ADVANCED FTIR SPECTROSCOPY OF PROTEIN STRUCTURAL DYNAMICS By ANUPAMA JAGADEESH THUBAGERE Bachelor of Engineering Visveswaraiah Technological University Bangalore, India 2005 Submitted to the Faculty of the Graduate College of the Oklahoma State University in partial fulfillment of the requirements for the Degree of MASTER OF SCIENCE December, 2007 ADVANCED FTIR SPECTROSCOPY OF PROTEIN STRUCTURAL DYNAMICS Thesis Approved: Dr. Aihua Xie Thesis Adviser Dr. Bruce Ackerson Dr. Bret Flanders Dr. Yin Guo Dr. A. Gordon Emslie Dean of the Graduate College ii ACKNOWLEDGEMENTS I wish to express my gratitude towards the advice, support and encouragement given by Dr. Aihua Xie during past two years. Her patience and kindness have guided me through my course study, research, and my thesis. I would also like to thank Dr. Bret Flanders, Dr. Yin Guo and Dr. Bruce Ackerson for serving on my committee and reviewing this thesis and Dr. Paul Westhaus for his comments and advice through the course of my graduate studies. I wish to extend my appreciation to Dr. Beining Nie for her invaluable guidance and help for completing the computational work analyzed in this thesis, Sandip Kaledhonkar for helping me collect the rapid scan FTIR data and to Edward Manda for his help with the Gaussian calculations. I want to extend my special regards to Dr. Lorand Kelemen who initiated the salt dependence studies on PYP. I also wish to acknowledge the friendship, help and support of many graduate students and department secretaries, especially including Susan Cantrell, Cindi Raymond, Warren Grider, Karthik Bhatt, Nikhil Mirjhankar, Ratnakar Deole, Sayali Saykhedkar and Purvi Patel. Finally, I would like to thank my parents, my brother and my dear companion Ashwin Gopinath for their understanding, love and encouragement. iii TABLE OF CONTENTS Chapter Page I. INTRODUCTION ................................................................................................. 1 1.1 Unraveling protein structure and its structural dynamics ...................................... 1 1.2 Experimental methods for protein structure determination ................................... 7 X ray Crystallography.................................................................................... 7 Nuclear Magnetic Resonance spectroscopy.................................................... 9 Fourier Transform Infrared spectroscopy .................................................... 13 1.3 FTIR spectroscopy for studying protein structural dynamics.............................. 16 1.4 From Infrared Spectra to Protein Structure......................................................... 17 II. IMPACT OF SALTS ON THE STRUCTURAL DYNAMICS OF PHOTOACTIVE YELLOW PROTEIN 2.1 The Hofmeister salts .......................................................................................... 20 Molecular properties of water....................................................................... 21 Solvent properties of Water.......................................................................... 23 Hydration shells of Ions ............................................................................... 24 Effect of hydrated ions on proteins............................................................... 28 The Photocycle of PYP ................................................................................ 29 2.2 Materials and Methods....................................................................................... 36 2.3 Results and Discussion ...................................................................................... 39 Salts affect the O-H stretching band of Water............................................... 39 Effect of salts on the photocycle of PYP ...................................................... 41 Light induced FTIR difference spectra of PYP ............................................. 48 -Suppressed structural dynamics.......................................................... 51 -Altered proton transfer pathway ......................................................... 54 Effect of Hofmeister cations on PYP............................................................ 55 2.4 Conclusions ....................................................................................................... 59 III. VIBRATIONAL STRUCTURAL MAKERS FOR STRUCTURAL CHARACTERIZATION OF TYROSINE IN PROTEINS 3.1 Introduction....................................................................................................... 62 3.2 Tyrosine in the active site of proteins................................................................. 63 3.3 Vibrational Structural Markers........................................................................... 69 iv 3.4 Methods and materials ....................................................................................... 71 Computational method....................................................................................... 71 Experimental method......................................................................................... 73 3.5 Results............................................................................................................... 74 Locating bands sensitive to hydrogen bonding interactions ................................ 75 3.6 Discussion ......................................................................................................... 77 2-dimentional Infrared spectroscopy .................................................................. 91 3.7 Conclusions ....................................................................................................... 99 IV. SUMMARY AND REMARKS ....................................................................... 100 4.1 Hofmeister series and protein structural dynamics ........................................... 100 4.2 Vibrational structural marker for hydrogen bonding of tyrosine ....................... 102 REFERENCES...................................................................................................... 106 APPENDIX........................................................................................................... 119 v LIST OF TABLES Table Page Table 2.1 The ionic radius and hydration number of anions and cations from the Hofmeister series ....................................................................... 26 Table 3.1 Calculated hydrogen-bonding properties of a phenol group interacting with neutral groups.................................................................82 Table 3.2 Calculated hydrogen-bonding properties of a phenol group interacting with positively charged groups...............................................88 Table 3.3 Calculated hydrogen-bonding properties of a phenol group interacting with negatively charged group.................................................89 Table 3.5 Vibrational modes of C-O stretching and O-H bending of p-cresol in different solvents .................................................................... 96 Table 3.6 Correrelations between the C-O stretching frequency and the hydrogen-bonding status of tyrosine side-chain groups in proteins......... 98 vi LIST OF FIGURES Figure Page Figure 1.1 The chromophore of GFP ......................................................................... 4 Figure 1.2 Hydrogen bonding interaction in proteins ................................................. 5 Figure 1.3 The chromophore binding pocket of PYP.................................................. 6 Figure 1.4 Number of protein structures deposited in the protein data bank................ 8 Figure 1.5 Structural representations of X-ray crystals of Myoglobin, GFP and Bacteriorhodopsin................................................................... 9 Figure 1.6 Structural representation of NMR structures tendamistat and PYP .............................................................................................. 10 Figure 1.7 Vibrations responsible for Amide I and Amide II in FTIR Spectra ................................................................................................ 14 Figure 1.8 Layout of time-resolved FTIR rapid-scan spectroscopy........................... 16 Figure 2.1 Representation of the structure of a single water molecule ...................... 22 Figure 2.2 Hydrogen bonding of water molecules .................................................... 22 Figure 2.3 Representation of dissolution of salt ions in water................................... 23 Figure 2.4 The complete hydration shell of Na+ ion ................................................. 24 Figure 2.5 Listing of the Hofmeister series ............................................................. 28 Figure 2.6 Structural representations of PYP, SR II and Rhodopsin from the protein data bank .............................................................................. 30 Figure 2.7 WT PYP UV/vis spectrum...................................................................... 35 Figure 2.8 Schematic representation of photocycle of PYP ...................................... 33 Figure 2.9 FTIR data of υO-H frequency of water with high salt concentration ................................................................................... 41 Figure 2.10 UV/vis data of WT PYP in 4.0 M NaCl solution ................................... 45 Figure 2.11 FTIR absorption spectra of WT PYP in varying NaCl concentrations .............................................................................. 47 Figure 2.12 Comparison of pB-pG spectra of PYP in solution, crystal and 4.0 M NaCl.................................................................................... 49 Figure 2.13 Comparison of pB-pG spectra of WT PYP in varying NaCl concentrations ............................................................................. 51 Figure 2.14 Comparison in 3-D of pB-pG spectra in varying NaCl concentrations ..................................................................................... 53 Figure 2.15 Representation of the pCA chromophore of PYP showing the hydrogen bonding interactions.............................................................. 54 Figure 2.16 Comparison of pB-pG spectra of WT PYP in 0.0 M salt with 4.0 M CsCl, 4.0 M KCl, 4.0 M NaCl and 4.0 M LiCl ........................... 56 vii Figure 2.17 Comparison in 3-D of pB-pG spectra of WT PYP in 4.0 M CsCl, 4.0 M KCl, 4.0 M NaCl and 4.0 M LiCl ............................ 57 Figure 2.18 Comparison of the Hofmeister ions based on their ionic radii and their effect on bulk water ................................................................ 58 Figure 3.1 Representation of the tyrosine molecule.................................................. 63 Figure 3.2 Structural representation of PYP, GFP and bacteriorhodopsin from the protein data bank...................................................................... 67 Figure 3.3 Representation of the different hydrogen bonding properties of tyrosine ............................................................................................... 70 Figure 3.4 Representation of the model compound for Gaussian calculations of vibrational frequencies of tyrosine..................................................... 71 Figure 3.5 ................................................................................................................ 73 Figure 3.6 The calculated vibrational frequencies of υO-H and υC-O and δCOH of tyrosine when hydrogen bonded with H2O......................... 76 Figure 3.7 FTIR data representing the experimental evidence of the υC-O and δCOH frequency of tyrosine.................................................... 77 Figure 3.8 2-D correlations of the υC-O and δCOH modes and 2-D correlations of υC-O and υO-H frequency of tyrosine ............................. 93 Figure 3.9 Experimental evidence for 2-D correlations of υC-O and δCOH modes of tyrosine hydrogen bonding in proteins. .................. 94 viii Chapter I Introduction 1.1 Unraveling Protein Structure and its Structural Dynamics The three-dimensional structure of a protein is crucial to its function (Creighton 1993). Protein structure is essential for correct function because it allows molecular recognition (Whisstock and Lesk 2003). Even though proteins are all composed of only 20 amino acids, each protein has a 3 dimensional native structure (physiologically folded) that is specified by its primary structure, so that it has a unique set of characteristics (Voet and Voet 2004). The 3D structure of a protein and its ability to carry out its correct biological function are very tightly linked such that small structural defects can lead to a number of protein folding diseases (Carugo 2007). These include genetic diseases such as cystic fibrosis and sickle cell anemia, which are caused by single residue deletion and mutation respectively, rendering the protein incapable of its normal function (Baker 2000, Caughey and Lansbury 2003). Misfolded proteins are also the cause of several neurodegenerative diseases like the Alzheimer's disease which is mainly caused due to the accumulation of abnormally folded A-beta in the human brain (Dobson 1999, Dobson 2002). Although amyloid beta monomers are soluble and harmless, they undergo a dramatic conformational change at sufficiently high concentration to form a beta sheet-rich tertiary structure that aggregates to form amyloid fibrils that deposit outside neurons in dense formations causing dementia (Ohnishi and Takano 2004). 1 Similarly, deposits of aggregated prion protein are observed in brain tissues from humans when the healthy PrP proteins in the nerve cells are mutated due to the presence of infected prions causing the mad cow’s disease (Murphy 2002). Although found in cattle, this infectious protein can be transmitted to humans through consumption of the infected meat. Hence, understanding the structure-function relations of proteins is the key to understanding theses ailments. The protein 3-D structure not only gives us an insight into the functioning of individual proteins but helps us classify them into families, giving us a better understanding of their structure-function relationship. For example, hemoglobin, myoglobin and cytochorme c all belong to the family of globular transport proteins (Creighton 1993). The means by which these transport proteins recognize the molecules they carry is a knowledge that is procured from their 3D structure (Frauenfelder and Wolynes 1985). G –protein coupled receptors also known 7-transmembrane receptors are a large family of transmembrane receptor proteins that activate cellular responses to an external stimulus. Some proteins from this family instigate visual response (rhodopsin); some are sensory signal mediators (adenosine), and some regulate the immune system (Dohlman, Thorner et al. 1991). Another family is the PAS domain proteins which are signaling modules monitoring changes in light, oxygen and over all energy of a cell. To date more than 2000 proteins have been identified as belonging to this family (Taylor and Zhulin 1999). It is also equally important to understand the protein-protein interactions, particularly the formation of multi-protein complexes which influence the functional integrity of the cell (Sanchez and Sali 1997). Protein-protein interactions are crucial to 2 understanding the overall communication network in the biological system. Communication amongst cells is very important and signaling proteins are their means to communicate. This signaling takes place when the protein undergoes conformational change (amongst other changes) that influences its interaction with the neighboring protein. Proteins are dynamic and not static systems (Adam and Gibbs 1965). The structural dynamics of protein dictates its function (Creighton 1993). The conformational changes of a protein need to be addressed as a critical issue to understand its functions. Even though a single protein executes a large number of motions, not all of them are coupled to its function and it becomes imperative to pick the important conformational changes that lead to functioning (Frauenfelder, Sligar et al. 1991). Also, proteins need a medium, a solvent to function. In the presence of some additional elements in the solvent, the conformational changes and hence the structural dynamics of the proteins can get affected (Fenimore, Frauenfelder et al. 2004) Proton transfer reactions in proteins Protons are the most mobile atoms in proteins and even though they are covalently bonded, their movement is not restricted. Proton transfer reactions also lead to changes in electrostatic properties of proteins thus underlining their importance in biological process (Nagle and Tristram-Nagle 1983). The proton transfer event is dependent on the affinity of the proton to stay with the donor or the acceptor. The proton prefers to stay with the structure that has lower energy. The proton is a strong acidic species and it readily interacts with basic groups of amino acids in proteins. If these sites are appropriately localized, a proton transfer can 3 occur between them. The interactions are stronger with the involvement of highly basic side chain amino acids (arginine, lysine and histidine). These basic groups are normally protonated and are often a part of a bridge that stabilizes the tertiary structure of proteins. However, proton transfer reactions involving lesser basic/acidic amino acids (tyrosine, glutamic acid and aspartic acid) also play an important role. Proton transfer is a mechanism used by many proteins for information transfer (Kulhanek, Schlag et al. 2003). In proteins like the Green Fluorescent Protein (discussed below), protonation of the chromophore is the key mechanism for signaling. Figure 1.1: The chromophore of GFP consisting of Ser65-Tyr66Gly67. GFP absorbs blue light at 395 nm (protonated) and fluoresces green (509 nm, deprotonated) by giving up its proton and then returns to its normal state by absorbing a proton. The bioluminescent Green Fluorescent Protein (GFP) is found in the jelly fish Aequorea victoria. The chromophore is formed by the cyclization of an internal Ser65Tyr66-Gly67 tripeptide (van Thor, Pierik et al. 1998a) wherein the deprotonated phenolate of Tyr66 at the active site is the cause of fluorescence (Cubitt, Heim et al. 1995). GFP can exist in at least two spectroscopically distinct states: GFP395 and GFP480, with peak absorption at 395 and 480 nm, respectively, presumably resulting from a change in the protonation state of the phenolic ring of its chromophore. The photoconversion of GFP involves a proton transfer event initiated by the deprotonation of the phenolic chromophore which leads to the rearrangement of the hydrogen bonding network in the protein and protonation of Glu222 (Yoo, Boatz et al. 2001). 4 Another example is the light-driven proton pump protein for solar energy transduction, bacteriorhodopsin (Figure 1.4). The protonated Schiff base of the retinal chromophore forms a hydrogen bond with Asp85. With the absorption of light, the retinal chromophore undergoes a trans to cis photoisomerization. This leads to the proton transfer from the Schiff base to Asp85 triggering the sequential proton transfer reactions for the proton to be pumped from the cytoplasmic side to the extracellular side (Heberle 2000, Lanyi 2004, Rothschild 1992). Hydrogen bonding interactions in proteins A hydrogen bond occurs when two electronegative atoms, such as nitrogen and oxygen, interact with the same hydrogen. The hydrogen is normally covalently attached to one atom, the donor, but interacts electrostatically with the other, the acceptor. This interaction is due to the dipole between the electronegative atoms and the proton (Nelson and Cox 2000). However, the hydrogen bond is part electrostatic (90%) and part covalent (10%) (Isaacs, Shukla et al. 2000), making them easier to cleave than covalent bonds but strong enough to participate in some of the most important biological processes. Figure 1.2: One example of hydrogen bonding between two oxygen atoms in a biomolecule. The hydrogen is shared between the two oxygens, forming a hydrogen bond. Hydrogen bonds are found in proteins for structural stability. Hydrogen bonding also plays an important role in determining the three-dimensional structures adopted by proteins where it is formed between the backbone oxygens and amide hydrogens, also 5 called peptide bonds (primary structure of proteins). They are also predominant in the secondary structures of proteins (α helices and β sheets) (Creighton 1993). The bonding between the amino acids of the same protein cause it to fold into a specific shape, which helps determine the molecule's physiological or biochemical role (Wang, Wales et al. 2006). The hydrogen bond dissociation energy in proteins is in the order of 10-40 kJ/mol. A typical value for protein folding energy is approximately, 40 kJ/mol, which is about one to four times the hydrogen bond dissociation. From this, we can see that even by breaking one hydrogen bond; we can impair the protein stability. Hydrogen bonds are a very important aspect of protein structure and function. One good example is the hydrogen bonding network of the bacterial blue light photoreceptor, photoactive Figure 1.3: The hydrogen bond network stabilizing the negative charge on the chromophore of photoactive yellow protein. yellow protein (PYP) from Ectothiorhodopsira halophila. On undergoes a blue light photocycle absorption, containing PYP several intermediates that is closely linked to the function of the protein (Imamoto, Kataoka et al. 1996, Meyer, Yakali et al. 1987). Its chromophore (pCoumaric acid) is stabilized by two hydrogen bonds in the initial state, one with Glu46 and the other with Tyr42 (Borgstahl, Williams et al. 1995a, Getzoff, Gutwin et al. 2003). In the Tyr42→Phe mutant, hydrogen bonds of the phenolic oxygen of Tyr42 with the phenolic oxygen of the chromophore are lost. This leads to reduced protein stability and possibly a less rigid protein (Brudler, Meyer et al. 2000a). 6 A protein’s function is also dependent on its dynamics. With limited movement, there is very little room for conformational change and hence suppressed activity. Therefore it becomes imperative to study the protein structure-function dynamics in order to get a lucid view of any biological system. 