Molecular Mechanisms of Interactions between Mono-layered

Subscriber access provided by UNIV OF SOUTHERN INDIANA

Article

Molecular Mechanisms of Interactions between Mono-layered Transition Metal Dichalcogenides and Biological Molecules Minyu Xiao, Shuai Wei, Junjie Chen, Jiayi Tian, Charles L. Brooks, E. Neil G. Marsh, and Zhan Chen J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.9b03641 • Publication Date (Web): 03 Jun 2019 Downloaded from http://pubs.acs.org on June 3, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of the American Chemical Society

Molecular Mechanisms of Interactions between Mono-layered Transition Metal Dichalcogenides and Biological Molecules Minyu Xiao1, Shuai Wei1, Junjie Chen1, Jiayi Tian1, Charles L. Brooks III,1,2 E. Neil G. Marsh1, Zhan Chen1* 1

Department of Chemistry, 2Department of Biophysics, University of Michigan, Ann Arbor,

Michigan, 48109, USA *

Corresponding author: Zhan Chen ([email protected])

1 ACS Paragon Plus Environment

Journal of the American Chemical Society 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract Single layered two-dimensional (2D) materials such as transition metal dichalcogenides (TMDs) showed great potential in many microelectronic or nanoelectronic applications. For example, because of its extremely high sensitivity, TMDs-based biosensor becomes a promising candidate for the next generation label-free detection. However, very few studies have been conducted on understanding the fundamental interactions between TMDs and other molecules including biological molecules, making the rational design of TMDs-based sensors (including biosensors) difficult. This study focuses on the investigations of the fundamental interactions between proteins and two widely researched single layered TMDs, MoS2 and WS2 using a combined study with linear vibrational spectroscopy attenuated total reflectance-FTIR and nonlinear vibrational spectroscopy sum frequency generation vibrational spectroscopy, supplemented by molecular dynamics simulations. It was concluded that a large surface hydrophobic region in a relatively flat location on the protein surface is required for the protein to adsorb onto a mono-layered MoS2 or WS2 surface with preferred orientation. No disulfide bond formation between cysteine groups on the protein and MoS2 or WS2 was found. The conclusions are general and can be used as guiding principles to engineer proteins to attach to TMDs. The approach adopted here is also applicable to study interactions between other 2D materials and biomolecules.


Page 2 of 26

Page 3 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Introduction Microelectronic devices have become smaller in size over the years. Two-dimensional mono-layered materials such as graphene and transition metal dichalcogenides (TMDs) like MoS2 and WS2 have been integrated into advanced microelectronics. Such microelectronics have wideranging applications in light emitting diodes, organic solar cells, and biosensors.1-6 In these applications, the graphene, MoS2, or WS2 monolayer interacts extensively with other molecules. Therefore, to improve the performance of such devices it is necessary to understand such interactions in molecular detail. For example, the demand for the next generation label-free biosensors has increased drastically in recent years. Applications such as blood testing, customized diagnostics, and forensic analyses would all benefit from more sensitive, faster, smaller and portable biosensors that require less sample. Single layered TMDs have shown great potential to be excellent electronic materials capable of interfacing with biological molecules, thereby facilitating label-free biosensing4,7 as well as many other applications. Graphene, the most widely used 2D material, has been extensively researched for various applications including field-effect transistors, sensors and solar applications.8-10 Unlike graphene, TMDs exhibit unique band structures,11,12 allowing not only photon adsorption/emission,1,13 but also switching effects for charge carrier transport.2,5 This unique property makes TMDs promising components of many advanced devices including both optical sensors and transistor-based FET sensors. However, the design of TMD-based optical sensors and transistor-based sensors requires thorough understanding of the surface properties of these materials and their interactions with other molecules.



