和频光谱在生物膜等生物界面中的应用 In situ molecular level studies on membrane related peptides and proteins in real time using sum frequency generation vibrational spectroscopy.pdf

In situ molecular level studies on membrane related peptides and proteins in real time using sum frequency generation vibrational spectroscopy Shuji Ye, Khoi Tan Nguyen, Stéphanie V. Le Clair, Zhan Chen * Department of Chemistry, University of Michigan, 930 North University Ave., Ann Arbor, MI 48109, USA article info Article history: Received 4 November 2008 Received in revised form 11 March 2009 Accepted 13 March 2009 Available online 21 March 2009 Keywords: Sum frequency generation SFG Cell membrane Antimicrobial peptide Membrane protein Lipid bilayer Melittin G proteins Alamethicin Tachyplesin I abstract Sum frequency generation (SFG) vibrational spectroscopy has been demonstrated to be a powerful technique to study the molecular structures of surfaces and interfaces in different chemical environments. This review summarizes recent SFG studies on hybrid bilayer membranes and substrate-supported lipid monolayers and bilayers, the interaction between peptides/proteins and lipid monolayers/bilayers, and bilayer perturbation induced by peptides/proteins. To demonstrate the ability of SFG to determine the orientations of various secondary structures, studies on the interactions between different peptides/proteins (melittin, G proteins, alamethicin, and tachyplesin I) and lipid bilayers are discussed. Molecular level details revealed by SFG in these studies show that SFG can provide a unique understanding on the interactions between a lipid monolayer/bilayer and peptides/proteins in real time, in situ and without any exogenous labeling. 2009 Elsevier Inc. All rights reserved. 1. Introduction Cell membranes play a crucial role in many biological functions of cells. They govern all interactions between cells and their environments, such as the exchange of information and ions/molecules between the inside and outside of the cells. The cell membrane can be quite complex and dynamic, with a thickness of several nanometers. A cell membrane mainly consists of a lipid bilayer, which is an arrangement of lipids in which the acyl chains of each layer interact through hydrophobic interactions and the hydrophilic head groups face the inside and outside of the cell. A variety of peptides and proteins are also embedded inside or associated with the cell membranes, helping to fulfill various cellular functions (Katsaras and Gutberlet, 2001; Mateo et al., 2006; Yeagle, 2005). It is important to study the structures and kinetics of membrane embedded/associated peptides/proteins in order to understand their functions. Results from these types of studies can help in the rational design of molecules that can more effectively mediate or interfere with various cellular events in the desired manner. Different experimental tools have been used to study peptides/ proteins in the membrane environment, and excellent results have been obtained. For example, surface plasmon resonance spectroscopy measurements allow for the determination of the peptide/ protein coverage on membranes (Besenicˇar et al., 2006; Devanathan et al., 2006; Salamon et al., 1997). Neutron reflection studies have also permitted the determination of the amount of adsorbed peptides/proteins (Fragneto-Cusani, 2001; Haas et al., 2007; Kucˇerka et al., 2007). Atomic force microscopy techniques, on the other hand, are very useful in studying the packing and surface ordering of membrane-bound proteins (Alessandrini and Facci, 2005; Brasseur et al., 2008; Engel and Gaub, 2008; Johnston, 2007; Laflamme et al., 2008; Richter et al., 2006). Ellipsometry, in contrast, can be utilized to follow proteins’ adsorption kinetics onto membranes and determine the adsorbed protein film thickness (Faiss et al., 2008). Excellent review articles have been published that summarize the applications of these techniques (and others) to the study of the interactions between peptides/proteins and lipid membranes (Cabiaux, 2004; Lee, 2005; McIntosh and Simon, 2006). Even though some molecular level information can be acquired by probing the interactions between cell membranes and proteins/peptides using these analytical tools, further