1.2 Experimental methods for protein structure determination Traditionally, a protein's 3-D structure was determined using one of two techniques: Xray crystallography or Nuclear Magnetic Resonance (NMR) spectroscopy. X-ray Crystallography The landmark achievement in the field of X-ray crystallography was the solvation of the crystal structure of the sperm whale myoglobin by Max Perutz and Sir John Cowdery Kendrew which won them the Nobel Prize in Chemistry in 1962 (Kendrew, Bodo et al. 1958). Even though crystal structures of water soluble proteins began to be solved in the late 1950's, it was not until 1983 that a successful mechanism for crystallizing membrane protein was put forth by Michel, Deisenhofer and Huber (Michel 1983) for which they got the Nobel Prize in 1988. The first membrane protein to be crystallized was the photosynthetic reaction center (1985). Initially, intrinsic membrane proteins posed a challenge to crystallize because they require detergents or other means to solubilize them in isolation, and such detergents often interfere with crystallization. Such membrane proteins are a large component of the genome and include many proteins of great physiological importance, such as ion channels and receptors (Lundstrom 2006). Over recent years many more membrane proteins have been crystallized (Figure 1.3B) and their structures elucidated by X-rays, 7 some of the most important ones being PS-I (Fromme and Mathis 2004)and bacteriorhodopsin (Grigorieff, Ceska et al. 1996). A Figure 1.4: Number of protein structures and membrane protein structures deposited annually in the Protein Data Bank (PDB). (A) The total number of structures deposited in the PDB per year. The data are taken from the PDB website (The RCSB Protein Data Bank ), as of December 2005; the PDB currently holds 31,248 protein structures in total. (B) The number of unique membrane protein structures solved for the years indicated. The data are taken from (Membrane proteins of known structure),as of December 2005. This figure has been reproduced from (Gao and Cross 2005). B X-ray crystallography is the most powerful method to obtain high resolution structural information on water soluble proteins, and has been crucial in understanding protein structure and function. Proteins up to 107 Da (1 Da=1 g/mol) can be crystallized; the largest structures solved to date have been various viruses and the complete ribosome. The use of synchrotron radiation for spectroscopy and diffraction has been realized by an ever-growing scientific community, beginning in the 1960s and 1970s. As synchrotron radiation provides an intense light source, devices that enhanced the intensity of synchrotron radiation were built. Third-generation synchrotron radiation sources were conceived and optimized to produce bright X-rays (Hendrickson and Ogata 1997). These 8 have been applied to probe structural changes during protein function of proteins like lysozyme (Chayen, Boggon et al. 1996) and PYP (Genick, Borgstahl et al. 1997). A B C Figure 1.5: (A) Myoglobin, was the first protein to be crystallized (PDB code: 1MBN) (B) Green Fluorescent protein (PDB code: 1EMA) (C) Bactreiorhodopsin (PDB code: 1C3W) Since 1958, over 39000 X-ray crystal structures of proteins, nucleic acids and other biological molecules have been determined (The RCSB Protein Data Bank ). X-ray crystallography is now used routinely by scientists to determine 3-D and 4-D molecular structure of proteins (Hajdu and Andersson 1993), to study the enzymatic catalytic sites and to study the protein interaction with pharmaceutical interaction and the changes that might be advisable to improve it (Scapin 2006). Nuclear Magnetic Resonance Spectroscopy In the past 10 years, NMR has proven to be a powerful alternative to X-ray crystallography for the determination of protein structures. The field of protein NMR was pioneered, among others, by Kurt Wuthrich, who shared the Nobel Prize in Chemistry in 2002 (Wuthrich 2003). Although the primary work was started in 1958 (Kowalsky 1962, 9 Saunders and Wishnia 1958), protein NMR techniques have been progressively used and improved in both academia and the biotech industry. The basic principle involved is a physical phenomenon based upon the quantum mechanical magnetic properties of an atom’s nucleus. All nuclei that contain odd numbers of protons and neutrons have an intrinsic magnetic moment and angular momentum. The most commonly measured nuclei are hydrogen-1 (the most receptive isotope at natural abundance) and carbon-13, although nuclei from isotopes of many other elements (e.g. 15 N, 17 O) can also be observed (Wuthrich 2001). In this technique, a sample is immersed in a magnetic field and bombarded with radio waves. These radio waves encourage the nuclei of the molecule to resonate, or spin. As the positively charged nucleus spins, the moving charge creates a magnetic moment. The thermal motion of the molecule further creates a torque that makes the magnetic moment "wobble". When the radio waves hit the spinning nuclei, they tilt even more, sometimes flipping over. These resonating nuclei emit a unique signal that is then picked up on a special radio receiver and translated using Fourier Transform algorithm. By measuring the frequencies at which different researchers can nuclei flip, determine molecular structure (McDonald and Phillips 1967). Figure 1.6: (A) The structure of the tendamistat protein using NMR technique by the Wuthrich group (Billeter, Schaumann et al. 1990). (B) The first NMR structure of PYP with 26 structures (Dux, Rubinstenn et al. 1998). 10 The first protein structure solved using the NMR technique by the Wuthrich group was of the alpha amylase inhibitor protein, Tendamistat (PDB code: 3AIT), consisting of 74 amino acids (Billeter, Schaumann et al. 1990). A unique characteristic of the NMR technique is that of providing useful information for describing the properties of unfolded or partially folded proteins (Rico M, 2004). NMR has the advantage over crystallographic techniques in that experiments are performed in solution as opposed to a crystal lattice that hinders protein movement. However, a protein of even a relatively modest molecular weight (14 kDa) has a very large number of chemically different protons, so that their resonances accumulate in a very narrow spectral width. This means that signal overlapping is a very serious problem if we want to observe isolated signals. Obtaining really strong magnetic fields in order to distinguish protons of individual elements is a difficult task. The amount of overlap of the lines decreases as the strength of the magnetic field increases. The most sensitive NMR spectrometer is currently at the EMSL lab, Washington, USA. This facility provides a 1000 MHz (23.5 T) magnetic field strength operated at 1.8 K. This frequency reflects the resonance frequency for protons in that field strength (Illman 1994). However, most of the labs have access to about only 500 MHz instruments, the one at Oklahoma State University being 600 MHz. It is also very difficult to characterize membrane proteins due to their sluggish movement. This reduces the sensitivity of the method for studying membrane proteins. Another issue with NMR is that time-resolved studies cannot be performed. This restricts the researcher from observing protein functions using real time data. The principles that make NMR possible tend to make this technique very time consuming and limit the application to small and medium-sized molecules (Wishart and 11 Sykes 1994). Due to this restriction, there have been only about 6000 NMR protein structures available so far (The RCSB Protein Data Bank). X-ray crystallization also has its drawbacks. The major rate limiting step is the arrangement of molecules in three dimensional crystals. Proteins may not be functional in crystals as their movement is impeded due to its crystal lattice. The lack of adequate water molecules is also a concern for functioning as the protein is not in its native, well hydrated environment (Makinen and Eaton 1973, Makinen and Fink 1977, Xie, Kelemen et al. 2001a). X-ray crystallography also does not have the means to recognize proton transfer nor is it sensitive to hydrogen bonding. It has to be used in conjunction with the neutron scattering technique to detect protons (Lu and Thomas 1998). However, this technique requires huge crystals and there are very few sources of neutrons around the world for diffraction experiments and the data collection can stretch into weeks or even months (Durbin and Feher 1996). Another major drawback associated with this technique is that the crystallization of proteins is a difficult task. Extreme care should be taken to see that the crystals must be of high quality (Blundell and Patel 2004). Crystals are formed by slowly precipitating proteins under conditions that maintain their native conformation or structure. These exact conditions are difficult to attain and have to be strictly monitored in order to attain a well ordered crystal of reasonable dimensions. Time-resolved X-ray crystallography has enabled researchers to “watch” proteins in action at the picosecond timescale. This mechanism was used to study a small globular protein, Myoglobin (Schotte, Soman et al. 2004). Even though time-resolved 12 measurement is possible with X-ray crystallography, it is both expensive and not economically feasible (Ng, Getzoff et al. 1995, Parak 2003). The lack of water molecules also provides a non-native environment for the protein functioning and the crystal lattice also impedes its function. Hence there arises a need to develop a method that is both time-resolved and that is sensitive to probe important functional characteristics of proteins like proton transfer pathway and hydrogen bonding network. This will give us an opportunity to “watch” proteins function in their native environment by providing us with real time data. This will also enable us to get a better understanding of the protein structural dynamics. Infrared Spectroscopy Although X-ray crystallography and NMR spectroscopy provides high resolution structures, it becomes more important to the have ability to monitor structural changes in response to a physiological stimulus in its native environment (Tamm and Tatulian 1997). A technique that is gaining importance in the field of protein structural dynamics is FTIR spectroscopy. The main advantages of FTIR spectroscopy lies in the fact that it the technique is very sensitive, requiring only sub-milligram quantities for sample preparation and detection; conformational states can be measured in aqueous environments; and the physiological conditions of a sample can be varied in situ (Braiman and Rothschild 1988, Krimm and Bandekar 1986). Although fairly nascent, it is a well developing technique which can be used to identify molecules by analysis of their constituent bonds. Infrared spectroscopy is based on the absorption of electromagnetic radiation by matter due to different vibrational modes of these chemical bonds. A group of atoms in a molecule (e.g. CH2) may have 13 multiple modes of oscillation caused by the stretching and bending motions of the group as a whole. If an oscillation leads to a change in dipole in the molecule, then it will absorb a photon which has the same frequency (Goormaghtigh, Cabiaux et al. 1990). FTIR spectroscopy provides information about the secondary structure content of proteins, unlike X-ray crystallography and NMR spectroscopy which provide information about the tertiary structure. Each compound has a characteristic set of absorption bands in its infrared spectrum. Characteristic bands found in the infrared spectra of proteins and polypeptides include the Amide I and Amide II. These arise from the amide bonds that link the amino acids. The absorption associated with the Amide I band leads to stretching vibrations of the C=O bond of the amide, absorption associated with the Amide II band B A Figure 1.7: The vibrations responsible for the Amide I and Amide II bands in the infrared spectra of proteins and polypeptides. (A) The Amide I band is due to carbonyl stretching vibrations while (B) the Amide II is due primarily to NH bending vibrations. leads primarily to bending vibrations of the N—H bond (Figure 1.7). Because both the C=O and the N—H bonds are involved in the hydrogen bonding that takes place between the different elements of secondary structure, the locations of both the Amide I and Amide II bands are sensitive to the secondary structure content of a protein. Time-resolved FTIR difference spectroscopy is a powerful method for structurefunction studies of proteins: high structural sensitivity (proton transfer, hydrogen bonding 14 perturbation, and secondary structural changes), excellent time-resolution (up to 10 ns), and accessible to proteins in both crystalline and solution environments. The technique has also been used to study an important model for membrane proteins. In depth studies of bacteriorhodopsin, which is a proton pump in the membrane, has been made possible by this powerful technique (Braiman, Mogi et al. 1988a, Rothschild 1992). FTIR difference spectroscopy is sensitive to functionally important structural transitions, including proton transfer (Xie, Hoff et al. 1996a, Xie, Kelemen et al. 2001a) chromophore isomerization and protonation (Xie, Kelemen et al. 2001a) and hydrogen bonding interactions (Nie, Stutzman et al. 2005). Therefore, this technique is ideally suited for assessing the impact of protein crystallization on the functionality of proteins. Time-resolved rapid scan FTIR spectroscopy Time-resolved rapid-scan FTIR spectroscopy is largely differently from conventional FTIR systems: (1) each scan can be measured and stored separately, allowing time-resolved capability. (2) The moving mirror is designed to move very fast (200 kHz) and with minimum friction using air-bearing design. The time-resolution is determined by how fast the moving mirror takes to complete a full scan. It takes only 98.7 ms for a full scan at 4.5 cm−1 spectral resolution. (3) The internal clock of an FTIR spectrometer is used to trigger laser flashes and data collection in order to achieve the best possible time-resolution. For a repetition rate of 0.5 Hz and 4.5 cm−1 spectral resolution, the data collection time is approximately 2 hours. 15 Figure 1.8: The layout of time-resolved rapid-scan spectroscopy Our lab uses an innovative technique to improve the time-resolution of rapid-scan FTIR by a factor of 4 using 4-way splitting method. The 4-way splitting means that a full scan is divided into 4 equal paths (Q1 to Q4), that each yields in principle identical information. We then combine these four quarters right after laser excitation from four experiments to generate the first full scan interferogram, and Fourier transform to an infrared difference spectrum. 1.3 FTIR for protein structural dynamics Proteins lack activity in the absence of water. The Photoactive Yellow Protein (PYP) is a blue-light sensor first identified in Halorhodospira halophila. The presence of a light-active chromophore, p-coumaric acid, allows the sensing of blue light (Meyer 1985). Upon photon absorption, the protein undergoes a photocycle, linked to the isomerisation of the chromophore (Xie, Hoff et al. 1996a). During the photocycle, transient intermediates are formed at different time scales. The signaling state of PYP is 16 formed within microseconds and the return to the ground state is on a sub-millisecond time scale. However, PYP requires ~500 water molecules to function as a photoreceptor. Its function is impeded by the lack of water in when crystallized (Hoff, Xie et al. 1999), the number of water molecules available for each PYP molecule falls to ~350. In addition, salts in crystallization solution may dehydrate proteins. The power of the water molecule lies in its large permanent electric dipole moment, its ability to form hydrogen bonds, and its freedom to reorient to stabilize charged groups. Dissolving salts in water has profound effects on the hydrogen bond network structure and dynamics of water (Cacace, Landau et al. 1997a). Our main aim is to understand under what circumstances and to what extent crystallization salts impede protein function in salt solutions. The properties of salts are often studied as part of the classical Hofmeister series (Cacace, Landau et al. 1997a). A range of frequently used crystallization salts from the Hofmeister series (Hofmeister 1888) are employed to study the effects of high concentrations of salts on the dynamics of equilibrium protein fluctuations and the extent of functionally important structural transitions of proteins. Using time resolved rapid scan FTIR spectroscopy to measure light induced structural changes of photoactive yellow protein at low and high salt concentrations is the crux of Chapter II. We study the effect of a series of salt concentrations with considerations given to pH, type of salts and the concentrations of salts. 1.4 From Infrared Spectra to Protein Structure Proteins are made of amino acids and each of them has a characteristic set of absorption bands in the infra red spectrum. Tyrosine residues are often found at the active 17 sites of proteins. The pKa value of tyrosine side chain group is 10.5 (Nelson and Cox 2000) so that these phenol groups are mostly protonated in proteins at steady states. The pKa values of these groups may change dramatically at the active sites of proteins in functional intermediate states, resulting in temporary or permanent deprotonation of these groups and therefore proton transfer between tyrosine residues and other key amino acids. Hydrogen-bonding interactions play key roles in regulating the pKa values of ionizable groups and driving proton transfer. FTIR difference spectroscopy is a sensitive technique to study structural changes in proteins. Virtually each chemical group in a protein contributes to the vibrational spectrum and especially changes in hydrogen bonding and proton transfer event lead to significant signals in the IR difference spectrum, due to changes in intensity, bandwidth, and frequency of the vibrational mode involved. Since protons and hydrogen bonds cannot be directly observed in proteins by X-ray crystallography and NMR spectroscopy, FTIR spectroscopy yields information that is complementary to that obtained by these two high resolution structural techniques. Hydrogen bonding is a fundamental for protein structure and function. Breaking a single hydrogen bond may impair the stability of a protein. Changes in hydrogen bonding interactions of proteins during function represent a major theme representing dynamic structure in the active site of proteins. Time-resolved FTIR spectroscopy is a powerful technique to probe transient structures of proteins. In order to extract specific, quantitative structural information from time-resolved FTIR spectra, we report the development of 2D vibrational structural markers for probing hydrogen bonding interactions of Tyrosine. Using density function based first principle calculations, we 18 identified two vibrational modes, the C-O stretching and the OH bending is sensitive to hydrogen bonding interactions. When the C-O stretching frequency is inadequate to distinguish zero and two hydrogen bonds, two dimensions of C-O stretching frequency combined with OH bending or stretching frequency may be used to make such distinction. This vibrational structural marker is based on ab initio computational studies and supported by experimental data. Our lab has already been active in developing a structural marker for probing hydrogen bonding status of protonated Asp and Glu residues (Nie, Stutzman et al. 2005). Further development of these markers are expected to enhance the power of timeresolved infrared difference spectroscopy for structural characterization of functionally important intermediate states of proteins, and consequently for understanding the functional mechanism of proteins. 19 Chapter II Impact of Salts on the Structural Dynamics of Photoactive Yellow Protein 2.1 The Hofmeister Salts Franz Hofmeister (1850-1922) was an early protein scientist, and is famous for his studies of salts that influence the solubility and conformational stability of proteins. Hofmeister was the first to propose that polypeptides were amino acids linked by peptide bonds, although this model of protein primary structure was independently and simultaneously conceived by Emil Fischer (Voet and Voet 2004). First proposed in 1888, the Hofmeister series (Hofmeister 1888) is a qualitative ordering of ions based originally on their ability to salt-out proteins from aqueous solution (Cacace, Landau et al. 1997a, Collins and Washabaugh 1985). However, it is now known that there are other physical phenomenon like enzyme activity (Pinna, Bauduin et al. 2005), protein stability (Broering and Bommarius 2005) and protein crystallization (Ducruix, Guilloteau et al. 1996) are also influenced the Hofmeister series. Salts affect in widely different manners the properties of biological macromolecules such as their stability, solubility, and biological activity (Samoilov, Yashkich.Vi et al. 1972, Vonhippe.Ph and Schleich 1969). At low concentrations (below 0.01 M), salts can stabilize proteins through nonspecific electrostatic interactions, dependent only on the ionic strength of the medium (Tanford 1961). At high concentrations (0.01 M to 1 M), however, salts exert specific effects on proteins which depend on the nature of the salt and its concentration, resulting in either the stabilization 20 or denaturation of proteins. The solubility of the protein, i.e., salting in or salting out (either precipitation or crystallization) is also affected (Arakawa and Timasheff 1984). In a three-component system such as protein-water-salt, knowledge of the preferential interactions of the proteins with the solvent components can give us an understanding into the manner by which additives affect the solubility and stability of proteins. Although the Hofmeister series has been studied extensively for its effect on the solubility and stability of proteins, here we offer new insights on the effects of these salts on structural dynamics and functionality of proteins. This chapter reveals a novel aspect of the classical Hofmeister series: the impact of salts on the structural dynamics motions of proteins, including those needed for functionally important conformational changes, by kosmotropic and chaotropic salts. Time-resolved FTIR spectroscopic techniques are employed to probe the effects of impaired water activity on the structural dynamics of proteins, including equilibrium fluctuations and functionally important transitions, due to the presence of salts from the Hofmeister series. Molecular properties of water Water has held a strange fascination for scientists over the years because of its inane quality of being a universal solvent. Water has a higher melting point, boiling point and heat of vaporization than most liquids (Nelson and Cox 2000) which gives water great internal cohesion. This strong intermolecular attraction is due to the structure of water molecule. The H2O molecule has a bent geometry with an O-H bond distance of 0.958 Å and an H-O-H bond angle of 104.5° (Figure 2.1). Each of its two hydrogen atoms shares an electron pair with the oxygen atom. The two unshared electron pairs of the oxygen atom give it a localized partial negative charge and the strong electro 21 negativity of oxygen gives the two hydrogen nuclei partial positive charges. This charge separation results in the water molecule having a dipole moment of 1.85 debye units. This separation of charges also accounts for an electrostatic attraction between the water molecules, also known as the hydrogen bond. Figure 2.1: A water molecule with one oxygen bonded to two hydrogens via covalent bonds of length 0.958 Å each, separated by an angle of 104.5° and having a dipole moment of 1.85 debye units. 