The interactions of graphene surfaces with other molecules, including biomolecules, have been widely studied and are relatively well understood. Single-layered graphene exhibits strong ππ interactions with aromatic functional groups, and strong dipole-dipole interactions with polar molecules.9,14-17 These properties make graphene well suited to detect aromatic molecules and molecules with strong dipoles. However, the surface properties of TMDs, including commonly used MoS2, and WS2 TMDs, are still incompletely characterized, hampering their use as optical and electronic sensors in micro- or nano-electronic devices. Recent studies have shown the potential of TMD-based electronic sensors for detecting biological molecules such as prostate specific antigen (PSA),3,18 glucose4 and dsDNA4. However, although various mechanisms have been hypothesized for how TMDs interact with biological molecules, very few experimental studies have been performed to test these hypotheses. Recent computational simulations suggested that lysozyme binds to the MoS2 surface in a well-defined orientation,19 whereas β-sheet protein (e.g., YAP65) denatures on the MoS2 surface.20 Experimental data suggests that disulfide bonds may form between MoS2 and thiol groups of other molecules.21 The lack of systematic fundamental understanding of TMDs’ interfacial interactions is partially due to the lack of appropriate analytical tools to probe the interface between TMD monolayer and the monolayer of biomolecules. Here, we report a systematic investigation of the interactions between several proteins and TMDs formed from MoS2 and WS2 using sum frequency generation (SFG) vibrational spectroscopy and attenuated total reflectance (ATR)-FTIR spectroscopy, supplemented by molecular dynamic (MD) simulations. SFG, a second-order nonlinear optical spectroscopy, has been extensively applied to study surfaces and buried interfaces in situ at the molecular level.22-33 We have applied SFG to study conformations and orientations of peptides and proteins at various 4 ACS Paragon Plus Environment

Page 4 of 26

Page 5 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


surfaces and interfaces.27,34-39 By combining SFG with ATR-FTIR spectroscopy, we have quantified the orientational distribution of immobilized peptides40 and enzymes, and determined the orientation of membrane protein complexes.35,41 Materials and methods Protein samples. β-glucosidase (β-Glu, PDB: 3AHX), 6-phospho-β-galactosidase (D308C) (βGal (D308C), β-Gal PDB: 2PBG) and haloalkane dehalogenase (A141 C) (HLD (A141C), PDB: 1MJ5) were over-expressed in E. coli, purified, and characterized as described previously.42-44 All enzymes were diluted to 5 μM in pH 7.4 PBS buffer before use. TMD surface preparation and AFM. MoS2 and WS2 surfaces were prepared by drop casting MoS2 and WS2 solutions onto appropriate supporting substrates (CaF2 right angle prism for SFG, ZnSe ATR crystal for ATR-FTIR, Si wafer for AFM measurements). MoS2 and WS2 solutions were purchased from Graphene Supermarket and were used without further purification. Both MoS2 and WS2 solutions were diluted 10 times with ethanol and were sonicated for 15 min before use. After drop casting MoS2 or WS2 onto surfaces, the samples were allowed to dry and then further sonicated for 15 min in ethanol to eliminate any loosely bonded multilayered TMD residues on the supporting substrate. SFG spectroscopy and ATR-FTIR spectroscopy. Detailed SFG theories have been published before22-31 and will only be briefly discussed here: SFG is a second-order nonlinear optical effect where two incoming photons are spatially and temporarily overlapped at the interface, and the third photon is generated with the sum of the two incoming photons’ energies. For this study, the two incoming photons are a visible photon (532nm) and an infrared photon. The infrared beam frequency is tunable so that chemical selectivity can be 5 ACS Paragon Plus Environment


achieved. The commercial SFG spectrometer used in this study is made by EKSPLA, the details of which have been extensively introduced in our previous publications.27,34-47 ATR-FTIR is a linear vibrational spectroscopy, and its high sensitivity originates from its multiple bounces of the infrared beam within the ATR crystal, allowing signal amplification. Using the polarized incoming infrared beam, one can measure the dichroic ratio Rp/s to quantify the orientation of target molecules.48,49 The ATR-FTIR instrument used in this study is a Nicolet 6700 FT-IR spectrometer with ATR accessories, made by Thermo Scientific. Clean right-angle CaF2 prisms and ZnSe ATR crystals were used for depositing MoS2 and WS2 for SFG spectroscopic and ATR-FTIR spectroscopic studies respectively. The MoS2 or WS2 sample surface was placed in contact with a protein solution (i.e., the HLD (A141C), β-gal (D398C) or -Glu solution) for 60 min. Then the protein solution was replaced by PBS buffer (pH 7.4) for several times to remove the loosely bound protein molecules. SFG spectra were collected from the interface between CaF2 (with adsorbed proteins on MoS2 or WS2) and buffer with ssp (s-polarized SFG signal, s-polarized input visible beam, p-polarized IR beam) and ppp polarization combinations of the input and output laser beams. ATR-FTIR spectra were collected from the ZnSe ATR crystal (with adsorbed proteins on MoS2 or WS2)/buffer interface with s or p polarized IR beam. Protein orientation analysis using SFG and ATR-FTIR We demonstrated that a combined study with SFG and ATR-FTIR can more accurately determine the orientation of a biological molecule such as a protein at an interface.35,41 Two orientation angles, a tilt angle θ and a twist angle ψ, are needed to define the orientation of a protein at an interface (Figure 1).34 Because a protein can have a very complicated structure, the


