Revealing the Effect of Protein Weak Adsorption to Nanoparticles on

Jan 18, 2017 - Revealing the Effect of Protein Weak Adsorption to Nanoparticles on the Interaction between the Desorbed Protein and its Binding Partne...
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Revealing the Effect of Protein Weak Adsorption to Nanoparticles on the Interaction between the Desorbed Protein and its Binding Partner by Surface-Enhanced Infrared Spectroelectrochemistry Li Liu, Li Zeng, Lie Wu, and Xiue Jiang* State Key Laboratory of Electroanalytical Chemistry, Changchun Institute of Applied Chemistry, Chinese Academy of Sciences, Changchun 130022, People’s Republic of China S Supporting Information *

ABSTRACT: In recent years, the properties of protein corona have attracted intense interest in the field of nanobio interface, but a long-ignored research issue is how the desorbed proteins suffering from conformational change upon weak association with nanoparticles affect their functional properties when further interacting with their downstream protein partners. In this Article, surfaceenhanced infrared absorption spectroscopy (SEIRAS) and electrochemical cyclic voltammetry were used to study the adsorption and redox properties of the soluble cytochrome c (cyt c) on 11-mercaptoundecanoic acid (MUA) selfassembled monolayer (SAM) after weakly binding to and then desorbed from nano-TiO2. For the first time, our study reveals that the weak interaction between cyt c and nano-TiO2 induces the protein to undergo a heterogeneous conformational change. More importantly, the cyt c with a largely unfolded conformation exhibits a weaker interaction with its binding partner mimics than the native-like cyt c but a faster adsorption rate even at a concentration that is much lower than that of native-like cyt c. Correspondingly, the cyt c with a large unfolding shows a greatly positive-shifted formal potential (Ef) relative to the native-like protein possibly due to the disruption of the pocket structure of heme in the vicinity of Met80. These properties could enable the largely unfolded cyt c to undergo a favorable binding but an unavailable electron transfer to cytochrome c oxidase even in the presence of high-concentration native cyt c, probably causing the disruption of electron flow.

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particle surface on their further interaction with downstream proteins, cellular receptors, or the other key structures in signal transduction. Cytochrome c (cyt c) is a roughly spherical heme protein with a diameter of ∼3.4 nm in mitochondrion in eukaryotes,11 transporting electrons from cytochrome bc1 complex to cytochrome c oxidase and eventually driving ATP synthesis.12 Its electron transfer function has been extensively studied by performing direct electrochemistry of cyt c adsorbed on alkanethiols-modified self-assembled monolayers (SAMs) with terminal carboxylate groups as its binding partner mimics.13,14 Therefore, cyt c is a good model protein for studying the effect of weak interaction between protein and nanomaterials on the binding of the desorbed protein to its biological partner with 11-mercaptoundecanoic acid (MUA) SAMs as its binding partner mimics. TiO2 nanomaterials (nano-TiO2) are finding an increased application in the food industry, medicine industry and daily life, so their potential toxic effects in humans and animals have been intensively and extensively investigated.15,16 Given the ample information on its physicochemical properties and toxic effect,17−20 nano-TiO2 has general significance as a

dvances in nanotechnology have accelerated the translation of nanomaterials into different areas of industry, technology, and medicine in the past decades, which increases the chance of human exposure to nanoparticles in our daily life. The high surface free energy of nanoparticles will drive progressive adsorption of biomolecules onto their surfaces to form hard corona (strongly associated and long-lived biomolecular boundary layer) and soft corona (the loosely associated and rapidly exchanging biomolecules that attach to either the nanoparticle surface in short time or to the hard corona for a longer time1) when they are in a biological milieu.2 The interaction of nanoparticles with biomolecules, such as proteins, not only provides a new biological identity to the nanoparticles but also disturbs the conformation of the proteins following the formation of soft or hard corona. Few doubt that both aspects are linked to biological impact.3 Therefore, studies on nanoparticle−protein interactions have received considerable attention, including the quantification of protein adsorption kinetics,4,5 disclosure of the biological effect of corona,6,7 and identification of the composition of hard corona.8,9 More recently, professor Dawson’s group has mapped the functional motifs of transferrin-coated polystyrene nanoparticles.10 Nevertheless, a long-ignored issue is the biological impact of the desorbed proteins suffering from conformational changes upon loose association with a nano© 2017 American Chemical Society

Received: May 20, 2016 Accepted: January 18, 2017 Published: January 18, 2017 2724

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730

Article

Analytical Chemistry model nanomaterial to evaluate the possible disturbance to protein conformation and function after weak interaction with nanomaterials. Electrochemical cyclic voltammetry is a label-free characterization technique. It can identify the different electron transfer processes of the cyt c with varied degree of unfolding.21 Surface-enhanced infrared absorption spectroscopy (SEIRAS) has been proved to be an extremely sensitive technique to gain the molecular information at interface even when the surface is immersed in water by taking advantage of distance-dependent enhancement effect.22 It can detect in situ the adsorption process of sample species at the liquid/solid interface in realtime. On the basis of the spectral change of interfacial water molecules induced by the sample adsorption, the intermolecular interaction can be revealed in situ and in real-time.23 Combining both information from cyclic voltammetry and SEIRAS, it can disclose adsorption-dependent electron transfer property.24 Moreover, potential-induced SEIRA difference spectroscopy can identify minor structural changes in biomolecules induced by electron acceptance and donation,25−27 and the possible electron-transfer mechanism of cyt c at the interface.24 In this work, we study the interaction of cyt c with its electron-transfer partner mimics of MUA SAMs after weakly binding to and then disengaging from the surface of nano-TiO2, and the subsequent effect on its electron transfer function at the molecular level by combining SEIRAS with cyclic voltammetry. It is found that the cyt c exhibits heterogeneity in conformation after weakly binding to nanoTiO2. The species with a large unfolding exhibit a weak interaction with its binding partner mimics but a fast adsorption rate, which results in a greatly positive-shifted formal potential (Ef) relative to native-like protein.

