Catalytic performance and molecular dynamic simulation of immobilized CC bond hydrolase based on carbon nanotube matrix

Colloids and Surfaces B: Biointerfaces 116 (2014) 365–371

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Catalytic performance and molecular dynamic simulation of immobilized C C bond hydrolase based on carbon nanotube matrix Hao Zhou, Yuanyuan Qu ∗ , Chunlei Kong, Duanxing Li, E. Shen, Qiao Ma, Xuwang Zhang, Jingwei Wang, Jiti Zhou State Key Laboratory of Industrial Ecology and Environmental Engineering (MOE), School of Environmental Science and Technology, Dalian University of Technology, Dalian 116024, China

a r t i c l e

i n f o

Article history: Received 16 October 2013 Received in revised form 31 December 2013 Accepted 13 January 2014 Available online 23 January 2014 Keywords: Carbon nanotube Immobilization C C bond hydrolase Interaction mechanism Molecular dynamics

a b s t r a c t Carbon nanotube (CNT) has been proved to be a kind of novel support for enzyme immobilization. In this study, we tried to find the relationship between conformation and catalytic performance of immobilized enzyme. Two C C bond hydrolases BphD and MfphA were immobilized on CNTs (SWCNT and MWCNT) via physical adsorption and covalent attachment. Among the conjugates, the immobilized BphD on chemically functionalized SWCNT (BphD-CSWCNT) retained the highest catalytic efficiency (kcat /Km value) compared to free BphD (92.9%). On the other hand, when MfphA bound to pristine SWCNT (MfphASWCNT), it was completely inactive. Time-resolved fluorescence spectrum indicated the formation of static ground complexes during the immobilization processes. Circular dichroism (CD) showed that the secondary structures of immobilized enzymes changed in varying degrees. In order to investigate the inhibition mechanism of MfphA by SWCNT, molecular dynamics simulation was employed to analyze the adsorption process, binding sites and time evolution of substrate tunnels. The results showed that the preferred binding sites (Trp201 and Met81) of MfphA for SWCNT blocked the main substrate access tunnel, thus making the enzyme inactive. The “tunnel-block” should be a novel possible inhibition mechanism for enzyme-nanotube conjugate. Crown Copyright © 2014 Published by Elsevier B.V. All rights reserved.

1. Introduction Enzymes are considered as attractive catalysts for various chemical reactions in mild conditions. However, the use of enzymes in industrial application are limited by several factors, mainly the high cost, low reusability and easy deactivation in harsh conditions [1,2]. Immobilization technology has been proved to be the most popular and efficient approach in lengthening the life time of an enzyme [3–5]. Recently, the interest in nanotechnology has provided a wealth of nanoscaffolds to support enzyme

Abbreviations: AFM, atomic force microscopy; CD, circular dichroism; CNT, carbon nanotube; DMF, dimethylformamide; EDC, N-ethyl-N -(3-(dimethyllamino) propyl) carbodiimide hydrochloride; MD, molecular dynamics; MES, 2-(Nmorpholino) ethanesulfonic acid; MWCNT, multi wall carbon nanotube; NHS, N-hydroxysuccinimide; PNPA, p-nitrophenyl acetate; PNPBen, p-nitrophenyl benzoate; SASA, solvent accessible surface area; SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis; SWCNT, single wall carbon nanotube; TEM, transmission electron microscopy; XPS, X-ray photoelectron spectroscopy. ∗ Corresponding author at: School of Environmental Science and Technology, Dalian University of Technology, Dalian 116024, China. Tel.: +86 411 8470 6251; fax: +86 411 8470 6252. E-mail address: [email protected] (Y. Qu). http://dx.doi.org/10.1016/j.colsurfb.2014.01.018 0927-7765/Crown Copyright © 2014 Published by Elsevier B.V. All rights reserved.

immobilization. The common used nanomaterials include metal oxides (ZnO, Fe3 O4 ), mesoporous silicas (SBA-15), carbon nanotube and graphene oxide [6–10]. The nanoscale structures of these materials can reduce diffusion limitation and maximize the surface area to increase enzyme loading, which overcomes the disadvantages of traditional immobilization matrix [11]. Among various types of nanomaterials, carbon nanotubes (CNTs) possess unique thermal, mechanical and biocompatibility characteristics, which makes it a promising material for supporting enzymes [12,13]. Recently, attachment of proteins on CNTs has been well-studied, and a variety of covalent and adsorption methods have been explored to immobilize proteins onto pristine or functional CNTs [14,15]. Asuri and co-workers reported that covalently attached subtilisin Carlsberg onto oxidized multi-walled carbon nanotubes (MWCNTs) loaded larger amount of protein molecules than that onto oxidized single-walled carbon nanotubes (SWCNTs), while the relative activities had no obvious difference [16,17]. However, there were limited studies comparing the effects of different immobilization methods and types of CNTs on the catalytic performances of CNT-enzyme conjugates. Therefore, the relationship among the catalytic performance of immobilized enzyme, the type of CNTs and immobilization methods need to be further investigated.

