.5 nm is present; however, its contribution to the signal varies between 20 and 50% within the experimental error. To shed some light onto this bimodal

presented in dotted lines in Figure 3A. For all spectra, the gtensor used was constant, g = [2.0087 2.006 2.0022]. The CWEPR spectrum of Atox1 was simulated with a correlation time of 2 × 10−9 s, and an isotropic electron−electron interaction (ωee) of 4 MHz (corresponds to a distance of 2.35 nm), line width 1.8 G, aN =16.5 G. Addition of Cu(I), in a ratio of Atox1:Cu(I) of 1:1, did not affect the CW-EPR spectrum. However, addition of the CTR1 c-terminal domain (pep1), either in the presence of Cu(I) or without it, caused a slight increase in the tumbling rate to 1 × 10−10 s, and decreased the line width to 1.3 G. Moreover, the spectrum was simulated without the presence of an isotropic electron−electron interaction. This suggests that the presence of the CTR1 cterminal domain slightly increased the mobility of the spin label attached to Atox1 but also reduced the presence of the isotropic electron−electron interaction between the two spin labels attached to the Atox1 dimer. Addition of excess Cu(I) to the Atox1-CTR1 c-terminal domain up to a ratio of 1:1:6 of Atox1:pep1:Cu(I) has no influence on the CW-EPR spectrum, as shown in Figure 3B. Figure 3C presents the CW-EPR spectra of the various peptides, listed in Table 1, in the presence of labeled Atox1. When pep1 was added to the Atox1 solution, a decrease in the electron−electron dipolar interaction between the doubly labeled Atox1 dimer occurred, as well as a slight increase in the mobility of the spin label. However, when pep3/pep4/pep5 was added to the Atox1 solution, a spectrum characterized by an exchange interaction appeared (marked by arrows), suggesting that the presence of these peptides folds the 5836



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Figure 5. Q-band DEER signals and corresponding distance distributions for Atox1 in the presence of pep1, pep3, pep4, and pep5.

(MBD) of the ATP7B protein were previously reported by single molecule FRET.12 The addition of the CTR1 c-terminal domain to the Atox1 solution in a ratio of 1:1 also resolves a bimodal distribution of 2.5 ± 0.4 and 4.2 ± 0.3 nm. The distribution around 2.5 nm, however, was found to be narrower in the presence of pep1, also observed in the clear time-domain modulations in the DEER signal. This suggests that Atox1 is less flexible, and is more localized, while interacting with pep1. These results complement the CW-EPR data, which showed a slight reduction in the dipolar interaction when pep1 was added to the Atox1 solution. Moreover, the sweep width of the echodetected spectrum at 80 K was narrower by 5 ± 0.5 G in the presence of pep1 than the sweep width of the echo-detected spectrum of Atox1 (data not shown). This suggests that, when pep1 was added to the solution, there was less contribution of smaller distances 4.0 nm) were absent in the presence of pep3/pep4/pep5. DEER experiments were also performed on Atox1 in the presence of Cu(I) (1:1:1 Atox1:pep1:Cu(I)) and pep3/pep4/

pep5 and are presented in Figure 6. The presence of Cu(I) did not significantly affect the distance distributions that were obtained without it and are presented in Figure 5. This is also consistent with the CW-EPR data, which did not reveal any conformational changes to Atox1 as a function of Cu(I) (Figure 3B). The geometrical state that is not associated with the crystal structure remains the dominant configuration state also in the presence of Cu(I). For pep3 the distance distribution obtained is 2.8 ± 0.8 nm, for pep4 it is 2.7 ± 0.8 nm, whereas for pep5 it is 2.8 ± 0.7 nm. Probing the Interaction between Labeled CTR1 cTerminal Domain (pep2) and Labeled Atox1 by DEER at 80 K. In order to explore the interprotein interaction between the CTR1 c-terminal domain and the dimer Atox1, the cysteine residue of the CTR1 c-terminal was also labeled with a MTSSL, and was named pep2. Figure 7A presents a model including the inter interaction between pep2 and Atox1 and the intra interaction within the two spin labels attached to the Atox1 dimer. The DEER signal should be composed of both the inter and intra dipolar interactions. However, the studied system is a transient system, where CTR1 transfers copper to the Atox1, 5838



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Figure 7. (A) Model for the interaction between the spin-labeled CTR1 c-terminal domain (pep2) and the Atox1 dimer. (B) Q-band DEER signals and corresponding distance distributions for Atox1 in the presence of pep1 and pep2. The insets show the corresponding Fourier-transformed spectra.