104.5° As the water molecule is nearly tetrahedral, each molecule can form up to 4 hydrogen bonds with the neighboring molecules (Figure 2.2). At any given instant in liquid water at room temperature each water molecule is believed to form hydrogen bonds with an average of 3.4 molecules. Essentially, the hydrogen atom is being shared unequally between two electronegative atoms. The atom to Figure 2.2: One water molecule can form up to 4 hydrogen bonds with the neighboring waters having an average life time of 1012 s, which makes the structure of water so dynamic(Voet and Voet 2004) which the hydrogen is covalently bound is the hydrogen donor; the other electronegative atom is the acceptor. In biological systems, the atoms participating in hydrogen bonding are oxygen and nitrogen. The distance between two hydrogen bonded atoms varies from 2.6 Å to 3.1Å. The hydrogen bond is part electrostatic (90%) and part covalent (10%) (Isaacs, Shukla et al. 2000). Hydrogen bonds are also directional, they are the strongest when the bonded molecules are oriented to allow maximum electrostatic interaction, i.e., when the hydrogen bonding angle is 180°. 22 Solvent properties of water The solvent properties of water are unique owing to the electric dipole of the molecule. An ion immersed in water attracts the oppositely charged ends of the solvent dipole (Figure 2.3). The ion is thereby surrounded by several concentric shells of oriented solvent molecules and is said to be hydrated. Dissolution of an ionic solute in water disturbs the hydrogen bonding between water molecules by causing a distinct change in the structure of liquid water since the positive and negative ions of the solute are surrounded by a hydration shell of water dipoles. The hydrated ions have geometry and properties somewhat different from the hydrogen bonded water molecules in the essence that they are more highly ordered and regular in structure. This effect is clearly visible in the solubility of biological molecules (such as proteins) in water. Water has a high dipole moment of 1.85 debye units and its dielectric constant is about 80. This determines the ability of water to ionize a salt on its dissolution in water. The partially positive hydrogens tend to orient themselves towards the anion and the partially negative oxygens orient themselves towards the cation as shown in the figure below. At this point, the salt is said to be hydrated. Figure 2.3: The water molecules separate ions on dissolution. The distance of separation and the high dielectric constant prevents the salt from coalescing and keeps the ions hydrated. 23 This distance of separation and the high dielectric strength of water decreases the force of attraction between the oppositely charged ions (Coulombs’ law) and prevents the salt from coalescing. Hydration shells of ions Dissolution of common salt or sodium chloride (NaCl) in water yields a solution containing the ions Na+ and Cl –. Owing to its high polarity, the H2O molecules closest to the dissolved ion are strongly attached to it, forming what is known as the inner or primary hydration shell (Figure 2.4). Positively charged ions such as Na+ attract the negative (oxygen) ends of the H2O molecules, as shown in Figure 2.3. The ordered structure within the primary shell creates, through hydrogen-bonding, a region in which the surrounding waters are also somewhat ordered; this is the outer hydration shell, or cybotactic region. Outer hydration shell (semi-ordered water) Inner hydration shell (chemiabsorbed and ordered water) Bulk water (random arrangement) Figure 2.4: The complete hydration of Na+ ion has three shells of water molecules surrounding it. The most strongly ordered water molecules are the first shell, followed by semi ordered water molecules and then the bulk waters that are not affected by the cation. The water molecules from the first and second shell are continuously replaced by the water molecules from the bulk media [www.chem1.com/acad/sci/wat-images/hydrated.gif]. The structure of the solvation shells of ions in bulk aqueous solutions has been investigated with NMR, x-ray and neutron diffraction (Kameda, Saitoh et al. 1993, Yamaguchi, Niihara et al. 1997). Unfortunately, these techniques are not capable of 24 providing information on the dynamics of aqueous solvation shells, because the timescales involved in these techniques are much longer than the typical lifetimes of the solvation structures. Hence, most of the data available is mainly from molecular dynamics simulations (Chandra, Uchimaru et al. 2000). Recently, the use of femtosecond infrared spectroscopy has enabled researchers to obtain information on the hydrogen bond dynamics of liquid water (Bakker, Kropman et al. 2005, Woutersen, Emmerichs et al. 1997). The value of spectral diffusion time of the hydration shells of certain anions (Cl−, Br−, and I−) was measured using this technique and was found to be is 20–50 times longer than for bulk liquid water hence showing relatively slow dynamics (Bakker, Kropman et al. 2005). Although ions have a small effect on the hydrogen bonds outside the anionic solvation shell (Samoilov, Yashkich.Vi et al. 1972), it is well known that ions can make liquid water much more viscous. This increase in viscosity has been considered to be one of the prime indications for the strong structure-making effect of ions like Na+. However, it should be realized that viscosity is a macroscopic property that represents the average behavior of a large number of water molecules in an aqueous solution. This means that an aqueous salt solution should not be viewed as a homogeneous liquid with a modified but uniform intermolecular interaction, but rather as a colloidal suspension of inert particles in pure liquid water, with the particles formed by the ions and their first hydration shells. Outside the first hydration shell, the ion is observed to have little effect on the strength of the hydrogen-bond interactions between the water molecules (Zhang and Cremer 2006). 25 The number of water molecules in the hydration shells of ions varies with the size and the charge distribution of the ion. The ionic radius is defined using two criteria: (1) The first criterion is a minimum density criterion. The cation (anion) radius is chosen to be the distance between the cation (anion) site and the minimum of the superposed electron density long the internuclear line. (2) The second criterion is a minimum overlap criterion. For any two touching spheres, centred at the cation and anion sites, some electrons belonging to the cation will overlap into the anion sphere, and vice versa. The ionic radii are defined as the radii of those spheres for which the sum of these two overlaps is a minimum. Electrons which lie outside both spheres are neglected. Ionic radius Hydration number Ion References: (Caminiti, (Å) Licheri et al. 1978, Caminiti, Li+ 0.6 4-6 Na+ 0.95 4-6 Paschina et al. 1979) (Domene and Sansom 2003) (Maslen 1967, Ramos, Cation K+ 1.33 2-5 Cs+ 1.69 2-3 F- 1.27 - Cl- 1.81 6-9 Br- 1.95 6-7 SO42- - 8 Anion Table 2.1: The ionic radius and the hydration number of some commonly studied anions and cations. The hydration number is the total number of water molecules in the first shell of hydration. 26 Barnes et al. 2000) Kosmotropes and Chaotropes There can be two kinds of ions, ones that have strong interaction with water which increase the structuring of water also called order makers (structure-makers) or kosmotropes. They are stabilizing solutes that increase the amount of water interacting with a macromolecule. The other kind of ions decreases the structuring of water and is hence called disorder-makers (structure-breakers) or chaotropes (Vorontsov and Novakovskaya 2007). Kosmotropes are stabilising solutes which increase the amount of water interacting with a macromolecule whereas chaotropes break down and weaken hydrogen bonding in a macromolecular structure, thereby decreasing the order of water and increasing its surface tension leading to destabilisation of the macromolecular structure (Zhao 2006). The different anions and cations can be divided into these two categories based on the charge they carry, the effect they have on water and the radii of the ion. Large singly charged ions with low charge density exhibit weak interactions with water than water itself and thus interfering less with the hydrogen bonding of the surrounding water are chaotropes. And small multiple charged ions that exhibit stronger interactions with water molecules than water itself and therefore capable of breaking water-water hydrogen bonds are kosmotropes. The radii of singly charged chaotropic ions are greater than 1.06 Å for cations and greater than 1.78 Å for anions(Collins 1995) thus the hydrogen bonding between water molecules is more broken in the vicinity of ionic kosmotropes than ionic chaotropes. 27 Figure 2.5: The Hofmeister series classified according to their effect on water molecules. Kosmotropic ions are structure makers, they form weak hydrogen bonds with the water molecules and chaotropes are structure breakers, they form strong hydrogen bonds with the water molecules, thus creating more order in the water structure (Cacace, Landau et al. 1997a). The kosmotropes and chaotropes also have a large effect on the surface tension of water, solubility of hydrocarbons, salting out and salting in of proteins, protein stability and protein denaturation that are discussed in the following section. Effect of hydrated ions on proteins Currently more and more intermediate structures of proteins are being studied using time-resolved or trapping crystallographic techniques. However, very few functional tests have been performed. There is no routine method available to assess the functionality of proteins in crystals. In addition, there is a lack of fundamental understanding on how crystallization of proteins may alter their function. This lack of understanding makes it difficult (i) to resolve disputes regarding crystallization effects, and (ii) to raise the much needed awareness that routine tests are crucial prior to all crystallographic studies of functional intermediates of proteins. Therefore, it is urgent to study the functionality of proteins in crystals, and to gain a deep understanding on the crystallization effects on protein structural dynamics. Chaotropic ions from the Hofmeister series tend to denature proteins. Chaotropic agents increase the solubility of non-polar substances in water consequently their ability 28 to denature proteins stems from their ability to disrupt hydrophobic interactions (Voet and Voet 2004). The number of water molecules necessary for hydrating an ion depends on the charge and the radius of the ion. Conversely, kosmotropic agents stabilize proteins and strengthen hydrophobic forces thus increasing the tendency of water to expel proteins. To further study the effect of hydrated ions on proteins, we employ a model protein, the Photoactive Yellow Protein with a regenerative photocycle. We study the effect of concentrated salt solution on the structural dynamics and kinetics of the protein. The Photocycle of PYP The Photoactive Yellow Protein (PYP) was first described by Terry E.Meyer in 1985. He purified the yellow protein from a halophilic phototropic bacterium Ectothiorhodospira halophila. It was first isolated and purified from salt encrusted mud taken from the shores of Summer Lake, Lake county, Oregon (Raymond and Sistrom 1967, 1969). It is a small (14 kDa), water-soluble, cytoplasmic protein that is also found in several other halophilic purple phototrophic bacteria (Meyer, Fitch et al. 1990, Meyer, Yakali et al. 1987). This unicellular prokaryotic grows under strict anaerobic conditions at temperatures of up to 50 C (Imhoff 1984). It grows optimally at 11 to 22% NaCl, but growth even occurs at 32% NaCl makes E. halophila the most halophilic Eubacteria known. The E. halophila cells swim by the motion of their bipolar flagella and their migration is induced by light, i.e., they exhibit the property of phototaxis. They move away from blue light, favoring green light (Sprenger, Hoff et al. 1993). Hence the 29 negative phototaxis is wavelength dependent and might be due to the fact that the wavelength of blue light is close to the UV region. Photoactive Yellow Protein is an excellent model for photoreceptor proteins and its biological function is similar to that of sensory rhodopsins (particularly SR II). Although, structurally different (Figure 2.6), they are both involved in similar function: signal transduction. Rhodopsins are found in both unicellular and complex organisms, their functions ranging from proteins allowing sight (visual rhodopsins with 214 residues) to light-activated proton pumps (Bacteriorhodopsin with 248 residues). Interestingly, (a) (c) (b) Fig 2.6: (a) Photoactive Yellow Protein with 125 amino acids; (b) Sensory Rhodopsin II with 217 amino acids; (c) Rhodopsin (Animal vision) with 326 amino acids. Bacteriorhodopsin is found in Halobacterium salinarum, an archaebacterium also found in salt lakes. Since rhodopsins are membrane proteins, they are not water soluble unlike PYP which is highly water soluble. Hence PYP, with only 125 amino acids is a compact yet a complete model system for a photoreceptor protein. PYP is also a prototype of PAS domain proteins (Procopio, Lahm et al. 2002). PAS is an acronym formed from the names of the proteins in which the PAS motive was 30 first recognized: the Drosophila period clock protein (PER), the vertebrate aryl hydrocarbon receptor nuclear translocator (ARNT) and the Drosophila single minded protein (SIM) (Nambu, Lewis et al. 1991). Proteins containing PAS domains are predominantly involved in signal transduction, some being receptors or transcriptional regulators. PAS domain proteins are found in all three divisions of cellular life (Bacteria, Archaea and Eucarya). Herg potassium Channel N-terminus (Morais, Barber et al. 1998) in animals and LOV2 (Light, Oxygen or Voltage) (Crosson and Moffat 2001) a photoreceptor domain from plants have structures solved from the PAS family. These two are amongst several hundreds of proteins that exhibit the same fold as PYP (Fig). PYP has a α/β fold, consisting of a central antiparallel β-sheet with six strands and flanked on both sides by loops and helices. It was the first protein from the PAS domain family for which the 3D structure was elucidated and consequently, it was proposed that the Photoactive Yellow Protein is a structural prototype in the PAS domain containing proteins (Pellequer, Wager-Smith et al. 1998). The photoactivity of PYP is expressed in the form of a photocycle. In photoactive proteins, the chromopore is usually at the heart of the functional characteristics that have to do with the absorption of photons. In the dark state, PYP absorbs a photon of proper wavelength (a blue photon), structural changes occur in the protein that leads to a signaling state that can be read by the bacteria it resides in. Once this process is complete, the protein returns to its dark adapted state that brings the cycle to a full circle. This self regenerative cycle requires that the protein be in a well hydrated form. It does not require the presence of additional co-factors or proteins to complete this photocycle. Since the function of the chromophore of PYP is to catch photons from the visible region of the 31 electro magnetic spectrum; UV/Vis spectroscopy can be employed to understand the basic absorption properties of PYP. The bright yellow color of PYP arises from a single, broad (60 nm) absorption band with a maximum at 446 nm (ε = 45 mM-1 cm-1 ; Fig.) (Meyer 1985). The most notable characteristics of the UV/Vis spectra obtained are the absorption bands at 278nm and 446nm. The ratio of these two peak heights gives the purity index of the protein, where a lower value than 0.50 is considered to be pure. 0.6 Absorption [OD] 0.5 0.4 0.3 0.2 pCoumaric acid 0.1 0.0 250 300 350 400 450 500 Wavelength [nm] Figure 2.7: The UV/Vis spectra of wt PYP at pH 7 and in 50mM NaDPO4 buffer showing a maximum peak at 446nm from the pCA chromophore and another peak at 278nm from the tyrosine and tryptophan residues close to the chromophore. The basic photocycle can be depicted as a simple scheme of only three species when put into terms of essential photocycle stages, as shown in Figure 2.8. In the ground state or dark adapted state, pG, the chromophore is deprotonated and the isomerization state is trans. The phenolate oxygen of the chromophore is stabilized by a hydrogenbonding network involving Tyr42, Glu46 and Cys69 as shown in Figure 2.15 (Borgstahl, Williams et al. 1995b).The second species pR is spectrally red shifted with respect to the ground state and is formed on a nano-second time scale (Ujj, Devanathan et al. 1998). 32 Here, the choromophore is still deprotonated but it is isomerized to cis. The third species pB, is spectrally blue shifted with respect to the ground state and is formed on a microsecond time scale (Meyer, Yakali et al. 1987). This species is presumed to the signaling state of the photoreceptor and is considered relatively stable to allow the signal to be processed by the organism. The chromophore accepts a proton from Glu46 and is protonated (Xie, Hoff et al. 1996a, Xie, Kelemen et al. 2001a) while it is still in the cis configuration. The protein is allowed to subsequently return to its ground state wherein the chromophore is deprotonated and is reisomerized to trans configuration on a millisecond time scale. Over the years the photocycle has become more and more complex with more intermediate steps being discovered by researchers. The short-lived intermediates refer to the pG to pR transition and the long-lived intermediates refer to the signaling states formed in the last two basic steps of the photocycle, pR to pB and pB to pG. The formation of pR is accompanied by the chromophore isomerization to cis. During the course of this isomerization, at least two short lived states named I0 and I‡0were identified by pico second transient absorption spectroscopy in 1998 (Devanathan, Brudler et al. 1999, Ujj, Devanathan et al. 1998). I0 is formed on a picosecond time scale and decays with a relaxation time of ~220 ps to I‡0. I‡0 then decays with a relaxation time of 3 ns to pR. During the pG to pR transition, there is very little structural change in the protein. There is no change in the position of the aromatic ring of the chromophore (Genick, Soltis et al. 1998, Xie, Hoff et al. 1996a). However the isomerization is facilitated by rotating the thiol ester carbonyl. 33 The chromophore configuration changes from C7 =C8-trans C9-S-cis to C7=C8-cis C9-Strans. The formation of pR requires 120-160 kJ.Mol-1 (van Brederode, Gensch et al. 1995) for photoisomerization. PYP in the pG state acquires 268 kJ Mol-1 by absorbing a photon at 446nm. This energy difference is used to drive the rest of the photocycle. The formation of the signaling states (pB→pG) involves the protonation of the chromophore and an overall structural change in the protein. In the transition from pRpB, the first event is the protonation of the chromophore resulting in the formation of pB’. Xie and others predicted the formation of pB’ in 1996 and provided experimental proof in 2001 with FTIR data (Xie, Hoff et al. 1996a, Xie, Kelemen et al. 2001a). They showed that protonation of the chromophore and the ionization of Glu46 occurred simultaneously, while the protein secondary structural change follows later (indicating the pB formation). Following the formation of pB’, the protein undergoes large conformational changes resulting in the formation of pB. A protein quake, driven by the negative charge on Glu46 after its deprotonation, occurs during the pB’ to pB transition (Xie, Kelemen et al. 2001a). The transition from pB’ to pB is spectrally silent. During the recovery step from pB to pG , deprotonation of the chromophore happens before its reisomerization to trans. Thus, there is another intermediate proposed PYPN by Demchuk in 2000. Its absorption spectra is similar to the absorption spectra of the ground state. However, due to the lack of experimental evidence, this idea is still under dispute. 34 Signaling state Signal Transduction Flagellar motion Negative phototaxis Figure 2.8: The schematic representation photocycle of PYP for receptor activation (Xie, AH., 1996). (i) in the pG to pR photoreaction, pCA is photoisomerized from a 7-trans 9-scis to a 7-cis 9-s-trans configuration, the global conformation of PYP remains unchanged; (ii) in the pR to pB′ transition, a direct proton transfer occurs from Glu46 to pCA, while the global conformation is still unchanged; (iii) the alteration in the distribution of buried charges in pB′ triggers a large conformational change, leading to the formation of pB, the likely signaling state; (iv) recovery of pG from pB involves pCA reisomerization, deprotonation of pCA, reprotonation of Glu46. This figure has been reproduced from (Xie, Hoff et al. 1996a). 35 2.2 Materials and Methods Sample preparation A mixture of de-ionized water and Deuterium oxide (mixed in equal parts) was used to prepare a 4.0 M sodium chloride (CAS 7647-14-5) solution. The amount of salt necessary to make the 4.0 M solution was carefully weighted out and dissolved in the solvent consisting of 50% H2Oand 50% D2O. The same procedure was repeated for ammonium chloride (CAS 12125-02-9), ammonium sulphate (CAS 7783-20-2) and magnesium sulphate (CAS 10034-99-8). For measuring the salt dependence on protein using rapid-scan FTIR spectroscopy, the concentration of the PYP sample used was ~8 mM. To prepare PYP at the desired pH, deuteration and with the right salt concentration, we wash the protein sample using 4.0 M NaCl solution. This salt solution is first prepared at a higher concentration of 4.2 M and then a calculated amount is mixed with 50 μL, 1.0 M sodium phosphate buffer. Hence our final solution is 4.0 M NaCl with 50 mM sodium phosphate buffer and the pH adjusted to 7.00 ± 0.05 using NaOD. Our main goals for washing PYP are (1) To reduce H2O contamination from liquid sample as well as exchangeable hydrogen atoms in PYP; (2) To adjust the sample pH to 7.0; and (3) To make sure that there is sufficient number of salt ions. We start by adding 390 μL of the 4.0 M NaCl buffer solution to 10 μL of 8 mM PYP, the resulting protein concentration is then 0.2 mM. Therefore, the buffer to protein ratio is 250:1; hence one wash is sufficient for pH adjustment. (2) The H2O contamination is 10 μL /400 μL = 2.5%. A second wash with the 4.0 M NaCl solution is needed to further reduce the 36 water contamination. Even though the samples are prepared with utmost care, some water contamination persists during washing and loading of the sample. The same procedure was repeated for 0.5 M, 1.0 M and 2.0 M NaCl and also for 4.0 M KCl, 4.0 M CsCl. Each solution sample was made by sandwiching 2.7 μL of the PYP solution between two CaF2 plates 25 mm in diameter and 2 mm thick using a 12 μm spacer. The sample cell is placed in a custom-made sample holder for measuring UV-VIS absorption spectra. The dark state PYP in the 250-550 nm range was measured using a UV-Vis (Cary 300) spectrometer for each sample. FTIR spectroscopy: A Bruker IFS 66v/s FTIR spectrometer with a Michelson interferometer was utilized for collecting the single beam spectra of water with salts. Three samples of empty, salt in 50% H2O and 50% D2O and just CaF2 windows were loaded in a custommade sample exchanger driven by step motor driver (Si3540, Applied Motion products, USA). Then the sample chamber was purged with nitrogen gas to remove water vapor and is maintained at a constant temperature of 300K using water circulating temperature controller (RTE 111 D3, NESLAB Instruments, Inc., USA). The optics chambers of the spectrometer were evacuated to eliminate water vapor along optical path. The step motor driver is externally triggered by the spectrometer in rapid-scan mode to synchronize the data collection and changing sample. In this way, the quality of infrared spectra was largely improved due to little environmental change between the samples and the background (empty). The signal to noise ratio was large after 256 averages. The scanning rate was chosen to be 40 kHz and the spectral resolution to be 2 cm-1. 