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As a novel type of cofactor-free biocatalysts, C C bond hydrolases catalyze a range of reactions including reverse Claisen condensation of 1,3-diketones, hydrolytic cleavage vinylogous 1, 5-diketone and reverse Friedel–Crafts acylation [18,19]. In addition, meta-cleavage product hydrolase MhpC, a member of C C bond hydrolases, could catalyze C C bond formations in organic media, which are among the most important fundamental reactions in organic synthesis [20]. For these reasons, C C bond hydrolases may have wide applications in biocatalytic processes. Surprisingly, there were only two cases reported by our group concerning the immobilization of C C bond hydrolases [21,22]. We firstly immobilized MfphA into mesoporous silica SBA-15 with relatively low loading amount (about 35 mg g−1 ) [21]. Afterwards, another C C bond hydrolase BphD was immobilized onto SWCNT with higher loading amounts, suggesting that CNTs should be a more suitable matrix for C C bond hydrolase immobilization [22]. Therefore, further studies should be performed to compare the catalytic performance of these two immobilized C C bond hydrolases based on different CNTs. On the other hand, in order to design a highly efficient immobilized enzyme, it is important to investigate the dynamics and thermodynamics properties of bio-nano systems, especially the interaction between nanoparticles and the binding site/function units of proteins [23]. Various spectroscopic technologies, such as steady-state and time-resolved fluorescence spectroscopy, circular dichroism (CD), Raman spectroscopy and UV–vis can provide useful information on the conformation change of protein and the driving forces for protein adsorption [24,25]. Besides these experimental methodologies, molecular simulation technologies, such as docking and molecular dynamics, have been used to identify the atomic details of possible interaction sites, binding energy and interaction mechanisms [26–30]. In the present study, we investigated the catalytic performance of different C C bond hydrolase-CNT conjugates. Then, steady-state/time-resolved fluorescence spectroscopy and CD were employed to reveal the binding mechanism and secondary structures of enzymes. Moreover, molecular dynamics simulation of SWCNT-MfphA was carried out in order to find the possible inhibition mechanism of SWCNT. This study should help us to improve understanding of the interactions between enzymes and CNTs.

was removed. Residue was washed by phosphate buffer 3 times to ensure complete removal of free enzyme. The amount of enzyme loaded onto the CNTs was determined by the difference between the concentration of enzyme in solution before and after adsorption using Bradford method [34]. 2.3. Covalent immobilization of enzymes onto CNTs The procedure of enzymes covalently attached to nanotubes was modified based on our previous report [22]. To add the carboxyl group into carbon nanotube, 200 mg CNTs were suspended in 400 ml mixture of H2 SO4 and HNO3 (3:1, v/v) and sonicated for 8 h. The suspension was diluted with ultrapure water and washed by filtering through 0.22 ␮m polycarbonate membrane repeatedly. The nanotube film on polycarbonate membrane was transformed into MilliQ water and sonicated again to remove residual acid. Then the acid treated CNTs were dispersed in 2-(N-morpholino) ethanesulfonic acid (MES) buffer (pH 6.2) and mixed with an equal volume of 400 mM NHS. The mixture was sonicated for 30 min followed by addition of 20 mM EDC to initiate the coupling of NHS to the carboxylic groups and the mixture was stirred at 150 rpm for 1 h. Finally, they were dried in a vacuum to obtain soluble powder and were characterized by X-ray photoelectron spectroscopy (XPS). Samples were analyzed using an ESCALB 250Xi X-ray photoelectron spectrometer (Thermo Scientific, USA) with a monochromatic Al K␣ X-ray source. Spectral analysis included a background subtraction and peak separation was performed using a least-squares curve-fitting program XPSpeak 4.1 (http://www.uksaf.org/software.html). Then, 1 ml functionalized >CNTs dispersion (1 mg/ml in phosphate buffer) was mixed with 5 ml enzymes solution (120 ␮g/ml), which was shaken at room temperature for 8 h at 150 rpm. All the other procedures were the same with those of physical adsorption. Both enzyme-loaded CNTs were washed and resuspended using phosphate buffer, and then subjected to SDS-PAGE in order to ensure the successful immobilization of the protein. 2.4. Atomic force and transmission electron microscopy

2. Experimental 2.1. Chemicals and m aterials SWCNT (purity 95%, length 5–15 ␮m, diameter < 2 nm) and MWCNT (purity 97%, length 5–15 ␮m, diameter < 10 nm) were purchased from Shenzhen Nanotech Port Co., Ltd. (Shenzhen, China). N-hydroxysuccinimide (NHS) and N-ethyl-N’-(3-(dimethyllamino) propyl) carbodiimide hydrochloride (EDC) were purchased from Sinopharm (Shanghai, China). All the other chemicals in this work were of analytical grade and used without further purification. Ultrapure water (18.2 M cm−1 resistivity) was used for preparation of all solutions and dispersions. Two C C bond hydrolases MfphA and BphD were overexpressed in recombinant Escherichia coli and purified according to our previous work [31,32]. 2.2. Physical adsorption of enzymes onto CNTs Adsorption of enzyme onto CNTs was performed using reported protocol [33]. In general, CNTs were sonicated in dimethylformamide (DMF) for 30 min and then DMF was removed gradually by repeated washing with 200 mM phosphate buffer (pH 7.0). 1 ml CNTs dispersion in the buffer (1 mg/ml) was mixed with 5 ml enzymes solution (120 ␮g/ml), and the mixture was shaken at room temperature for 8 h at 150 rpm. The CNT-enzyme suspensions were centrifuged at 10,000 rpm and the supernatant