and the Atox1 transfers the copper ion to the ATP7A/B. Such systems are characterized by a weak interaction between the initial proteins to their targets, as previously mentioned.28 Hence, not all the Atox1 proteins that are detected by the EPR are coupled to the CTR1 c-terminal domain, and the probability of exciting an inter dipolar interaction between pep2 and Atox1 is reduced. Hence, the contribution of the inter dipolar interaction should be less than the contribution of the intra dipolar interaction to the DEER time domain signal. Figure 7B presents the DEER signal of Atox1+pep2 in the presence of Cu(I), and the corresponding distance distributions. For comparison, the signal of Atox1+pep1+Cu(I) is also presented in Figure 7B. The addition of pep2 to the Atox1 solution also presents a bimodal distribution, similar to the addition of pep1 to Atox1 solution. However, the distribution around 2.5 nm is much broader than in the presence of pep1. This is also observed in the Fourier transformed spectra (insets in Figure 7B), where in the presence of pep1 two clear signals appear at 0.6 and 2.5 MHz, corresponding to distances of about 4.4 and 2.7 nm, respectively. However, in the presence of pep2, the signal around 2.5 MHz is not well resolved due to the broad distribution. This suggests that the CTR1 c-terminal and the Atox1 dimer closely interact between them, and that at least one of the interdistance distributions between pep2 and the Atox1 dimer should be within the 2.5 ± 1.0 nm distribution. CD Characterization. Figure 8A shows the CD spectra of Atox1 and Atox1 in the presence of pep1 or pep3 with Cu(I) (in a 1:1:1 ratio Cu(I):peptide:Atox1). Atox1 has an overall βαββαβ fold structure, and its CD spectrum is characterized by a negative peak at 220 nm. The CD spectra of pep1 and pep3 are identical and show a negative peak at 198 nm, corresponding to a random coil structure (see Figure S4A, Supporting Information). Addition of pep1/pep3 to the Atox1 solution caused a change in the region of 190−210 nm, as expected. In order to explore the thermal stability of Atox1 with pep1 and pep3, CD spectra were recorded as a function of temperature. Figure 8B presents the CD spectra of Atox1 in the presence of Cu(I) (ratio 1:2 Cu(I):Atox1) at various temperatures. As the temperature increased, unfolding of the Atox1 occurred, causing a disappearance of the 220 nm peak,

Figure 8. (A) CD spectra for Atox1+Cu(I), Atox1+Cu(I)+pep1, and Atox1+Cu(I)+pep3 at 20 °C. (B) CD spectra for Atox1+Cu(I) at various temperatures from 20 to 90 °C.

and formation of a negative peak at 205 nm. The changes at 220 and 205 nm for Atox1+Cu(I) and Atox1+Cu(I) in the presence of pep 1 and pep3 are presented in parts A, B, and C of Figure 9, respectively, and a linear fit was applied. For Atox1 in the presence of Cu(I), the thermal stability is the lowest, and 5839



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one has not yet been resolved. Moreover, the data presented here suggests that the dimer nature of Atox1 is preserved while interacting with the c-terminal domain of CTR1. The Atox1, however, assumes a more confirmed structure in the presence of the CTR1 c-terminal domain, yielding a narrower distance distribution function in the EPR data. Chen et al. found by FRET that the Atox1 interacts with the metal binding domain no. 4 of the ATPases Wilson disease protein (WDP) in two geometries as well, which can interconvert dynamically.12 Where FRET can follow the Cu(I) transient from Atox1 to the MBD in real time and obtained kinetic information, EPR cannot directly observe such a mechanism but rather inform on structural and dynamics changes that occur on the Atox1 while interacting with a target protein. Chen et al. suggested that the ability of Atox1 and MBD4 to form multiple interaction complexes with different interaction geometries has functional significance. First, it increases the probability of complex formation when Atox1 and WDP encounter through diffusion inside cells. The formed complex may proceed to accomplish Cu(I) transfer or, if unproductive, convert to another complex for Cu(I) transfer. Second, it raises the possibility of Atox1 interacting with two WDP MBDs simultaneously and hence cooperative efforts among WDP MBDs for Cu(I) transfer. The two geometrical states assumed by the Atox1 were also preserved in the presence of Cu(I), as was observed here by EPR. Moreover, we found that while in the presence of the WTCTR1 c-terminal domain the Atox1 favors two geometrical states and in the presence of the mutated CTR1 c-terminal domain only one geometrical state is assumed by the Atox1. Kinetic CD measurements showed that the c-terminal CTR1 domain without the last three amino acids reduced the thermal stability of the Atox1. This suggests that the last three amino acids of the CTR1 cterminal domain are important for the following: (1) maintaining the crystal structure geometry of the Atox1 in addition to the second conformational state, (2) obtaining a more constrained structure of the Atox1 protein, and (3) increasing the thermal stability of the Atox1. Maryon et al.11 reported using 64Cu uptake assay in HEK293 cells, in which the intracellular side of the hCTR1, especially the last three amino acids, HCH, are not essential for copper permeation but are important for copper regulation of the copper entry rate. Together with the data obtained in this research, this suggests that the last three amino acids of the intracellular hCTR1 are important for a proper interaction and folding mechanism with the target protein, as well as for cellular copper transfer and regulation. We have recently reported on the interaction between the CTR1 c-terminal domain and a general copper binding site, represented by a methionine segment.28 There as well, the methionine segment assumed a more confirmed structure in the presence of the CTR1 c-terminal domain. Moreover, the interaction between the methionine segment and pep3−pep5 resulted in CW-EPR spectra also dominated by an exchange interaction, which is different from the interaction that was observed between the methionine segment and pep1. This suggests that the methionine segment can indeed represent a good general copper binding compared to other copper metallochaperones. This cysteine to metal coordination was also recently found to be important to the coordination of the cisplatin drug.54