prepared by ultrasonically dispersing 10 mg of nano-TiO2 in 2 mL of PBS (10 mM, pH 7) for 5 min were mixed respectively with 0.5 mL of 15 μM and 0.6 mL of 68.7 μM cyt c (Sigma) dissolved in PBS (10 mM, pH 7), obtaining two mixtures at the ratios of nano-TiO2 (mg) to cyt c (nmol) as 1:1.5 (sample 1) and 1:20.6 (sample 2), respectively. Both mixtures were stirred vigorously for 2 h, and then centrifuged at 12 000g for 10 min or at 8000g for 10 min for the sample 1 and 2, respectively. The stirring time was determined based on the fact that the obtained supernatant shows an unchanged UV−vis spectrum of cyt c. The inductively coupled plasma-optical mass spectrometry (ICP-MS) (DIONEX ICS-1000, USA) analysis indicates that both supernatants contain no more than 2.4 ppb (μg/L) nano-TiO2. At the ratio of 1:1.5, the obtained supernatant and nano-TiO2/cyt c complex layer are denoted as cyt c (1) and complex (1), respectively. At the ratio of 1:20.6, they are denoted as cyt c (2) and complex (2), respectively. The complexes were regarded as nano-TiO2 cyt c corona (bound) and the supernatants as unbound cyt c including most of the desorbed cyt c after weak interaction with nano-TiO2. The sample of highly diluted cyt c (2) (HD cyt c (2)) was prepared by 90-fold dilution of cyt c (2). For comparison, the sample of native cyt c (N-cyt c) was handled in exactly the same manner as that used in the preparation of cyt c (2) except the absence of nano-TiO2, and by highly diluting N-cyt c the HD N-cyt c was obtained. The concentration of protein was determined by UV−vis adsorption spectroscopy (LAMBDA 25 spectrometer, PerkinElmer) using a 1 mm path length cell and with the absorptivity reported by Margoliash and Frohwirt;29 the average of 4 scans was applied for each concentration measurement. Circular Dichroism (CD) spectra of protein were recorded on a JASCO J-810 CD spectrometer (Hitachi, Japan) using a 1 mm path length cell; 100 scans and 4 scans were applied for sample spectra measurements at low and high concentrations, respectively. The detection limits of native cyt c PBS (10 mM, pH 7) are 8.8(±0.84) × 10−8 M for UV−vis measurement and 2.1(±0.5) × 10−7 M for CD measurement. Si Substrate Treatment and Au Film Preparation. One of the three square flat surfaces of a triangular Si prism was polished with 1.0 μm Al2O3 slurry and then washed thoroughly with deionized water. The cleaned Si substrate was treated by immersion in 40 wt % aqueous solution of NH4F for 1 min. Then, a thin gold film was prepared on it by chemical deposition.30 The flat surface of the treated Si substrate was exposed to a 1:1:1 volume mixture of (1) 0.03 M NaAuCl4, (2) 0.3 M Na2SO3 + 0.1 M Na2S2O3 + 0.1 M NH4Cl, and (3) 2.5 vol % HF solution for 90 s. After rinsing in deionized water, the Au/Si prism was transferred into a 0.1 M H2SO4 solution. The gold film was cleaned by several oxidation−reduction scans in the range of −0.1 and 1.4 V (vs Ag/AgCl). After electrochemical cleaning, the Au/Si prism was tightened to a poly(trifluorochloroethylene) cell with a Viton O-ring and a Cu plate keeping electric contact with the gold film. SEIRAS Monitoring of MUA SAMs Formation and the Subsequent Adsorption of Cyt c Samples. To monitor the formation of MUA (Sigma) SAMs, a reference spectrum was first scanned in ethanol. With the addition of 0.1 mM MUA into the ethanol, sample spectra were recorded. At the adsorption time of 90 min, the MUA SAM was obtained.31 Then, the MUA/Au film was washed with ethanol, deionized water and 10 mM PBS (pH 7) sequentially. After that, 500 μL of 10 mM PBS (pH 7) was added onto the MUA/Au and scanned to obtain a reference spectrum. Following the addition