Atomic force microscopy (AFM) images for MfphA-SWCNT and MfphA-CSWCNT were obtained using PicoPlus II (Agilent, USA) in the tapping mode. A dilute suspension of the CNT-enzyme composite was prepared in phosphate buffer (200 mM, pH 7.0), and 30 ␮l of samples were dropped on a freshly cleaved mica surface. Transmission electron microscopy (TEM) images for CNTs and CNT-enzyme conjugates were captured using a FEI Tecnai T12 transmission electron microscope operated at 120 kV. 2.5. Enzymatic assay procedure The esterase activities of MfphA and BphD were measured using p-nitrophenyl acetate (PNPA) and p-nitrophenyl benzoate (PNPBen) as substrate, respectively. For a typical activity assay, 40 ␮M stock solution of substrate was added into sodium phosphate buffer (200 mM, pH 7.0), and the reaction was initiated by adding free or immobilized enzyme. The reaction rate was measured at 405 nm using spectrophotometer (JASCO V-560, Japan). An appropriate control with a heat-inactivated enzyme (incubated at 100 ◦ C for 10 min) was set for each assay. Measured reaction rates were verified to be linear at least for 2 min. For kinetic assays, appropriate equations were fitted to the initial velocities determined at different substrate concentrations (from 2 to 100 ␮M) using the non-linear regression function of Origin 8.0.

H. Zhou et al. / Colloids and Surfaces B: Biointerfaces 116 (2014) 365–371

2.6. Fluorescence measurement For the fluorescence measurement, carbon nanotube suspensions were added into the protein solutions in sequence. Both the concentrations of carbon nanotube and proteins were calculated by weight for consistency. The concentrations of carbon nanotube were adjusted to 0, 5, 10, 20 and 40 ␮g/ml, respectively. The concentration of protein was 40 ␮g/ml. The systems were excited at 280 nm, and the emission wavelength was adjusted from 285 to 500 nm with a scanning speed of 2400 nm/min. Excitation and emission slit widths were both set to 500 V. Fluorescence lifetime measurements were performed with a Horiba Jobin Yvon FluoroMax-4 TCSPC (Horiba Jobin Yvon, France) using the time correlated single photon counting method. Each fluorescence decay profile was fitted using two and three exponential function to find the best fit. Excitation was performed at 295 nm with a nano-LED. Enzymes were fixed at a concentration of 20 or 40 ␮g/ml. 2.7. Circular dichroism spectroscopy The secondary structures of free C C bond hydrolases and the CNT-enzyme conjugates were monitored by CD spectroscopy (JASCO J-810, Japan). CD measurements were operated over the range 200–260 nm at a scan rate of 500 nm/min. The spectra were measured in a temperature-controlled 1 mm path length cell, and each spectrum was the average of six scans. All CD measurements were performed at 25 ◦ C in 5 mM Tris–HCl (pH 8.0) buffer. The protein concentrations of free and immobilized enzymes were 150 ␮g/ml. The contents of ␣-helix were estimated using Eq. (1): %␣ − helix =

222 − 3000 −390

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3.0 was used for the calculation of tunnels during the 15 ns MD simulation of SWCNT-MfphA conjugate [37]. Totally 300 snapshots in an interval of 50 ps were extracted from the overall MD trajectory and used to analyze the time evolution of tunnels. Tunnel occupy percentage was defined as the snapshots numbers ratio of containing tunnel and total. Visual analyses of tunnels were carried out using PyMOL 0.99 and R. 3. Results and discussion 3.1. Characterization of functionalized CNTs by XPS In order to immobilize the protein covalently, the CNTs needed to be functionalized by carboxylation firstly. XPS was employed to give the quantitative information on the elemental composition of the oxidized SWCNT and MWCNT. As shown in Fig. SI 1, after treated with HNO3 /H2 SO4 , the total oxygen amount of SWCNT increased from 1.68% to 3.96%. Meanwhile, the total oxygen amount of MWCNT increased from 1.05% to 6.91%. These results indicated CNTs had been oxidized. Furthermore, the C1s and O1s spectra of samples were analyzed. Three main peaks were extracted from the C1s spectra. The peaks centered at 284.5 and 285.6 eV were due to sp2 C C bonds of graphitic carbon and defects of CNTs surfaces. Also, the peak centered at 289.2 eV was contributed by carboxyl groups of oxidized CNTs [38]. There was a main peak around 532 eV in the O1s spectra for all the samples, which also suggested the existence of carboxyl or carbonyl group [39]. The signal for carboxyl group in pristine CNTs might be due to the impurity introduced during chemical vapor deposition. From the results of XPS, it could be ensured that large amount of carboxyl groups have been successfully added to the surfaces of CNTs.

