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Chapter 35

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Characterizing the Structure of Surface-Immobilized Proteins: A Surface Analysis Approach Joe E. Baio,1,2 Tobias Weidner,1,2 and David G. Castner*,1 1National ESCA and Surface Analysis Center for Biomedical Problems (NESAC/BIO), Departments of Bioengineering and Chemical Engineering, University of Washington, Seattle, Washington, U.S.A. 2Max Planck Institute for Polymer Research, Mainz, Germany *E-mail: [email protected]

There are many techniques that allow surface scientists to study interfaces. However, few are routinely applied to probe biological surfaces. The work presented here demonstrates how detailed information about the conformation, orientation, chemical state, and molecular structure of biological molecules immobilized onto a surface can be assessed by electron spectroscopy, mass spectrometry, and nonlinear vibrational spectroscopy techniques. This investigation began with the development of simple model systems (small proteins, and peptides) and has evolved into a study of more complex – real world systems. Two model systems based on the chemical and electrostatic immobilization of a small rigid protein (Protein G B1 domain, 6kDa) were built to develop the capabilities of time-of-flight secondary ion mass spectrometry (ToF-SIMS), near edge X-ray absorption fine structure spectroscopy (NEXAFS) and sum frequency generation (SFG) spectroscopy as tools to probe the structure of surface immobilized proteins. ToF-SIMS sampled the amino acid composition of the exposed surface of the protein film. Within the ToF-SIMS spectra, an enrichment of secondary ions from amino acids

© 2012 American Chemical Society In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


located at opposite ends of the proteins were used to describe protein orientation. SFG spectral peaks characteristic of ordered α-helix and β-sheet elements were observed for both systems and the phase of the peaks indicated a predominantly upright orientation for both the covalent and electrostatic configurations. Polarization dependence of the NEXAFS signal from the N 1s to π* transition of the peptide bonds that make up the β-sheets also indicated protein ordering at the surface.

Introduction Wolfgang Pauli once lamented that “God made the bulk; the surface was invented by the devil.” His frustration, implied by this famous quote, stems from the inherent difficulties associated with characterizing and explaining the heterogeneous nature of surfaces. Since Pauli’s time, surface scientists, in their quest for novel explanations of surface phenomena have made some headway (1). This progress has been made in part due to the development of a range of techniques that involve bombarding the surface with photons, electrons or ions (2). Yet, most of these techniques have been developed to look at simple systems – single, small molecules interacting with a model surface. For example, fifty years after Pauli’s death, surface scientists are still trying to characterize the structure of a single water molecule interacting with a Pt 111 surface (3). In comparison to these non-organic systems - biological molecules are orders of magnitude more complex. Protein–surface mediated phenomena are directly influenced by the conformation, orientation, activity, organization and surface concentrations of the adsorbed proteins (4). Then combine this with the fact that most traditional surface analysis tools perform only under ultra high vacuum conditions (2) and accurately characterizing the structure of a surface bound biomolecule quickly becomes a Herculean affair. Proteins at a surface or interface mediate most biological interactions. The capsulation of an implanted surface, affinity chromatography of a protein, cellular signaling or the analyte capture performance of a biological sensor - are all influenced by the structure of proteins at a surface (4–7). Yet, of the 70,000+ protein structures solved and uploaded to the protein database, not a single one describes the structure of the protein at a surface (8). This despite research that has demonstrated that the structure of a protein changes when it comes in contact with a surface (9–11). Therefore, do these reported crystal structures accurately define the structure of the protein at the surface? For example, hydrophobic effects between the protein and a substrate may drive the protein to expose hydrophobic domains– inducing conformational changes (12, 13). Typically, changes in conformation are then probed by assessing the activity of the surface, where a qualitative view of protein orientation and conformation can be provided by some sort of binding assay. Observed changes in binding are then assumed to be a direct result of orientation or the exposure of different domains within the 762 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


protein (9, 11, 14, 15). Yet, a detailed, atomic-level picture of protein orientation or structural changes induced by the surface still does not exist. In response to this explosion of interest centered around the construction of biological based immunosensors, chemists have proposed a range of possible protein immobilization schemes based on coordination complexes (16–19), ligand-reception (20), covalent conjugation (19, 21–26), hydrophobic/hydrophilic driving forces (27–29) and electrostatic interactions (30–32). The ability of these biological devices to bind specific targets is directly related to the accessibility of capture groups at the sensor surface (14). For devices based on proteins and antibodies, these immobilization schemes must preserve the conformation of the protein and successfully orient binding sites so that they are accessible. To avoid a trial and error approach, and truly the design these devices at the molecular level, high-resolution techniques are needed to assess the structure, the activity and the orientation of these proteins. While methods like x-ray diffraction (XRD) and nuclear magnetic resonance (NMR) do provide angstrom level resolution of atomic positions of proteins in crystals or solutions (33, 34) – they do not provide the sensitivity required to characterize the structure of monolayer or sub-monolayer concentrations of proteins interacting with a surface. As a result, a single technique that provides a high-resolution picture of complex organic molecules at a surface, is still elusive. Currently, biological assays like ELISA provide qualitative insights into changes in activity of surface bound proteins (35). Scanning probe techniques can provide images and force curves of proteins unfolding at a surface (36). Optical techniques like surface plasmon resonance (SPR) can monitor changes at a surface by providing quantitative measures of kinetics (37, 38). Yet, even together, these techniques all fail to fully bridge the current gap between characterizing simple molecules on non-organic surfaces to providing structural details of proteins at a complex biointerface. The overarching goal of the work presented here - is the development of a suite of high-resolution surface analytical techniques to fully explore the structure, orientation and ordering of proteins at an interface. The hope is that the complementary techniques developed here will provide molecular information that can then be applied to the design and characterization of biomaterial surfaces. The strategy we have adopted is to began with the development of some simple model systems (small proteins and peptides), use these systems to test our surface analytical methods, and then evolve into a study of some more complex, real world systems.