details regarding such molecular interactions need to be elucidated. Several spectroscopic techniques that can probe detailed structural information have been applied to study membrane-related peptides and proteins, leading to in-depth understanding of such molecular interactions. For example, Attenuated Total Reflectance Fourier Transform Infrared (ATR-FTIR) measurements can provide 1047-8477/$ - see front matter 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.jsb.2009.03.006 * Corresponding author. Fax: +1 734 647 4865. E-mail address: zhanc@umich.edu (Z. Chen). Journal of Structural Biology 168 (2009) 61–77 Contents lists available at ScienceDirect Journal of Structural Biology journal homepage: www.elsevier.com/locate/yjsbi

vibrational spectra (or fingerprints) of membrane proteins and peptides. However, the technique may suffer from a lack of intrinsic surface sensitivity and therefore the signals from the molecules in the cell membrane environment might be confused with those in the bulk environment, e.g., in the bulk solution. Also, ATR-FTIR only gives one vibrational measurable of the tilt angle <cos2 h> for orientation determination of a membrane protein or peptide (Tamm and Tatulian, 1997). Nuclear magnetic resonance (NMR) spectroscopy is another powerful spectroscopic technique that is widely used to characterize peptide/protein-lipid interactions (Bader et al., 2003; Bechinger, 1999, 2004; Lindblom and Grobner, 2006; Naito and Kawamura, 2007; Wang, 2008). Solution and solid-state NMR have been successfully used to determine the sitespecific secondary structures and dynamics of membrane-bound proteins (Andronesi et al., 2005; Dürr et al., 2007; Fernández et al., 2001). Both solution and solid-state NMR can also be used to study several aspects of the lipid-protein interactions, such as looking at the proteins’ effects on the lipid dynamics and determining which parts of the protein are interacting with the head group or tail region of the lipids (Dvinskikh et al., 2007; Lee et al., 2008). However, due to NMR’s low sensitivity, large amounts of protein/ peptide are required (milligram quantities), sometimes isotope-labeled proteins are needed, and experimental time can be very long. Difficulties can also be encountered in sample preparation (i.e., high concentrations of sample can cause membrane proteins to aggregate). The different model membranes used in NMR also make it complicated to create an asymmetric lipid bilayer (i.e., controlling the composition of each leaflet). Over the last two decades, sum frequency generation (SFG) vibrational spectroscopy has been developed into a very powerful and highly versatile spectroscopic tool for surface and interfacial studies (Anglin and Conboy, 2008; Bain, 1995; Baldelli, 2008; Belkin and Shen, 2005; Chen, 2007; Dreesen et al., 2004a,b; Fourkas et al., 2007; Gautam and Dhinojwala, 2002; Gopalakrishnan et al., 2006; Gracias et al., 1999; Holman et al., 2004; Hopkins et al., 2005; Iwahashi et al., 2008; Kim et al., 2008; Koffas et al., 2004; Li et al., 2008; Ma and Allen, 2006; Moore and Richmond, 2008; Opdahl et al., 2004; Richmond, 2002; Rupprechter and Weilach, 2008; Shen and Ostroverkhov, 2006; Shultz et al., 2002; Stiopkin et al., 2008; Voges et al., 2007; Yang et al.,2002; Ye et al., 2008). SFG is a vibrational technique that is intrinsically surface-sensitive, requires small amounts of sample, and with which the experiments can be done in situ and in real-time (Allen et al., 2000; Bain, 1995; Belkin and Shen, 2005; Buck and Himmelhaus, 2001; Chen, 2007; Chen and Chen, 2006; Chen et al., 2002b, 2005a; Eisenthal, 1992; Gopalakrishnan et al., 2006; Gracias et al., 1999; Hopkins et al., 2005; Koffas et al., 2004; Lambert et al., 2005; Miranda and Shen, 1999; Moore and Richmond, 2008; Opdahl et al., 2004; Perry et al., 2006; Richmond, 2001, 2002; Shen, 1989; Shen and Ostroverkhov, 2006; Shultz et al., 2000; Tadjeddine and Peremans, 1996; Wang et al., 2005a; Williams and Beattie, 2002; Zhuang and Shen, 1996). SFG permits the identification of interfacial molecular species (or chemical groups), and also provides information about the interfacial structure, such as the orientation and the orientation distribution of functional groups on the surface. SFG