(1)

where  222 is the molar ellipticity of the protein at 222 nm [35]. 2.8. Computational methods The structure of zigzag SWCNT with diameter of 1.0 nm was generated by TubeGen3.4 (http://turin.nss.udel.edu/research/ tubegenonline.html). The structure of MfphA was constructed through homology modeling according to the previous work [31]. Molecular dynamics (MD) simulation was performed with GROMACS 4.5.5 using g53a6 force field [36]. The initial separation of the geometric center of the SWCNT and MfphA was set as 3.5 nm, and the minimum distance between SWCNT and MfphA was 0.8 nm. The SWCNT-MfphA conjugate was solvated with SPC water, and chlorine ions were added by replacing the water molecules to neutralize the system charge. Protonation states of titratable groups Arg, Lys, His, Asp and Glu were estimated using online server PDB2PQR at pH 7.0 (http://kryptonite.nbcr.net/pdb2pqr 1.8/). The systems were subjected to steepest descent and conjugate gradient energy minimization. The 15 ns NPT (isothermal–isobaric ensemble) MD simulation was performed with a time step of 2 fs. CAVER

3.2. Protein loading capacity and morphology of the conjugates After preparing the carboxylation CNTs, two C C bond hydrolases were absorbed or covalently immobilized onto the CNTs. Totally, eight conjugates were obtained through immobilization. There were obvious difference on both loading amounts and catalytic efficiencies of the conjugates. For both SWCNT and MWCNT, the loading amount of protein through covalent attachment was higher than that by physical adsorption. The maximum loading amounts for BphD and MfphA were 508 and 425 mg g−1 CNT, respectively (Table 1). It was noticed that the lowest loading amount of MfphA-CNTs was 8-folds higher than that of MfphA-SBA-15 [21], which suggested that CNTs was more efficient alternative immobilization matrix for C C hydrolase. In order to prove that the protein had been immobilized on various CNTs, SDS-PAGE, TEM and AFM were performed. After washing 3 times by phosphate buffer, the conjugates were boiled to release protein from CNTs, and then the released protein was detected by SDS-PAGE (Fig. SI 2). It was showed that all the samples exhibited a single band, which suggested that the protein was successfully immobilized. TEM images of bare SWCNT and MfphA-SWCNT

Table 1 The apparent kinetic parameters and loading amounts of immobilized enzymes. Values were means ±standard deviations of results from three experiments. Conjugates

Loading amount (mg g−1 CNT)

Km (␮M)

Vmax (␮M s−1 )

Vmax /Km (s−1 )

Free MfphA MfphA-SWCNT MfphA-MWCNT MfphA-CSWCNT MfphA-CMWCNT Free BphD BphD-SWCNT BphD-MWCNT BphD-CSWCNT BphD-CMWCNT

– 301 ± 15 281 ± 22 391 ± 32 425 ± 27 – 276 ± 19 335 ± 31 508 ± 37 347 ± 24

15.0 ± 2.1 ND 15.4 ± 1.7 14.6 ± 2.9 12.6 ± 2.3 3.4 ± 0.7 52.2 ± 9.1 30.8 ± 7.4 2.7 ± 1.1 64.2 ± 11.4

4.5 ± 0.7 ND 0.10 ± 0.02 0.29 ± 0.09 0.14 ± 0.05 2.6 ± 0.4 0.93 ± 0.12 0.78 ± 0.21 1.9 ± 0.2 0.62 ± 0.17

0.30 ND 0.007 0.019 0.011 0.74 0.018 0.025 0.69 0.010

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conjugates showed that the nanotubes existed in MfphA-SWCNT became thicker and rougher than the bare one (Fig. SI 3). It was demonstrated that the protein molecules had been attached onto the CNTs, which was in agreement with SDS-PAGE results. Furthermore, clearer morphology was presented by AFM, which exhibited the “coarse-grained” three-dimensional structure for both MfphA-SWCNT and MfphA-CSWCNT (Fig. 1). The size of free MfphA (dimer) was about 7.6 nm × 4.1 nm × 4.2 nm according to our previously calculated results [21]. As shown in Fig. 1(B), the height of globular structure in SWCNT was about 30 nm, indicating the MfphA molecules might be aggregated when they were immobilized onto SWCNT. Fig. 1(C) showed two different globular structures in MfphA-CSWCNT. The heights of these structures were 6 nm and 19 nm (Fig. 1(D)). This result indicated both single MfphA and multiple MfphA molecules existed on CSWCNT. A dimension of 6 nm might suppose a slight unfolding happened when MfphA was covalently immobilized on CSWCNT. 3.3. Catalytic performance of conjugates Although the loading capacities of CNTs for both enzymes were similar, the catalytic activities were different. Among the MfphA-CNT conjugates, MfphA-SWCNT lost activity completely. The other three conjugates, i.e. MfphA-MWCNT, MfphA-CSWCNT and MfphA-CMWCNT retained less than 30% of specific activity relative to that of free enzyme. In contrast, the BphD-CNT conjugates were still highly active, especially BphD-CSWCNT, which has a comparable activity to free BphD (data not shown). Furthermore, catalytic kinetics of enzyme-CNT conjugates on ester substrates was performed to investigate the interaction between C C bond hydrolase and CNTs. All of the conjugates followed the conventional Michaelis–Menten kinetics when PNPA and PNPBen were used as substrates. The Km values of all the MfphA-CNT conjugates to p-nitrophenyl acetate were similar to that of the free MfphA (except for MfphA-SWCNT, which has no detectable activity), while the Vmax values dropped 15.8 to 43.5 folds compared with that of the free MfphA (Table 1). The reduction of Vmax might due