The Protein G B1 Model System This initial work involved inducing the B1 domain of Protein G into two different orientations by creating two versions of the protein by site directed mutagenesis (39). This small barrel shaped protein is 3nm in height and contains just a single alpha helix and four anti-parallel beta-sheets. To immobilize this B1 domain, we took advantage of the large body of work in the literature, where researchers have reported reproducible multiplexed biomolecular surfaces by 763 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


attaching proteins and DNA (40, 41) via bioactive ligands (16–18, 20, 22–25, 42, 43), Originally, a single cysteine was introduced onto the exposed loop at either end of the protein (V21C and T11C) (39). This cysteine presented a thiol group which drove binding to a maleimide-oligo(ethylene glycol)-functionalized (MEG) substrate (Figure 1). On both of these substrates we expected that the two variants of this protein should induce itself into two different end-on orientations.

Figure 1. Protein G B1 immobilization schemes: A. Protein G B1 variants, V21C and T11C, with cysteines introduced at opposite ends of the protein, were immobilized via the cysteine thiol onto maleimide-oligo(ethylene glycol)-functionalized gold. B. The charge variant of Protein G B1, D4′, was immobilized via electrostatic interactions onto amine and carboxyl functionalized gold.

We then characterized these different systems with a suite of surface analysis tools. We set out to probe ordering of secondary structures within these protein films by Sum Frequency Generation spectroscopy (SFG); determine the geometry of specific bonds within these surface bound proteins with Near Edge X-ray Adsorption Fine Structure (NEXAFS) spectroscopy; and asses the orientation induced by these conjugation schemes with Time-of-Flight Secondary Ion Mass Spectrometry (ToF-SIMS).

Characterizing Ordering of Secondary Structure Just like other vibrational spectroscopic techniques N-C=O, N-H, C-H, and O-H vibrational modes observed within SFG spectra are all used to identify secondary structures (ie. α-helices, β-sheets and β-turns) and amino acid side chains. However, the SFG selection rules dictate that these N-C=O, N-H, C-H, 764 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


and O-H vibrational modes will only start to appear if secondary structures, amino acid side chains, or water are ordered at the interface (44–47). During an SFG experiment, one incident photon source is kept at a fixed frequency within the visible range, while the other is a tunable or broad-band IR source. When the sum of the frequencies of the incident photons is equal to a frequency of the resonance modes of the specimen, a non-linear susceptibility term, X(2), exhibits a sudden change in magnitude (48). The square of the magnitude of X(2) is proportional to the intensity of the reflected summed beam. No change in the SFG response will be observed in a medium with inversion symmetry (X(2)=0) (48), but inversion symmetry is always broken at interfaces. As a result of these selection rules, we expect that any signal observed in the amide I stretching region will only originate from ordered secondary structures within this surface immobilized protein. This B1 domain contains just a single alpha helix and four anti-parallel betasheets and within the amide I SFG spectra (Figure 2A), collected from the two cysteine mutants bound to the maleimide functionalized Au, we observe three peaks at 1626, 1645 and 1675cm-1 (39). The peak near 1645 cm-1 is characteristic of ordered alpha helices while the two peaks at 1626 and 1675 cm-1 originate from beta-sheets. However, in contrast, the spectrum collected from a protein film made up of the wild-type version of the protein does not contain these spectral features (Figure 2A). Thereby implying that without the thiol group, inserted by the cysteine mutation, this protein adsorbs onto the maleimide with a random distribution of orientations (39).

Figure 2. A. SFG amide I spectra of Protein G B1 wild type and cysteine mutants (V21C and T11C) on maleimide-oligo(ethylene glycol)- functionalized gold (39). B. SFG amide I spectra of a monolayer of D4′ Protein G B1 on amine functionalized gold (51). The amide I peak near 1645 cm-1 is characteristic of ordered α-helices, while those near 1630 and 1680 cm−1 are characteristic of ordered β-sheet structures. These features are absent from data taken from a monolayer of wild-type protein on the same substrate. 765 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