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EXPERIMENTAL SECTION Synthesis and Characterization of Nano-TiO2. NanoTiO2 was synthesized as described elsewhere.28 Briefly, 10 mL of pure TiCl4 was first dissolved in 40 mL of deionized water generated using a Millipore Milli-Q system (Billerica, MA) and then mixed with 12.5 mL of 2 M NaOH solution under vigorous agitation. The mixture was then refluxed for 1h. White precipitation occurred immediately as soon as the solution was refluxed. After washing with deionized water and ethanol three times respectively and then being dried at 60 °C, the titania powders were obtained. The structure, morphology and size of the as-synthesized nano-TiO2 were characterized by X-ray diffraction (XRD) that was performed on a Bruker D8 Advance X-ray diffractometer (German, Cu Kα radiation, λ = 1.5406 nm), by transmission electron microscopy (TEM) that was performed on an H-600 electron microscope (Hitachi, Japan) operated at 75 keV, and by atomic force microscopy (AFM) that was recorded on a Digital Instrument Nanoscope IIIa Multimode system (Santa Barbara, CA) with a silicon cantilever using the tapping mode. The hydrodynamic diameter and electrophoretic mobility of the nano-TiO2 in 10 mM phosphate buffer solution (PBS, pH 7) were determined using a dynamic light scattering (DLS) method (ZEN 3600, Malvern Instrument, UK). Nitrogen adsorption−desorption measurements were made on an ASAP 2020-Physisorption Analyzer (Micrometritics, USA). The specific surface area was calculated using Brunauer−Emmett−Teller (BET) equation at P/P0 < 0.3. Preparation and Characterization of the Treated Cyt c Samples. Either unbound or bound cyt c samples were prepared in two ways: 1 and 0.4 mL of 5 mg/mL nano-TiO2 2725

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730

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Analytical Chemistry of each cyt c sample in the PBS, sample spectra were recorded for 30 min so that the protein adsorption reached saturation and a cyt c-adsorbed layer was formed onto the MUA/Au surface. For each spectrum, 512 scans were collected with a spectral resolution of 4 cm−1. Electrochemical Measurements and Redox-Induced SEIRA Difference Spectroscopy of Various Cyt c/MUA Films. Cyclic voltammograms (CVs) were recorded with a CHI 830A electrochemical workstation (CH Instruments, Austin, TX). The Au/Si substrate was used as a working electrode, a Pt wire and an Ag/AgCl electrode with saturated KCl solution as counter and reference electrodes, respectively. The roughness factor of the nanostructured Au film was determined to be 2.0, using the Au-oxide reduction charge density method. The used buffer of 10 mM PBS (pH 7) was purged with high-purity nitrogen for at least 10 min prior to electrochemical measurements to remove the dissolved oxygen. To obtain the redox difference spectrum of the electric couple at 0.01 or 0.02 V, a reference SEIRA spectrum was taken from the N-cyt c/MUA or cyt c (2)/MUA or HD cyt c (2)/MUA film at −0.1 V, where the adsorbed native or native-like cyt c was fully reduced, and the sample spectrum was acquired at 0.1 V, after which the adsorbed cyt c was fully oxidized. To obtain the redox difference spectra of the electric couple at 0.18 or 0.2 V, a reference SEIRA spectrum was taken from the cyt c (2)/ MUA or HD cyt c (2)/MUA film at 0.05 V, where the adsorbed cyt c with a largely unfolded conformation was fully reduced, and the sample spectrum was acquired at 0.4 V, after which it was fully oxidized. The whole procedure was repeated nine times, and the difference spectra were averaged to improve the signal-to-noise ratio.

Committee on Powder Diffraction Standards (JCPDS) and no other minor phase peaks are detected. Figure 1B exhibits a typical TEM image of the as-prepared TiO2 nanowires. It can be seen that the super thin nanowires (3−4 nm in diameter) align into flat-cable-like “stacks” with their long axis parallel to each other. The self-assembled TiO2 flat cables have a length of 400−800 nm and a width of 70−130 nm (hereinafter referred to as “nano-TiO2”). AFM image shows that the height of the nano-TiO2 is ∼4 nm (Figure 1D). The hydrodynamic diameter measured by DLS is used to evaluate the stability of nano-TiO2 in 10 mM PBS (pH 7) over time with and without cyt c (Figure 1C). There is no obvious change in the hydrodynamic diameter of either nano-TiO2 or nano-TiO2/cyt c complex in 10 mM PBS (pH 7) within 3 h although the hydrodynamic diameter of the complex (∼1000 nm) is obviously larger than that of the unbound nano-TiO2 (∼300 nm). It is worth to note that the hydrodynamic size of the nonspherical TiO2 flat cables is much smaller than their actual length (TEM result) since the hydrodynamic diameter defines a sphere according to the intensity fluctuation of scattered light caused by the Brownian motion of particles.32 The electrophoretic mobility of the nanoTiO2 in 10 mM PBS (pH 7) is −3.28 ± 0.12 mV averaged from seven measurements and its BET specific surface area was measured to be 86.7 m2/g. The typical nitrogen adsorption− desorption isotherm of nano-TiO2 is shown in Figure S1. Far-UV CD Measurement of the Conformation of Cyt c Desorbed from Nano-TiO2. Through vigorously stirring the mixture of nano-TiO2 and native cyt c in 10 mM PBS (pH 7) and then centrifuging the mixture, we obtained the cyt c (1)/ complex (1) and cyt c (2)/complex (2) at the ratios of nanoTiO2 (mg) to cyt c (nmol): 1:1.5 and 1:20.6, respectively. In terms of concentration of protein in the supernatant, the cyt c (1) is 0.4(±0.034) μM and the cyt c (2) is 35(±0.26) μM. In terms of protein coverage, the complex (1) is 1 cyt c molecule/ (10 nm)2 nano-TiO2 and the complex (2) is 2.15 cyt c molecules/(10 nm)2 nano-TiO2. To disclose the conformational change of cyt c after weakly binding to nano-TiO2, farUV CD measurements of cyt c (1) and cyt c (2) were carried out since the technique is regarded as a sensitive probe for protein secondary structure.33 Figure 2 shows the far-UV CD spectra of background, 0.4(±0.034) μM cyt c (1), 35(±0.26) μM cyt c (2), 35(±0.14) μM native cyt c, as well as diluted cyt c (2) and diluted native cyt c in the same concentration as the cyt