to the substrate’s inability to reach the active site, when MfphA was immobilized on CNTs. It meant that the inhibition mechanism of CNTs for MfphA was noncompetitive inhibition. However, the catalytic performance of BphD-CNTs conjugates was different from MfphA-CNTs conjugates. It was indicated that BphD-SWCNT, BphD-MWCNT and BphD-CMWCNT had much higher Km and lower Vmax values than free BphD. Whereas the Km and Vmax values of BphD-CSWCNT were similar with free BphD. The inhibition mechanism of CNTs for BphD had been supposed in our previous molecule docking studies. It indicated that CNTs could insert into the interface of adjacent subunits of BphD and disrupt their ␲–␲ stacking interactions [22]. 3.4. Spectrophotometric assays for enzyme-CNT conjugates It was reported that nanomaterial can change the secondary structure of enzyme or enter into the active pocket, thus the enzyme catalytic efficiency could be affected [33,40]. Spectroscopic methods, including fluorescence spectroscopy and CD, were used to explain the different catalytic performance of CNT-enzyme conjugates. In order to explore the CNT binding sites on C C bond hydrolases, we analyzed changes in intrinsic fluorescence spectra with increasing CNT concentrations (from 0 to 40 ␮g/ml). Fig. SI 4 showed the intrinsic fluorescence of all the samples were quenched in varying degrees, while the max of free MfphA (331 ± 3 nm) and BphD (330 ± 2 nm) did not change with CNTs addition. When max was around 330 nm, the dominant fluorophores were tryptophan residues which located in non-polar environments and possibly interacting with polar neighbors [41]. In general, quenching of protein intrinsic fluorescence could be caused by excited-state reaction, molecule rearrangement, energy transfer, ground state complex formation and collisional quenching [42]. The different mechanisms of quenching are usually classified as either dynamic quenching or static quenching, which can be distinguished by fluorescence lifetime measurements. As shown in Table 2, the fluorescence lifetime data of free C C bond hydrolases and enzymeCNT conjugates fitted well to the sum of a three exponential

Fig. 1. AFM images of MfphA-SWCNT conjugates. (A) MfphA-SWCNT conjugate. (B) The height change along the SWCNT (direction of the axis). (C) MfphA-CSWCNT conjugate. (D) The height change along the CSWCNT (direction of the axis).

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Fig. 2. CD spectra of C C bond hydrolase in the absence or presence of CNTs. (A) CD spectra of free BphD and BphD-CNT conjugates. a: Free BphD, b: BphD-SWCNT, c: BphD-MWCNT, d: BphD-CSWCNT, e: BphD-CMWCNT. (B) The content of ␣-helix in free BphD and BphD-CNT conjugates. Bars indicate the standard deviation from six scans. (C) CD spectra of free MfphA and MfphA-CNT conjugates. a: Free MfphA, b: MfphA-SWCNT, c: MfphA-MWCNT, d: MfphA-CSWCNT, e: MfphA-CMWCNT. (D) The contents of ␣-helix in free MfphA and MfphA-CNT conjugates. Bars indicate the standard deviation from six scans.

decay with a 2 value around 1.20. The intrinsic fluorescence decay profiles of MfphA at 334 nm were similar with different concentrations of SWCNT and CSWCNT. The decay times were 1.02 to 1.25 ns for  1 , 2.04 to 2.50 ns for  2 , and 4.08 to 5.01 ns for  3 . Meanwhile, those profiles of BphD and its CNT conjugates exhibited similar results. Since dynamic quenching mechanism was not expected to have a stable lifetime, the static quenching mechanism was supposed for the intrinsic fluorescence quenching of BphD and MfphA (i.e. a ground state surface complex was formed). This result was in agreement with the quenching mechanism of MWCNTs on the intrinsic fluorescence of bovine serum albumin [24]. Table 2 The fluorescence decay profiles for free C C bond hydrolases and enzyme-CNT conjugates. The concentrations of protein were 40 mg L−1 for MfphA-SWCNT2, MfphA-CSWCNT2, BphD-SWCNT2 and BphD-CSWCNT2. All the other samples were 20 mg L−1 . Conjugates

 1 (ns)

 2 (ns)

 3 (ns)

2

Free MfphA MfphA-SWCNT1 MfphA-SWCNT2 MfphA-CSWCNT1 MfphA-CSWCNT2 Free BphD BphD-SWCNT1 BphD-SCWNT2 BphD-CSWCNT1 BphD-CSWCNT2