The phase of these peaks, with respect to the Au nonresonant background, can also shed some light on the overall orientation of these secondary structures. Previous work examining a helical peptide adsorbed onto functionalized Au demonstrate that when the spectral feature at 1645 cm-1 is out of phase with the nonresonant Au background (peak amplitude is negative) the peptide backbone is oriented parallel to the substrate (49). The phase of the peaks found in Figure 2A are in phase with the nonresonant background (positive amplitude) therefore we can conclude that that helix within the B1 domain is pointing in an orthogonal direction to the surface. Based on the location of the cysteine mutation we expect that the helix orientation should be more upright for the T11C compared to the V21C system, which is consistent with the SFG spectra (Figure 2A) that show a more prominent amide resonance at 1645 cm-1. This qualitative view of orientation can be expanded to include a more quantitative analysis by comparing spectra across different polarization combinations (i.e., ppp versus ssp) (50–52). For example, Nguyen et. al., probing the orientation of a α-helix inserted within a membrane, demonstrated that changes in tilt angle across the long axis of the helix can be directly calculated from the dichroism of the signal strength between the ppp and ssp spectra (50). NEXAFS, as a technique, can offer detailed information about the bonding environment of molecules at a surface and provide additional information about the orientation and order of bonds within a protein film (19, 39, 51). During a NEXAFS experiment, polarized synchrotron x-rays are absorbed by electrons at the core levels exciting photoelectrons. The resulting holes at the core levels are filled by an electron at a higher energy level, which induces the emission of either an Auger electron or a photon. As these electrons travel to the surface they typically encounter an inelastic scattering processes. Therefore, detection of partial electron yield is dictated by the electron scattering cross section and for organic thin films is typically ~10nm (52). The orientation and tilt angles of ordered molecular bonds can be determined by simply following the change in the x-ray absorption as the incident angle of the electric field vector of the x-rays is varied. Based on this polarization dependence, groups have used NEXAFS to determine tilt angles of specific bonds within DNA oligomers, model peptides and proteins immobilized to a surface (16, 17, 19, 24, 25, 39, 43, 51, 53). To complement the SFG characterization, ordering of the two immobilized Protein G variants were also described by the polarization dependence of the π* feature, within the carbon and nitrogen K-edges (Figure 3). Partial electron yield NEXAFS N K-edge spectra collected from the two Protein G variants can be found in Figure 3 (39). Assuming a well ordered protein film - any observable polarization dependence of the π* feature, at 400.6 eV, can be related to the ordering of the amide bonds within the protein backbone. Tilt angles of the π* molecular orbitals were then calculated from the magnitude of this polarization dependence. The estimated tilt angles of inner β-strands were 40-50° for both variants - one variant is more tilted than the other. If we assume that both the helix and the β-sheets point in the same direction, then these tilt angles of the β-strands are consistent with the SFG results which demonstrate that the helix is also pointing upright, with respect to the substrate (39). 766 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


Figure 3. Partial electron yield NEXAFS spectra of the nitrogen K-edge for the two cysteine variants (V21C and T11C) immobilized onto maleimide-oligo(ethylene glycol)-functionalized gold, acquired at angles of 20° and 70°, along with the difference between 70°, 55° and 20° spectra. The prominent dichroism observed at 400.6 eV in the difference spectra (70°-20°) is attributed to peptide bonds of the four anti-parallel β-sheets. Reproduced with permission from ref. (39). Copyright 2010 American Chemical Society.

Characterizing Protein Orientation Following the protein adsorption step any changes in overall end-on orientation, induced by our immobilization schemes, were characterized by ToF-SIMS. ToF-SIMS involves bombarding a surface with a pulsed primary ion beam that sputters molecular fragments. The primary ion hits the surface inciting a collision cascade and within this energized region many processes are occurring, including post emission ionization, recombination, etc (54). The small fraction of these fragments ( equimolar mixture (“V + T”) > T11C, is expected for end-on orientations of the two mutants. Reproduced with permission from ref. (39). Copyright 2010 American Chemical Society.

Similarly, if amino acids are asymmetrically distributed about the protein – intensities of secondary ions originating from these amino acids can be directly related to conformation and orientation (32, 63–65). The B1 domain of Protein G has an asymmetric distribution of amino acids. The N-terminus is rich in tyrosine while at the opposite end of the protein, the C-terminus, is rich in leucine and asparagine (Figure 4). ToF-SIMS data from the T11C and V21C variants showed an enrichment of secondary ions originating from asymmetric amino acids (Asparagine: 70, 87, and 98 m/z; Leucine: 86 m/z; Tyrosine: 107 and 136 m/z) concentrated in the opposite end of the protein from the cysteine (Figure 769 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


4). For a semi-quantitative assessment of orientation, we created a ratio relating the intensities of these secondary-ions originating from either end of the protein (Figure 5) (26, 39). Observed changes in this ratio, for the two variants on both substrates, indicate two distinct end-on orientations. This was in spite of the fact that the thickness of this protein layer is similar to the SIMS sampling depth. These intensity ratios were also compared with 50:50 mixtures of the variants and with nonspecifically immobilized proteins in random orientations. Additionally, we explored a range of conjugation protocols and found that for immobilization onto the MEG substrates, orientation was enhanced by increasing both the pH (7.0 to 9.5) and salt concentration (0 to 1.5 M NaCl) of the protein-buffer solution (Figure 5) (39). These ratios illustrate a transition from a randomly oriented protein film at a neutral pH (Figure 5; pH 7) to an oriented film as we increase the salt concentration to 1.5M NaCl (Figure 5; NaCl). The addition of the NaCl at pH 7 may inhibit the charge-charge interactions of adjacent proteins, thereby, improving the packing and orientation of the film.