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RESULTS AND DISCUSSION Structural Characterization of Nano-TiO2. The XRD pattern of the as-synthesized TiO2 is shown in Figure 1A. All peaks can be well indexed to tetragonal rutile phase TiO2 in the Powder Diffraction File (PDF No. 21-1276) published by Joint

Figure 2. Far-UV CD spectra of 0.4(±0.034) μM cyt c (1), 35(±0.26) μM cyt c (2), and 35(±0.14) μM native cyt c in 10 mM PBS (pH 7). For comparison, the spectra for diluted native cyt c and diluted cyt c (2) in the same concentration as the cyt c (1) are presented. Background spectrum was obtained with 2.4 μg/L nano-TiO2 suspension in 10 mM PBS (pH 7).

Figure 1. XRD pattern (A), TEM image (B), size stability (C) and AFM image (D) of the as-prepared nano-TiO2. (C) Stability of the nano-TiO2 and nano-TiO2/cyt c complex in 10 mM PBS (pH 7) over time within 3 h from the viewpoint of hydrodynamic diameter measured by DLS. 2726

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730

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Analytical Chemistry c (1). The background spectrum was obtained by scanning 10 mM PBS (pH 7) containing 2.4 ppb nano-TiO2, the maximum concentration of nano-TiO2 in the supernatants of the complex (1) by ICP-MS analysis. For native cyt c, the 222 nm dichroic band is associated with α-helical n-π amide transitions and the 209 nm dichroic band is corresponding to the π−π* amide transitions.34 For the cyt c (2), the CD intensity at 222 nm exhibits almost no change relative to that of native cyt c at the same concentration without and even with high salt concentration (Figure S2). However, for the cyt c (1), the CD intensity at 222 nm is much less than those of both the diluted cyt c (2) and the diluted native cyt c at the same concentration. Relative to native cyt c, there is a 34 ± 2% decrease in the intensity at 222 nm, suggesting a large disturbance to protein secondary structure.35 According to the report,36 the conformational change of cyt c might be induced mainly by the hydrophobic interaction with nano-TiO2. SEIRA Spectroelectrochemistry Study of Electron Transfer Function and Adsorption Properties of TiO2Treated Cyt c Samples on MUA SAMs. To reveal the effect of weak interaction with nano-TiO2 on the electron transfer function of cyt c, we investigated redox properties of cyt c (1) and cyt c (2) adsorbed on MUA SAM/Au surfaces by cyclic voltammetry, respectively. The cyt c (1) displays a pair of redox peaks at Ef of 0.23 V (cyt c (1) (0.23 V)) (Figure 3A, black)

even after several potential scans, which was greatly positively shifted compared to Ef of 0.02 V obtained from native cyt c (Figure 3A, red), and the peak-to-peak separation (ΔE) between oxidation and reduction potential was also enlarged from 0.046 V (native cyt c) to 0.054 V (cyt c (1)), indicating a more difficult oxidization process likely due to a largely unfolded conformation. The cyt c (2) exhibits two pairs of redox peaks: one pair at Ef of 0.02 V with ΔE of 0.05 V and the other pair at Ef of 0.18 V with ΔE of 0.18 V (Figure 3B, black). The separation between Ef of the two electric couples is 0.16 V and thus the two redox processes of cyt c (2) correspond to two species of cyt c. The electric couple of 0.18 V is unstable, disappearing completely after several potential scans (Figure 3B, green). Given the similar Ef, it is reasonable to deduce that the cyt c (2) corresponding to the electric couple of 0.02 V, cyt c (2) (0.02 V), should have native-like conformation, while the cyt c (2) corresponding to the electric couple of 0.18 V, cyt c (2) (0.18 V), should possess a largely unfolded conformation, similar to the conformation of cyt c (1). Although far-UV CD shows almost no change in the conformation of cyt c (2) possibly due to the lower concentration of largely unfolded cyt c than the detection limit of CD measurement, we can clearly identify heterogeneous cyt c (2) with varied degree of unfolding by electrochemical cyclic voltammetry because the largely unfolded cyt c has a faster adsorption rate than native-like cyt c (discussed later) although a rather low concentration in the bulk phase of cyt c (2). To further reveal the binding properties between heterogeneous cyt c and its electron transfer partner mimics, we respectively investigated the adsorption of cyt c (1) and cyt c (2) on MUA SAMs by SEIRAS, considering that the terminal carboxylate groups of MUA have always been used as cyt c binding partner mimics for a long time.13,25,31 The adsorption of cyt c (1) results in the appearance of very weak amide I and II bands as well as the positive band assigned to OH stretching vibration of water within the saturation adsorption time of 30 min (Figure 3C), which is significantly different from the adsorption of native cyt c. For native cyt c, the strong electrostatic interaction with MUA results in displacement of the water molecules being preadsorbed on the carboxylate groups so as to produce a strong negative band of water at ∼3500 cm−1 (Figure 3D, red). For cyt c (1), the adsorptioninduced positive water bands could be explained based on two considerations. First, the large degree of unfolding results in the enhanced hydration degree of protein due to the exposure of nonpolar residues buried in the interior of a protein,37,38 which could enable the cyt c (1) to carry more water molecules onto the MUA SAM upon adsorption than native cyt c. Second, the weak electrostatic interaction with MUA due to the interference with the domain structure surrounding the adsorption sites of cyt c (1) (demonstrated later) could not effectively displace the preadsorbed water molecules. For cyt c (2), the initial adsorption also results in weak amide I and II bands, as well as positive OH stretching vibration of water, similar to the adsorption of cyt c (1). Then, the positive water band gradually turns into a relatively strong negative peak at ∼3500 cm−1 and a flattened positive band at ∼3300 cm−1 with adsorption time (Figure 3D, black). Associated with cyclic voltammogram of cyt c (2) and referring the SEIRA behaviors of native cyt c and cyt c (1), it is possible to propose that the appearance of positive water band corresponds to the adsorption of largely unfolded cyt c (2), cyt c (2) (0.18 V), while the appearance of negative water band corresponds to the adsorption of native-like cyt c