1.02 1.07 1.02 1.25 1.02 1.54 1.58 1.55 1.62 1.54

2.04 2.15 2.05 2.50 2.04 3.08 3.11 3.09 3.34 3.09

4.08 4.30 4.12 5.01 4.08 6.18 6.32 6.19 6.43 6.18

1.22 1.15 1.17 1.26 1.20 1.22 1.24 1.19 1.25 1.17

The CD method was used to reveal the relationship between secondary structure and catalytic performance (Fig. 2). The CD spectra of both free BphD and MfphA exhibited two negative double humped peaks at 208 nm and 222 nm, which was characteristic of ␣-helical structure (Fig. 2(A and C)) [25]. The ␣-helix structure contents of free BphD and MfphA were about 45% and 43.5%, respectively. After the protein conjugated with different CNTs, the ␣-helix structure contents of all the immobilized BphD were generally comparable or even higher (in the case of BphD-CSWCNT) than that of free BphD (Fig. 2(B)). Therefore, high catalytic efficiency of BphD-CSWCNT may be attributed to the stability of ␣-helix structure. Compared with free MfphA, the immobilized MfphA lost 5% to 26% of ␣-helix structure, indicating the CNTs could induce partial change of MfphA conformation (Fig. 2(D)). However, there seemed no necessary relationship between protein secondary structure and catalytic efficiency. For example, MfphA-SWCNT was completely deactivated during immobilization process with slight change of ␣helix content. Zhang et al. reported that functionalized CNTs could inhibit the activity of ␣-chymotrypsin by specific binding without significant change of secondary structure [43], which could be a good interpretation for inactivation of MfphA studied in this paper. 3.5. Atomic details of the interaction between MfphA and SWCNT As mentioned above, among eight CNT-enzyme conjugates, MfphA-SWCNT exhibited no hydrolytic activity at all, in which

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Fig. 3. Molecular dynamics simulation of MfphA-SWCNT conjugate. (A) Timecourse curve (I) of minimum distance between SWCNT and backbone of MfphA, and timecourse curve (II) of solvent accessible surface area SASA (SASAcomplex -SASAprotein -SASASWCNT ). (B) Represent binding mode of MfphA and SWCNT between 10–15 ns (obtained through cluster analysis of Gromacs). (C) Time evolution of the secondary structure of the residues in MfphA-SWCNT. (D) Time-evolution of tunnels in MfphA. The active-site residues, together with Trp-201 and Met-81 were shown in sticks. The tunnels were clustered and two top-ranked tunnel clusters were shown. The presence/absence of tunnels in each snapshot was shown in bottom. If the tunnel exists in a snapshot, it appears as a white strip, otherwise it appears as a blue strip. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

␣-helix content of MfphA was only slightly decrease. Therefore, it could be concluded that MfphA inactivation should be resulted by the atomic details of the interaction between MfphA and SWCNT. As a powerful approach, molecular dynamic (MD) simulation can not only be used to identify the possible contact sites and secondary structure change, but also reveal the possible inhibition mechanism of SWCNT. Firstly, the absorption process of MfphA on the surface of SWCNT was simulated by MD method. The dynamics of minimum distance (between backbone atom of MfphA and SWCNT) with time was shown in Fig. 3(A) (curve I). During 0–15 ns simulation, the minimum distance firstly increased from 0.8 to 1.8 nm at 2 ns, and then gradually decreased and fluctuated around 0.4 nm. The increase of minimum distance within 2 ns could be explained that the protein molecule rotated and translated in order to find a favorable binding conformation with SWCNT. Meanwhile, the SASA (solvent accessible surface area, SASAcomplex -SASAprotein -SASASWCNT ) was plotted in Fig. 3(A) as a function of simulation time (curve

II). The SASA gradually decreased during the whole simulation timescale, demonstrating that the hydrophobic surface of MfphA and SWCNT overlapped during the adsorption process. This phenomenon indicated that hydrophobic interactions indeed played an important role in the immobilization process. Thus, the simulated system was ready for further analysis, including binding site, secondary structure and tunnels of MfphA. Fig. 3(B) showed the typical binding mode of carbon nanotube and MfphA (obtained by cluster analysis of conformations between 10–15 ns). There were two regions, i.e. residues 75–80 and 196–201, involved in the interaction with carbon nanotube (Fig. 3(B)). In general, four types of interactions controlled the binding of proteins to CNTs, which were the vdW interaction (especially ␲–␲ stack), hydrophobic interaction, amphiphilicity interaction and electrostatic interaction [44]. In the case of MfphA-SWCNT conjugate, Met-81, Ala-196 and Trp-201 were hydrophobic residues, which could bind with carbon nanotube through hydrophobic interactions. Meanwhile, aromatic residues Trp-201 and Tyr-79