37 Time resolved rapid scan FTIR spectroscopy: The steady state FTIR absorption spectra and the light-induced FTIR difference absorption changes were also measured using a IFS 66v/s FTIR spectrometer (Bruker, Germany). The spectral resolution was 2 cm-1 for absorption and 4.5 cm-1 for difference absorption in the range 4000-900 cm-1. The PYP photocycle was triggered using laser pulses with pulse duration of 4 ns and energy of 3 mJ at 475 nm. The laser repetition was 10 Hz. The laser beam size at the sample was 6 mm in diameter. The light induced FTIR absorption changes were measured with 15 ms time resolution using the rapid scan method. Each of the spectra is the average of 400 measurements (scans).The 4 ns long actinic light flash for the time-resolved IR measurements was provided by a YAG laser (Surelite II-10, Continuum, USA) pumped OPO (Surelite OPO, Continuum, USA) tuned to 475 nm. We can improve the time-resolution of rapid-scan FTIR by a factor of 4 by using the quadruple split method. The 4-way splitting means that a full scan is divided into 4 equal paths (Q1 to Q4), that each yields in principle identical information. However, the data obtained, in reality, shows some noticeable differences. These differences arise from the slight variation in the zero path difference of the interferogram. We have been able to overcome this technical difficulty by repeating the experiment four times, such that the laser flash excites the sample at the start of each of the 4 quarters, respectively. We then combine these four quarters right after laser excitation from four experiments to generate the first full scan interferogram which is then Fourier transformed to an infrared difference spectrum. 38 2.3 Results and Discussion Salts affect the O-H stretching frequency of water The O-H stretch mode of the water molecules was excited using a mid-infrared signal. A mixture of 50% H2O and 50% D2O was used to ensure that the signal from H2O is not saturated. As a starting point for our studies, we selected two salts, NH4Cl and (NH4)2SO4 having a common cation, NH4+. It is a well known fact that the NH4+ ion in solution behaves in a way very similar to water. Spectroscopic results (NMR and IR absorption) show that the “NH4+ ion fits into the tetrahedral structure of liquid water”, and “the ammonium ion does not influence the structure of the solvent” (Vollmar 1963). As a consequence, “a solution of an ammonium salt may be considered, to a good approximation, as a solution of anions in water” (Caminiti, Licheri et al. 1978). The Full Width Half Maxima (FWHM) of pure water is about 294 cm-1. The FWHM narrows down to 272 cm-1 for 4.0 M NH4Cl and broadens to 314 cm-1 for 2.0 M (NH4)2SO4. The broadness of the peak in the absorption spectra is an indication of the structuring of the molecules. An ordered structure like ice will have a sharp absorption band when compared to liquid water. The same reasoning can be used to understand the change in bandwidth of υOH of water. The chaotropic Cl- structurally orders the water molecules around it that is indicated by the narrowing of half width (Figure 2.9A). There is much conclusive work done regarding Cl- ions; the existence of well defined hydration shells around Cl- ions has been strongly supported and there is general agreement in the literature also in the description of these ionic environments (Magini, Paschina et al. 1982). 39 The OH stretch frequency of pure H2O- D2O mixture seen at 3411 cm-1 is blue shifted to 3431 cm-1 with the addition of NaCl. The presence of NH4Cl in the aqueous mixture also yielded the same shift to 3431 cm-1. The anion in both these compounds is Cl-. This anion forms a strong hydrogen bond with the water molecule as the acceptor, structuring the water molecule. This is seen as a blue shift in the O-H stretching frequency. The result is concurrent with the frequency shift band at 3431 cm-1 for ammonium chloride (NH4Cl). We conclude from this that the shift to higher frequency is due to the presence of the common anion Cl- (Figure 2.9B). The interaction of the anion with the water molecule is stronger than the interaction between water molecules. This interaction with Cl- breaks the hydrogen bond between water molecules and makes the structure more ordered leading to the increase in the vibrational frequency. Observing the full width half maximum (FWHM) shift in wavelength also shows an interesting result. The FWHM of υO-H narrows from 294 cm-1 for pure water to 272 cm-1 for NH4Cl and 263 cm-1 for NaCl. This again shows the affect of the chaotropic anion Cl- with the water molecules leading to stronger vibrational signal as indicated by the narrowed peaks. 40 1.0 1.0 A B Pure water Pure water Absorption [OD] Absorption [OD] 0.8 4.0 M NH 4Cl 2.0 M (NH4)2SO4 0.6 0.4 0.0 3800 3600 3400 3200 3000 -1 0.6 0.4 41 C 0.8 2.0 M(NH4 )2 SO4 2.0 M MgSO4 0.6 0.4 0.2 3600 3400 -1 Wavenumber [cm ] 0.0 3800 3600 3400 3200 3000 Wavenumber [cm ] Pure water Absorption [OD] 4.0 M NaCl -1 Wavenumber [cm ] 0.0 3800 4.0 M NH 4Cl 0.2 0.2 1.0 0.8 3200 3000 Figure 2.9: (A) The υO-H of the infrared absorption spectra of pure water (Black) is compared with 4.0 M NH4Cl (green) and 2.0 M (NH4)2SO4 (red) in water. The FWHM of υO-H narrows from 294 cm-1 for pure water to 272 cm-1 for NH4Cl and broadens to 314 cm-1 for (NH4)2SO4 indicating the effect of the the chaotropic Cl- and the kosmotropic SO42- respectively. (B) The υO-H of the infrared absorption spectra of pure water (Black) is compared with 4.0 M NaCl (green) and 4.0 M NH4Cl (red) in water. The FWHM of υO-H narrows from 294 cm-1 for pure water to 272 cm-1 for NH4Cl and 263 cm-1 for NaCl indicating the effect of the common chaotropic anion, Cl-. (C) The υO-H of the infrared absorption spectra of pure water (Black) is compared with 2.0 M (NH4)2SO4 (green) and 2.0 M MgSO4 (red) in water. The FWHM of υO-H broadens from 294 cm-1 for pure water to 314 cm-1 for (NH4)2SO4 and 323 cm-1 for MgSO4 indicating the effect of the common kosmotropic anion, SO42-. On the contrary, the investigations carried out on oxy-anion hydration are few and, in addition, their results are not in agreement. The complication in studying the SO42- ion is that (1) The weakness of the oxy-anion-water hydration; (Musinu, Paschina et al. 1982) (2) Experimental evidence indicates that the O…O intermolecular peak is slightly shifted to distances longer than pure water implying the bonds that are formed are weaker than those in pure water. (Brown 1976) In fact, the hydration of the oxy anions involves O--.H-O interactions in which O-O distances are superimposed on and cannot be distinguished from the H2O-H2O distances usually present in aqueous solutions. However, Musini and group ((Musinu, Paschina et al. 1982) have shown that the sulphate ion has its own shell of hydration with approximately 8 water molecules. The weak interaction of the sulphate ions with the water molecules is seen as a broadening of the half width in case of (NH4)2SO4 (Figure 2.9A). A closer look at the peak positions of the salt solutions yields more conclusive proof of the water interactions with the halide anion and the oxy anion (Figure 2.9B). The effect of the anion SO4 - was studied in order to understand the kosmotropic salts (Figure 2.9C). However, SO4- is a special case of kosmotropic salts as they can also be classified as oxy anions. The interaction of these oxy anions with water molecules as explained before. This can be seen as the inhomogeneous broadening the peaks when compared with the υOH of pure water. The OH stretching frequency red shifts from 3411 cm-1 to 3408 cm-1 in 2.0 M (NH4)2SO4 solution and to 3398 cm-1 in the presence of 2.0 M MgSO4. SO4- forms weak hydrogen bonds with the water molecules. This causes the 42 downshift in wavenumber. The interaction of the anion with the water molecule is weaker than the interaction between water molecules. This interaction with SO4- breaks the hydrogen bond between water molecules and makes the structure less ordered leading to the decrease in the vibrational frequency (Figure 2.9C). The FWHM of υO-H broadens from 294 cm-1 for pure water to 314 cm-1 for (NH4)2SO4 and 323 cm-1 for MgSO4 indicating the effect of the common kosmotropic anion, SO42-. This again implies that the interaction of SO42- with the water molecules is much weaker than the interaction between water molecules themselves leading to weaker vibrational signal as indicated by the narrowed peaks. The effect of cations are not discussed in these two cases as both Na+ and NH4+ are weak kosmotropes and it is clear from the FTIR studies (Figure 2.9B and 2.9C) that the effect on water molecules is more due to the anion rather than the cation. The purpose of selecting these particular ions is for the reason that they have high biological significance in various life forms. Effect of salts on the photocycle of PYP Photoactive Yellow Protein does not require additional co-factors or proteins to complete its photocycle. This self regenerative cycle only requires that the protein be in a well hydrated form. Dehydration of proteins in general leads to loss of biological functions of proteins (Hoff, Xie et al. 1999, Rupley and Careri 1991). For PYP, it is found that ~500 water molecules per protein molecule are needed for normal kinetics and structural dynamics (Hoff, Xie et al. 1999). The total water content in crystals is low: on average, protein crystals contain approximately 50% of solvent by volume (Creighton and Freedman 1993). Furthermore, crystallization solvent contains a significant fraction 43 of precipitation agent. Hence the proteins can be dehydrated in crystals due to limited water content. In addition, salts in crystallization solution may dehydrate proteins. The power of the water molecule lies in its large permanent electric dipole moment, its ability to form hydrogen bonds, and its freedom to reorient to stabilize charged groups. Dissolving salts in water has profound effects on the hydrogen bond network structure and dynamics of water (Cacace, Landau et al. 1997a). The mean lifetime of bound water varies dramatically from picoseconds for Cs+ to a month for Cr3+. At high concentrations of crystallization salt, a large percent of water becomes bound water. If the lifetime of bound water is long, they lose their ability to escort surface charge movements of proteins during structural transitions. Therefore, even without a crystal lattice, proteins can be dehydrated in the high concentration salt solutions typically used in protein crystallization (Gilliland, Tung et al. 2002). This is an overlooked but important effect of crystallization on protein dynamics. The properties of salts are often studied as part of the classical Hofmeister series. We employed a range of frequently used crystallization salts from the Hofmeister series to study the effects of high concentrations of salts on the dynamics of equilibrium protein fluctuations and the extent of functionally important structural transitions of proteins. In order to understand the effects of both anions and cations, we chose a series of salts with a common cation (Na+) and a series with common anion (Cl-). The visible absorption spectra of the WT PYP solution samples in different salt solutions and that of the sample with no salt were measured from 250 nm to 550 nm. The spectra was baseline corrected (Figure 2.10). UV-Vis data indicates the purity and the 44 intactness of PYP after washing 4.0 M NaCl solution. The purity index (ration of peaks at 280 nm/446 nm) was found to be 0.46. But, in order to understand the structural implications better, we need to use time-resolved FTIR measurements. 0.5 Absorption [OD] wtPYP in 4.0 M NaCl Figure 2.10: UV/Vis data of wt PYP indicating the absorption maxima at 446 nm. The 4.0 M salt solution does not affect the purity index of the protein, which was calculated to be 0.46. 0.4 0.3 0.2 0.1 0.0 250 300 350 400 450 500 Wavelength [nm] The IR absolute absorption spectrum is a very effective tool to compare the ground state of PYP in different solutions and in the crystal state. Among others the IR absorption spectra provides information about the protein backbone structure (Amide I) the state of the chromophore and the functionally important carboxylic side chains of the protein. The IR absorption spectra of our solution samples were baseline corrected and then normalized for comparison according to the area of their Amide I absorption band. The second derivatives of the absorption spectra were calculated to obtain the positions of the heavily overlapping constituting bands. In our measurements the IR absorption of WT PYP in different concentration of NaCl solutions were found practically identical in the Amide I region (Figure 2.11). The normalized spectra of the salt dependent samples overlap almost perfectly. The second derivative of these spectra confirmed that the positions of the constituting bands are the same at the different NaCl concentrations. The main IR bands within the Amide I region 45 are at 1612 cm-1, 1635 cm-1, 1644 cm-1, 1661 cm-1 and at 1687 cm-1 wavenumber for all the salt concentrations. The 1497 cm-1 and the 1726 cm-1 bands, determined by the second derivative among other major bands, have already been assigned to the chromophore and to the protonated Glu46 group, respectively. This is a clear indication that in the initial pG state, salt solutions do not play any significant role in the IR absorption of the protein. The structure of PYP remains unaffected even in the presence of high salt concentration of 4.0 M NaCl. The absorption of the Amide I band is uniform in the presence of varying salt concentrations. The 1497 cm-1 that is assigned to the ring vibration of the phenolic chromophore is unchanged. The other major fingerprint band at 1726 cm-1 that is assigned to the protonated Glu46 also remains unchanged in the presence of high concentration of salt (Figure 2.11). 46 0.8 wtPYP with 0 M NaCl Absorption [OD] wtPYP with 4 M NaCl wtPYP with 2 M NaCl wtPYP with 1 M NaCl wtPYP in 0.5 M NaCl 0.6 0.4 0.2 0.002 1402 1440 1471 1455 1515 1498 1537 1584 1688 -0.002 1551 0.000 1726 1635 -0.004 1644 2nd Derivative of Absorbance 0.0 -0.006 1800 1700 1600 1500 -1 1400 1300 Wavenumber [cm ] Figure 2.11: (A) Infrared absorption spectra of wt PYP in varying salt concentrations indicate that the amide I band is independent of the presence of salt. The protein in the pG state is not affected by the presence of salt (B) The second derivative shows the overlap of the chromophore (1515 cm-1) and the Glu46 ionization band (1726 cm-1). 47 Light induced FTIR difference spectra of PYP In the pG state, the OH group of Glu46 is hydrogen bonded to the negatively charged phenolic oxygen of pCA consistent with the X-ray structure of PYP (Borgstahl, Williams et al. 1995a, Xie, Hoff et al. 1996a). In pG, the pCA chromophore is deprotonated and in the 7-trans 9-s-cis conformation. This buried negative charge on pCA is delocalized over the phenolic oxygen and ring, and is stabilized by specific electrostatic interaction with the positively charged Arg52, as well as by hydrogen bonding interactions with the OH groups of Glu46 and Tyr42 in the pCA binding pocket. Upon absorption of a blue photon, the pCA chromophore is photoisomerized into a 7-cis 9-s-trans configuration, leading to the formation of pR. Glu46 donates a proton to the phenolic oxygen of pCA, forming a putative unstable pB’ state. This local proton transfer event neutralizes the pCA, generates a new buried unstable negative charge of COO- of Glu46, and terminates the pCA-/Arg52+ interaction. This unstable negative charge triggers a large conformational change leading to the formation of a long lived signaling state and hence receptor activation which is repressed by the absence of COO- formation in the E46Q mutant (Xie, Kelemen et al. 2001a). The initial dark state of pG is recovered from pB at a longer time scale (250 ms) through a series of steps involving the reiosomerization of pCA to 7-trans 9-s-cis conformer, deprotonation of pCA and reprotonation of Glu46 (Xie, Hoff et al. 1996a). PYP is a highly water soluble protein and in our FTIR samples, the protein is fully hydrated with 4800 water molecules per PYP (closer to physiological conditions). When the hydration level is reduced to ~350 water molecules per PYP, the large structural changes are fully suppressed (Hoff, Xie et al. 1999). The hydration level in P63 crystals is 48 ~500 water molecules per PYP. This partial dehydration can reduce the extent of structural changes. wt PYP in solution wt PYP in 4.0 M NaCl wt PYP in P6 3 crystals 1571 1515 1740 1497 1689 1609 0.0 1726 Diff Absorption [mOD] 4.0 Figure 2.12: pB - pG infrared spectra of PYP in solution (black), P63 crystals (green) and in 4.0 M NaCl (red). The data for crystal studies was previously done (Xie, Kelemen et al. 2001b). 1641 -4.0 1750 1700 1650 1600 -1 1550 1500 Wavenumber [cm ] The complete photocycle of PYP is accompanied by four major interactions between the anionic chromophore and Glu46. (1) Isomerization of the pCA from 7-trans 9-s-cis conformer to 7-cis 9-s-trans conformer which takes place on a time scale of ps-ns. (2) Intramolecular proton transfer from Glu46 to the anionic pCA chromophore which takes place on a time scale of 250 μs leading to the depratonation of Glu46. (3) The negative charge of COO- of Glu46 triggering a large conformational change resulting in the formation of the long lived signaling state (2 ms). (4) Reisomerization of pCA from 7-cis 9-s-trans to 7-trans 9-s-cis and reprotonation of Glu46. In the P63 crystals, the lack of water molecules induces suppressed dynamics, and the number of steps that has to be retraced by the PYP in order to get back to its global 49 conformation is fewer. The chromophore reisomerization, deprotonation of pCA leads to the resetting of the global dark state of the PYP molecule. The time required by the photoreceptor to complete its photocycle is much lesser in high salt concentration (crystallized state) than in salt-free solution. This partial dehydration of PYP molecule is mimicked by the dissolution of PYP in high salt concentration wherein the ions compete with the protein for the water molecules. Figure 2.12 shows the suppressed structural changes in the P63 crystals due to the lack of water molecules. The partial dehydration due to the presence of salts leads to suppressed conformational changes that can be further observed in the light induced FTIR difference spectra of PYP. The difference absorption spectrum is the dark (pG state) minus the light (pB state) spectra. The difference spectra are measured using the Rapid Scan method and show a strong variation with salt concentration. The spectra are normalized to the 1497 cm-1 band which has effectively the same amplitude throughout pG and pB. The spectrum measured on 0 M salt concentration shows the typical pB-pG difference spectrum with the spectral markers for conformational change (1624 cm-1, 1689 cm-1), chromophore protonation (1497 cm-1, 1515 cm-1) and Glu46 deprotonation (1726 cm-1) (Xie, Hoff et al. 1996a). With increasing salt concentration from 0 M to 4.0 M at this time point the Amide I band intensities decrease, so does that of the 1689 cm-1 band. Besides the amplitude drop the position of the negative Amide I difference band observed at 1641 cm-1 at 0 M salt shifts to 1637 cm-1 in high salt concentration and the positive Amide I band from 1624 cm-1 to 1621 cm-1. The chromophore bands do not change significantly, only a 1.5 cm-1 shift to lower frequencies is observed. The amplitude of the 50 so far unassigned bands change with the salt concentration: the 1610 cm-1, 1550 cm-1 and 1624 1440 cm-1 negative and the 1565 cm-1 and 1660 cm-1 positive bands. 0.0 M NaCl 0.5 M NaCl 1.0 M NaCl 1571 1515 1669 1740 2.0 M NaCl 4.0 M NaCl 1162 1253 1302 1497 1689 1609 0 1726 Diff Absorption [milli OD] 5 1641 -5 1800 1700 1600 1500 1400 -1 1300 1200 Wavenumber [cm ] Figure 2.13: Infrared difference absorption spectra of wt PYP in varying salt concentrations indicating the ionization status of Glu46 (1740 cm-1) which is protonated in the absence of salt and tends to get ionized in the presence of high salt concentration. This implies the suppression of conformational changes in the presence of high salt concentration. Suppressed structural dynamics of PYP The IR difference absorption spectra collected after the laser flash are better studied with the aid of 3-dimensional plots gives us information about wavelength, difference absorption as well as the time scale of the photocycle. These spectra all have similarly large Amide I bands at 1624 cm-1 and at 1641 cm-1 when compared to the chromophore bands. The amplitude of the 1690 cm-1 band has also decreases. The large Amide I and 1690 cm-1 do bands indicate that larger conformational changes take place at this time. Although the listed spectral features are consistent with the pB-pG difference 51 spectra of WT PYP there are alterations at high salt concentrations from the pB-pG spectrum of the salt free sample. The amplitude of the 1624 cm-1 positive and the 1641 cm-1 negative bands are somewhat smaller at high salt concentration. For the lowest 0.5 M concentration the maximum amplitude for the 1624 cm-1 band is 5.2 mOD, for the highest 2.5 M one it is only 0.4 mOD. Also the positive 1660 cm-1 band is virtually missing from the 2 M and the 4 M sample. This suggests that the signaling state of high salt containing PYP sample has slightly different conformation than that of 0 M salt. Combining this information with the kinetics of the Amide I band amplitudes we can say that at the end of the photocycle a pB-like state is formed at every salt concentration but the population of this state is significantly lower at higher concentrations (Figure 2.13). 