Figure 3. (A, B, F) CVs for various cyt c-adsorbed films from 0.4(±0.034) μM cyt c (1), 35(±0.26) μM cyt c (2), and HD cyt c (2) (∼0.4 μM) in 10 mM PBS (pH 7) onto the MUA SAMs (black) and CVs after several potential scans of the cyt c (2)/MUA and HD cyt c (2)/MUA films (green); (C, D, E) SEIRAS monitoring of the adsorption processes of the individual cyt c samples within 30 min (black) from 1st to 8th: 1, 4, 8, 12, 16, 20, 24, and 30 min. In CVs, the cross and its nearby number denote the position and value of the formal potential for the local electric couple. For comparison, the CV of native cyt c/MUA and the SEIRAS adsorption of native cyt c at the concentration of 10(±0.14) μM onto the MUA SAMs are presented in A, B, and D (red). 2727

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730

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Analytical Chemistry (2), cyt c (2) (0.02 V). On the basis of the sequence of appearance and the evolution of the positive and negative water peaks, it can be deduced that the adsorption rate of cyt c (2) (0.18 V) is faster than that of cyt c (2) (0.02 V) although it undergoes weaker interaction with the MUA SAMs. Nevertheless, the strong adsorption ability could enable the nativelike cyt c (2) (0.02 V) to displace the adsorbed species of cyt c (2) (0.18 V) with the proceeding of adsorption so as to disturb the adsorption stability of cyt c (2) (0.18 V), leading to its electrochemical instability. On the other hand, the steric hindrance formed by rapid adsorption of cyt c (2) on MUA SAMs at a high concentration will greatly limit the interaction space of largely unfolded cyt c, thus destabilizing the adsorption of cyt c (2) (0.18 V). To validate this hypothesis, we diluted cyt c (2) up to 90 times and tested the adsorption and electrochemical properties of the highly diluted cyt c (2) (HD cyt c (2)) at the MUA SAMs. Different from cyt c (2) (Figure 3D), the initial adsorption of the HD cyt c (2) produces a positive water band with the intensity at ∼3300 cm−1 larger than that at ∼3500 cm−1 (Figure 3E), suggesting an alternative rearrangement of interfacial water molecules possibly because of the different adsorption orientation or/ and conformation of cyt c (2) at an extremely low concentration. As the adsorption proceeded, the positive water band at ∼3500 cm−1 turns into negative and then gradually increases downward while the positive water band at ∼3300 cm−1 gradually increases upward (Figure 3E), suggesting that the adsorption rate of the largely unfolded cyt c is still faster than that of the native-like cyt c even at an extremely low concentration. In terms of electrochemistry, the HD cyt c (2)/MUA also exhibits two pairs of redox peaks: one pair with Ef of 0.01 V and ΔE of 0.044 V (HD cyt c (2) (0.01 V)) and the other pair with Ef of 0.2 V and ΔE of 0.05 V (HD cyt c (2) (0.2 V)), as shown of the black line in Figure 2F. As expected, the electric couple of HD cyt c (2) (0.2 V) has a better reversibility and stability than that of cyt c (2) (0.18 V) (Figure 3B). The electric couple of 0.2 V shifted to 0.23 V (HD cyt c (2) (0.23 V)) with ΔE of 0.046 V after several potential scans instead of disappearance (Figure 3F, green). Since high dilution will decrease the adsorption rate of both species, HD cyt c (2) (0.2 V) have enough time and space to effectively interact with MUA, leading to a more stable adsorption and an alternative rearrangement of interfacial water molecules, and thus a better electrochemical reversibility and stability. The shift of Ef from 0.18 V (Figure 3B, black) to 0.2 V (Figure 3F, black) might suggest an adsorption-induced further unfolding of the largely unfolded cyt c (2). The shift of Ef from 0.2 to 0.23 V (Figure 3F), the same Ef of cyt c (1), after several potential scans might suggest the electric field-induced further unfolding in the conformation of HD cyt c (2) (0.2 V). Interestingly, when the corresponding measurements were carried out at physiological conditions or using mitochondria-targeted carbon nanotubes (CNTs), the similar adsorption and electrochemistry phenomena were also observed in Figures S3 and S4. Thus, proteins desorbed from a nanomaterial after a weak interaction maintain altered conformation and redox properties over the time course of the experiments could be a universal effect. Figure 4 shows the normalized redox-induced SEIRA difference spectra of native cyt c, cyt c (2), and HD cyt c (2) adsorbed on the MUA SAMs in the region of 4000 to 1620 cm−1. For native or native-like cyt c, the background spectrum was taken at −0.1 V and the sample spectrum was measured at