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may provide ␲–␲ stacking interactions to stabilize the carbon nanotube-enzyme complex. Therefore, the main driving forces for binding SWCNT were hydrophobic interaction and ␲–␲ stacking. After analyzing the binding process and possible binding site, we investigated the possible inhibition mechanism. Fig. 3(C) showed that when MfphA interacted with SWCNT, its secondary structure was not significantly changed. The calculated percentage composition of ␣-helix was 37%, which was in accordance with CD result. Therefore, completely deactivation of MfphA in MfphA-SWCNT conjugate seemed not due to the secondary structure change. Meanwhile, active sites of MfphA were deeply buried into the hydrophobic core. The substrate-enzyme interaction should not to be disrupted by SWCNT, which was supported by the similar Km value of free MfphA and MfphA-CNTs conjugates. Therefore, the inhibition mechanism in MfphA-SWCNT would be also different with “poison mechanism” of CNT to WW domains [40]. As well known, the tunnels were spatial structures for transporting substrate to the buried active site of protein, which was the first condition for catalytic reaction [45]. However, it is common that protein conformation can change in water, which should determine the presence or absence of a specific tunnel. Therefore, it is necessary to find out the effects of immobilization on the tunnels. Fig. 3(D) showed two top-ranked tunnels in MfphA. In the first 2 ns, these two tunnels had lower occupy percentage than that in the following 13 ns, indicating interaction with SWCNT could affect the global conformation of MfphA. It showed that two of the binding residues, i.e. Trp-201 and Met-81, were located at the entrance of tunnel 2 and formed a “gate” to control the substrate diffusion and product release. Therefore, when MfphA was immobilized onto SWCNT, the main substrate access tunnel of MfphA could be blocked. This atomic interaction should be the reason for MfphA inactivation. Meanwhile, it also suggested that tunnel 2 was the main tunnel in MfphA. We thought physical/chemical characterization of different CNTs would significantly affect the possibility of blockage. For example, existence of carboxyl groups on the surface of CNTs would enhance the electrostatic interactions between enzyme and CNTs, thus modulates the orientation of enzyme. Therefore, the attachment sites of MfphA on CSWCNT might be different with that on SWCNT. 4. Conclusions A comprehensive study on CNT-based immobilized C C bond hydrolases was performed to compare the catalytic performance at different conditions (immobilized method, type of CNT and type of enzyme). The BphD-CSWCNT retained comparable catalytic efficiency with free BphD, while MfphA-SWCNT lost its activity completely. Enzymatic kinetic studies suggested the inhibition type for MfphA-CNT was noncompetive inhibition, while for BphD-CNT was a mixed inhibition. Steady and time-resolve fluorescence spectra indicated the formation of static ground complex during the immobilized process. Molecular dynamics simulation on MfphASWCNT gave information about the adsorption process, binding sites and secondary structure change. Moreover, a novel inhibition mechanism (blocking the “gate” of substrate tunnel) was proposed, which should give deep insight into the interaction between enzyme and nanomaterial.

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Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.colsurfb. 2014.01.018. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29]

[30] [31] [32] [33] [34] [35] [36]

[37]

[38] [39] [40]

Acknowledgements

[41]

The authors gratefully acknowledge the financial supports from the National Natural Science Foundation of China (Nos. 21176040; 51078054) and Program for New Century Excellent Talents in University (No. NCET-13-0077). We also want to acknowledge Dr. Qian Zhang of Dalian institute of chemical physics in assistance of fluorescence lifetime measurement.

[42] [43] [44] [45]