Page 6 of 26

Page 7 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


experimental data obtained from limited measurements may correspond to many possible orientation angle combinations. With more experimentally measured orientational parameters, more accurate orientation information can be obtained. The combined use of SFG and ATR-FTIR can lead to better determined orientation results compared to the use of SFG or ATR-FTIR alone. In this research, both SFG and ATR-FTIR will be used to examine the molecular interactions between MoS2 or WS2 and protein molecules. For an SFG experiment, the generated SFG signal is proportional to the square of the second order nonlinear susceptibility: (2) 2

𝐼 ∝ |𝜒𝑒𝑓𝑓 |

(eq. 1)

( 2) components can be measured using different polarizations of the Different  eff ( 2) input/output beams such as ssp and ppp. Such  eff components can be correlated to the second

order nonlinear susceptibility defined in the x-y-z lab-fixed frame (which has 27 tensor components). This second order nonlinear susceptibility is proportional to the number of the molecules N, molecular orientation angles θ (tilt angle) and ψ (twist angle), and the hyperpolarizability tensor () of the molecule defined in the molecular frame. Therefore the measured χeff components can also be related to N, θ, ψ and : (2)

𝜒𝑒𝑓𝑓 ∝ (𝑁, 𝜃, 𝜓, 𝛽𝑒𝑓𝑓 )

(eq. 2)

For complicated biological molecules like proteins, the hyperpolarizability tensor elements for one type of secondary structure (e.g., α-helical structure) of the molecule can be further calculated by adding up the individual α-helical segments according to the bond additivity model described previously:27 7 ACS Paragon Plus Environment


∗ 𝑑𝛼ℎ𝑒𝑙𝑖𝑥,𝑛

𝛽𝑖𝑗𝑘,ℎ𝑒𝑙𝑖𝑥,𝑛 = (

𝑑𝑄

𝑑𝜇ℎ𝑒𝑙𝑖𝑥,𝑛

)

𝑖,𝑗

×(

# 𝑜𝑓 ℎ𝑒𝑙𝑖𝑐𝑒𝑠 1 2𝜋 ∫ (𝑅 2𝜋 0

𝛽𝑖𝑗𝑘,𝑝𝑟𝑜𝑡𝑒𝑖𝑛 = ∑𝑛=1

𝑑𝑄

)

𝑘

∗ 𝛼ℎ𝑒𝑙𝑖𝑥,𝑛 ∗ 𝑅𝑇 )(𝑅 ∗ 𝜇ℎ𝑒𝑙𝑖𝑥,𝑛 )𝑑𝜓

Page 8 of 26

(eq. 3)

(eq. 4)

In the above equation, α represents the Raman polarizability derivative, μ is the IR ( 2) transition dipole moment, and R represents the Euler rotation matrix. In SFG experiments,  eff ( 2) components such as χssp and χppp can be measured. With a measured  eff component ratio (e.g.,

χssp/χppp ratio), the value “N” can be canceled. With the calculated  components from the protein crystal structure, orientation information θ (tilt angle) and ψ (twist angle) can be obtained. For ATR-FTIR, the measured dichroic ratio (measured by the s and p polarized spectra) can be used to deduce protein’s orientation information: 𝑅𝑝/𝑠 =

𝐴∥ 𝐴⊥

=

𝐸𝑥2 𝐸𝑦2

〈𝑀2 〉 𝐸𝑧2

+ 〈𝑀𝑧2 〉 𝑦

𝐸𝑦2

(eq. 5)

Where A// and A are the absorbance of the α-helical amide I mode in the parallel and perpendicular polarizations. Ex, Ey, and Ez are the electric field amplitudes and can be derived for the experimental conditions used. Mx, My and Mz represent the transition dipole components of a helix in the lab-fixed frame, which are related to the transition dipole moments in the molecular frame and the orientation of the helix.35 As the treatment used in SFG data analysis, by using the bond additivity model, the overall transition dipole moment of the protein can be calculated. Therefore with measured dichroic ratio, known Ex, Ey, and Ez for a particular experiment, and the calculated transition dipole moments in the molecular frame, the orientation of the protein can be deduced. 8 ACS Paragon Plus Environment