Electrostatic Conjugation This initial Protein G B1 model system, based on the cysteine-maleimide bond, helped us develop and highlight the capabilities of ToF-SIMS, NEXAFS spectroscopy and SFG as tools to probe the structure of surface immobilized proteins. Information about the orientation was provided by the intensities of secondary-ions originating from amino acids asymmetrically distributed within the protein’s three-dimensional structure. NEXAFS and SFG experiments illustrated how we can define the geometry of molecular bonds, thus, complementing the ToF-SIMS characterization of overall protein orientation. So with this newly constructed toolbox in hand – we expanded this work to include proteins induced into different orientations by pairing electrostatic dipoles within the protein to charged substrates (Figure 1B). This expansion was focused around the same Protein G B1 domain, but instead of inserting cysteines, a charge distribution was created within the protein. Negatively charged amino acids are uniformly distributed throughout wild-type Protein G B1, so at one end of the protein a neutral region was created by replacing negatively charged amino acids with neutral residues (Figure 1B). This mutant (D4′) was then immobilized onto two oppositely charged substrates (COO- and NH3+ functionalized gold) (51). Within the amide I SFG spectra, acquired for the D4′ variant immobilized onto a NH3+ functionalized surface, are spectral features related to ordered α-helicies (1645 cm-1) and β-sheets (1630 and 1680 cm-1) within the amide I SFG spectrum (Figure 2b) (51). This implies that the electrostatic interaction between the protein and the surface drives the protein into an ordered monolayer. This also corresponded to the observed polarization dependence of the N1s to π* transition, within the N K-edge NEXAFS spectra, related to the β-sheet peptide bonds present within the protein film. Finally, ToF-SIMS data, taken from the D4′ adsorbed onto COO- and NH3+ functionalized gold demonstrated a well-defined separation between the two samples. The observed two-fold increase in the 770 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


ratio of secondary-ion intensities originating from opposite ends of the protein indicates opposite orientations of the Protein G B1 fragment on the two different surfaces (see Figure 6) (51). Again, a charge distribution was created at opposite ends of the protein by substituting specific negatively charged amino acids with neutral residues. As a result, asparagine was no longer asymmetrically distributed within the protein and peaks originating from asparagine were not included in the calculation ratio of secondary-ion intensities.

Figure 6. ToF-SIMS peak ratios calculated as the sum of intensities of secondary ions from methionine (62 and 105 m/z) and tyrosine (107 and 136 m/z) divided by the sum of intensities of secondary ions from the leucine/isoleucine residues (86 m/z). Error bars represent the standard deviation across fifteen analysis spots over three distinct samples. Reproduced with permission from ref. (51). Copyright 2012 American Chemical Society. 771 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


Future Approaches: Amino Acid Labeling Strategies Information about the end-on orientation and secondary structure of proteins immobilized onto surfaces is, as discussed above, useful for a number of practical bioengineering problems. However, important phenomena regarding the molecular mechanisms of protein–surface interactions and the structure of surface proteins can only be understood in detail by probing individual side chains with Ångstrom resolution. We have shown that isotope labels at specific protein sites in combination with SFG spectroscopy is a promising route towards high-resolution protein structures on surfaces (66, 68). Deuteration of C–H bonds leads to a red shift of nearly 800 cm-1 of that particular resonance. Therefore one can measure SFG spectra of the deuterium labeled amino acid without spectral confusion with C-H containing side chains. Established procedures for SFG orientation analysis of aromatic and aliphatic groups can then be applied to probe the side chain orientation. In an earlier study, we have determined the orientations of the entire set binding side chains of an amphiphilic model peptide containing lysine and leucine side chains on a polystyrene surface in situ (66). We also determined the orientation of individual phenylalanine side chains in the binding domain of statherin on its native mineral hydroxyapatite (53). Both studies allowed the determination of both tilt and torsion angles for the respective side chains. One challenge in the context of single amino acid detection is the extremely low surface density of labeled species. The surface area of a small 15 aminoacid peptide is on the order of 400 Å2. This means a single side chain has an approximately 15-20 times lower surface density than terminal groups in typical self-assembled monolayers with a footprint of around 20-27 Å2 per molecule. For larger proteins the surface density of individual labeled sites quickly drops to a hundredth of a monolayer or less. To test the feasibility of extending our labeling approach from peptides to proteins, we collected SFG spectra of deuterium labeled tyrosine (Tyr45) and isoleucine (Ile6) sites in the T11C mutant of the B1 domain of protein G (v.s.). Again, T11C was immobilized via cysteine onto a MEG self-assembled monolayer (SAM) on gold (Figure 7). T11C covers a surface area of ca. 800 Å2, leading to a surface density of the labeled amino acids of the equivalent of 2-3% of a monolayer. The aliphatic and aromatic groups have spectrally distinct resonance positions, labeling aromatic and aliphatic species in a single protein causes no spectral overlap and two species can be probed in a single experiment. A C–D stretching range SFG spectrum collected in ppp polarization of the surface bound protein is shown in figure 7. There is a clear signature of both the aromatic tyrosine and the aliphatic isoleucine above and below 2230 cm-1, respectively. Isoleucine resonances are visible near 2150 cm-1, assigned to the methylene Fermi resonance and near 2220 cm-1, assigned to the asymmetric CD3 stretching mode. From the positive polarity of the peaks we can conclude that the methyl vibrations are in phase with the non-resonant SFG gold background. It has been shown that a constructive interference of methyl resonances with the gold background signal is indicative of isopropyl methyl groups pointing towards the surface (67, 68). From the absence of a symmetric CH3 mode we can conclude 772 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