L. Cao, L. van Langen, R.A. Sheldon, Curr. Opin. Biotechnol. 14 (2003) 387. A. Liese, L. Hilterhaus, Chem. Soc. Rev. 42 (2013) 6236. M.L. Verma, C.J. Barrow, M. Puri, Appl. Microbiol. Biotechnol. 97 (2013) 23. R. DiCosimo, J. McAuliffe, A.J. Poulose, G. Bohlmann, Chem. Soc. Rev. 42 (2013) 6437. S. Cantone, V. Ferrario, L. Corici, C. Ebert, D. Fattor, P. Spizzo, L. Gardossi, Chem. Soc. Rev. 42 (2013) 6262. M.H. Liao, D.H. Chen, Biotechnol. Lett. 23 (2001) 1723. S. Chakraborti, T. Chatterjee, P. Joshi, A. Poddar, B. Bhattacharyya, S.P. Singh, V. Gupta, P. Chakrabarti, Langmuir 26 (2010) 3506. M. Hartmann, Chem. Mater. 17 (2005) 4577. M. Vinoba, M. Bhagiyalakshmi, S.K. Jeong, Y. Yoon, S.C. Nam, Colloids Surf. B: Biointerfaces 90 (2012) 91. J.L. Zhang, F. Zhang, H.J. Yang, X.L. Huang, H. Liu, J.Y. Zhang, S.W. Guo, Langmuir 26 (2010) 6083. S.A. Ansari, Q. Husain, Biotechnol. Adv. 30 (2012) 512. M.F. Cabral, L.F. Sgobbi, E.M. Kataoka, S.A.S. Machado, Colloids Surf. B: Biointerfaces 111 (2013) 30. W. Feng, P.J. Ji, Biotechnol. Adv. 29 (2011) 889. I.V. Pavlidis, T. Vorhaben, T. Tsoufis, P. Rudolf, U.T. Bornscheuer, D. Gournis, H. Stamatis, Bioresour. Technol. 115 (2012) 164. H.Y. Zhou, Q.X. Mu, N.N. Gao, A.F. Liu, Y.H. Xing, S.L. Gao, Q. Zhang, G.B. Qu, Y.Y. Chen, G. Liu, B. Zhang, B. Yan, Nano Lett. 8 (2008) 859. P. Asuri, S.S. Karajanagi, E. Sellitto, D.Y. Kim, R.S. Kane, J.S. Dordick, Biotechnol. Bioeng. 95 (2006) 804. P. Asuri, S.S. Bale, R.C. Pangule, D.A. Shah, R.S. Kane, J.S. Dordick, Langmuir 23 (2007) 12318. A.C. Ruzzini, G.P. Horsman, L.D. Eltis, Biochemistry 51 (2012) 5831. E. Siirola, A. Frank, G. Grogan, W. Kroutil, Adv. Synth. Catal. 355 (2013) 1677. C. Li, M. Hassler, T.D.H. Bugg, ChemBioChem 9 (2008) 71. H. Zhou, Y.Y. Qu, Y.B. Bu, X.L. Li, C.L. Kong, Q. Ma, Q. Zhang, X.W. Zhang, J.T. Zhou, ChemPlusChem 77 (2012) 293. E. Shen, Y.Y. Qu, H. Zhou, C.L. Kong, Q. Ma, X.W. Zhang, J.T. Zhou, Chin. J. Catal. 34 (2013) 723. M. Mahmoudi, I. Lynch, M.R. Ejtehadi, M.P. Monopoli, F.B. Bombelli, S. Laurent, Chem. Rev. 111 (2011) 5610. X.C. Zhao, R.T. Liu, Z.X. Chi, Y. Teng, P.F. Qin, J. Phys. Chem. B 114 (2010) 5625. J.T. Cang-Rong, G. Pastorin, Nanotechnology 20 (2009) 255102. M. Calvaresi, S. Hoefinger, F. Zerbetto, Chem. Eur. J. 18 (2012) 4308. W. Feng, X.C. Sun, P.J. Ji, Soft Matter 8 (2012) 7143. J.W. Shen, T. Wu, Q. Wang, Y. Kang, Biomaterials 29 (2008) 3847. V.E. Kagan, N.V. Konduru, W.H. Feng, B.L. Allen, J. Conroy, Y. Volkov, I.I. Vlasova, N.A. Belikova, N. Yanamala, A. Kapralov, Y.Y. Tyurina, J.W. Shi, E.R. Kisin, A.R. Murray, J. Franks, D. Stolz, P.P. Gou, J. Klein-Seetharaman, B. Fadeel, A. Star, A.A. Shvedova, Nat. Nanotechnol. 5 (2010) 354. M.F. Cabral, J.D. Barrios, E.M. Kataoka, S.A.S. Machado, E. Carriho, C.D. Garcia, A.A. Ayon, Colloids Surf. B: Biointerfaces 103 (2013) 624. A. Li, Y.Y. Qu, J.T. Zhou, F. Ma, H. Zhou, S.N. Shi, Process Biochem. 45 (2010) 94. H. Zhou, Y.Y. Qu, C.L. Kong, Y.G. Wu, K. Zhu, J. Yang, J.T. Zhou, Biotechnol. Lett. 34 (2012) 1107. S.S. Karajanagi, A.A. Vertegel, R.S. Kane, J.S. Dordick, Langmuir 20 (2004) 11594. M.M. Bradford, Anal. Biochem. 72 (1976) 248. R.W. Woody, Methods Enzymol. 246 (1995) 34. S. Pronk, S. Páll, R. Schulz, P. Larsson, P. Bjelkmar, R. Apostolov, M.R. Shirts, J.C. Smith, P.M. Kasson, D. van der Spoel, B. Hess, E. Lindahl, Bioinformatics 29 (2013) 845. E. Chovancova, A. Pavelka, P. Benes, O. Strnad, J. Brezovsky, B. Kozlikova, A. Gora, V. Sustr, M. Klvana, P. Medek, L. Biedermannova, J. Sochor, J. Damborsky, PLoS Comput. Biol. 8 (2012) e1002708. T. Ramanathan, F.T. Fisher, R.S. Ruoff, L.C. Brinson, Chem. Mater. 17 (2005) 1290. J.P. Metters, M. Gomez-Mingot, J. Iniesta, R.O. Kadara, C.E. Banks, Sens. Actuat. B: Chem. 177 (2013) 1043. G.H. Zuo, Q. Huang, G.H. Wei, R.H. Zhou, H.P. Fang, ACS Nano 4 (2010) 7508. K.K. Khan, S. Mazumdar, S. Modi, M. Sutcliffe, G.C. Roberts, S. Mitra, Eur. J. Biochem. 244 (1997) 361. Y.J. Hu, Y. Liu, L.X. Zhang, R.M. Zhao, S.S. Qu, J. Mol. Struct. 750 (2005) 174. B. Zhang, Y.H. Xing, Z.W. Li, H.Y. Zhou, Q.X. Mu, B. Yan, Nano Lett. 9 (2009) 2280. M. Calvaresi, F. Zerbetto, Acc. Chem. Res. 46 (2013) 2454. L. Biedermannová, Z. Prokop, A. Gora, E. Chovancová, M. Kovács, J. Damborsky, R.C. Wade, J. Biol. Chem. 287 (2012) 29062.