Page 9 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Since SFG and ATR-FTIR measured different parameters,35 the combined method can be used to better determine the protein orientation at interfaces.35 More details on how to deduce protein orientation using SFG and ATR-FTIR were published and highlighted in the Supporting Information. (Figure S1)

Figure 1 (a) Illustration of HLD in its (θ = 0°, ψ = 0°) orientation and (b) tilt angle θ and twist angle ψ defined in lab-fixed coordination frame (x, y, z)

Results and discussion



Figure 2 The MoS2 surface (deposited on CaF2) was placed in contact with (a) β-Glu (b) β-Gal (D308C) or (c) HLD (A141C) solution for 60 min, and then the protein solution was replaced by buffer for several times to wash off loosely bound proteins. SFG ppp spectrum was then collected from each MoS2/buffer interface.

As discussed above, it has been reported that disulfide bonds may form between MoS2 and thiol groups of other molecules.21 If enzymes could be selectively attached to MoS2 TDMs through cysteine residues, this would provide a valuable method for interfacing biological molecules with electronic devices. To examine whether it may be possible to selectively immobilize enzymes by this method, we studied the possible immobilization of three different enzymes β-Glu (wildtype), β-Gal-D308C and HLD-A141C on MoS2 surface. Each of these enzymes possesses a single surface-reactive cysteine residue and in previous studies we have covalently immobilized each on maleimide-terminated self-assembled monolayer (SAM) surfaces in well-defined orientations.4244

A MoS2 surface (deposited on CaF2) was incubated with solutions of either β-Glu, β-GalD308C, or HLD-A141C for 60 min, and the surface was then washed several times with buffer (containing 5% tween 20 surfactants) to remove non-covalently bound protein. SFG ppp-polarized spectrum was then collected from each MoS2/buffer interface (Figure 2). Figure 2 shows that no 10 ACS Paragon Plus Environment

Page 10 of 26

Page 11 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


SFG signal could be detected for either the β-Glu or β-Gal-D308C enzymes whereas HLD-A141C exhibited a well-defined SFG spectrum which was successfully collected from HLD (A141C) on MoS2. The absence of SFG signal from β-Glu and β-Gal-D308C could be due to the enzymes not being adsorbed to MoS2 or the adsorbed proteins being randomly oriented on MoS2, thereby canceling out their spectral contributions. In either case this is evidence that the two proteins do not interact with the MoS2 surface via disulfide bonds, otherwise they should adopt certain preferred orientations to generate SFG amide I signal. More discussion on the absence of SFG signals will be presented below. Further ssp-polarized SFG spectrum was collected from HLD-A141C adsorbed on MoS2 (Figure 3). In both SFG ssp- and ppp-polarized spectra, the signals are dominated by a peak around 1650 cm-1, contributed by the alpha-helices in the enzyme. These SFG signals indicate that the adsorbed HLD-A141C adopts a preferential orientation on the MoS2 surface. Unlike graphene, it is difficult to prepare a large monolayer MoS2 sheet covering the entire CaF2 surface. Instead the MoS2 monolayers form islands on a substrate surface, which can be observed using AFM (Supporting information, Figure S3). Therefore control experiments were performed to show that no SFG signals could be detected from the CaF2 prisms lacking MoS2 coating after incubation with proteins. These results confirmed that the SFG spectra arise from HLD adsorbed on MoS2, rather than CaF2. Next, s- and p-polarized ATR-FTIR spectra were collected from HLD-A141C adsorbed on a MoS2-coated ZnSe ATR crystal (Figure 3). The control experiment results show that no HLDA141C adsorbed to the ATR crystal after buffer washing (Supporting Information, Figure S4). This confirms that the ATR-FTIR signals were contributed by the HLD-A141C molecules adsorbed on MoS2. 11 ACS Paragon Plus Environment