has been applied to study the structure and orientation of biomolecules, such as lipids (Anderson et al., 2006; Anglin et al., 2007; Anglin and Conboy, 2008; Chen et al., 2007b; Doyle et al., 2004; Harper and Allen, 2007; Kim et al., 2001; Levy and Briggman, 2007; Liu and Conboy, 2004a,b, 2005a,b, 2007; Lobau et al., 1999; Ma and Allen, 2006, 2007; Nickolov et al., 2006; Ohe et al., 2004; PetralliMallow et al., 1999; Sovago et al., 2007; Watry et al., 2003; White et al., 2006), and peptides/proteins (including membrane-related proteins/peptides) (Chen and Chen; 2006; Chen et al., 2005a,b; Clarke et al., 2005; Dreesen et al., 2004a,b; Evans-Nguyen et al., 2006; Humbert et al. 2006; Kim and Cremer, 2001; Kim et al., 2002, 2003; Kim and Somorjai, 2003; Knoesen et al., 2004; Mermut et al., 2006; Rocha-Mendoza et al., 2007; Sartenaer et al., 2007; Wang et al., 2003a,b,c, 2005b, 2006; York et al., 2008). Planar substrate-supported lipid bilayers have been widely used as a model to mimic cell membranes. Their suitability for biological studies has been extensively tested. These lipid bilayers are readily prepared by directly depositing lipid monolayers or bilayers onto the substrates such as glass, mica, quartz, and silicon surfaces using Langmuir-Blodgett method or vesicle fusion method (Kalb et al., 1992; Steinem et al., 2000; Tamm and McConnell, 1985; Tamm, 1988; Tamm and Tatulian, 1997; Thompson and Palmer, 1988). In addition, many research groups are also employing different strategies to improve the properties of supported lipid bilayers (i.e., using ultrathin polymer to support lipid bilayers) (Sackmann, 1996; Tanaka and Sackmann, 2005; Zhao and Tamm, 2003). Previous studies have indicated that planar substrate-supported lipid bilayers can offer several advantages over other model membranes, such as free-standing lipid bilayers, solvent-free lipid bilayers, or phospholipid vesicles. Planar substrate-supported lipid bilayers are unilamellar and geometrically well defined. They can maintain excellent mechanical stability without losing their fluid nature. These advantages of substrate-supported bilayers make it possible to carry out experiments that probe structural and dynamic properties of membranes and protein-lipid interactions, using the surface analytical techniques mentioned above (Castellana and Cremer, 2006; Kalb et al., 1992; McConnell et al., 1986; Sackmann, 1996; Tamm and McConnell, 1985; Tamm, 1988; Tamm and Tatulian, 1997; Tanaka and Sackmann, 2005). In this paper, we will first present a brief introduction of the theoretical background needed to understand SFG, and then summarize recent studies on the interactions between lipid membranes (monolayers and bilayers, focusing especially on substrate-supported lipid bilayers) and biomolecules monitored by SFG in real time and in situ. The names of lipids mentioned in this paper and their respective abbreviations are listed in Table 1. 2. Theoretical background of SFG Although it has only been about twenty years since the first SFG spectra were recorded by Shen (Guyotsionnest et al., 1987; Hunt et al., 1987; Zhu et al., 1987), SFG is now being used by a growing number of research groups for a variety of applications, including polymer and biological interface studies. Many excellent review Table 1 The full names of lipids and their abbreviations that appear in this paper. Abbr. Full Name POPC 1-Palmitoyl-2-Oleoyl-sn-Glycero-3-Phosphocholine POPG 1-Palmitoyl-2-Oleoyl-sn-Glycero-3-[Phospho-rac-(1-glycerol)] (Sodium Salt) DMPC 1,2-Dimyristoyl-sn-Glycero-3-Phosphocholine d-DMPC 1,2-Dimyristoyl-D54-sn-Glycero-3-Phosphocholine-1,1,2, 2-D4-N,N,N-trimethyl-D9 DMPC-d54 1,2-Dimyristoyl-D54-sn-Glycero-3-Phosphocholine DPPC 1,2-Dipalmitoyl-sn-Glycero-3-Phosphocholine d-DPPC 1,2-Dipalmitoyl-D62-sn-Glycero-3-Phosphocholine-1,1,2, 2-D4-N,N,N-trimethyl-D9 DPPG 1,2-Dipalmitoyl-sn-Glycero-3-[Phospho-rac-(1-glycerol)] (Sodium Salt) d-DPPG 1,2-Dipalmitoyl-D62-sn-Glycero-3-[Phospho-rac-(1-glycerol)] (Sodium Salt) DSPC 1,2-Distearoyl-sn-Glycero-3-Phosphocholine d-DSPC or DSPCd83 1,2-Distearoyl-D70-sn-Glycero-3-Phosphocholine-1,1,2, 2-D4-N,N,N-trimethyl-D9 DSPC-d70 1,2-Distearoyl-D70-sn-Glycero-3-Phosphocholine DSPG 1,2-Distearoyl-sn-Glycero-3-[Phospho-rac-(1-glycerol)] (Sodium Salt) 62 S. Ye et al. / Journal of Structural Biology 168 (2009) 61–77