Figure 9. Temperature effect at 205 nm (black squares) and 220 nm (gray squares) for (A) Atox1+Cu(I), (B) Atox1+Cu(I)+pep1, and (C) Atox1+Cu(I)+pep3. The linear fit slopes at 205 nm are 0.154 deg·cm2· 10−3/(dmol·°C) for Atox1, 0.09 deg·cm2·10−3/(dmol·°C) for Atox1+pep1, and 0.105 deg·cm2·10−3/(dmol·°C) for Atox1+pep3.

the rate of change of the CD spectra as a function of temperature is the highest. The presence of pep1 and pep3 stabilizes the Atox1 a bit, and the rate of change at 220 nm is smaller; however, in the presence of pep1, the rate is 10% less than with pep3. This suggests a better thermal stability of the Atox1 in the presence of pep1 rather than pep3.

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DISCUSSION Herein, we probed the specific interaction between the CTR1 c-terminal domain and the metallochaperone Atox1 by EPR spectroscopy and CD. EPR data presented here showed that the Atox1 consists of two conformational states as a dimer. One is consistent with the reported crystal structure, and another 5840



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(5) Robinson, N. J.; Winge, D. R. Copper Metallochaperones. Annu. Rev. Biochem. 2010, 79, 537−562. (6) Rosenzweig, A. C. Copper Delivery by Metallochaperone Proteins. Acc. Chem. Res. 2001, 34, 119−128. (7) Arnesano, F.; Banci, L.; Bertini, I.; Huffman, D. L.; O’Halloran, T. V. Solution Strcuture of the Cu(I) and Apo Forms of the Yeast Metallochaperone, Atx1. Biochemistry 2001, 40, 1528−1539. (8) Wernimont, A. M.; Huffman, D. L.; Lamb, A. L.; O’Halloran, T. V.; Rosenzweig, A. C. Structural Basis for Copper Transfer by the Metallochaperone for the Menkes/Wilson Disease Proetin. Nat. Struct. Biol. 2000, 7, 766−771. (9) Nevitt, T.; Ohrvik, H.; Thiele, D. J. Charting the Travels of Copper in Eukaryotes From Yeast to Mammals. Biochem. Biophys. Acta 2012, 1823, 1580−1593. (10) Kaplan, J. H.; Lutsenko, S. Copper Transport in Mammalian Cells: Special Care for Metal with Special Needs. J. Biol. Chem. 2009, 284, 25461−25465. (11) Maryon, E. B.; Molloy, S. A.; Ivy, K.; Yu, H.; Kaplan, J. H. Rate and Regulation of Copper Transport by Human Copper Transporter 1 (hCTR1). J. Biol. Chem. 2013, 288, 18035−18046. (12) Keller, A. M.; Benitez, J. J.; Klarin, D.; Zhong, L.; Goldfogel, M.; Yang, F.; Chen, T.-Y.; Chen, P. Dynamic Multibody Protein Interactions Suggest Versatile Pathways for Copper Trafficking. J. Am. Chem. Soc. 2012, 134, 8934−8943. (13) Benitez, J. J.; Keller, A. M.; Ochieng, P.; Yatsunyk, L. A.; Huffman, D. L.; Rosenzweig, A. C.; Chen, P. Probing Transient Copper Chaperone- Wilson Disease Protein Interactions at the SingleMolecule Level with Nanovesicle Trapping. J. Am. Chem. Soc. 2008, 120, 2446−2447. (14) Ralle, M.; Lutsenko, S.; Blackburn, N. J. Copper Transfer to the N-terminal Domain of the Wilson Disease Protein (ATP7B): X-ray Absorption Spectroscopy of Reconstituted and Chaperon-Loaded Metal Binding Domains and their Interaction with Exogenous Ligands. J. Inorg. Biochem. 2004, 98, 765−774. (15) Rodriguez-Granillo, A.; Crespo, A.; Estrin, D. A.; WittungStafshede, P. Copper-Transfer Mechanism From the Human Chaperone Atox1 to a Metal-Binding Domain of Wilson Disease Protein. J. Phys. Chem. B 2010, 114, 3698−3706. (16) Wernimont, A. K.; Yatsunyk, L. A.; Rosenzweig, A. C. Binding of Copper(I) by the Wilson Disease Protein and its Copper Chaperone. J. Biol. Chem. 2004, 279, 12269−12276. (17) Tumer, Z.; M?ller, L. B.; Horn, N. Mutation Spectrum of ATP7A, the Gene Defective in Menkes Disease. Adv. Exp. Med. Biol. 1999, 448, 83−95. (18) Scushan, M.; Bhattacherjee, A.; Ben-Tal, N.; Lutsenko, S. A