Figure 4. Redox-induced SEIRA difference spectroscopy study of the conformation and hydration degree of cyt c in various cyt c-adsorbed films. The redox-induced SEIRA difference spectra of native cyt c/ MUA (black) and cyt c (2) (0.02 V)/MUA (red) at the sample potential of 0.1 V with the reference potential of −0.1 V, for cyt c (2) (0.18 V)/MUA (green) and HD cyt c (2) (0.2 V)/MUA (blue) at 0.4 V with the reference of 0.05 V, in the normalized size according to the intensity of 1693(−) cm−1. For comparison, the difference spectra of the MUA SAM between at −0.1 and 0.1 V (cyan) and between at 0.4 and 0.05 V (magenta) are present.

+0.1 V. For cyt c (2) (0.18 V) or HD cyt c (2) (0.2 V), the background spectrum was taken at 0.05 V and the sample spectrum was measured at 0.4 V. In the difference spectrum of native cyt c (Figure 4, black), the bands at 1693(−) and 1674(+) cm−1 are assigned to the type III β-turns of the reduced and oxidized states, respectively, while the bands at 1667(−) and 1660(+) cm−1 are assigned to the type II β-turns or α-helix of the reduced and oxidized states, respectively.25 The native-like cyt c (2) (0.02 V) (Figure 4, red) shows similar peak shape and position in this region in addition to 1 cm−1 hypsochromic shift for the oxidized β-turn II or α-helix, suggesting a very small change in protein conformation. In the case of cyt c (2) (0.18 V) (Figure 4, green), it is notable that significant changes occur in the peak shape and position for both the reduced and oxidized β-turn III besides 1 cm−1 hypsochromic shifts for both the oxidized and reduced β-turn II or α-helix, suggesting a large degree of unfolding in β-turn III structure. For the HD cyt c (2) (0.2 V) (Figure 4, blue), the difference spectrum of β-turn II or α-helix is still distinguishable, but the difference spectrum of β-turn III almost disappears, suggesting a further unfolding of these secondary structures possibly due to its effective interaction with MUA under the highly diluted condition. Thus, from the spectral point of view, it is confirmed that cyt c structure is heterogeneous after interacting with a small amount of nanoTiO2 at the molecular level. In native cyt c, the type III β-turns contain the residues 14−17 and 67−70. Since the residues 13, 72, and 86 are known as the electrostatic interaction sites of cyt c with MUA,25 the large degree of unfolding in the β-turn III structure could directly interfere with the domain structure surrounding 13 and/or 72, thus reducing the electrostatic interaction or changing the interaction mode. On the other hand, the large disturbance to the β-turn III could affect the interaction between heme and Tyr67, and thus interfere with the pocket structure of the heme and its axial ligand species since the Tyr67 OH is H-bonded to the S atom of the Met80 heme ligand,39,40 which might be one of the reasons causing the positive shift of Ef. In the case of cyt c (1)/MUA, the difference spectrum is indistinguishable in the amide band. Since the cyt c 2728

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730

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Analytical Chemistry

Figure 5. Comparison of the CVs in 10 mM PBS (pH 7.0) for the various cyt c-adsorbed films: (A) complex (1)-3/MUA (black) and cyt c (1)/ MUA (red) and (B) complex (2)-3/MUA (black) and HD cyt c (2)/MUA (red).

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CONCLUSION In summary, using SEIRAS combined with electrochemical cyclic voltammetry, we can identify heterogeneous cyt c species after weak interaction with and then disengagement from nanoTiO2 including native-like cyt c and largely unfolded cyt c (Scheme 1). The large disturbance to the structure of type III