papers have summarized details about SFG theory and research (Allen et al., 2000; Bain, 1995; Belkin and Shen, 2005; Buck and Himmelhaus, 2001; Chen, 2007; Chen and Chen, 2006; Chen et al., 2002b, 2005a,; Eisenthal, 1992; Gopalakrishnan et al., 2006; Gracias et al., 1999; Hopkins et al., 2005; Koffas et al., 2004; Lambert et al., 2005; Miranda and Shen, 1999; Moore and Richmond, 2008; Opdahl et al., 2004; Perry et al., 2006; Richmond, 2001, 2002; Shen, 1989; Shen and Ostroverkhov, 2006; Shultz et al., 2000; Tadjeddine and Peremans, 1996; Wang et al., 2005a; Williams and Beattie, 2002; Zhuang and Shen, 1996), which will not be repeated here. In a typical SFG experiment, two pulsed laser beams, one with a fixed frequency in the visible frequency range (xVis) and one with a tunable frequency in the infrared frequency range (xIR), are overlapped spatially and temporally on the sample (Fig. 1A). The SFG signal is generated at the sum frequency of the two input beams by the nonlinear process, xSF = xVis + xIR. Therefore, the SFG process can be simply viewed as a combination of infrared (IR) absorbance and Raman scattering, as shown in the energy diagram in Fig. 1B. The intensity of the sum frequency signal is proportional to the square of the vibration’s second-order nonlinear susceptibility vð2Þ eff (Bain, 1995; Buck and Himmelhaus, 2001; Chen and Chen, 2006; Chen et al., 2002b, 2005a; Eisenthal, 1992; Lambert et al., 2005; Miranda and Shen, 1999; Richmond, 2001; Shen, 1984, 1989; Tadjeddine and Peremans, 1996; Wang et al., 2005a; Williams and Beattie, 2002; Zhuang et al., 1999). Different components of vð2Þ eff can be probed using different polarization combinations of the input and output laser beams. From such measurements, orientation information of surface molecules and functional groups can be deduced (Gautam and Dhinojwala, 2001; Hirose et al., 1992a,b, 1993; Kim and Somorjai, 2003; Shen, 1984; Wang et al., 2001). More details about SFG theory and data analysis can be found in Appendix A. The selection rules of SFG make the technique surface sensitive. As we stated above, the SFG signal intensity is proportional to the square of the vibration’s second-order nonlinear susceptibility vð2Þ eff . vð2Þ eff vanishes when a material has inversion symmetry under the dipole approximation (Bain, 1995; Buck and Himmelhaus, 2001; Chen and Chen, 2006; Chen et al., 2002b, 2005a; Eisenthal, 1992; Gautam and Dhinojwala, 2001; Hirose et al., 1992a,b, 1993; Lambert et al., 2005; Miranda and Shen, 1999; Richmond, 2001; Shen, 1984, 1989; Tadjeddine and Peremans, 1996; Wang et al., 2001, 2005a; Williams and Beattie, 2002; Zhuang et al., 1999). The majority of bulk materials exhibits inversion symmetry, thus they do not generate SFG signals. However, the presence of an interface causes a break in the symmetry, and therefore, molecules on surfaces or at interfaces can generate SFG signal. Therefore, SFG is an intrinsically surface/interface specific technique, different from linear vibrational spectroscopic techniques such as ATR-FTIR and Raman scattering. 3. Recent SFG studies on membrane-related peptides/proteins In this section, we will summarize recent SFG studies on membrane-related peptides and proteins. Such studies have demonstrated that SFG can provide a unique understanding of the interactions between a lipid monolayer/bilayer and peptides/proteins without any exogenous labeling, in real time and in situ. 3.1. SFG studies on lipid monolayers, substrate-supported lipid bilayers and hybrid bilayer membranes As mentioned above, it has been extensively shown that substrate-supported lipid bilayers are valid models for cell membranes (Castellana and Cremer, 2006; Kalb et al., 1992; McConnell et al., 1986; Sackmann, 1996; Steinem et al., 2000; Tamm and McConnell, 1985; Tamm, 1988; Tamm and Tatulian, 1997; Tanaka and Sackmann, 2005; Thompson and Palmer, 1988; Zhao and Tamm, 2003). SFG has been applied to investigate the structures of these lipid bilayer systems (Anglin et al., 2007; Anglin and Conboy, 2008; Chen et al., 2007b; Doyle et al., 2004; Levy and Briggman, 2007; Liu and Conboy, 2004a,b, 2005a,b, 2007; Petralli-Mallow et al., 1999), with