52 A Time [s ] W ave numb Time C [s ] num be W ave WT PYP in 2.0 M NaCl numb Wave -1 ] r [cm D Diff. Absorbance [milli OD] Diff. Absorbance [milli OD] 53 Time [s ] Diff. Absorbance [milli OD] -1 ] er [cm WT PYP in 1.0 M NaCl B WT PYP in 0.5 M NaCl Diff. Absorbance [milli OD] Diff. Absorbance [milli OD] WT PYP in 0.0 M salt -1 ] er [cm Time [s] W ave numbe -1 ] r [cm WT PYP in 4.0 M NaCl E Figure 2.14: 3-dimentional plots show the effect of varying salt concentrations on the PYP photocycle. Starting with 0 M salt, the effect is more pronounced in the form of suppressed conformational change as the salt concentration is increased from 0.5 M to 4.0 M. Time [s ] Wave e numb -1 ] r [cm Altered proton transfer pathway The vibrational frequency of C═O stretching mode is sensitive to the hydrogen bonding interaction of the carbonyl oxygen with a hydrogen bond donor(s) or hydroxyl group with a hydrogen bond acceptor(s). Without any hydrogen bond, the C═O stretching is high, in the region of 1760 cm-1 (Nie, Stutzman et al. 2005). When hydrogen bonded, the C═O stretching frequency downshifts to a lower frequency. This frequency can be as low as 1710 cm-1 (Nie, Stutzman et al. 2005). Therefore, the C═O stretching frequency of Glu46 can be employed as a spectral probe of the hydrogen bonding interaction between Glu46 and the pCA chromophore. The difference spectrum of PYP in 0.5 M NaCl solution shows the appearance of a positive band at 1740 cm-1 (Figure 2.13). The amplitude of this band increases with increasing salt concentration. This is the frequency at which the protonated carboxyl groups absorb. The only neutral carboxyl group in WT PYP is Glu46. Therefore we assign the 1750 cm-1 positive band to Glu46. The 1726 cm-1 vibration was earlier assigned to protonated Glu46 in pG state (Xie, Hoff et al. 1996a). The negative band at this position means that the protonated Glu46 either become deprotonated or its environment changed therefore its frequency shifted. This feature together with the 1740 cm-1 band suggests that the COOH of Figure 2.15: PYP binds a unique pCA chromophore that is stabilized by the side chain groups of Glu46 and Tyr42 and the backbone of Cys69 via hydrogen bonding interactions. Glu46 group remains protonated but its frequency shifts in the presence of salt. The position of these two carboxylic bands is 54 independent of the salt concentration. According to our previous results (Xie, Hoff et al. 1996a), Glu46 is the proton donor for the anionic chromophore and is negatively charged in the signaling state (pB state). The appearance of the 1740 cm-1 band is a clear indication of the protonation of Glu46 in the pB state. Although this suggests that the Glu46 is no longer the proton donor to the chromophore, the shifting of the 1497 cm-1 band to 1515 cm-1 is an implication of the protonation of the chromophore ring (unpublished results). The phenolic oxygen of the pCA chromophore gets a proton from a different source in order to complete the photocycle. PYP binds a unique pCA chromophore that is stabilized by the side chain groups of Glu46 and Tyr42 and the backbone of Cys69 via hydrogen bonding interactions (Figure 2.13). At this stage, we can only speculate about the source of photon for the chromophore in the circumstance that Glu46 is unable to do so. Effect of Hofmeister cations on PYP The Hofmeister salts also have a profound effect on the proteins as individual ions. To study the anionic effect, several chloride salts with the same concentration were used. The difference spectra obtained (Figure 2.16) shows the same frequency shift of the protonation of Glu46 from 1726 cm-1 to 1740 cm-1. The chromophore protonation bands (1497 cm-1 and 1515 cm-1) also overlap. This gives us a clear indication of the effect of the Cl- anion on the structural changes of PYP during the photocycle. However, the effect of the different cations can be seen in the decrease in amplitude of the Amide I band. The effect on the conformational change while comparing the amplitude and the kinetics in the Amide I bands is seen by 4.0 M NaCl, followed by 4.0 M KCl and then 4.0 M CsCl. 55 KCl is believed to have a higher stabilizing (salting out) effect on protein when compared to NaCl and CsCl which have a higher destabilizing (salting in) effect on the protein. However, we see that the effect of NaCl on the photocycle of PYP is much higher than the effect of the same concentration of KCl. 4.0 M NaCl suppresses the conformational change and accelerates the kinetics of the photocycle (Figure 2.17) more than the equivalent concentration of KCl. This effect is dependent on the radii of the ions, i.e, the charge distribution of the cation. However, it is interesting to note that the effect of Li+ is similar to the effect of Na+. 1624 0.0 M salt 4.0 M CsCl 4.0 M KCl 4.0 M NaCl 1740 1515 1669 1571 4.0 M LiCl 1162 1253 1497 1302 1609 1689 0 1726 Diff Absorption [milli OD] 5 1641 -5 1800 1700 1600 1500 1400 -1 Wavenumber [cm ] 1300 1200 Figure 2.16: Infrared difference absorption spectra of wt PYP in different salts from the Hofmeister series having a common anion Cl- indicating the ionization status of Glu46 (1740 cm-1) which is protonated in the absence of salt and tends to get ionized in the presence of high salt concentration. This implies the suppression of conformational changes in the presence of high salt concentration underlying the impact of the chaotropic chloride anion. 56 [s] er numb W ave WT PYP in 4.0 M KCl 57 C Time [s] Diff. Absorbance [milli OD] WT PYP in 4.0 M NaCl Time -1 [cm ] -1 ] r [cm numbe W ave Diff. Absorbance [milli OD] Diff. Absorbance [milli OD] Time Diff. Absorbance [milli OD] A WT PYP in 0.0 M salt [s] B er numb W ave -1 [cm ] WT PYP in 4.0 M CsCl D Time [s] -1 ] er [cm numb W ave Figure 2.17: 3-dimensional plots indicate the effect of high concentration salts on the photocycle of PYP. The effect of the anion is predominant. But the variation in the suppression of the conformational change is attributed to the effect of the cations which follow the Hofmeister series. Na+ has a stronger effect on the conformational change as compared to K+. Cs+ has the least effect. The above phenomenon can be studied with a microscopic perspective by comparing the entropy of water molecules near monovalent ions to that of water molecules in bulk solution as determined by thermodynamic (Krestov 1962) or dynamic measures such as NMR (Endom, Hertz et al. 1967). The Figure shown is a plot (reproduced from (Collins 1995)) of the entropy of water near monovalent ions (Li+, Na+, K+, Cs+) as calculated from the entropy of hydration of the ion (from dissolving the ion in water) versus the ionic radius of the ion (Krestov 1962). A negative ΔS (upper portion of Fig) indicates tightly bound water that is less mobile than bulk water, whereas a positive ΔS (lower portion of Figure 2.18) indicates loosely held water that is more mobile than bulk water. Figure 2.18: The abscissa is the crystal radii of the ions in angstrom. The entropy of pure water minus the entropy of water near the ion is plotted against the radii. Positive values of ΔS indicate water is more mobile than bulk water. Negative values of ΔS indicate water that is less mobile than bulk water (Krestov 1962). Increasing ion size (decreasing ion charge density) is associated with increasing mobility of nearby water molecules. If this mobile, loosely held water is immediately adjacent to the ion, as suggested by x-ray and neutron diffraction data (Skipper and Neilson 1989), then the horizontal line in Fig. indicating ΔS = 0 separates the strongly hydrated ions (above the line) from weakly hydrated ions (below the line). The ionic radius of Cs+ is greater than K+ and Na+. This implies that the water surrounding Cs+ and 58 K+ are more loosely bound than around Na+ which shows higher suppression of conformational change in during the photocycle of PYP. Hence water molecules have higher mobility in the presence of Cs+ ion than in the presence of K+ or Na+ ions. The effect of Li+ and Na+ on the suppression of structural dynamics of PYP was observed to be similar. This could be due to the fact that their hydration shells have the same number of water molecules (Table 2.1). However, from the above figure it is clear that Li+ binds water more tightly than NaCl. This effect has to be further investigated before drawing any conclusions. Also, since this transition from weak to strong hydration occurs at a larger size for anions than for cations, the anions must be more strongly hydrated than the cations since anions begin to immobilize adjacent water molecules at a lower charge density than do cations. 2.4 Conclusions Suppressed structural dynamics The Amide I infrared absorption bands overlap after normalization (Figure 2.11). The second derivative of the IR Amide I band shows that the main absorption bands do not shift with changing salt concentration as long as the protein is in dissolved state. These results indicate that the environment of the chromophore and the secondary structure of the protein backbone are not changing at different salt levels. Additional indication for the state of the chromophore besides the absorption spectrum are the infrared absorption band positions assigned to the chromophore in the absolute IR spectra and in the light-induced difference spectra . These bands positioned at 1498 cm-1 and 1515 cm-1 are unvaried when the salt concentration is changed (Figure 2.13). The 59 1726 cm-1 band in the IR absorption spectrum and the negative band at the same position in the IR difference spectra has been assigned to the COOH vibration of Glu46. Because of the H-bond between them, this vibration frequency is sensitive to the distance between COOH of Glu46 and the oxygen on the chromophore ring just as it was reported (Brudler, Rammelsberg et al. 2001). Therefore the position of this band may indirectly indicate a change in the environment of the chromophore. In spite of the salt concentration increase in the PYP solution samples; this band position is also unchanged. Altered proton transfer pathway The 1725 cm-1 vibration was earlier assigned to protonated Glu46 in pG state (Xie, Hoff et al. 1996b). The negative band at this position means that the protonated Glu46 either become deprotonated or its environment changed therefore its frequency shifted. This is so because this vibration can be depleted either by deprotonation of Glu46 or with the shift of the frequency of this vibration. Both processes result in the same amount of absorption drop at 1725 cm-1 (Figure 2.13 and 2.16). The 2-D spectra of wt PYP in different salts from the Hofmeister series show large amide I difference signals (Figure 2.16). The amide I band is suppressed in accordance with the strength of the cationic interaction with the water molecules in the solution in a pattern that follows the Hofmeister series. The degree to which the amide I signals are suppressed provide a direct indication of the extent to which these salts suppress the functional structural changes. These experiments demonstrate that functional conformational changes indeed can be suppressed by the presence of precipitating agents that are frequently used for protein crystallization. This would establish that the presence of these salts is a major factor in the suppression of large structural changes in protein 60 crystal. In addition, the data reveals a novel Hofmeister effect in addition to the widely studied effects on protein solubility and stability: the suppression of functional protein conformational changes. PYP in high salt concentrations experiences accelerated kinetics, altered proton transfer pathway and suppressed conformational changes. We find that the presence of salts even at a low concentration of 0.5 M is sufficient to alter the active site proton transfer pathway and suppress conformational changes. Such changes may arise from suppressed mobility of water molecules in the presence of salts. There is a large coupling between the deprotonation of Glu46 and large structural changes. This supports the proposed mechanism that the negative charge on Glu46 serves as the “electrostatic epicenter” that triggers and drives large amplitude protein quake leading to signaling state (pB) formation. Our data also demonstrates that the structure of a transient protein state observed by X-ray crystallography is the same as its counterpart in solution. Time resolved FTIR spectroscopy offers a sensitive and direct method to detect and determine any differences involved. 61 Chapter III Vibrational Structural Makers for Structural Characterization of Tyrosine in Proteins 3.1 Introduction Proteins are dynamic in nature. In order to understand how a protein performs its function based on laws of physics, it is critical to probe and investigate functionally important structural transitions of the protein. Time-resolved Fourier Transform infrared spectroscopy FTIR offers excellent time resolution (picoseconds to seconds), and contains extensive structural information. The real challenge is how to extract structural information from time resolved infrared data. We will report computational methods for developing vibrational structural markers of tyrosine. Using density function theory (DFT) based first principle computational studies combined with experimental data, we found that it is possible to unambiguously determine if the hydroxyl group with the phenolic ring in tyrosine is a hydrogen bonding partner to the nearby amino acids. In addition, we show that it possible to determine the number and nature of hydrogen bonding interactions of a phenolic group in proteins using a combination of C-O stretching and O-H bending frequencies (2D vibrational spectroscopy). The average occurrence of tyrosine in proteins is about 3.2% (Voet and Voet 2004) and they are most often found at the active sites of proteins. The pKa value of tyrosine side−chain group is 10.5 (Voet and Voet 2004) so that these phenol groups are mostly protonated in proteins at steady states. The pKa values of these groups may 62 change dramatically at the active sites of proteins in functional intermediate states, resulting in temporary or permanent deprotonation of these groups and therefore proton transfer between tyrosine residues and other key amino acids. Hydrogen-bonding interactions play key roles in regulating the pKa values of ionizable groups and driving proton transfer. Proteins are made of long chains of α-amino acid sequences. Each of 20 different amino acids consists of a carboxylic (-COOH), an amino (-NH2) groups and a –R group. R-groups distinguish one amino acid from another and bring about different structural properties of proteins. The R Figure 3.1: Tyrosine molecule with phenol functional group group in case of Tyrosine is a phenol ring that is capable of forming 2 hydrogen bonds (one as a donor and the other as an acceptor). The – OH group can also lose a proton ionizing the tyrosine residue. Hence it becomes possible for us to address two fundamental elements in protein structure and function using the same model: (1) Hydrogen bonding; (2) Proton transfer. (The details of proton transfer mechanism are outside the scope of this chapter). 3.2 Tyrosine at active site Tyrosine residues are of functional importance in proteins which can be categorized into those involved in signal transduction (BLUF domain proteins, Green fluorescent protein and photoactive yellow protein); in energy transduction (Bactriorhodopsin and Photosystem II) and in enzymatic reactions (InhA or enoyl-ACP reductase). These proteins have tyrosine residues in their active sites and experimental 63 evidence shows that hydrogen bonding plays a crucial role in the structural dynamics in the proteins hence being responsible for the functional dynamics as well. Proteins involved in Signal transduction The photoactivation mechanism of the BLUF (Blue light sensing using FAD, blue light photoreceptor) domain proteins which are found in heterotrophic and photosynthetic bacteria, involves the signaling state formation via light-driven electron and proton transfer from the conserved Tyr to flavin adenine dinucleotide (Gauden, Grinstead et al. 2007). The best characterized of these proteins is AppA from the purple photosynthetic bacterium Rhodobacter sphaeroides. AppA functions as an anti-repressor of photosynthetic gene expression responding to high levels of blue light and oxygen scarcity (Braatsch, Gomelsky et al. 2002). Signal transduction in BLUF proteins is initiated by blue light absorption by flavin (an active cofactor of BLUF domain) and subsequent conversion from the dark-adapted state (dark state) to the light-induced signaling state (light state). The flavin isoalloxazine ring is surrounded by several polar amino acids, forming H-bond networks. The Tyr21 residue in the vicinity of flavin is conserved among BLUF domains and mutagenesis studies of this Tyr residue shows that it is indispensable for the proper photoreactions (Laan, van der Horst et al. 2003). The oxidation of Tyr21 by the excited singlet-state FAD* and subsequent formation of a TyrFADH radical pair are essential mechanisms of the BLUF reaction (Gauden, van Stokkum et al. 2006). Thus, Tyr plays a key role in the electron transfer mechanism. Another well studied system is the photoactive yellow protein (PYP) from Ectothiorhodopsira halophila, again a blue-light bacterial photoreceptor. On light absorption, PYP undergoes a photocycle containing several intermediates that is closely 64 linked to the function of the protein (Imamoto, Kataoka et al. 1996, Meyer, Yakali et al. 1987). Its chromophore (p-coumaric acid) is deprotonated in the ground state. Tyr42 is crucial for stabilizing the native conformation of the chromophore through hydrogen bonding interaction in the receptor state (pG) (Borgstahl, Williams et al. 1995a, Getzoff, Gutwin et al. 2003). In the Tyr42->Phe mutant, hydrogen bonds of the phenolic oxygen of Tyr42 with the phenolic oxygen of the chromophore are lost. This leads to reduced protein stability and possibly a less rigid protein that gives rise to a second PYP population with an altered chromophore conformation as shown by UV/ visible and FT Raman spectroscopy (Brudler, Meyer et al. 2000a). Green Fluorescent Protein (GFP) is a bioluminescence protein from the jelly fish Aequorea victoria. The chromophore is formed by the cyclization of an internal Ser65Tyr66-Gly67 tripeptide (van Thor, Pierik et al. 1998a) wherein the deprotonated phenolate of Tyr66 at the active site is the cause of fluorescence (Cubitt, Heim et al. 1995). GFP can exist in at least two spectroscopically distinct states: GFP395 and GFP480, with peak absorption at 395 and 480 nm, respectively, resulting from a change in the protonation state of the phenolic ring of its chromophore. When GFP is expressed in Escherichia coli, its chromophore is mainly present as the neutral species. UV and visible light convert GFP from this neutral form into the anionic form (van Thor, Pierik et al. 1998b). The equilibrium between these states is governed by a hydrogen bond network that permits proton transfer between the chromophore and neighboring side chains. The photoconversion of GFP involves a proton transfer event initiated by the deprotonation of the phenolic chromophore which leads to the rearrangement of the hydrogen bonding network in the protein and protonation of Glu222 (Yoo, Boatz et al. 2001). 65 Proteins involved in Energy transduction Bacteriorhodopsin is a purple membrane protein, a well-studied light-driven proton pump for solar energy transduction. The protein has 11 residues of tyrosine, of which only Tyr185 is structurally active. It gains a proton in the first photoreaction, bR>K630 and loses it in the second stage, bR->M412 (Braiman, Mogi et al. 1988c). This residue is responsible for stabilizing Asp212 via a strong hydrogen bond in bR550 (Rothschild, Braiman et al. 1990) and M412 intermediate state (Ames, Ros et al. 1992). The mutation of Tyr-Phe eliminates this (Braiman, Mogi et al. 1988b, Rothschild and Marrero 1982), hence giving conclusive proof of the importance of hydrogen bond of the tyrosine residue. 66 A B Y66 G67 H148 Y92 + Y145 S65 E222 C Y185 RET D212 K126 Figure 3.2: Proteins with tyrosine at the active site. (A)Photoactive Yellow Protein, PDB code: 1NWZ; (B) Green Fluorescent Protein, PDB code: 1EMA; (C)Bacteriorhodopsin, PDB code: 1C3W 67 Photosystem II (PS II) of plants and cyanobacteria is the site of light-induced oxidation of water to molecular oxygen, an essential step in photosynthesis. A tyrosine (TyrZ, D1-Tyr161) of the D1 polypeptide of PS II is a redox intermediate between the chlorophylls that constitute the primary donor of PS II and the Mn cluster responsible for water oxidation. Models have been recently presented in which TyrZ is directly involved in proton or hydrogen atom abstraction from water (Force, Randall et al. 1995, Gilchrist, Ball et al. 1995). There is a second redox-active tyrosine (TyrD) on the homologous polypeptide D2 of PS II that is stable in its oxidized form (Faller, Debus et al. 2001). The proton generated by the TyrD radical is thought to remain in its vicinity having an electrostatic influence on the location and potential of the chlorophyll cation, P+. This effect is believed to be important for the kinetics of TyrZ oxidation and to provide a significant thermodynamic boost to the enzyme. In addition, through its electrostatic influence, TyrD(H+) may confine the highly oxidising cation P+ to the chlorophyll nearest to TyrZ, thereby accelerating TyrZ oxidation (Rutherford, Boussac et al. 2004). In wild-type PS II, TyrD* is hydrogen bonded to the side chain of D2- His189 (Rodriguez, 1987) (Hienerwadel, Boussac et al. 1997). TyrZ* is also hydrogen bonded (Force, Randall et al. 1995, Un, Tang et al. 1996) and the hydrogen bond donor(s) to TyrZ is D1-His190 (Berthomieu, Hienerwadel et al. 1998). Proteins involved in enzymatic reactions InhA, the enoyl-ACP reductase from Mycobacterium tuberculosis catalyses the NADH dependent reduction of long chain trans-2-enoyl-ACP fatty acids (Rozwarski, Vilcheze et al. 1999). The catalytic mechanism of InhA involves the role of two conserved amino acids, Tyr158 and Lys165. This enzyme is inhibited by the anti- 68 tubercular drug isoniazid. Hence studying the catalytic mechanism of this enzyme and identifying the amino acid residues involved in catalysis can give an insight into the development of novel anti-tubercular drugs. Tyr156 is conserved within the enoyl reductase family and is involved in the chemical steps of the reduction reaction. It functions as a hydrogen bond donor and as an acceptor hence provides electrophilic stabilization of the transition state for the reaction by hydrogen bonding to the carbonyl of the substrate. The crystal structure (Rozwarski, Vilcheze et al. 1999) also supports this theory of tyrosine being the elctrophilic catalyst. Replacement of Tyr156 by Phenylalanine results in 24 fold decrease in kcat, hence establishing the catalytic function of the tyrosine residue (Parikh, Moynihan et al. 1999). Tyrosine can also be phosphorylated by protein kinases and is a key step in signal transduction and regulation of enzymatic activity. It is a precursor of the neurotransmitters epinephrine, norepinephrine and dopamine, all of them extremely important in the brain and transmits nerve impulses and prevents depression. These examples show that buried tyrosine side−chain group may change their protonation states during the functional processes of proteins. In this chapter, we focus on developing a vibrational structural marker for probing the hydrogen-bonding status of buried neutral tyrosine side−chain group, including those that change their protonation states during the functional processes. 