With the known crystal structure from the PDB file, we can calculate the SFG signal responses (e.g., ssp and ppp signal strengths) contributed by all alpha-helical structures in a protein as a function of the protein orientation (defined using tilt and twist angles). Similarly, we can also calculate the orientation dependent s and p ATR-FTIR signal strengths from alpha-helical structures in the protein. Here we assume that HLD (A141C) will not have substantial conformational changes after adsorption to MoS2 (which is confirmed by MD simulation results presented later). Therefore we can calculate the orientation dependent SFG and ATR-FTIR signal strengths of the interfacial HLD (A141C) using the HLD crystal structure, shown by orientation dependent maps (Figure S1). Detailed orientation dependent map generation can be found in previous publications and the supporting information.34,35 SFG ppp- and ssp-polarized spectra were fitted using the standard SFG spectral fitting formula27 and the fitting parameters are displayed in Tab S1. The SFG fitted χppp/χssp ratio was 1.55, which can be used to compare to the calculated orientation dependent χppp/χssp ratio to deduce the possible orientation angle regions. The ATR-FTIR fitting result (Tab S2) indicates that the measured dichroic ratio for alpha-helical components in HLD is 1.74, which can also be applied to compare to the calculated orientation dependent ATR-FTIR dichroic ratio to deduce the protein orientation. Since the final orientation of the interfacial HLD on MoS2 should satisfy both SFG and ATR-FTIR measurements, the overlapped possible orientation angle regions deduced from SFG and ATR-FTIR should be the final results, which are shown with a heat map (Figure 3c). Clearly, the most likely orientations of HLD (A142C) on MoS2 should be either around ~40°, ~90° (tilt angle, twist angle), or 140°, ~260° (tilt angle, twist angle). The zero position (0° tilt angle, 0° twist angle) of the protein is shown in Figure 1a.


Page 12 of 26

Page 13 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 3 (a) SFG ppp and ssp spectra and (b) ATR-FTIR s and p spectra collected from HLD (A142C) adsorbed on MoS2 surfaces in buffer (after placing the MoS2 surface in contact with HLD solution for 60 min, and replacing the protein solution with buffer for several times to wash off loosely bound proteins); (c) Heat map indicating most likely orientations of HLD (A141C) on MoS2 surface.

Figure 4 (a-d) Orientations of HLD (141C) adsorbed on MoS2 deduced using coarse grain MD simulation with different initial poses of the protein (rotation axis: residue109-residue 159). To help distinguish between these two orientations and gain further insights into the nature of the protein-MoS2 interaction, coarse-grain molecular dynamics (MD) simulations of HLDA141C adsorbed on MoS2 were performed. General details of the methodology have been



published previously and specific information related to this study is included in the Supporting Information. For coarse-grained MD simulation, as shown in the Supporting Information, only one parameter related to the surface energy is needed to describe the surface. When different initial orientations of the protein were used in the simulation, different final simulation results were obtained. Figure 4 shows the four initial orientations of HLD-A141C on the MoS2 surface (prepared by rotating 90° along x-axis for each different initial orientation) and the final simulation results. Figures 4a, 4d show that under these two initial orientations, HLD (A141C) could adsorb onto the MoS2 surface with the same orientation. Figures 4b and 4c show that HLD (A141C) cannot adsorb onto the MoS2 surface under these two initial orientations. The most stable orientation identified by the MD simulations is shown in Figures 4a and 4d. Such an orientation was then compared with the protein orientation deduced from the SFG and ATR-FTIR experiments (Figure 5). This comparison indicates that the orientation with 140°, 260° (tilt angle, twist angle) matches the simulation data, suggesting that this orientation is more likely orientation.

Figure 5 (a) The stabilized HLD (A141C) structure on MoS2 surface plotted in the ribbon form obtained by MD simulation; (b) Experimentally deduced HLD (A141C) orientation on MoS2 at 140°, 260° (tilt angle, twist angle). 14 ACS Paragon Plus Environment

Page 14 of 26

Page 15 of 26 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 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Numerous studies have shown that proteins often denature on hydrophobic surfaces including polymers, self-assembly monolayers etc.50-52 This denaturation process generally involves the transient exposure of the hydrophobic protein core, due to protein breathing motions, and subsequent physisorption onto the hydrophobic surface. The MoS2 surface hydrophobicity was investigated53,54 and in ambient condition (which is our experimental condition) it was measured to be relatively hydrophobic, with a water contact angle between 80 and 90 degrees,53 thus it is interesting that the simulated adsorption of HLD (A141C) to MoS2 did not lead to the denaturation of the protein due to exposure of the hydrophobic core to the surface. To further understand the interaction dynamics of HLD (A141C) on the MoS2 surface, we examined the HLD (A141C) - MoS2 interactions in more detail with the MD simulation trajectory. Figure 6 shows the representative snapshots from the simulation trajectory (on the left) and their corresponding hydrophobicity maps of the top-down view (looking from the interface side) of HLD (A141C) (on the right) for intermediate/transition states. As shown in Figure 6, HLD (A141C) was adsorbed to the surface shortly (

Molecular Mechanisms of Interactions between Mono-layered

Recommend Documents