(1) and the HD cyt c (2) (0.23 V) have the same Ef, we consider that the former has a similar conformation to the latter. In the difference spectrum of native cyt c/MUA, it is also observed a redox-induced negative water peak at ∼3300 cm−1 (Figure 4, black), while it is hardly resolved for the MUA SAM under the same switch of electric field (Figure 4, cyan), suggesting a decrease in the hydration degree of cyt c from the reduced to oxidized states. Notably, the intensity of this negative water peak is almost the same as those of native-like cyt c (Figure 4, red) and HD cyt c (2) (0.2 V) (Figure 4, blue) but much smaller than that of cyt c (2) (0.18 V) (Figure 4, green), suggesting a larger change in the hydration degree of the latter cyt c. From the energy point of view, the redoxinduced rearrangement of water molecules is a major determinant of the total reorganization free energy (λ) for cyt c in its redox process.41 Its increase in degree will inevitably enlarge the λ of the redox process, causing the increased ΔE and the irreversibility. Thus, a reduced change in the redoxinduced water rearrangement resulted in a relatively reversible redox process for native cyt c, native-like cyt c and HD cyt c (2) (0.2 V). The interaction of nano-TiO2 cyt c corona with MUA was also studied by electrochemical cyclic voltammetry. The nanoTiO2/cyt c complexes prepared with sample 1 and sample 2 were first washed with 10 mM PBS (pH 7.0) for 3 times, named complex (1)-3 and complex (2)-3, respectively. Then, the complex (1)-3 and complex (2)-3 were incubated with the MUA SAMs in 10 mM PBS (pH 7.0) to obtain complex (1)-3/ MUA and complex (2)-3/MUA, respectively. After that, the obtained complex (1)-3/MUA underwent potential scanning in a renewed 10 mM PBS (pH 7.0) and exhibits a pair of redox peaks at Ef of 0.23 V with ΔE of 0.06 V (Figure 5A), consistent with that of the electric couple of the cyt c (1)/MUA (Figure 3A). The complex (2)-3/MUA also shows two pairs of redox peaks: one pair at Ef of 0.2 V with ΔE of 0.055 V and the other pair at Ef of 0.01 V with ΔE of 0.05 V (Figure 5B), consistent with those of two electric couples of the HD cyt c (2)/MUA (Figure 3B). The consistence suggests that the soft corona protein could interact with its downstream partner after being desorbed from the complex, and the disturbance to protein conformation following the weak interaction is irreversible on the time scale of these experiments and under the experimental conditions used.

Scheme 1. Adsorption of Heterogeneous Cyt c with Different Degree of Unfolding after Weak Interaction with Nano-TiO2 on MUA SAMs

β-turns results in weak but rapid adsorption of the cyt c on MUA SAMs at low concentration even in the presence of large amount of native-like cyt c. After forming a stable adsorption, such cyt c exhibits a positively shifted formal potential, which could favor an accumulation of the obtained cyt c (Fe2+), causing the disruption of electron flow and the exclusion of the cyt c from electron transport chain. This study points out a possible impact for the desorbed proteins suffering from conformational change due to their weak adsorption onto nanoparticles on their interactions with biological components. It is of great significance to the study of nanobio interface. 2729

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730

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Analytical Chemistry

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(14) Monari, S.; Ranieri, A.; Bortolotti, C. A.; Peressini, S.; Tavagnacco, C.; Borsari, M. Electrochim. Acta 2011, 56, 6925−6931. (15) Tong, T. Z.; Wilke, C. M.; Wu, J. S.; Binh, C. T. T.; Kelly, J. J.; Gaillard, J. F.; Gray, K. A. Environ. Sci. Technol. 2015, 49, 8113−8123. (16) Zhao, X.; Sheng, L.; Wang, L.; Hong, J.; Yu, X.; Sang, X.; Sun, Q.; Ze, Y.; Hong, F. Part. Fibre Toxicol. 2014, 11, 47. (17) Sayes, C. M.; Wahi, R.; Kurian, P. A.; Liu, Y.; West, J. L.; Ausman, K. D.; Warheit, D. B.; Colvin, V. L. Toxicol. Sci. 2006, 92, 174−185. (18) Allouni, Z. E.; Høl, P. J.; Cauqui, M. A.; Gjerdet, N. R.; Cimpan, M. R. Toxicol. In Vitro 2012, 26, 469−479. (19) Chen, X.; Cheng, B.; Yang, Y.; Cao, A.; Liu, J.; Du, L.; Liu, Y.; Zhao, Y.; Wang, H. Small 2013, 9, 1765−1774. (20) Zhang, X.; Li, W.; Yang, Z. Arch. Toxicol. 2015, 89, 2207−2217. (21) Fedurco, M.; Augustynski, J.; Indiani, C.; Smulevich, G.; Antalik, M.; Bánó, M.; Sedlák, E.; Glascock, M. C.; Dawson, J. H. J. Am. Chem. Soc. 2005, 127, 7638−7646. (22) Joo, J.; Uchida, T.; Cuesta, A.; Koper, M. T. M.; Osawa, M. Electrochim. Acta 2014, 129, 127−136. (23) Wu, L.; Zeng, L.; Jiang, X. J. Am. Chem. Soc. 2015, 137, 10052− 10555. (24) Liu, L.; Zeng, L.; Wu, L.; Jiang, X. J. Phys. Chem. C 2015, 119, 3990−3999. (25) Ataka, K.; Heberle, J. J. Am. Chem. Soc. 2004, 126, 9445−9457. (26) Lin, S.; Jiang, X.; Wang, L.; Li, G.; Guo, L. J. Phys. Chem. C 2012, 116, 637−642. (27) Liu, L.; Wu, L.; Zeng, L.; Jiang, X. Chinese Physics B 2015, 24, 12801. (28) Liu, L.; Wang, N.; Guo, L. RSC Adv. 2012, 2, 2809−2814. (29) Margoliash, E.; Frohwirt, N. Biochem. J. 1959, 71, 570−578. (30) Miyake, H.; Ye, S.; Osawa, M. Electrochem. Commun. 2002, 4, 973−977. (31) Jiang, X.; Ataka, K.; Heberle, J. J. Phys. Chem. C 2008, 112, 813− 819. (32) Nickel, C.; Angelstorf, J.; Bienert, R.; Burkart, C.; Gabsch, S.; Giebner, S.; Haase, A.; Hellack, B.; Hollert, H.; Hund-Rinke, K.; et al. J. Nanopart. Res. 2014, 16, 2260. (33) Kelly, S. M.; Price, N. C. Curr. Protein Pept. Sci. 2000, 1, 349− 384. (34) Myer, Y. P. J. Biol. Chem. 1968, 243, 2115−2122. (35) Pinheiro, T. J. T.; Elöve, G. A.; Watts, A.; Roder, H. Biochemistry 1997, 36, 13122−13132. (36) Pan, H.; Qin, M.; Meng, W.; Cao, Y.; Wang, W. Langmuir 2012, 28, 12779−12787. (37) Kamal, J. K. A.; Zhao, L.; Zewail, A. H. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 13411−13416. (38) Zhang, L.; Yang, Y.; Kao, Y.; Wang, L.; Zhong, D. J. Am. Chem. Soc. 2009, 131, 10677−10691. (39) Ying, T. L.; Wang, Z. H.; Lin, Y. W.; Xie, J.; Tan, X. S.; Huang, Z. X. Chem. Commun. 2009, 4512−4514. (40) Casalini, S.; Battistuzzi, G.; Borsari, M.; Bortolotti, C. A.; Di Rocco, G.; Ranieri, A.; Sola, M. J. Phys. Chem. B 2010, 114, 1698− 1706. (41) Paltrinieri, L.; Borsari, M.; Ranieri, A.; Battistuzzi, G.; Corni, S.; Bortolotti, C. A. J. Phys. Chem. Lett. 2013, 4, 710−715.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.6b01964. Additional information about nitrogen adsorption− desorption isotherm of nano-TiO2, far-UV CD detection of conformational changes of the cyt-c-treated by nanoTiO2 at physiological condition, adsorption, and electrochemistry of the cyt-c-treated by nano-TiO 2 at physiological condition and treated by mitochondriatargeted CNTs-COOH (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Phone: +86 431 85262426. Fax: +86 431 85685653. E-mail: [email protected]. ORCID