3.3 Vibrational structural markers Hydrogen bonding is an important element in protein structure and function. The hydrogen bond dissociation energy in proteins is in the order of 10-40kJ/mol. A typical value for protein folding energy is approximately 40kJ/mol which is about one to four 69 times the hydrogen bond dissociation. From this, we can see that even by breaking one hydrogen bond; we can impair the protein stability. The phenol group of tyrosine can form up to two hydrogen bonds with a neighboring hydrogen bond donor and acceptor as shown below. The phenolic oxygen and the hydroxyl hydrogen of a phenol group may each form one hydrogen bond. It is therefore energetically preferable for tyrosine to form as many hydrogen bonds as possible. A B C Figure 3.3: Tyrosine molecule with (A) one hydrogen bond with phenolic oxygen as acceptor; (B) one hydrogen bond with hydroxyl hydrogen as donor (C) two hydrogen bonds, one as acceptor and other as donor The Vibrational Structural Marker (VSM) described here offers a generally applicable spectroscopic tool for probing the hydrogen bonding status of tyrosine and can be extended to other phenolic groups of amino acids present in different proteins. Vibrational structural markers need to exhibit two general properties. (i) The markers should be well resolved, and should not overlap significantly with other signals. (ii) The signal should allow clear experimental conclusions to be drawn on functionally important structural changes in the protein, i.e, the marker should be sensitive to distinct structural changes in the functional group (like deprotonation). It is challenging to monitor the hydrogen bond states of tyrosine residues for functional intermediate states of proteins and for proteins that their X-ray crystal structures are not available. In this chapter we report that the C−O stretching frequency is a sensitive infrared structural marker for detecting and monitoring hydrogen-bonding 70 status of tyrosine side−chain group. When the C−O stretching frequency is inadequate to distinguish zero and two hydrogen bonds, two dimensions of C−O stretching frequency combined with O−H bending or stretching frequency may be used to make such distinction. This vibrational structural marker is based on ab initio computational studies and supported by experimental data. The applications of vibrational structural markers are expected to enhance the power of time-resolved infrared difference spectroscopy for structural characterization of functionally important intermediate states of proteins, and consequently for understanding the functional mechanism of proteins. 3.4 Experimental and Computational methods to determine VSM Computational method A 4-propyl-phenol molecule was employed to model the side−chain of neutral Tyr residue in proteins, as shown below. Both energy and vibrational frequency calculations were performed in vacuum using ab initio methods based on density function theory (Frisch, Frisch et al. 2003). Water molecules and molecules that model protein Figure 3.4: Model compound for calculation of vibrational frequencies of tyrosine backbone and side−chain groups of polar and charged amino acids were utilized to serve as hydrogen-bond donor and/or acceptor for Gaussian03 (Frisch, Frisch et al. 2003) calculations of hydrogen-bonding properties of tyrosine interacting with these molecules. The notation, OH−X, represents that the phenolic oxygen is hydrogen bonded to a proton−donor molecule. Similarly, OH−Y indicates that the hydroxyl hydrogen is hydrogen bonded to a proton−acceptor molecule (Y). The notation, OH−XY denotes that 71 both the phenolic oxygen and hydrogen are hydrogen bonded, one molecule being the donor(X) and the other being the acceptor(Y). To calculate the hydrogen bond dissociation energy defined as ΔE=EPHE + EAA – EPHE,AA PHE stands for the 4-propyl-phenol molecule and AA for amino acid hydrogen bonding partners. The structure of the model compound for tyrosine was first constructed in ChemDraw, the co-ordinates generated in Chem 3D and the initial structure optimized using an empirical method (PM3). The geometry was then optimized from first principles using Gaussian 03 using B3LYP/6-31G(d) after which the energy was calculated using B3LYP/6-311+G(2d,p). The resulting output energy is in atomic unit (au), which is converted into kJ/mol with 1au=2619.6 kJ/mol (Nie, Stutzman et al. 2005). Vibrational frequency calculations were carried out in several steps. First, the structure was optimized using B3LYP/6-31G (d) method. Then all the force constants for vibrational motion were calculated using the same method and the vibrational modes were computed. Calculated frequencies are systematically higher than experimental values. A scaling factor is recommended for each type of computational method to compensate for the over estimation of force constants (Foresman and Frisch 1996). The scaling factor varies with the method and for B3LYP/6-31G (d) the recommended scaling factor is 0.9613 (Foresman and Frisch 1996), representing ~3.9% reduction from the calculated vibrational frequencies (Nie, Stutzman et al. 2005). Finally GaussView was 72 employed to view the vibrational motions of any chosen mode from the output file of Gaussian03. A B C Figure 3.5: (A) Initial structure from ChemDraw; (B) Geometry optimized using empirical method (PM3) from Chem3D (C) Geometry optimized using DFT in Gaussian03 There are 3 × N – 6 calculated vibrational frequencies corresponding to 3 × N – 6 vibrational motions, where N is the number of atoms in each structure. A Fortran90 code was written (Nie 2006) to generate a 2D data set (wavenumber vs. intensity of vibrational frequency) using Gaussian function: ( x − xν ) 2 Y (ν ) = I (ν ) * exp( − ) d2 where I(ν) is the intensity of the vibrational frequency Xν in Gaussian03 output and FWHM (full width at half maximum) is 2 ln 2d where d was set to be 2 cm−1. Experimental Method A Bruker IFS 66v/s FTIR spectrometer with a Michelson interferometer was utilized for rapid-scan measurements. Three samples of empty, tyrosinol in H2O and tyrosinol in D2O were loaded to a custom-made sample exchanger driven by step motor driver (Si3540). Then the sample chamber was purged with nitrogen gas to get rid of water vapor and is maintained at a constant temperature of 300K using water circulating temperature controller (Neslab RTE 111). The optics chambers of the spectrometer were evacuated to eliminate water vapor along optical path. The step motor driver is externally triggered by the spectrometer in rapid-scan mode to synchronize the data collection and 73 changing sample. In this way, the quality of infrared spectra was largely improved due to little environmental change between the samples and the background (empty). The signal to noise ratio was large after 256 averages. The scanning rate was chosen to be 40 kHz and the spectral resolution to be 2 cm−1. Identify the number and the type of hydrogen-bond interactions from protein crystal structures The pdb (protein data bank) files for high resolution crystal structures of proteins were obtained from the RCBS Protein Data Bank. To determine the number and the type of hydrogen bonding interactions for each buried phenol group, we first examined and identified all the plausible hydrogen-bond donors and/or acceptors within the hydrogen bonding distance of a tyrosine side−chain group using PyMOL. Next, we checked the distance, bond angle and dihedral angles between the potential hydrogen bonding partners. A hydrogen bond is expected to meet all three criteria: less than 3.2 Å for the hydrogen-bond length, 120 ± 20º for the CZ−O…X angle, and 150 to 180° or 0 to 30° for the CE−CZ−O…X dihedral angle. X stands for a heavy atom that is hydrogen bonded to a phenol group. Finally we examined the compatibility of hydrogen bonding interaction. For example, it is impossible for a hydrogen-bond acceptor to form a hydrogen bond with another hydrogen-bond acceptor. 3.5 Results 2D Infrared Probe for Hydrogen Bonding of Tyr As the first step for assigning the vibrational structural marker for hydrogen bonding of tyrosine, we calculate the vibrational frequencies of the isolated model molecule and its subsequent cases of hydrogen bondings: 74 1. Tyrosine forming hydrogen bond with water molecule as proton donor, 2. Tyrosine forming one hydrogen bond with water molecule as acceptor, 3. Tyrosine forming two hydrogen bonds with two water molecules as donor and acceptor. The comparision of calculated vibrational frequencies are as shown in Figure 3.6 and the band assignment of tyrosine side chain is shown in Table. The vibrational frequencies of tyrosinol in H2O and D2O obtained using FTIR spectroscopy are also listed. Locating bands that are sensitive to hydrogen-bonding interactions In the fingerprint region, there are two bands of OH−Y (Figure 3.6) that their frequencies largely shift due to hydrogen-bonding interactions: 1255 cm−1 down−shifted to 1235 cm−1 and 1162 cm−1 up−shifted to 1170 cm−1. Using GaussView 2.1 for Windows (Gaussian, Inc.), we identified that 1255 cm−1 was attributed to C−O stretching and 1162 cm−1 to O−H bending modes. When the hydroxyl hydrogen of tyrosine forms a hydrogen bond with the water molecule as proton donor (OH−Y in Figure 6c), the frequency of C−O stretching up-shifted from 1255 cm−1 to 1271 cm−1 and the frequency of O−H bending also up-shifted from 1162 cm−1 to 1221/1211 cm−1. The 1221/1211 cm−1 doublet is due to coupled O−H bending and C−H bending on the main chain. Noticeably there is one more frequency shifted from 1327 cm−1 to 1350 cm−1 in the Figure 6c that is attributed to C−C stretching on the ring coupled O−H bending. This frequency is insensitive to the hydrogen-bonding interaction in Figure 3.6. The vibrational modes of 1327 cm−1, 1255 cm−1, and 1162 cm−1 are shown in Figure 3.6. 75 3500 1272 1350 1163 1171 1234 3100 -1 2900 1600 1353 D 1500 1400 1255 3361 3300 1222 1211 Intensity 3408 C C D 1327 B B Intensity 3612 1236 1327 3606 1255 A A 1300 1200 1100 -1 Wavenumber [cm ] Wavenumber [cm ] Figure 3.6: The calculated vibrational frequencies in the region of 2870 – 3630 cm−1 1030 – 1650 cm−1 (A) of isolated 4-propyl-phenol (B); one hydrogen bond OH-X (C); one hydrogen bond OH-X (D); and two hydrogen bonds OH-X with water molecule(s) . In principle, the frequency 1327 cm−1 of C−C stretching on the ring coupled O−H bending may be considered as a vibrational structural marker of the type of hydrogenbonding interaction of tyrosine: it shifts less than 2 cm−1 for OH−X and it up-shifted to 1350 – 1357 cm−1 for OH−Y. However, this frequency is not a good marker for probing hydrogen-bonding status of tyrosine side−chain for three reasons: (1) the frequency shift of 2 cm−1 is so small between isolated tyrosine and OH−X that it is difficult to distinguish zero hydrogen bond and one hydrogen-bonding interaction of OH−X; (2) the frequency shift upon deuteration is insensitive to hydrogen- bonding interaction; (3) The intensity of this frequency in experimental data is small in both absorption and second derivative spectra(Figure 3.7). Therefore, we focused on the C−O stretching and O−H bending 76 frequencies that are sensitive to the hydrogen- bonding interaction in the fingerprint region. 0.8 Absorbance A 0.6 Figure 3.7: (A) The absorption spectrum (B) and the second derivative of L-tyrosinol in H2O at pH =0.5 (black) and in D2O at pH*=0.2 (red). The C-O stretching band at 1264 cm-1 is clearly visible and so is the O-H bending band at 1246 cm-1. 0.4 0.2 0.0 1055 1048 1178 1264 1258 1246 1454 1517 1429 1615 1599 1589 Second Derivative B 1700 1600 1500 1400 1300 Wavenumber [cm-1] 1200 1100 1000 3.6 Discussion To explore, evaluate, and determine the qualification of the C−O stretching and O−H bending frequency as vibrational structural markers for detecting the hydrogenbonding status of tyrosine, we performed a range of calculations of tyrosine side−chain group hydrogen-bonding interacting with water, protein backbone, and polar and charged side−chain groups of amino acids. These side−chain groups include the possible hydrogen bonding partners for Tyrosine in any given protein system. We used 10 polar side−chain groups (Ser, Thr, Cys, Met, Asn/Gln, Tyr, Asp/Glu, His, Lys, and Arg), 2 77 negatively charged side−chain groups (Tyr and Asp/Glu) and 3 positively charged side−chain groups (Arg, His, and Lys). Tables 3.1 and Table 3.2 show the computational results as well as the structures of molecules that are employed to model the different side−chain groups of polar and charged amino acids. For the convenience of discussion, we will refer these model compounds as their corresponding amino acids. Table 3.1 shows the calculated hydrogen-bonding properties of tyrosine side−chain group interacting with water, backbone, and neutral polar amino acids. There is a geometric component involved in hydrogen bonds, and for single donor acceptor systems, such as N-H---O and O-H---O, the strongest hydrogen bonds are collinear (Creighton and Freedman 1993). Electrostatic calculations suggest that deviation of 20° from linearity leads to a decrease in binding energy of approximately 10% (Thompson and Pimentel 1960). For an isolated phenol group without any hydrogen-bonding interactions, the calculated C–O stretching frequency is 1255 cm−1. However, when the phenolic oxygen of tyrosine forms one normal hydrogen bond with a polar group (OH−Y), all six computational results (from Arg to His) show that the C–O stretching frequency is red– shifted from 1255 cm−1 to 1235–1239 cm−1. The strength of these hydrogen bonds is in the range of 8.7–16.7 kJ/mol. In the case of hydrogen-bonding interaction, Asp/Glu–1, this frequency is further red–shifted to 1230 cm−1 and the hydrogen-bonding strength is stronger (23.2 kJ/mol). This is due to an additional weak hydrogen bond between carbonyl oxygen of Asp/Glu and C–H on the ring of tyrosine (Desiraju and Steiner 1999). When the hydroxyl hydrogen of tyrosine forms one normal hydrogen bond with a polar group (OH−X), nine computational results (from His to Asp/Glu) show that C–O 78 stretching frequency is blue–shifted from 1255 cm−1 to 1264–1277 cm−1. The strength of these hydrogen bonds is in the range of 23.0–38.4 kJ/mol, much stronger than that of OH−Y (8.7–16.7 kJ/mol), indicating OH−X type of hydrogen-bonding interaction is more stable than OH−Y type of hydrogen-bonding interaction. When a phenol group forms two hydrogen bonds, the C–O stretching frequency is 1252 – 1255 cm−1, fairly close to the C–O frequency of isolated phenol group. It is expected to be very close to the C–O stretching frequency of a phenol group with zero hydrogen bond because of combined hydrogen bonding effects on the C–O stretching frequency: average red shift of 18 cm−1 for OH−X and blue shift of 16 cm−1 for OH−X. The average hydrogen-bond strength is 22.9 kJ/mol per hydrogen bond for two hydrogen-bonding interactions. The results discussed above are based on well-formed hydrogen bond(s) with neutral polar groups. Noticeably when two tyrosine side−chain groups form one hydrogen bond, the hydrogen-bond strength is stronger than that of OH−Y and weaker than OH−X due to the coupling of two phenol groups. The C–O stretching frequency therefore is further red−shifted to 1231 cm−1 and less blue−shifted to 1261 cm−1. There are four special cases in Table 3.1 and 3.3 that need to be addressed. The first special case is the hydrogen-bonding interactions with Cys and Met, shown in Table 3.1. The hydrogen-bond strength for Cys−A is 5.1 kJ/mol, much weaker than other OH−Y type of hydrogen-bonding interactions (average of 12.7 kJ/mol). The hydrogenbond strength for Cys−B and Met−B is 14−17 kJ/mol, also much weaker than other OH−X type of hydrogen-bonding interactions (average of 30.7 kJ/mol). The hydrogenbond dissociation energy of these hydrogen-bonding interactions is approximately half of 79 the average of calculated hydrogen-bond dissociation energy for OH−Y and OH−X. This is due to the fact that sulfur atom is large so that the hydrogen bond is long and weak, and that the hydrogen-bond strength of OH−X is weaker interaction than that of OH−X. Therefore, this hydrogen-bonding interaction is classified as half hydrogen bond or weak hydrogen bond. The C−O stretching frequencies for Cys−A, Cys−B, and Met−B are 1243 cm−1, 1261 cm−1, and 1259 cm−1, respectively. These frequency shifts are less but in consistency with the trend of C−O stretching frequency shift due to hydrogen-bonding interactions. The second special case is the hydrogen-bonding interactions with neutral Arg and Lys, shown in Table 3.1. The pKa value of Arg and Lys side−chain is 12.5 and 10.5 (Nelson and Cox 2000). Therefore they are normally charged in proteins. The case that Arg or Lys is neutral is fairly rare. Therefore, we do not include hydrogen-bonding interactions with neutral Arg and Lys into our classification of vibrational structural marker for probing hydrogen-bonding status of tyrosine side−chain group. We will address the special case that the hydrogen-bond partner is positively charged Arg+, Lys+, and His+ below. The third special case is dealing with two deformed hydrogen bonds. Asn/Gln and protonated Asp/Glu may take part in hydrogen-bonding interactions both as proton donor and proton acceptor. When a phenol group forms two hydrogen bonds with these groups as illustrated in Table 3.1, these two hydrogen bonds are deformed due to geometrical constraints. Their hydrogen bond angles are 158°/137° for Asn/Gln and 145°/154° for Asp/Glu, largely deviated from the optimal hydrogen-bond angle of 170° − 180°. Such 80 deformations lead to reduced hydrogen-bond dissociation energy, ~35.3 kJ/mol. In comparison with 22.9 kJ/mol per hydrogen bond for two well-formed hydrogen-bonding interactions (2 H2O in Table 3.1), the hydrogen-bond dissociation energy for two deformed hydrogen bonds, 35.3 kJ/mol, is approximately 1.5 times of 22.9 kJ/mol. With proteins having double acceptor systems, as in tyrosine residues, bifurcated hydrogen bonds with non-linear angles are preferred. The occurrence of hydrogen bonds in protein structure has been extensively reviewed by Baker and Hubbard (1984). They found that 90% of N-H---O bonds in proteins lie between 140 and 180°, and that they are centered around 158°C. For C=O---H, the range is more broadly distributed between 90° and 160° and centered around 129°. Therefore, we classify such two deformed hydrogen bonds (Asn/Gln and Asp/Glu) as forming 1.5 hydrogen bonds. The fourth special case is that a phenol group forms hydrogen bond(s) with a charged side−chain group. Most charged groups in proteins are solvent exposed. However, buried charged groups have been found in the active sites of proteins. Therefore, we examined the hydrogen-bonding interactions of tyrosine side−chain with positively charged side−chains (Arg+, Lys+, and His+) and with negatively charged side−chains (COO− of Asp/Glu or a negatively charged phenolic oxygen of Tyr). 81 TABLE 3.1 Calculated hydrogen-bonding properties of a phenol group interacting with neutral groups H-bond H-bond H-bond † † No. of Structure Amino Acid* Length Angle H-bond† (Å) Energy‡ νC-O νO-H Bending νO-H Stretching (ε = 1 /2 /4) (cm-1) (cm-1) (cm-1) (deg) (kJ/mol) 0 N/A N/A N/A 1255.5 1162.9 3606.1 CYS − A ~0.5 3.65 150.8 5.1/0.09/-3.9 1243.2 1162.6§ 3607.7 ARG − A 1 3.11 174.4 11.9/8.2/4.4 1239.1 1164.4§ 3609.0 1 3.06 172.1 10.7/12.0/6.6 1236.4 1164.1§ 3609.6 1 2.9 159.9 11.9/9.7/3.7 1234.3 1161.1 3614.4 82 TYR ASN/GLN − A THR − A 83 SER − A 1 2.91 162.8 11.2/5.4/-0.9 1236.0 1166.2§ 3612.9 BB – A 1 3.09 171.4 13.1/9.3/4.5 1235.9 1164.6§ 3608.8 H2O – A 1 2.93 158.8 14.8/8.7/3.5 1235.5 1166.8§ 3612.4 HIS − A 1 3.01 173.8 16.7/11.3/5.4 1235.8 1170.2§ 3611.8 TYR − A 1 2.87 167.2 18.5/11.3/4.6 1230.8 1166.0§ 3614.1 1 2.83 167.2 23.2/13.6/4.6 1229.8 1155.0 3614.7 HIS − B 1 2.85 173.8 38.1/31.3/23.2 1274.8 1230.2 3236.5 LYS − B 1 2.81 168.2 34.9/29.0/23.1 1276.1§ 1244.6 3105.7 ASP/GLU − A ASN/GLN − 1 2.77 165.4 26.9/18.2/6.6 1273.1 1227.3 3312.0 1 2.82 179.0 23.0/17.9/12.8 1271.8 1221.5 3408.4 1221.3 3427.8 1 2.81 158.4 27.7/11.3/-3.8 1268.9§ 1210.6 3383.8 B H2O – B ASP/GLU – B 84 SER − B 1 2.79 172.7 24.1/17.7/10.7 1267.6 1225.7 3370.7 THR − B 1 2.78 167.7 26.8/23.1/15.7 1266.2 1228.9 3364.4 BB – B 1 2.77 171.0 33.6/12.4/6.4 1265.5 1227.6 3367.8 1 2.84 173.6 24.0/11.3/-3.8 1263.5§ 1222.1§ 3447.5 MET − B ~0.5 3.42 167.8 14.4/8.55/1.36 1261.4 1197.5 3440.7 TYR − B 1 2.87 167.2 18.5/11.3/4.6 1260.9 1201.0 3497.6 ASP/GLU − B CYS − B ~0.5 ARG 1.5 ASN/GLN ASP/GLU 85 2 H2O 2 SER 3.35 149.1 2.72 157.4 3.03 144.8 2.72 157.5 1.5 2.94 136.6 2.72 145.0 1.5 2.77 154.0 2.77 173.1 2 2.89 164.7 2.76 173.1 2 2.87 * 17.1/8.5/1.1 1258.7 1194.6 3452.0 48.4/35.8/25.1 1265.8 1249.6 3028.2§ 33.0/24.0/11.4 1257.5§ 1235.9 3221.9 37.5/19.4/3.56 1245.9 1221.3 3338.9 3252.9 45.8/35.7/27.5 1254.7 1233.8 3361.0 39.3/30.5/22.0 1251.6 1237.1 3332.2 166.0 The structures presented here are optimized using B3LYP/6-31G(d) method. Color codes for atoms: black for carbon atoms, red for oxygen atoms, blue for nitrogen atoms, yellow for sulfur atoms, and white for hydrogen atoms. The dashed lines represent hydrogen bonds. His−A indicates that the phenolic oxygen of a phenol group forms a hydrogen bond while His−B designates that the hydroxyl hydrogen of a phenol group forms a hydrogen bond. The same notation is used for other amino acids. BB is the abbreviation of backbone. † The hydrogen-bond length is measured between the heavy atoms of a pair of hydrogen-bond donor and acceptor. When two hydrogen bonds are formed, the upper value is for hydroxyl hydrogen and the bottom value is for phenolic oxygen. ‡ The vibrational frequencies were calculated using B3LYP/6-31G(d) method on optimized structures. The energy was computed using B3LYP/6-311+G(2d,p) method. § The vibrational frequency was the average of two coupled vibrational modes that their frequencies are next to each other. 86 In the case that a phenol group forms a hydrogen bond with a positively charged side−chain of His or Lys (His+ or Lys+ in 1.2), the strength of such a hydrogen bond is very strong, 60 − 71 kJ/mol. This is approximately three times as strong as the average hydrogen-bond strength of a phenol group interacting with one polar neutral hydrogenbonding partner, ~22.9 kJ/mol per hydrogen bond. When a phenol group is hydrogen bonded to a positively charged side−chain of Arg, two deformed hydrogen bonds are formed with the phenolic oxygen. The hydrogen-bond dissociation energy for these two deformed bonds is 59 kJ/mol, similar to that of single hydrogen-bonding interactions with His+ or Lys+. The C–O stretching frequency is largely red–shifted from 1255 cm−1 to 1194–1204 cm−1. Therefore, a low C–O stretching frequency (1194 to 1204 cm−1) indicates strong hydrogen-bonding interactions with positively charged side−chain groups. In the case that the hydrogen-bond partner is negatively charged (Asp/Glu− or Tyr−), our calculations lead to proton movements. Starting from an initial structure for a hydrogen-bonding interaction between a phenol group and a COO− group or a Tyr−O− group, we found that the proton on tyrosine is shared between the two groups after structural optimization. Since negatively charged side−chains do not have hydrogen-bond donors, no hydrogen bond is formed with the oxygen of a phenol group. When the hydrogen bonding distance is fixed at 2.6A and then at 2.7A to curb proton transfer, independent bending of O-H disappears and we see it being coupled with COOstretching frequency at 1672 cm-1 (Nie, Stutzman et al. 2005). 