that the average orientation of the methyl units is strongly tilted (69). At higher wavenumbers there is a strong mode near 2290 cm-1 related to a v2 ring mode and two overlapping ring resonances at 2240 cm-1 and 2276 cm-1 visible in the spectrum which can be assigned to v7 and v13 ring vibrations, respectively. This data clearly confirms, that labeling of individual protein sites for surface protein structure analysis is a concept that can be extended to proteins.

Figure 7. A SFG spectrum of the labeled T11C protein immobilized onto maleimide-oligo(ethylene glycol)-functionalized gold acquired at ppp polarization. Signature peaks of both the aromatic tyrosine and the aliphatic isoleucine above and below 2230 cm-1, respectively. Isoleucine resonances are visible near 2150 cm-1, assigned to the methylene Fermi resonance and near 2220 cm-1, assigned to the asymmetric CD3 stretching mode.

773 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.

Summary


One major hurdle in the design of biomaterial interfaces it the accurate characterization of the protein – surface interactions. Here we have applied three techniques to probe and describe the orientation, chemical state, and molecular structure of proteins immobilized onto a surface. We believe that this set of tools has now reached a state of development where ToF-SIMS, NEXAFS, and SFG can be routinely applied to the characterization of protein films. The models systems described here provided straightforward examples of how overall protein orientation is characterized by ToF-SIMS and a molecular level picture of bond orientation is provided by NEXAFS and SFG experiments.

Acknowledgments The authors acknowledge support from NIH grants EB-2027 (NESAC/BIO), GM-074511 and DE-012554 during preparation of this manuscript as well as for some of the results reported in it. J.E.B. and T.W are grateful to the Max Planck Society for financial support of this work.

References 1. 2. 3.

4.

5.

6. 7. 8. 9.

Somorjai, G. Chemistry In Two Dimensions: Surfaces; Cornell University Press: Ithaca, New York, 1981. Vickerman, J.; Gilmore, I. Surface Analysis: The Principal Techniques, 2nd ed.; John Wiley and Sons: West Sussex, U.K., 2009. Arnadottir, L.; Stuve, E. M.; Jonsson, H. Adsorption of water monomer and clusters on platinum(111) terrace and related steps and kinks I. Configurations, energies, and hydrogen bonding. Surf. Sci. 2010, 604 (21-22), 1978–1986. Horbett, T. Proteins: Structure, Properties and Adsoption to Surfaces. In Biomaterial Science: An Introduction to Materials in Medicine; Ratner, B., Hoffman, A., Schoen, F., Lemons, J., Eds.; Academic Press: New York, 1996; pp 133−141. Nakanishi, K.; Sakiyama, T.; Imamura, K. On the adsorption of proteins on solid surfaces, a common but very complicated phenomenon. J. Biosci. Bioeng. 2001, 91 (3), 233–244. Hynes, R. Molecular-Biology of Fibronectin. Annu. Rev. Cell Biol. 1985, 1, 67–90. Hynes, R. O.; Yamada, K. M. Fibronectins - Multifunctional Modular Glycoproteins. J. Cell Biol. 1982, 95 (2), 369–377. RCSB Protein Data Bank. 2011. Butler, J. E.; Ni, L.; Nessler, R.; Joshi, K. S.; Suter, M.; Rosenberg, B.; Chang, J.; Brown, W. R.; Cantarero, L. A. The Physical And FunctionalBehavior Of Capture Antibodies Adsorbed On Polystyrene. J. Immunol. Methods 1992, 150 (1-2), 77–90. 774 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