a recent review article (Chen and Chen, 2006) summarizing these SFG studies. Here, we do not attempt to reiterate the details, but instead will give some brief discussions on how some fundamental questions regarding the interactions between peptides/proteins and lipid bilayers can be answered using SFG. For example, knowledge of the conformations of lipids in the bilayer, both before and after their interactions with antimicrobial peptides, aids in understanding the peptide’s mode of interaction. These modes of interaction can be generally separated into the carpet, barrel-stave and toroidal pore modes (Yang et al., 2001; Chen and Chen, 2006). Theoretically, each of these peptide/protein–lipid bilayer interaction modes should induce different conformational changes on the lipid bilayers, resulting in distinct SFG spectral changes. Therefore, using these kinds of SFG lipid studies, details regarding the interactions of antimicrobial peptides, and other peptides/proteins, with lipid systems may be elucidated. Before we go further to discuss the interactions between peptides/proteins and lipid bilayers, knowledge regarding the membrane model system, the lipid bilayer, is needed. When studying such interactions with SFG, substrate-supported lipid bilayers are used as the typical model for a cell membrane. Optical techniques using fluorescence labeling methods have previously been used to study the physical properties of such bilayers. Although these techniques offer important insights (e.g., about the flip-flop of lipids in a membrane, the transition temperature of a lipid system, and lipid domain segregation) (Bagatolli, 2006; Heberle et al., 2005; Lee, 2005), the incorporation of a fluorescent label on the lipids, or of a fluorescent probe inside the bilayer, may affect the behavior of ground state hυVis hυIR virtual state hυSF A B Fig. 1. SFG experimental geometry and energy diagram. (A) Total reflection geometry employed for the experiments described in Sections 3.3 and 3.4. A bilayer is immersed in a small reservoir with an approximate volume of 1.6–2.0 mL; (B) Simplified energy level diagram of vibrational sum frequency generation process. S. Ye et al. / Journal of Structural Biology 168 (2009) 61–77 63

the system and generate some artifacts in the experiments. An advantage of using SFG to study these types of supported bilayers is that it is not necessary to introduce any bulky labels. The only form of labeling required to study the bilayers is the introduction of isotope labeling. In order to monitor each leaflet of a lipid bilayer individually, it is necessary to have one leaflet deuterated, in order to break the inversion symmetry of the bilayer and obtain SFG signal. One important aspect of the behavior of lipids in cell membranes is their ability to translocate both laterally and across the membrane (flip-flop). With this in mind, Conboy and his colleagues have investigated some important physical properties of substratesupported lipid bilayers, such as the kinetics of the flip-flop and the transition temperatures of various lipid bilayers using SFG (Liu and Conboy, 2004a,b, 2005a,b). More recently, they investigated the asymmetric distribution of domains in lipid bilayers by carrying out SFG spectroscopic measurements of symmetric C–H stretching modes of the fatty acid methyl groups (Liu and Conboy, 2007). In this research, they correlated the intensity of the C–H symmetric stretch of the fatty acid methyl groups with the symmetry of the lipid bilayer, with the stronger intensity obtained when the bilayer became more asymmetric. The breakage in symmetry was caused by the dislocation of the gel and liquid-crystalline phase domains at the transition temperature. They have also done SFG studies on the lateral pressure dependence of the phospholipid transmembrane diffusion rate in supported lipid bilayers (Anglin and Conboy, 2008). Their results indicated that the kinetics of lipid flip-flop in these membranes show a strong lateral pressure dependence. Based on these data, they successfully determined the activation area for phospholipid flip-flop (Anglin and Conboy, 2008). Briggman and coworkers have employed hybrid bilayer membranes (HBM) as an alternative for supported lipid bilayers in their research (Anderson et al., 2004, 2006, 2007; Levy and Briggman, 2007; Petralli-Mallow et al., 1999). The HBM system studied has one single lipid layer deposited on top of a hydrophobic selfassembled monolayer (SAM). Briggman’s group has extensively studied the properties of HBMs and found that the transition temperature of the lipid layer depends greatly on the packing density and the crystallinity of the SAM layer underneath it (Anderson et al., 2007). They also carried out SFG studies on the effect of cholesterol on phospholipids (Levy and Briggman, 2007). Their research has shown that HBMs could be used as excellent model membranes for biological studies on peripherally-bound proteins. However, it is challenging to use HBMs as cell membrane models to study lipid flip-flop and some transmembrane proteins that extensively interact with the inner-leaflet. Many research groups also model cell membranes by using lipid monolayers. The fundamental properties such as transition temperatures and effects of cholesterol on monolayers have been investigated using SFG (Bonn et al., 2004; Ohe et al., 2007a,b; Roke et al., 2003). 3.2. SFG studies on the interactions between peptides/proteins and lipid monolayers/bilayers: C–H stretching frequency region Early SFG studies on the interactions between proteins/peptides and lipid monolayers, HBMs and supported lipid bilayers focused on the C–H stretching frequency region. Table 2 contains a list of some of those studies. One of the very first SFG studies on the interaction between proteins and lipid monolayers was done by Cremer and his colleagues (Kim et al., 2003). They investigated the orientation of gramicidin A in a DMPC monolayer using the SFG C–H stretching signals generated from the side chains of gramicidin A. Their results indicated that the orientation of gramicidin A was concentration dependent (Kim et al., 2003). Conboy and coworkers studied the effect of gramicidin A on the flip-flop of DSPC lipids in a substrate-supported phospholipid bilayer (Anglin et al., 2007). Using SFG and ATR-FTIR, they showed that gramicidin A induced rapid flip-flop of the DSPC lipids. In this study, the C–H stretching signals of the lipid’s hydrophobic chain were studied by SFG and the amide signal of gramicidin A was observed by ATR-FTIR. The detection of amide signal from the peptide using ATR-FTIR indicated that gramidicin A was bound to the bilayer. SFG signal of the lipid C–H stretches could be observed because an asymmetric bilayer (with one leaflet deuterated) was used in the study. The time-dependence of the SFG C–H signals can be used to monitor the flip-flop rate, because as the system becomes more symmetric (from interchange between outer- and inner-leaflet lipids), the signal should decrease. By studying the time-dependent C–H signals both with and without gramicidin A bound, they showed that gramicidin A induced a faster flip-flop rate than when no gramicidin A was added to the bilayer. Neivandt and coworkers examined a protein in the fibroblast growth factor (FGF) family, FGF 1, and its interaction with an HBM (Doyle et al., 2004). Using SFG, they showed that this protein caused deformation of the DSPG lipid layer even at very low concentrations. They also found that this process was reversible to a certain extent when the protein was washed off the HBM. Itoh and his colleagues investigated the interactions between antibiotic polymyxin B (PMB) and monolayers composed of DPPG, as well as DPPC (Ohe et al., 2004). By monitoring the SFG C–H stretching signals of the lipids’ side chains and the O–H stretching signals of water molecules, along with the measurements of the pressurearea isotherms, they found that this antibiotic peptide bound to the negatively charged lipids (DPPG), but not to the neutral lipids (DPPC). They also showed that the binding of PMB to the lipid monolayer affected the phase transition of the lipid monolayer. Because SFG can provide structural information such as functional group composition, orientation, and ordering at a surface or interface with a submonolayer sensitivity, it has also been applied to determine the average minimal inhibitory concentration (MIC) of antimicrobial peptides (or analogue oligomers) in membranes. Chen et al. used SFG to investigate