87 TABLE 3.2 The hydrogen bonding interactions of neutral Tyrosine with positively charged amino acid side chains H-bond H-bond H-bond † † No. of Structure Amino Acid* Length Angle H-bond† (Å) Energy‡ νC-O νO-H Bending νO-H Stretching (ε = 1 /2 /4) (cm-1) (cm-1) (cm-1) (deg) (kJ/mol) 88 HIS+ 1 2.78 167.3 59.9/30.1/12.0 1202.8 1167.4 3595.5 LYS+ 1 2.74 164.5 71.0/45.5/25.7 1198.0 1159.7§ 3593.8 2.90 149.3 ARG+ 2 58.7/31.9/15.7 1194.1 1177.8 3585.6 2.92 147.6 TABLE 3.3 The hydrogen bonding interactions of neutral Tyrosine with negatively charged amino acid side chains Structure Amino Acid* No. of H-bond† H-bond Length† (Å) H-bond Angle† (deg) H-bond Energy‡ (kJ/mol) νC-O (cm-1) νO-H Bending (cm-1) νO-H Stretching (cm-1) GLU- 1 2.55* 171.1 105.2 1257.6 2564.9§ 1652.3 GLU- 1 2.54* 169.8 106.2 1303.0 1489.9§ 2282.5 89 * The first structure is optimized with no constraints and the hydrogen bonding distance between the two oxygens involved in the bond is 2.55 Å. This shows a very unique shift in the O-H stretching frequencies when compared to the previous calculations. However, when the distance between the α carbons of the two groups are fixed, the hydrogen bond distance optimizes at 2.54 Å. The first calculation was done by the method previously mentioned. We observed that the frequencies of O-H bending and the O-H stretching get interchanged. However, when we fixed the distance between the hydrogen bonding partners, i.e., the distance between the hydrogen of tyrosine and oxygen of Glu-, the values were as tabulated. § indicates that the O-H bending frequency was observed to be highly coupled. Overall, the hydrogen-bond strength of tyrosine side-chain group is stronger for OH−X than that of for OH−Y with neutral polar groups. The hydrogen-bond strength of OH−X with positively charged side−chain groups is stronger than that of OH−X. Similar pattern was observed for the vibrational structural marker of protonated carboxylic group (Nie, Stutzman et al. 2005). These computational results support that the C–O stretching frequency is a sensitive vibrational structural marker for probing the hydrogen-bonding status of tyrosine side−chain group: it is 1255 cm−1 for zero hydrogen bond, 1230−1236 cm−1 for one hydrogen bond OH−X and 1264−1277 cm−1 for one hydrogen bond OH−X with neutral polar group, and 1194−1204 cm−1 for one hydrogen bond OH−Y with positively charged side−chain group. The case that a phenol group forms two deformed hydrogen bonds in proteins is fairly rare because of structural constraints. In fact, most hydrogenbonding interactions found in crystal structures of proteins are single hydrogen bond (Nie, Stutzman et al. 2005). In the case of single hydrogen-bonding interactions with Cys (1259 cm−1) or Met (1261 cm−1) or coupled Tyr–Tyr (1261 cm−1), and two hydrogenbonding interactions (1252–1255 cm−1), it is difficult to use the C–O stretching frequency 90 to distinguish them from zero hydrogen bond (1255 cm−1) and one hydrogen bond OH−X (1264−1277 cm−1). Therefore, additional vibrational information will be needed. Two-dimensional infrared spectroscopy As shown in Table 3.2 and 3.3, the O–H stretching and bending frequencies are also sensitive to hydrogen-bonding interactions. However, O–H stretching and bending frequencies can not be used to distinguish zero hydrogen bond from one hydrogenbonding interaction of OH−X with neutral polar group or with positively charged side– chain group. When both O–H frequency (either stretching or bending) and C–O stretching frequency are used in a two-dimensional (2D) plot (see Figure 3.5), six categories are well separated by either C–O stretching or O–H frequency or both: zero hydrogen bond (■), one hydrogen bond OH–X with neutral polar groups (□), one hydrogen bond OH–X with positively charged groups (●), one hydrogen bond OH–Y with neutral polar groups (○), one hydrogen bond OH–Y with Cys or Met or Tyr (▲), and two hydrogen bonds OH–XY with neutral polar groups (). Due to the fact that strong O–H absorption from solvents overlaps with the O–H stretching modes from the tyrosine side–chain groups into consideration, the O–H stretching frequency is understudied. In principle, this frequency range can provide valuable structural information for amino acid side–chains. The use of protein samples in hydrated film or crystal form may greatly reduce solvent absorption in this region. Under these conditions, 2D plot of O–H stretching and C–O stretching frequencies can be helpful. In other circumstances, 2D plot of O–H bending and C–O stretching frequencies is more suitable to provide information on hydrogen-bonding status of tyrosine side–chain groups in proteins. 91 With no hydrogen bond, the C–O stretching frequency is 1255cm-1, red-shifts to a range between 1240-1225cm-1 with the formation of one hydrogen bond as the acceptor and further red-shifts to the range of 1205-1190cm-1 with hydrogen bonding to a positive charge donor. The OH bending frequency however shifts with the formation of hydrogen bond with an acceptor. It blueshifts from 1162cm-1 for 0 hydrogen bond to 1185-1195 cm-1 with the formation of weak hydrogen bond with acceptor and further blueshifts to 12151230 cm-1 with the formation of a strong hydrogen bond with an acceptor. With the formation of two hydrogen bonds, OH bending frequency is again in the range of 1215-1230 cm-1 whereas there is a red-shift in the CO stretching frequency to 1260-1240cm-1. This might be an indication that the C–O stretching frequency is more sensitive to changes as the donor contributes more is much stronger than the CO stretching frequency. Examining this trend, we further estimate a blue shift in the O-H bending frequency with the formation a hydrogen bond with a negatively charged group to a region beyond 1230cm-1. However it is clear from table 3.3 that the OH bending frequency is coupled to the COO- stretching frequency. 92 3600 1220 O-H Stretching νOH [cm -1 ] C-O-H Bending δCOH [cm -1] 1240 1200 1180 3500 3400 3300 1160 1280 1260 1240 C-O Stretching 1220 νCO [cm 1200 -1 1180 3200 1280 ] 1260 1240 1220 1200 C-O Stretching ν CO [cm -1] Figure 3.8: (A)Two–dimensional correlations of O–H bending and C–O stretching modes (B) O–H stretching and C–O stretching modes for probing the specific type of hydrogen–bonding interactions in proteins: zero hydrogen bond (■), one hydrogen bond OH–X with neutral polar groups (□), one hydrogen bond OH–X+ with positively charged groups (●), one hydrogen bond OH–Y with neutral polar groups (○), one hydrogen bond OH–X with Cys or Met or Tyr (▲), and two hydrogen bonds OH–XY with neutral polar groups (). Experimental evidence for a vibrational structural marker To establish the C−O stretching frequency as a vibrational structural marker for probing the hydrogen-bonding status of tyrosine side–chain groups, experimental evidence will be required. A model compound of tyrosine side–chain group, p-cresol in various solvents including aprotic and apolar solvent, proton acceptor solvents, and proton donor solvents, had been studied using FTIR and Raman spectroscopy (Hienerwadel, Boussac et al. 1997, Takahashi, Okajima et al. 2007, Takeuchi, Watanabe et al. 1989). Takahashi et al, discuss the computational and experimental result for characterizing the tyrosine in TePixD, a BLUF domain protein. Using DFT calculations, they propose that the hydrogen bond between Tyr8-Gln50 is stronger due to the fact that the bond is of OH-X, i.e., Tyr8 being the donor. This is consistent with the crystal structure provided by (Kita, Okajima et al. 2005). 93 The results from literatures are summarized in Table 3.5 and 3.6. The vibrational modes of C−O stretching and O−H bending appear at 1255 cm−1 and ~1176 cm−1 for pcresol in CCl4, i.e., zero hydrogen-bonding PS II 1255 C-O-H Bending δCOH [cm-1] interaction. In proton-acceptor solvents, the PS II frequency of C−O stretching mode of p- 1250 cresol is up−shifted from 1255 cm−1 to 1245 TePixD 1263 − 1272 cm−1 and the frequency of 1240 O−H 1235 1275 1270 C-O Stretching νCO [cm-1] mode of p-cresol is up−shifted from 1176 cm−1 to 1199 − 1251 Human MnSOD 1230 1280 bending 1265 cm-1. In proton-donor solvents (strong Figure 3.9: Experimental evidence for two–dimensional correlations of O–H bending and C–O stretching modes for probing the specific type of hydrogen–bonding interactions in proteins: one hydrogen bond OH– His (●), one hydrogen bond OH–Gln(●), and two hydrogen bonds OH–XY with H2O and Gln (▲). acid), the frequency of C−O stretching mode of p-cresol is down−shifted from 1255 cm−1 to 1235 − 1240 cm−1. Furthermore, the Raman spectroscopy of L-tyrosine hydrochloride and glycyl-L-tyrosine hydrochloride showed that the C−O stretching frequency was 1232 – 1236 cm−1 and crystal structure analysis of the two chemicals identified the hydrogen bonding interactions to be OH–X (Takeuchi, Watanabe et al. 1989). In the solvents that serve as both proton donor and acceptor, the frequencies of C−O stretching and O−H bending modes of p-cresol are in the range of 1260 – 1240 cm−1. These experimental data agrees well with our computational data (Table 1.1). Experimental evidence of was also found in proteins. We have carefully searched for proteins for which their structures have been studied using both high-resolution X-ray crystallography and infrared/resonance Raman (RR) spectroscopy. The results are 94 summarized in Table 3.5. All tyrosine groups in Table 3.5 form one or two hydrogen bonds. There is more OH–X kind of hydrogen-bonding interactions than OH–Y. This is due to the fact that OH–X is stronger than OH–X by 18 kJ/mol (the difference of averaged hydrogen-bond dissociation energy. The corresponding C–O stretching frequency is in the range of 1265 –1279 cm−1 for OH–X and 1240 – 1249 cm−1 for OH– X. This frequency distribution qualitatively agrees well with our computational results. The case of tyrosine with zero hydrogen bond was not found. This is probably because buried phenol group forms hydrogen bond(s) easily. 95 Table 3.5 Vibrational modes of C−O stretching and O−H bending of p-cresol in different solvents Exp. νC−O Exp. νO−H H-Bond (cm−1) (cm−1) 0 1255 No. of Solvent* Solvent structure CCl4 CCl4 Ref.# 1177 1175 O Acetamide 1274 1240 3 1272 N/A 1 1271 1251 2 1269 1251 2 1268 1246 2 1267 N/A 1 1267 N/A 1 1267 1230 2 1265 1224 2 1263 1229 2 NH2 Triethylamine N H3C N 4-MeImH N H H3C N 1-MeIm N Pyridine N 1 O Ethyl acetate Diethyl ether H3C O CH3 H3C O CH3 O N,N’-DMF CH3 H N CH3 O 1,4-Dioxane O H3C + 1-MeImH + HN N 96 O Trifluoroacetic acid HO CF3 1240 N/A 1 1 TCL-acetic N/A 1235 N/A 2 Water H2O 1260 1240 1−2 1261 1244 2 1241* 1241* 3 2 OH 2-propanol H3C CH3 O Acetamide NH2 Abbreviations: 4-MeImH is 4-methylimidazole; 1-MeIm is 4-methylimidazole; N,N’DMF is N,N’-dimethylformamide. * CO stretch and OH bending are coupled. (1)(Takeuchi, Watanabe et al. 1989) (2)(Hienerwadel, Boussac et al. 1997) (3) (Takahashi, Okajima et al. 2007) 97 Table 3.6 Correlations between the C−O stretching frequency and the hydrogen-bonding status of tyrosine side−chain groups in proteins IR/Raman IR/Raman PDB No. of Freq. Freq. Phenol H-bond Role of H-bond Length Group of Partner Tyrosine (Å) His190 Acceptor His189(0) Acceptor Protein* H-bonds C−O (cm-1) O−H (cm-1) 1279 1255 PSII TyrZ 1275 1250 PSII TyrD 1274 N/A code Resolution 2.78 2AXT 3.00 Å 2.59 2AXT 3.00 Å CcO (P.D., 98 Tyr35 SER134 Acceptor 3.09 1QLE 3.00 Å oxidized) 1 1265 N/A SRII-HtrII Tyr199 Asn74 Acceptor 2.74 1H2S 1.90 Å 1265 1242 TePixD Tyr8(0) Gln50 Donor 3.2 1X0P 2.0 Å 1249 N/A AppA Tyr21 Gln63 Donor 2.52/2.55/2.59# 1YRX 2.30 Å 1240 N/A bR (K) Tyr185 Asp212 Donor 2.75 H2O Donor 2.8 Gln143 Acceptor 3.0 Human 2 1265 1232 Tyr34 MnSOD 1M0K 1.43 Å 1LUV 1.85 Å 3.7 CONCLUSION Our density function theory based ab initio computational studies on the hydrogenbonding properties of a tyrosine side–chain group show that the hydrogen-bonding interactions of OH−Y of a phenol group as the hydrogen-bond donor is stronger than that of OH−X. We identified that there is a strong correlation between the frequency of C−O stretching frequency and the hydrogen-bonding status of tyrosine side−chain that is hydrogen bonded to polar or positively/negatively charged amino acid side chains (Table 3.2 and 3.3). This correlation is further supported by available experimental evidence (Table 3.5 and 3.6). We provided both computational and experimental evidence that support the establishment of the C−O stretching frequency of tyrosine side chain as a vibrational structural marker for probing the hydrogen-bonding status: 1255 cm−1 for zero, 1230 to 1243 cm−1 for phenolic oxygen OH−X and 1265 to 1277 cm−1 for hydroxyl hydrogen OH−Y forming a strong hydrogen bond with neutral polar amino acid side−chain groups, and 1194 to 1204 cm−1 for hydroxyl hydrogen OH−X forming a strong hydrogen bond with positively charged amino acid side chain groups. A twodimensional infrared spectroscopy, C−O stretching vs. O−H bending (Figure 3.6), may be helpful to identify two hydrogen bonding interactions or weak OH−Y interaction from zero hydrogen bond when the marker of C−O stretching frequency is inadequate. 99 Chapter IV Summary and Remarks 4.1 Hofmeister series and Protein Structural Dynamics Protein structural dynamics play a key role in the functioning of the proteins (Creighton 1993). The dynamics of the protein are also dependent on the nature of the water molecules surrounding them. Indeed, they lack activity in the absence of water. In solution the proteins possess a conformational flexibility, depends on the activity of the water within its microenvironment; that is, the freedom that the water has to hydrate the protein (Parsegian 2002). The addition of small molecules like neutral salts to water has profound effects on the water structure. The Hofmeister series is concerned with effects of salts at high concentrations (> 0.1 M) and hence must be highly solvated, i.e., hydrogen bonded to water itself (Cacace, Landau et al. 1997b). These ions in turn compete with the protein for water molecules and form strong hydrogen bonds with several shells of water molecules around them (Broering and Bommarius 2005). The number of water molecules ordered around these ions depends on the size of the ion as well as its charge distribution (Neilson, Broadbent et al. 1993). In the analysis of the FTIR data of PYP, the signals in the Amide I region are the main focus. We observe that the large amide I difference signals for PYP are suppressed by the presence of high concentration (4.0 M) salts, in a pattern that follows the Hofmeister series. The degree to which the Amide I signals are 100 suppressed provide a direct indication of the extent to which these salts suppress the functional structural changes (Frauenfelder, Fenimore et al. 2006). Since the water molecules strongly bind to the ions of the salt, the protein molecules are partially dehydrated (Collins 1995). This affects the functioning of the protein molecule as studied in the case of PYP wherein the conformational change during photoreceptor activity was suppressed. PYP binds a unique pCA chromophore that is stabilized by the side chain groups of Glu46 and Tyr42 and the backbone of Cys69 via hydrogen bonding interactions. With the addition of salt, the protein environment is altered, that leads to suppressed structural dynamics of the photoreceptor and a deviation from the original proton pathway for neutralization of the anionic chromophore. The Hofmeister series have always been studied extensively for its effect on the solubility and stability of proteins. Our work offers new insights on the effect of these salts on the structural dynamics and functionality of proteins. Future outlook Water helps enable life-supporting biological functions such as protein folding or enzyme catalysis. The microscopic realm where water and proteins meet is very exciting and difficult to unravel. From a biologist’s point of view, water and proteins must interact on a nanosecond time scale, because that is how fast proteins move. But according to a physicist, this interaction would happen much faster - on the picosecond time scale because that is how fast water molecules move. However, the unique property of water molecules is that they slow down to a speed midway to connect with proteins. It is an essential biological interaction that has to work right every time. If the water moved too slowly, it could get in the way of proteins trying to meet - it would be a bottleneck in the 101 process. And if it moved too fast, it would not connect with the protein at all (Zhang, Kao et al. 2007). In the presence of salts, the water molecules preferentially bind to the salt ions (Collins 1995). This changes in speed of water molecules in the presence of salts and can be an important factor in the altering the protein dynamics. In case of PYP, the alternate proton donor for stabilizing the chromophore leading to the accelerated completion of the photocycle is yet to be unraveled and requires deeper investigation into the salt-water-protein interactions. Time resolved step scan FTIR spectroscopy with a higher time resolution can be used to probe the transient structures of the photocycle in order to understand the altered proton pathway accompanying the accelerated kinetics. 4.2 Vibrational Structural Marker for Hydrogen bonding of Tyrosine We have used time-resolved infrared difference spectroscopy to probe structural changes in functional processes of proteins and interactions of individual groups of the proteins with local environment. Some of the vibrational frequencies of the specific groups often shift upon change of local environment. The changes of local environment include water penetration, changes of hydrogen bonding interactions including formation or breaking of hydrogen bonds and changes of hydrogen-bond strength, and changes of hydrophobic interactions. At the active sites of some proteins where proton transfer occurs, the hydrogen bonding interactions are predominant (Ayala, Perry et al. 2005, Berthomieu, Hienerwadel et al. 1998, Brudler, Meyer et al. 2000b, Takahashi, Okajima et al. 2007). It is therefore necessary to establish vibrational structural markers to probing hydrogen-bonding status of key residues at the active site of proteins. 102 Tyrosine is an important amino acid and is found in the active sites of many proteins involved in signal transduction (Green Fluorescent Protein, Photoactive Yellow Protein), energy transduction (Bacteriorhodopsin, Photosstem II) as well as enzymatic functions (enoyl-ACP reductase). It can form a total of two hydrogen bonds, one as a donor and the other as an acceptor. Hydrogen bonds are a key element in proton transfer pathways linking protein functional dynamics. Like every unique amino acid, tyrosine has its own signature band in the infrared spectrum. We observe from experimental results that the C-O stretching and O-H bending combined with O-H stretching frequency is sensitive to the hydrogen bond formation. We have performed extensive computational studies based on density functional theory on the vibrational frequencies of the neutral Tyr side chain groups interacting with polar neutral side chain groups (Ser, Thr, Cys, Met, Asn/Gln, Tyr, Asp/Gln, His, Lys, and Arg), charged side chain groups (Arg+, Lys+, His+, COO- of Asp/Glu) water molecules, and backbone. Supported by experimental evidence, we have successfully established vibrational structural markers for detecting hydrogen-bonding status of Tyr (Chapter III). The C−O stretching frequency of Tyr side−chain (phenol) group is an excellent vibrational structural marker for probing hydrogen-bonding status: 1255 cm−1 for zero, 1230−1243 cm−1 for hydroxyl oxygen of the phenol group and 1265−1277 cm−1 for hydroxyl hydrogen of the phenol group forming one hydrogen bond with polar neutral groups, 1194−1204 cm−1 for hydroxyl hydrogen forming one strong hydrogen bond with positively charged side−chain groups, and to about 1303 cm-1 for hydroxyl hydrogen forming one strong hydrogen bond with negatively charged side−chain group. 103 The O-H stretching frequency of Tyr side-chain shifts from 3606 cm−1 for zero to 3607-3614 cm−1 for hydroxyl oxygen of the phenol group and 3236-3452 cm−1 for hydroxyl hydrogen of the phenol group forming one hydrogen bond with polar neutral groups, 3585-3595 cm−1 for hydroxyl hydrogen forming one strong hydrogen bond with positively charged side−chain groups and to about 2282.5 cm−1 for hydroxyl hydrogen forming one strong hydrogen bond with negatively charged side−chain group. The O-H bending frequency of Tyr side-chain shifts from 1162.9 cm−1 for zero to 1164-1229 cm−1 for hydroxyl oxygen of the phenol group and 1197-1265 cm−1 for hydroxyl hydrogen of the phenol group forming one hydrogen bond with polar neutral groups, 1159-1177cm−1 for h hydrogen forming one strong hydrogen bond with positively charged side−chain groups and to about 1489 cm−1 for hydroxyl hydrogen forming one strong hydrogen bond with negatively charged side−chain group. A two-dimensional infrared spectroscopy, C−O stretching vs. O−H stretching or C−O stretching vs. O−H bending, is helpful to identify the type of hydrogen−bonding interaction when the marker of C−O stretching frequency is inadequate. From this we can summarize the following: If Tyr (0) acts as a proton acceptor, the C-O stretching is shifted to a lower frequency. The stronger the hydrogen bond, the larger the shift. If the Tyr (0) acts as a proton donor, the most sensitive VSM is the O-H stretch or OH bending. Under extreme conditions, the O-H stretching can be shifted from 3600 cm-1 to 1300 cm-1, while the OH bending from 1180 cm-1 to 2500 cm-1. While interacting with negatively charged group, the C-O stretching shifts to a higher frequency, but again O-H bending and O-H stretching frequencies are more sensitive. 104 Future outlook Each IR spectrum has a wealth of information with each band telling a story of its own. Many interesting bands remain for further investigation. Our lab has been involved in understanding each fingerprint in order to be able to structure a protein by looking at the IR bands. This project was primarily started by Dr.Beining Nie while doing her doctoral research at the Oklahoma State University. We have successfully established vibrational structural markers for detecting hydrogen-bonding status of protonated Asp/Glu (Nie, Stutzman et al. 2005) and now for Tyrosine (Chapter III). 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Are ionic liquids kosmotropic or chaotropic? An evaluation of available thermodynamic parameters for quantifying the ion kosmotropicity of ionic liquids. J. Chem. Tech. Biotech. 