10. Plant, A. L.; Locasciobrown, L.; Haller, W.; Durst, R. A. Immobilization Of Binding-Proteins On Nonporous Supports - Comparison Of Protein Loading, Activity, And Stability. Appl. Biochem. Biotechnol. 1991, 30 (1), 83–98. 11. Baugh, L.; Vogel, V. Structural changes of fibronectin adsorbed to model surfaces probed by fluorescence resonance energy transfer. J. Biomed. Mater. Res., Part A 2004, 69A (3), 525–534. 12. Haynes, C. A.; Sliwinsky, E.; Norde, W. Structural And Electrostatic Properties Of Globular-Proteins At A Polystyrene Water Interface. J. Colloid Interface Sci. 1994, 164 (2), 394–409. 13. Phillips, D. C.; York, R. L.; Mermut, O.; McCrea, K. R.; Ward, R. S.; Somorjai, G. A. Side chain, chain length, and sequence effects on amphiphilic peptide adsorption at hydrophobic and hydrophilic surfaces studied by sum-frequency generation vibrational spectroscopy and quartz crystal microbalance. J. Phys. Chem. C 2007, 111 (1), 255–261. 14. Tajima, N.; Takai, M.; Ishihara, K. Significance of Antibody Orientation Unraveled: Well-Oriented Antibodies Recorded High Binding Affinity. Anal. Chem. 2011, 83 (6), 1969–1976. 15. Michael, K. E.; Vernekar, V. N.; Keselowsky, B. G.; Meredith, J. C.; Latour, R. A.; Garcia, A. J. Adsorption-induced conformational changes in fibronectin due to interactions with well-defined surface chemistries. Langmuir 2003, 19 (19), 8033–8040. 16. Cheng, F.; Gamble, L. J.; Castner, D. G. XPS, TOF-SIMS, NEXAFS, and SPR characterization of nitrilotriacetic acid-terminated self-assembled monolayers for controllable immobilization of proteins. Anal. Chem. 2008, 80 (7), 2564–2573. 17. Huang, Z. H.; Park, J. I.; Watson, D. S.; Hwang, P.; Szoka, F. C. Facile synthesis of multivalent nitrilotriacetic acid (NTA) and NTA conjugates for analytical and drug delivery applications. Bioconjugate Chem. 2006, 17 (6), 1592–1600. 18. Wegner, G. J.; Lee, N. J.; Marriott, G.; Corn, R. M. Fabrication of histidinetagged fusion protein arrays for surface plasmon resonance imaging studies of protein-protein and protein-DNA interactions. Anal. Chem. 2003, 75 (18), 4740–4746. 19. Baio, J.; Cheng, F.; Ratner, D.; Stayton, P.; Castner, D. Probing orientation of immobilized humanized anti-lysozyme variable fragment by time-of-flight secondary-ion mass spectrometry. J. Biomed. Mater. Res., Part A 2011, 97A (1), 1–7. 20. Jung, L. S.; Nelson, K. E.; Stayton, P. S.; Campbell, C. T. Binding and dissociation kinetics of wild-type and mutant streptavidins on mixed biotin-containing alkylthiolate monolayers. Langmuir 2000, 16 (24), 9421–9432. 21. Xia, N.; Hu, Y.; Grainger, D. W.; Castner, D. G. Functionalized Poly(ethylene glycol)-Grafted Polysiloxane Monolayers for Control of Protein Binding. Langmuir 2002, 18 (8), 3255–3262. 22. Cheng, F.; Gamble, L. J.; Grainger, D. W.; Castner, D. G. X-ray Photoelectron Spectroscopy, Time-of-flight Secondary Ion Mass Spectrometry and Principal Component Analysis of the Hydrolysis, 775 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.

23.

24.


25.

26.

27.

28.

29.

30. 31.

32.

33. 34. 35. 36. 37.

Regeneration and Reactivity of NHS-containing Organic Thin Films. Anal. Chem. 2007, 79, 8781–8788. Huang, W.; Wang, J. Q.; Bhattacharyya, D.; Bachas, L. G. Improving the activity of immobilized subtilisin by site-specific attachment to surfaces. Anal. Chem. 1997, 69 (22), 4601–4607. Lee, C. Y.; Gamble, L. J.; Grainger, D. W.; Castner, D. G. Mixed DNA/oligo (ethylene glycol) functionalized gold surfaces improve DNA hybridization in complex media. Biointerphases 2006, 1 (2), 82–92. Lee, C. Y.; Gong, P.; Harbers, G. M.; Grainger, D. W.; Castner, D. G.; Gamble, L. J. Surface coverage and structure of mixed DNA/alkylthiol monolayers on gold: Characterization by XPS, NEXAFS, and fluorescence intensity measurements. Anal. Chem. 2006, 78 (10), 3316–3325. Baio, J. E.; Weidner, T.; Samuel, N. T.; McCrea, K.; Baugh, L.; Stayton, P. S.; Castner, D. G. Multi-technique Characterization of Adsorbed Peptide and Protein Orientation: LK310 and Protein G B1. J. Vac. Sci. Technol., B 2010, 28 (4), C5D1–C5D8. Bos, M. A.; Kleijn, J. M. Determination of the Orientation Distribution of Adsorbed Fluorophores Using TIRF. 2. Measurements On Porphyrin And Cytochrome-c. Biophys. J. 1995, 68 (6), 2573–2579. Fraaije, J.; Kleijn, J. M.; Vandergraaf, M.; Dijt, J. C. Orientation of Adsorbed Cytochrome-c as a Function of the Electrical Potential of the Interface Studied by Total Internal-Reflection Fluorescence. Biophys. J. 1990, 57 (5), 965–975. Lee, J. E.; Saavedra, S. S. Molecular orientation in heme protein films adsorbed to hydrophilic and hydrophobic glass surfaces. Langmuir 1996, 12 (16), 4025–4032. Chen, S. F.; Liu, L. Y.; Zhou, J.; Jiang, S. Y. Controlling antibody orientation on charged self-assembled monolayers. Langmuir 2003, 19 (7), 2859–2864. Chi, Q. J.; Zhang, J. D.; Andersen, J. E. T.; Ulstrup, J. Ordered assembly and controlled electron transfer of the blue copper protein azurin at gold (111) single-crystal substrates. J. Phys. Chem. B 2001, 105 (20), 4669–4679. Wang, H.; Castner, D. G.; Ratner, B. D.; Jiang, S. Y. Probing the orientation of surface-immobilized immunoglobulin G by time-of-flight secondary ion mass spectrometry. Langmuir 2004, 20 (5), 1877–1887. Cavanagh, J. Protein NMR spectroscopy : principles and practice; Academic Press: San Diego, 1996. Rupp, B. Biomolecular crystallography : principles, practice, and application to structural biology; Garland Science: New York, 2009. Clark, M. ELISA Techniques. In Plant Molecular Biology; Weissbach, A., Weissbach, H., Eds.; Academic Press: New York, 1986; pp 742−766. Li, M. Q. Scanning probe microscopy (STM/AFM) and applications in biology. Appl. Phys. A: Mater. Sci. Process. 1999, 68 (2), 255–258. Apte, J.; Gamble, L.; Castner, D.; Campbell, C. Kinetics of leucine-lysine peptide adsorption and desorption at -CH3 and -COOH terminated alkylthiolate monolayers. Biointerphases 2010, 5 (4), 97–104.