Structural Model for the Copper ATPase ATP7B to Facilitate Analysis of Wilson Disease Causing Mutations and Studies of Tranport Mechanism. Metallomics 2012, 4, 669−678. (19) Boal, A. K.; Rosenzweig, A. C. Crystal Structure of Cisplatin Bound to Human Copper Chaperone. J. Am. Chem. Soc. 2009, 131, 14196−14197. (20) Portnoy, M. E.; Rosenzweig, A. C.; Rae, T.; Huffman, D. L.; O’Halloran, T. V.; Culotta, V. C. Strcuture-Function Analyses of the Atx1Metallochaperone. J. Biol. Chem. 1999, 274, 15041−15045. (21) Hussain, F.; Olson, J. S.; Wittung-Stafshede, P. Conserbed Residues Modulate Copper Release in Human Copper Chapeone Atox1. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 11158−11163. (22) Palm-Espling, M. E.; Andersson, D. C.; Bjorn, E.; Linusson, A.; Wittung-Stafshede, P. Determinants for Simultaneous Binding of Copper and Platinum to Human Chaperone: Atox1: Hitchhiking not Hijacking. PLoS One 2013, 8, e70473. (23) Aller, S. G.; Unger, V. M. Projection Structure of the Human Copper Transporter CTR1 at 6Å Resolution Structure Reveals a Compact Trimer with a Novel Channel-Like Architecture. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 3627−3632. (24) De Feo, C. J.; Aller, S. G.; Siluvai, G. S.; Blackburn, N. J.; Unger, V. M. Three-Dimensional Strcuture of the Human Copper Transporter hCTR1. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 4237−4242.

Finally, we showed that the CTR1 c-terminal domain interacts with the Atox1 dimer, where one of the distance distributions between the cysteine on the CTR1 c-terminal domain and Cys41 of one of the Atox1 monomers should be around 2.5 nm.

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CONCLUSIONS Exploring the interaction between Atox1 and the c-terminal domain of CTR1 by EPR spectroscopy and CD yielded several important findings advancing the understanding of the Cu import mechanism from the CTR1 c-terminal domain. It was revealed that the Atox1 keeps its dimer nature also in the presence of the CTR1 c-terminal domain. Two geometrical states, however, are assumed by the Atox1. One is similar to the geometrical state reported by the crystal structure, while the latter has not yet been constructed. In the presence of the CTR1 c-terminal domain, both states are assumed; however, the structure of Atox1 is less flexible in the presence of the CTR1 c-terminal domain. When the last three amino acids of the CTR1 c-terminal domain, HCH, are missing, or when the cysteine is mutated to methionine or alanine, only one geometrical state is assumed by the Atox1, which is not the crystal structure state. In addition, the flexibility of the Atox1 protein is lower in the presence of the wt-CTR1 c-terminal, and the thermal stability of the Atox1 is higher, compared to the mutated CTR1 c-terminal domain. The restriction of the Atox1 in the presence of the CTR1 c-terminal domain is similar to the behavior of a methionine segment, a general copper metal binding site, when it interacts with the CTR1 c-terminal domain.

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ASSOCIATED CONTENT

S Supporting Information *

Supporting figures for additional EPR and CD data. This material is available free of charge via the Internet at http:// pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: 972-3-7384329. Fax: 972-3-7384053. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This study was supported by the Israel Science Foundation, grant no. 280/12. The Elexsys E580 Bruker EPR spectrometer was partially supported by the Israel Science Foundation, grant no. 564/12.

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