the molecular interactions between a small antimicrobial oligomer and a single substrate-supported lipid bilayer with a hydrogenated leaflet (DPPG) and a deuterated leaflet (d-DPPG) (Chen et al., 2006). It was observed from the C–H and C–D stretching signals that the distal leaflet was disrupted at a very low peptide/lipid ratio, while the proximal leaflet remained intact below a threshold concentration very close to the MIC (0.8 mg/mL) value. The orientation of this oligomer was deduced by SFG C–H signals and the results indicated that the small antimicrobial oligomer acted as a ‘‘molecular knife” by disrupting primarily the outer leaflet of the bilayer at lower concentrations and further inserting into the entire bilayer at a certain critical (MIC) concentration. 3.3. SFG studies on the interaction between peptides/proteins and lipid bilayers: amide I frequency region 3.3.1. Detection of SFG amide I signal The SFG results presented in the previous section are obtained from SFG signals acquired mainly in the C–H stretching frequency region, which are comprised of signals from the lipids and protein side chains. The feasibility of detecting SFG amide I signals from interfacial proteins/peptides was demonstrated by Chen and his coworkers in 2003 (Wang et al., 2003b). Because the water bending mode does not contribute noticeable SFG signals, it is not necessary to perform a background subtraction to obtain the SFG amide I signals from interfacial proteins and peptides; instead, they can be detected directly (Clarke et al., 2005; Wang et al., 2003b, 2005c). In addition, proteins in the bulk solution do not generate SFG sig- 64 S. Ye et al. / Journal of Structural Biology 168 (2009) 61–77

nals, and thus SFG can selectively probe interfacial proteins/peptides. Moreover, SFG is able to provide more measurements than ATR-FTIR in studying the orientation of interfacial proteins/peptides. In SFG studies, our group has also adopted a near total reflection experimental geometry (Fig. 1A) that enables us to obtain very strong SFG amide I signals of interfacial proteins, which makes the data analysis easier and more accurate (Wang et al., 2003b). SFG amide I signals of proteins can be affected by the surface coverage, orientation, and secondary structures of the adsorbed proteins (Clarke et al., 2005; Wang et al., 2003b, 2005c). The amide I mode contains predominately the peptide C@O stretching bands. These C@O groups are held together by hydrogen bonds within the secondary structures and the frequency of the C@O stretch depends heavily on its hydrogen-bonded environment. The peak center of the amide I band, therefore, depends on the secondary structure adopted by the peptide/protein. The Chen group has successfully demonstrated that SFG amide I signals can be used to distinguish a-helical and b-sheet structures of peptides and proteins on polymers and lipid bilayers (Chen and Chen, 2006; Chen et al., 2005b, 2007a,c; Clarke et al., 2005; Wang et al., 2003b, 2005b,c, 2007, 2008). Recently, Somorjai’s group developed a new optical parametric amplifier (OPA) in the SFG spectrometer that can also create a tunable infrared light between 1500 and 2000 cm1 ; this IR beam can be used to study the amide I signals of interfacial peptides (York et al., 2008). Based on previous FTIR and Raman studies, SFG amide signals of different secondary structures can be assigned. Using antimicrobial peptides that are known to adopt a-helical structures in lipid bilayers, Chen et al. were able to detect SFG amide I signals of ahelical structures centered at around 1650 cm1 (Chen et al., 2007c). Using model b-sheet peptide tachyplesin I, Chen et al. also showed that SFG amide I signals from a b-sheet structure has characteristic peaks at 1635 and 1685 cm1 , corresponding to the B2 mode and B1/B3 modes, respectively (Chen and Chen, 2006). Recent results in our laboratory on alamethicin indicate that SFG amide I signals of 310 helical structures have peaks at around 1635 and 1670 cm1 . These results are indeed in good agreement with previous FTIR and Raman studies. More details regarding the interfacial structures of different secondary structure domains of proteins/peptides can be obtained by the careful data analysis of SFG amide I signals collected using different polarization combinations, which will be presented below. 