38. Green, R. J.; Frazier, R. A.; Shakesheff, K. M.; Davies, M. C.; Roberts, C. J.; Tendler, S. J. B. Surface plasmon resonance analysis of dynamic biological interactions with biomaterials. Biomaterials 2000, 21 (18), 1823–1835. 39. Baugh, L.; Nguyen, P.-C.; Baio, J. E.; Weidner, T.; Gamble, L. J.; Castner, D. G.; Stayton, P. S. Probing the Orientation of Surface-Immobilized Protein G B1 using ToF-SIMS, Sum Frequency Generation, and NEXAFS Spectroscopy. Langmuir 2010, 26 (21), 16434–16441. 40. Seidel, M.; Niessner, R. Automated analytical microarrays: a critical review. Anal. Bioanal. Chem. 2008, 391 (5), 1521–1544. 41. Laurent, N.; Voglmeir, J.; Flitsch, S. L. Glycoarrays - tools for determining protein-carbohydrate interactions and glycoenzyme specificity. Chem. Commun. 2008 (37), 4400–4412. 42. Nam, J. M.; Han, S. W.; Lee, K. B.; Liu, X. G.; Ratner, M. A.; Mirkin, C. A. Bioactive protein nanoarrays on nickel oxide surfaces formed by dip-pen nanolithography. Angew. Chem., Int. Ed. 2004, 43 (10), 1246–1249. 43. Nelson, K. E.; Gamble, L.; Jung, L. S.; Boeckl, M. S.; Naeemi, E.; Golledge, S. L.; Sasaki, T.; Castner, D. G.; Campbell, C. T.; Stayton, P. S. Surface characterization of mixed self-assembled monolayers designed for streptavidin immobilization. Langmuir 2001, 17 (9), 2807–2816. 44. Kim, J.; Opdahl, A.; Chou, K. C.; Somorjai, G. A. Hydrophobic-interactioninduced alignment of polymers at the solid/liquid interface studied by infrared-visible sum frequency generation. Langmuir 2003, 19 (23), 9551–9553. 45. Chen, X. Y.; Wang, J.; Sniadecki, J. J.; Even, M. A.; Chen, Z. Probing alphahelical and beta-sheet structures of peptides at solid/liquid interfaces with SFG. Langmuir 2005, 21 (7), 2662–2664. 46. Chen, X. Y.; Clarke, M. L.; Wang, J.; Chen, Z. Sum frequency generation vibrational spectroscopy studies on molecular conformation and orientation of biological molecules at interfaces. Int. J. Mod. Phys. B 2005, 19, 691–713. 47. Mermut, O.; Phillips, D. C.; York, R. L.; McCrea, K. R.; Ward, R. S.; Somorjai, G. A. In Situ Adsorption Studies of a 14-Amino Acid Leucine-Lysine Peptide onto Hydrophobic Polystyrene and Hydrophilic Silica Surfaces Using Quartz Crystal Microbalance, Atomic Force Microscopy, and Sum Frequency Generation Vibrational Spectroscopy. J. Am. Chem. Soc. 2006, 128 (11), 3598–3607. 48. Lambert, A. G.; Davies, P. B.; Neivandt, D. J. Implementing the Theory of Sum Frequency Generation Vibrational Spectroscopy: A Tutorial Review. Appl. Spectrosc. Rev. 2005, 40, 103–145. 49. Weidner, T.; Apte, J. S.; Gamble, L. J.; Castner, D. G. Probing Orientation and Conformation of α-Helix and β-Strand Model Peptides on Self-Assembled Monolayers Using Sum Frequency Generation and NEXAFS Spectroscopies. Langmuir 2010, 26 (5), 3433–3440. 50. Nguyen, K. T.; Soong, R.; Im, S. C.; Waskell, L.; Ramamoorthy, A.; Chen, Z. Probing the Spontaneous Membrane Insertion of a Tail-Anchored Membrane Protein by Sum Frequency Generation Spectroscopy. J. Am. Chem. Soc. 2010, 132 (43), 15112–15115. 777 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