Xiue Jiang: 0000-0002-0194-1553 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was financially supported by the National Science Foundation for Excellent Young Scholar of China (21322510), the National Natural Science Foundation of China (21675149), the Key Research Program of Frontier Sciences, CAS (QYZDY-SSW-SLH019), and the Science and Technology Innovation Foundation of Jilin Province for Talents Cultivation (Grants 20150519014JH, 20140520082JH).

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REFERENCES

(1) Miclãus, T.; Bochenkov, V. E.; Ogaki, R.; Howard, K. A.; Šutherland, D. S. Nano Lett. 2014, 14, 2086−2093. (2) Mahmoudi, M.; Lynch, I.; Ejtehadi, M. R.; Monopoli, M. P.; Bombelli, F. B.; Laurent, S. Chem. Rev. 2011, 111, 5610−5637. (3) Monopoli, M. P.; Aberg, C.; Salvati, A.; Dawson, K. A. Nat. Nanotechnol. 2012, 7, 779−786. (4) Cedervall, T.; Lynch, I.; Lindman, S.; Berggård, T.; Thulin, E.; Nilsson, H.; Dawson, K. A.; Linse, S. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 2050−2055. (5) Röcker, C.; Pötzl, M.; Zhang, F.; Parak, W. J.; Nienhaus, U. Nat. Nanotechnol. 2009, 4, 577−580. (6) Ge, C.; Du, J.; Zhao, L.; Wang, L.; Liu, Y.; Li, D.; Yang, Y.; Zhou, R.; Zhao, Y.; Chai, Z.; Chen, C. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 16968−16973. (7) Deng, Z. J.; Liang, M.; Monteiro, M.; Toth, I.; Minchin, R. F. Nat. Nanotechnol. 2011, 6, 39−44. (8) Tenzer, S.; Docter, D.; Kuharev, J.; Musyanovych, A.; Fetz, V.; Hecht, R.; Schlenk, F.; Fischer, D.; Kiouptsi, K.; Reinhardt, C.; Landfester, K.; Schild, H.; Maskos, M.; Knauer, S. K.; Stauber, R. H. Nat. Nanotechnol. 2013, 8, 772−781. (9) Walczyk, D.; Bombelli, F. B.; Monopoli, M. P.; Lynch, I.; Dawson, K. A. J. Am. Chem. Soc. 2010, 132, 5761−5768. (10) Kelly, P. M.; Åberg, C.; Polo, E.; O’Connell, A.; Cookman, J.; Fallon, J.; Krpetić, Ž .; Dawson, K. A. Nat. Nanotechnol. 2015, 10, 472− 479. (11) Kagan, V. E.; Borisenko, G. G.; Tyurina, Y. Y.; Tyurin, V. A.; Jiang, J.; Potapovich, A. I.; Kini, V.; Amoscato, A. A.; Fujii, Y. Free Radical Biol. Med. 2004, 37, 1963−1985. (12) Skulachev, V. P. FEBS Lett. 1998, 423, 275−280. (13) Suárez, G.; Santschi, C.; Martin, O. J. F.; Slaveykova, V. I. Biosens. Bioelectron. 2013, 42, 385−390. 2730

DOI: 10.1021/acs.analchem.6b01964 Anal. Chem. 2017, 89, 2724−2730