3.3.2. SFG studies on a-helical proteins/peptides in lipid bilayers 3.3.2.1. Data analysis for orientation determination using amide I band. As mentioned above, it has been shown experimentally that amide I peak centers are different for each type of secondary structure. Therefore, analyses of amide I signals provide insights into the protein backbone secondary structures. This section will focus on how to deduce the orientation of a-helical structures. Conventional polarized ATR-FTIR has been widely applied to study the orientation of a-helical structures of membrane peptides/proteins (Tamm and Tatulian, 1997). In ATR-FTIR studies, the tilt angle of the ahelices can be calculated from the order parameter (Sh), which is defined as: Sh ¼ 3 < cos2 h > 1 2 ; ð1Þ with h being the tilt angle between the helix’s principal axis and the surface normal. The bracket denotes the time and ensemble average. Theoretically, <cos2 h> can be determined from the measured intensity ratio in ATR-FTIR using p- and s-polarized IR light (Tamm and Tatulian, 1997). If we assume h to have the simplest delta distribution, the orientation of the helix can be determined from this intensity ratio. The orientation in reality can be more complicated and such a simple distribution may not be adequate enough to describe the orientation of the peptide/protein. For example, a helix may adopt two different orientations or have a broad orientation distribution. To characterize such a more complicated orientation distribution, more measured parameters are needed. For example, when the parameter Sh approaches zero, <cos2 h> is equal to 1/3, and thus there is always ambiguity in whether all helices have the same tilt angle around 54.7, or a completely random orientation, or other orientations/distributions in between. In order to determine which of the three cases is correct, more measurements using different spectroscopic methods would be required (Chen et al., 2007c). It has been shown that it is possible to deduce the orientation of functional groups, such as methyl, methylene, phenyl, and carbonyl, by measuring different tensor elements using SFG spectra collected with different polarization combinations (Briggman et al., 2001; Gautam et al., 2000; Hirose et al., 1992a,b, 1993; Oh-e et al., 2002; Tyrode et al., 2005). By analyzing the SFG amide I signals using group theory and projection operators, the Chen group has been able to investigate interfacial protein structures and deduce the orientation of a-helical peptides (Lee et al., 2006; Wang et al., 2007, 2008). They showed that both the amide I A mode and amide I E1 mode of an a-helix can contribute to SFG signals (Wang et al., 2008). Using the near total internal reflection geometry (Wang et al., 2003b), ssp and ppp amide I signals can be collected. These signals are mainly due to contributions from the vyyz and vzzz susceptibility components, respectively. The dependence of vyyz and vzzz susceptibility components on the molecular hyperpolarizability is described by the following equations: For the A mode: vA;xxz ¼ vA;yyz ¼ 1 2Ns ð1 þ rÞ < cos h > ð1 rÞ < cos3 h > bccc ð2Þ vA;zzz ¼ Ns r < cos h > þð1 rÞ < cos3 h > bccc ð3Þ For the E1 mode: vE1;xxz ¼ vE1;yyz ¼ Ns < cos h > < cos3 h > baca ð4Þ vE1;zzz ¼ 2Ns < cos h > < cos3 h > baca ð5Þ Table 2 SFG studies on the interactions between proteins/peptides and lipid monolayers, HBMs and supported lipid bilayers. Peptides or Proteins Membrane Studied region Reference Gramicidin A DMPC-d54 monolayer CH Kim et al. (2003) Polymyxin B DPPC monolayer, DPPG monolayer CH Ohe et al. (2004) FGF-1 DSPG hybrid bilayer membranes CH Doyle et al. (2004) Antimicrobial Oligomers DPPG/d-DPPG bilayer CH, CD, Amide I Chen et al. (2006) Antimicrobial peptides DPPG/d-DPPG bilayer CH, CD, Amide I Chen and Chen (2006) Gramicidin A DSPC/DSPC-d70 bilayer, DSPC-d83/DSPC-d83 bilayer CH, CD Anglin et al. (2007) Melittin dDPPG/dDPPG bilayer, dDPPG/DPPG bilayer CH, CD Chen et al., 2007b Melittin DPPG/DPPG Amide I Chen et al. (2007c) Heterotrimeric G Protein bc Subunit POPC/POPC bilayer, POPC/POPG bilayer Amide I Chen et al., 2007a S. Ye et al. / Journal of Structural Biology 168 (2009) 61–77 65