51. Baio, J. E.; Weidner, T.; Baugh, L.; Gamble, L. J.; Stayton, P. S.; Castner, D. G. Probing the Orientation of Electrostatically Immobilized Protein G B1 by Time-of-Flight Secondary Ion Spectrometry, Sum Frequency Generation, and Near-Edge X-ray Adsorption Fine Structure Spectroscopy. Langmuir 2012, 28 (4), 2107–2112. 52. Stöhr, J. NEXAFS Spectroscopy; Springer-Verlag: Berlin, 1992; Vol. 25. 53. Weidner, T.; Dubey, M.; Breen, N. F.; Ash, J.; Baio, J. E.; Jaye, C.; Fischer, D. A.; Drobny, G. P.; Castner, D. G. Direct Observation of Phenylalanine Orientations in Statherin Bound to Hydroxyapatite Surfaces. J. Am. Chem. Soc. 2012, 134 (21), 8750–8753. 54. Vickerman, J. C.; Brown, A. A.; Reed, N. M. Secondary ion mass spectrometry: principles and applications; Oxford University Press: New York, 1989. 55. Garrison, B. J.; Postawa, Z. Computational View of Surface Based Organic Mass Spectrometry. Mass Spectrom. Rev. 2008, 27, 289–315. 56. Jackson, J. E. Principal Components And Factor-Analysis .1. Principal Components. J. Qual. Technol. 1980, 12 (4), 201–213. 57. Graham, D. J.; Wagner, M. S.; Castner, D. G. Information from complexity: Challenges of TOF-SIMS data interpretation. Appl. Surf. Sci. 2006, 252 (19), 6860–6868. 58. Wagner, M. S.; Graham, D. J.; Ratner, B. D.; Castner, D. G. Maximizing information obtained from secondary ion mass spectra of organic thin films using multivariate analysis. Surf. Sci. 2004, 570 (1-2), 78–97. 59. Mantus, D. S.; Ratner, B. D.; Carlson, B. A.; Moulder, J. F. Static SecondaryIon Mass-Spectrometry of Adsorbed Proteins. Anal. Chem. 1993, 65 (10), 1431–1438. 60. Wagner, M. S.; Castner, D. G. Analysis of adsorbed proteins by static time-offlight secondary ion mass spectrometry. Appl. Surf. Sci. 2004, 231, 366–376. 61. Tidwell, C. D.; Castner, D. G.; Golledge, S. L.; Ratner, B. D.; Meyer, K.; Hagenhoff, B.; Benninghoven, A. Static ToF SIMS and XPS Characterization of Adsorbed Albumin and Fibronectin Films. Surf. Interface Anal. 2001, 31, 724–733. 62. Xia, N.; May, C. J.; McArthur, S. L.; Castner, D. G. Time-of-flight secondary ion mass spectrometry analysis of conformational changes in adsorbed protein films. Langmuir 2002, 18 (10), 4090–4097. 63. Lhoest, J. B.; Detrait, E.; van den Bosch de Aguilar, P.; Bertrand, P. Fibronectin adsorption, conformation, and orientation on polystyrene substrates studied by radiolabeling, XPS, and ToF SIMS. J. Biomed. Mater. Res. 1998, 41 (1), 95–103. 64. Lhoest, J. B.; Wagner, M. S.; Tidwell, C. D.; Castner, D. G. Characterization of adsorbed protein films by time of flight secondary ion mass spectrometry. J. Biomed. Mater. Res. 2001, 57 (3), 432–440. 65. Apte, J. S.; Collier, G.; Latour, R. A.; Gamble, L. J.; Castner, D. G. XPS and ToF-SIMS Investigation of alpha-Helical and beta-Strand Peptide Adsorption onto SAMs. Langmuir 2010, 26 (5), 3423–3432. 66. Weidner, T.; Breen, N. F.; Li, K.; Drobny, G. P.; Castner, D. G. Sum frequency generation and solid-state NMR study of the structure, orientation, 778 In Proteins at Interfaces III State of the Art 2012; Horbett, T., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2012.


and dynamics of polystyrene-adsorbed peptides. Proc. Natl. Acad. Sci. U.S.A. 2010, 107 (30), 13288–13293. 67. Ward, R. N.; Davies, P. B.; Bain, C. D. Orientation of Surfactants Adsorbed on a Hydrophobic Surface. J. Phys. Chem. 1993, 97 (28), 7141–7143. 68. Breen, N. F.; Weidner, T.; Li, K.; Castner, D. G.; Drobny, G. P. A Solid-State Deuterium NMR and Sum-Frequency Generation Study on the Side-Chain Dybamics of Peptides Adsorbed onto Surfaces. J. Am. Chem. Soc. 2009, 131 (40), 14148–14149. 69. Nishi, N.; Hobara, D.; Yamamoto, M.; Kakiuchi, T. Chain-length-dependent change in the structure of self-assembled monolayers of n-alkanethiols on Au(111) probed by broad-bandwidth sum frequency generation spectroscopy. J. Chem. Phys. 2003, 118 (4), 1904–1911.


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