81:877-891. 118 Appendix PART A: Operation protocol of Brilliant laser Daily startup procedure • Turn the control key on the front panel of the laser box to the “I” position if it is in “O”. (When key is turned to “I” position, both Power and Interlock LED should Lit on RB panel.) • (If it is in “I” already, skip this step) Wait about 15 minutes until the water temperature equilibrates for flashlamp operation. • Turn on the computer. Wait until Windows has started. • Turn on the motor driver. • Start the OPO control software “OPOTEK”. • Select file F:\opotek\opotek6x.ini • Select “Yes” to home the stepping motor. • Press “Start Flashlamp” button to start flashlamp. Wait for 30 minutes to warm up the laser. • Tune the OPO to a desired wavelength by selecting the target wavelength (475nm for wt PYP). • Select the desired percent of energy. Check the repetition rate. • Open the beam shutter of the laser. Wait 8 seconds before Q-switch control electronics enables the laser operation. 119 • Press the “Fire laser” button to start laser emission. • Wait for 20 minutes to warm up the harmonic generators before measuring the energy and phase matching adjustment. Shutdown procedure: • Stop the laser emission by pressing the “stop” buttons of the Q-Switch and flashlamp on the remote control box or by pressing the “Stop laser” button on the OPO control panel on the computer and then press “Return” and “Quit” to exit the software. • Turn off the motor drive. • Shut down the computer. • Close the beam shutter of the laser • Turn the control key on the front panel of the laser box to the “O” position for the weekend or long holidays. 120 PART B: Experimental Protocol for rapid-scan FTIR of wt-PYP in salt Checklist: • Warm up the laser according to laser startup procedure • Internal MCT Detector is cooled down • Vacuum on (EVAC. IF/ VENT SM on the FTIR) • Temperature controller is on • IR beam is not blocked (Check signal in OPUS-NT) • IR signal (no sample) at the centerburst should be in the range of ±0.75 and ADC count is around -12000 with Sample signal gain of 1 • Laser energy before sample (0.26 mJ/mm2, 7.45 mJ on 6 mm area) • Purge the N2 in the sample chamber (with higher flow rate 140 (3.62 L/min, glass ball) for ~5 minute and then reduce it to ~80 (2.21 L/min, glass ball). • Measurement parameters in OPUS are OK Checklist at the end of day: • Vent optics and turn off the vacuum pump • Turn off purging nitrogen gas. • Slow down the scanner velocity to 10 kHz step by step • Shut down the laser according to laser shutdown procedure • Turn off the temperature controller • Turn off the delay generators and power meter • Remove the samples from sample chamber and clean the windows (for BaF2 windows, need to clean the windows immediately after each day’s measurement) • Back up the experimental data on CD (two copies) at the end of experiments 121 Measurements: Experimental Parameters for Cary UV/Vis spectrum Range: 550-250 nm Double Beam Mode SBW: 2.0 nm Baseline Correction: ON Scan Rate: 600 nm/min Data Interval: 1.0 nm Mode: Absorption Experimental Parameters for absorption measurement D:\OPUS\MEAS\0_Experimental files\Rapidscan_3sample_exchanger). Xpm File:- abs_2cm_3sampleExchanger.XPM TRS File:- RPD_3sampleexchanger_2cm_1scan_40kHz.TRS Optic settings Detector: MCT internal Aperture: 6 mm Scanner Velocity: 40.0 kHz Sample Signal Gain: 4 (with no sample ADC count is 20200) Advanced settings Spectral resolution: 2 cm-1 Save data from 4000 cm-1 to 850 cm-1 122 (saved at Acquisition settings High Folding Limit: 7899.94 cm-1 Low Folding Limit: 0.00 cm-1 Wanted High Frequency: 6000 cm-1 Wanted Low Frequency: 800 cm-1 Acquisition mode: Double-sided forward backward FT settings Phase resolution: 16 cm-1 Phase Correction Mode: Mertz Apodization Function: Blackman-Harris 3-Term Zero filling factor: 4 Experimental Parameters for rapid-scan FTIR (saved at D:\OPUS\MEAS\0_Experimental files\RapidScan). XPM:- wtPYP_rpsc_norm_4.5m_200kHz.XPM TRS:- RPD_4S_norm_4.5cm_200kHz.TRS Do not use files from this directory. Make a Copy of these files to desired folder and then use them. Split Forward/Backward Inteferogram (for Rapidscan FB measurements) Split Doublesided Inteferogram (for quadruple splitting rapid-scan measurements) 123 Optic settings Detector: MCT internal Aperture: 6 mm Scanner Velocity: 200.0 kHz Sample Signal Gain: 4 Advanced settings Spectral resolution: 4.5 cm-1 Save data from 4000 cm-1 to 850 cm-1 Acquisition settings High Folding Limit: 5266.62 cm-1 Low Folding Limit: 0 cm-1 Wanted High Frequency: 5250 cm-1 Wanted Low Frequency: 800 cm-1 Acquisition mode: Double sided forward backward FT settings Phase resolution: 16 cm-1 Phase Correction Mode: Mertz Apodization Function: Blackman-Harris 3-Term Zero filling factor: 4 124 Delay Generators All the output signals are TTL (inverted normal) signals. FTIR FWD signal triggers the delay generator slave 1. Slave 1 A = T + 10ms B = A + 30μs D=T + 95ms Slave 2 A = T+10ms+272 μs B = A + 30μs C = T+ 10ms D = T+4.022s For quadruple splitting measurement, Change the delay time for the 1st, 2nd, 3rd, 4th measurements of 10 ms, 23 ms, 59 ms, and 72 ms to A on both delay generators. FWD cable of FTIR (trigger out) is connected to DG Slave 1’s “Ext Trig” (trigger in) Slave 1’s “To’ is connected to Slave 2’s “Ext Trig” Flash trigger cable is connected to DG Slave 1’s normal AB output Q-switch trigger cable is connected to DG Slave 2’s normal AB output Rapid-scan’s Trigger In cable is connected to DG Slave 2’s invert CD output because TKDA that triggers starting of the measurement responds to “high”, not rising edge. Here 272 μs is the Q-Switch delay, corresponding to 40% laser energy. The Q-Switch delay is in the range of 192 μs (100%) – 305 μs (5%). The 272μs delay time is to give an example of how delay generators are setup. 125 In order to test laser is working properly (i.e. while not doing any measurement), Slave 1 should be triggered internally and Slave 2 should be Externally Triggered. Experimental procedure: Daily procedure 1. Warm up the laser using laser protocol and change trigger mode in Opotek control software from internal to external. 2. Turn on Flashlamp and Q-switch with internal triggering using the remote control box and block the laser beam before it enters to FTIR chamber. 3. Cool down the MCT detector. 4. Evacuate FTIR optics compartment. 5. Turn on the delay generators and check the settings. 6. Load the protein sample in a small sample holder. 7. Measure the VIS absorption spectrum of the sample using Cary 300 UV/Vis spectrometer. Insert samples into FTIR sample chamber (Bruker IFS66v). 8. Start purging with higher flow rate 140 (3.62 L/min, glass ball) for ~5 minute and then reduce it to ~80 (2.21 L/min, glass ball). 9. Set the desired temperature. Wait an extra 10 min for equilibration after the sample reached the desired temperature. 10. Measure IR absorption spectrum. Check IR absorption, Amide I OD. Take special care for the water vapor bands. 11. Bloch the laser beam path by using a thick magazine. 12. Start Rapid-scan measurement. 126 13. Stop the flashlamp and Q-switch on the Remote control Box (RB). 14. Switch triggering mode of flashlamp from internal to external. 15. Switch triggering mode of Q-switch from internal to external. 16. Turn on the flashlamp and wait for 80 fires of flashlamp. And then turn on the Q-switch. 17. When the laser beam is visible, remove the block from the laser path letting the laser beam reach sample chamber. 18. Measure one rapid scan for testing; 10 loops, test band positions, decay time. 19. Check if the photocycle is complete (by checking if there is a spectra with 0 amplitude). 20. Load the experimental file for measuring rapid scan with increased number of loops. 21. After rapid-scan is complete each time (when Fourier transformation starts, the frequency of FWD has changed), turn off the Q-switch and the flashlamp immediately) 22. Repeat collecting several sets of measurements. 23. Measure IR absorption spectrum again. The absorption of 5% protein bleaching is acceptable. 24. Measure VIS absorption spectrum of the protein sample. Data analysis using macro programs: 1. Load all the rapid-scan measurement files in OPUS window. 2. Delete all the data files in C:\Program Files\OPUS\WORK\ directory. 127 3. Select “Run Macro” in the OPUS-NT “Macro” pull down menu. 4. Open the main macro program for rapid-scan data analysis “E:\OPUS_macro\macro_NT\RPDSCAN\RPD_NORM\updates\RPD_NORM.MTX” . 5. Select all the single beam spectra and drag them to the blank space labeled “Rapidscan(s)” in the popup window. 6. Specify the “First block to extract” to be 1 and “Last block to extract” (Find the last block number by showing parameters in “Trace/Multiple” of the rapid-scan OPUS files. 7. Specify the “No of data sets”. 8. Select “Calc mOD” and “Average”. 9. Click “OK”. 10. The difference absorption spectra will be loaded to the screen. They are saved in C:\Program Files\OPUS\WORK\ folder. 11. Copy and Paste them to your local directory and delete them from the WORK folder to prevent loading wrong set of data for next data analysis. 128 PART C: Selected Salts from Hofmeister Series Salt Chemical formula Molecular weight Sodium fluoride Sodium chloride Sodium bromide Sodium iodide NaF NaCl NaBr NaI 41.99 58.44 102.9 149.89 Solubility (% by weight) 3.97% 26.45% 48.6% 64.8% Lithium chloride LiCl 42.33 45.81% 13.26M Sodium chloride NaCl 58.44 26.45% 5.13M Potassium chloride KCl 74.55 26.22% 4.73M Cesium chloride CsCl 168.5 65.64% 9.1M Calcium chloride CaCl2 111.02 44.83% 3.19M Magnesium chloride MgCl2 95.31 35.9% NH4Cl 53.49 28.34% 5.5M (NH4)2SO4 132.14 43.3% 5M MgSO4 120.5 26.3% 2.56M NaH2PO4 119.99 48.68% Na2HPO4 141.96 10.55% Ammonium chloride Ammonium sulphate Magnesium sulphate Sodium phosphate monobasic Sodium phosphate dibasic Solubility (by volume) 0.95M 5.13M 9.2M 11.3M Protocol for protein sample preparation Prepare Neutral Buffer (as Stock): To prepare 500 μL, 1M phosphate buffer in D2O at pH*=6.6 (pH=7.0): 1M, 200 μL sodium phosphate dibasic (Na2HPO4) and 1M, 400 μL sodium phosphate monobasic (NaH2PO4·H2O) will be prepared. 132.5μL of 1M sodium phosphate dibasic 129 with 367.5 μL of 1M sodium phosphate monobasic will be mixed to obtain 1M phosphate buffer at pH 7.0. Since the two solutions are prepared at the same concentration, the final buffer concentration will also be 1M independent of the mixing ratio. Step 1: Prepare 200 μL, 1M of sodium phosphate dibasic (MW=141.96 g/mol). Mix and dissolve 28.3 mg of Na2HPO4 in 195 μL of D2O. (The estimated volume from previous experiments of sodium phosphate dibasic is 5 μL). Step 2: Prepare 400 μL, 1M of sodium phosphate monobasic (MW=137.99 g/mol). Mix and dissolve 55 mg of NaH2PO4·H2Oin 390 μL of D2O. (The estimated volume from previous experiments of sodium phosphate monobasic is 10 μL). Step 3: Prepare ~500 μL, 1M pH* 6.6 (pH=7.0) NaxD3-xPO4 buffer (as stock) Mix 132.5μL of 1M sodium phosphate dibasic with 367.5 μL of 1M sodium phosphate monobasic Na2HPO4 (26.5% by volume) + NaH2PO4⋅H2O (73.5% by volume) Step 4: Measure the pH value of the mixed buffer. If the pH* measured is lower than pH*6.6, add more sodium phosphate dibasic. If the pH* measured is higher than pH*6.6, add more sodium phosphate monobasic. 130 Step 5: Freeze the buffer mixture using liquid N2 and vacuum dry overnight at the core facility at NRC. - Turn motor on. - Place the frozen sample and balance with another tube of equivalent weight. - Start rotor - Turn the valve with arrow pointing up to evacuate the sample chamber. This procedure ensures that the H2O molecules are eliminated from the sample and the hydrogen atoms in the chemical are replaced by deuterium. Step 6: Add 500μL pure D2O when buffer is required for sample preparation. Step 7: Store this buffer in an O-ring sealed container and place the container in desiccators. Prepare Salt Solution(4M NaCl in D2O at pH 7) Step 1: Prepare 1 mL, 4M NaCl (MW=58.44 g/mol) in pure D2O. Mix and dissolve 233 mg of NaCl in ~950 μL of D2O. ** We need to manually compare the amount of D2O required to make 1mL of sample with 1mL plain water in a similar test tube. Step 2: Mix 1 mL, 4M NaCl with 50 μL, 1M neutral buffer so that the final buffer concentration in the sample is 50mM. Step 3: Check pH and if required add 2 μ drops of 1M NaOD to the salt solution to adjust the pH*=6.6 (pH=7.0). Prepare Salt Solution(4M KCl in D2O at pH 7) Step 1: Prepare 1 mL, 4M KCl (MW=74.551 g/mol) in pure D2O. Mix and dissolve 298 mg of KCl in ~950 μL of D2O. 131 ** We need to manually compare the amount of D2O required to make 1mL of sample with 1mL plain water in a similar test tube. Step 2: Mix 1 mL, 4M KCl with 50 μL, 1M neutral buffer so that the final buffer concentration in the sample is 50mM. Step 3: Check pH and if required add 2 μ drops of 1M NaOD to the salt solution to adjust the pH*=6.6 (pH=7.0). Prepare Salt Solution(4M LiCl in D2O at pH 7) Step 1: Prepare 1 mL, 4M LiCl (MW=42.329 g/mol) in pure D2O. Mix and dissolve 169 mg of LiCl in ~950 μL of D2O. ** We need to manually compare the amount of D2O required to make 1mL of sample with 1mL plain water in a similar test tube. Step 2: Mix 1 mL, 4M LiCl with 50 μL, 1M neutral buffer so that the final buffer concentration in the sample is 50mM. Step 3: Check pH and if required add 2 μ drops of 1M NaOD to the salt solution to adjust the pH*=6.6 (pH=7.0). Prepare Salt Solution(4M CsCl in D2O at pH 7) Step 1: Prepare 1 mL, 4M CsCl (MW=168.5 g/mol) in pure D2O. Mix and dissolve 674 mg of CsCl in ~950 μL of D2O. ** We need to manually compare the amount of D2O required to make 1mL of sample with 1mL plain water in a similar test tube. Step 2: Mix 1 mL, 4M CsCl with 50 μL, 1M neutral buffer so that the final buffer concentration in the sample is 50mM. 132 Step 3: Check pH and if required add 2 μ drops of 1M NaOD to the salt solution to adjust the pH*=6.6 (pH=7.0). Prepare Salt Solution(4M NaBr in D2O at pH 7) Step 1: Prepare 1 mL, 4M NaBr (MW=102.9 g/mol) in pure D2O. Mix and dissolve 411 mg of NaBr in ~900 μL of D2O. ** We need to manually compare the amount of D2O required to make 1mL of sample with 1mL plain water in a similar test tube. Step 2: Mix 1 mL, 4M NaBr with 50 μL, 1M neutral buffer so that the final buffer concentration in the sample is 50mM. Step 3: Check pH and if required add 2 μ drops of 1M NaOD to the salt solution to adjust the pH*=6.6 (pH=7.0). Prepare PYP to desired pH and deuteration This is achieved by repeated washes of PYP using proper buffer. The main goals for washing PYP are (1) To reduce H2O contamination (from liquid sample as well as exchangeable hydrogen atoms in PYP); (2) To adjust the sample pH to desired pH. Two washes are sufficient for Rapid scan FTIR. For time-resolved rapid-scan FTIR experiments: The first wash: Step 1: place ~390 μL of 4M salt solution in neutral buffer (pH=7.0) and 10 μL PYP sample in H2O (8 mM) in Y10 (Microcon) filter. • Set centrifugation speed to: 9000 rpm (RCF=6,610) 133 • Centrifugation time: 20 minutes Make sure that >50% of buffer solution is collected in the tube after passing the filter. Step 2: Set Increase the centrifugation speed: 12,000rpm (RCF=11750) • Centrifugation time: *20 minutes * Time varies with different salts. Check table below. Make sure > 390 μL of buffer added is collected in the tube. Comments: (1) After adding 390 μL of buffer to 10 μL of 8 mM PYP, the resulting protein concentration is 0.2 mM. Therefore, the buffer to protein ratio is 500:2 or 250:1. So one wash is sufficient for pH adjustment. (2) The H2O contamination is 10 μL /400 μL = 2.5%. A second wash is needed to further reduce the water contamination. The second wash: Step 1: place ~390 μL of 4M salt solution in neutral buffer (pH=7.0) and 10 μL PYP sample in H2O (8 mM) in Y10 (Microcon) filter. • Set centrifugation speed to: 9000 rpm (RCF=6,610) • Centrifugation time: 20 minutes Make sure that >50% of buffer solution is collected in the tube after passing the filter. Step 2: Set Increase the centrifugation speed: 12,000rpm (RCF=11750) • Centrifugation time: *20 minutes * Time varies with different salts. Check table below. 134 Make sure > 390 μL of buffer added is collected in the tube. Step 3: When ~390 μL of the buffer solution is collected in the test tube, dismantle the filter and connect it to the adapter and fix this adapter to a small 500 μL tube. Balance with an identical setup in the opposite end. Run centrifuge for 2 minutes at 5000 rpm. Step 4: Seal the test tube with the o-ring provided cap and freeze sample until ready to use for FTIR measurements. Salt solution (4.0 M) NaCl CsCl LiCl KCl NaBr Time to centrifuge (min) @ 9000 rpm @ 12000rpm (RCF=6,610) (RCF=11,750) 20 10 20 5 20 20 20 10 20 20 135 PART D The most notable characteristic of the PYP is the absorption maximum at 446nm and the purity index which is the ratio between the peak height at 278 nm and 446 nm. The purity index is a measure for the purity of the protein, where a value lower than 0.50 is considered pure. In each of the following cases, we found the purity index to range from 0.42-0.48. The figure below shows the UV Vis data of wild type PYP in varying concentrations (4.0 M, 2.0 M, 1.0 M and 0.5 M) of NaCl solution. To make the method more consistent, a stock solution of 4.2 M NaCl solution was prepared in D2O and the pH adjusted to 7.0 by using a small quantity (a few microdrops) of NaOD. This stock solution was then diluted by adding a calculated amount of D2O to obtain 2.0 M, 1.0 M and 0.5 M NaCl solution. The protein sample was washed using Eppendorf Microcentrifuge with the respective solution to obtain wt PYP sample in concentrated salt solution. wtPYP in 4.0 M NaCl 2.0 M NaCl 1.0 M NaCl 0.6 Absorption [OD] 0.5 M NaCl 0.5 0.4 0.3 0.2 0.1 0.0 250 300 350 400 Wavelength [nm] 136 450 500 The figure below shows UV-Vis data of wt PYP in different salts taken from the Hofmeister series. The protein sample was washed with the respective salt solution prepared in D2O and pH adjusted to 7.0 0.8 0.7 4.0 M NaBr 4.0 M CsCl 4.0 M KCl 0.6 4.0 M NaCl Absorption [OD] 4.0 M LiCl 0.5 0.4 0.3 0.2 0.1 0.0 250 300 350 400 450 500 -1 Wavenumber [cm ] The cation series from the Hofmeister salts Apart from doing the anion series (different chloride salts), we have performed FTIR measurements of wt PYP using the cation series (different sodium salts) from the Hofmeister salts. The figure below shows the effect of Cl- ion and Br- ion on the structural dynamics of the protein. The reduction in the amplitude of the Amide I band clearly indicates the suppression of the protein structural dynamics. However, the data obtained have to be further investigated in order to draw any further conclusions. We tried to do the FTIR measurements of wt PYP in 4.0 M NaI and 4.0 M NaF. Sodium fluoride (NaF) has very low solubility (3.9% by weight, CRC Handbook of Chemistry and Physics) which hindered us from preparing the sample at high concentration. Sodium 137 8.0 0.0 M NaCl 4.0 M NaCl 1624 1740 1515 1571 1669 4.0 1253 1641 -4.0 1162 1497 1302 1609 1689 0.0 1726 Diff Absorption [milli OD] 4.0 M NaBr 1800 1700 1600 1500 1400 -1 1300 1200 Wavenumber [cm ] iodide (NaI) has photoactive properties, due to which it bleaches the PYP when washed together. Due to these technical difficulties, the work with respect to the cation series is not complete but will be continued by our lab in the near future. The Effect of MgSO4 on wt PYP The effect of 2.0 M Magnesium sulphate on PYP was observed to be different from that of the other salts used. The pB-pG spectra of wt PYP in 2.0 M MgSO4 and 0.67 M MgSO4 are shown below. It is interesting to note that the peak at 1740 cm-1, which signifies the protonation of Glu46 is absent in the presence of MgSO4, while it was clearly observed in the presence of 4.0 M NaCl, 4.0 M KCl, 4.0 M LiCl and 4.0 M CsCl. This suggests that the Glu46 is the proton donor to stabilize the chromophore as the band at 1740 cm-1 disappears with the ionization of Glu46. Another important observation is the acceleration of the photocycle. From the 2-D spectra it is clear that the suppression of protein structural dynamics is a reduced effect in the presence of 2.0 M MgSO4 than in 138 the presence of the other chloride salts tested. The magnitudes of the Amide I band in the salt-free environment and in the presence of salt are very similar. However, it is interesting to note the acceleration of the photocycle in the presence of salt. The magnitude of the Amide I band is about 4.5 OD to begin with but quickly decays to zero, much faster than the salt free sample. We can do further step scan measurements understand the time and the dynamics of the photocycle better. 0.67 M MgSO 4 1515 1571 1669 4.0 1162 1497 1253 1689 0.0 1726 -4.0 1641 Diff Absorption [milli OD] 1624 Salt-free wt PYP 2.0 M MgSO 4 1800 1700 1600 1500 1400 -1 1300 1200 Wavenumber [cm ] Infrared difference absorption spectra of wt PYP in varying salt concentrations indicating the absence of ionization band of Glu46 (1740 cm-1) which is ionized even the presence of 2.0 M MgSO4.This also shows the suppression of conformational changes in the presence of high salt concentration. 139 Time [s ] Diff. Absorbance [milli OD] Diff. Absorbance [milli OD] WT PYP in 0.0 M salt nu Wave m ber WT PYP in 0.67 M MgSO4 Time -1 [cm ] [s ] nu Wave mber -1 [cm ] Diff. Absorbance [milli OD] 140 WT PYP in 2.0 M MgSO4 3-dimentional plots show the effect of varying salt concentrations on the PYP photocycle. Starting with 0 M salt, the effect is more pronounced in the form of suppressed conformational change and accelerated photocycle as the salt concentration is changed to 0.67 M MgSO4 and 2.0 M MgSO4. W ave numb -1 ] er [cm VITA Anupama Jagadeesh Thubagere Candidate for the Degree of Master of Science Thesis: ADVANCED FTIR SPECTROSCOPY FOR STRUCTURAL DYNAMICS OF PROTEINS Major Field: Photonics Biographical: Personal Data: Born in Bangalore, India, on September 30, 1983. Education: Graduated from Sri Aurobindo Memorial School, Bangalore in May 1999, received Bachelor of Engineering in Electrical and Electronics from Visveswaraiah Technological University, Bangalore in May 2005. Completed the requirements for the Master of Science degree with a major in Photonics at Oklahoma State University in December 2007. Professional Membership: Member of American Physical Society since 2007. Name: Anupama Thubagere Date of Degree: December, 2007 Institution: Oklahoma State University Location: Stillwater, Oklahoma Title of Study: ADVANCED FTIR SPECTROSCOPY OF PROTEIN STRUCTURAL DYNAMICS Pages in Study:140 Candidate for the Degree of Master of Science Major Field: Photonics Scope and Method of Study: The first role of water is to aid the functioning of proteins. Presence of a high concentration of salt reduces the solubility of proteins, so that it induces protein precipitation. Many times, protein precipitation does not lead to protein crystallization. Here we report our studies that salt solutions suppress or inhibit protein structural dynamics. Photoactive yellow protein (PYP), a bacterial blue light photoreceptor protein, is employed as a model system in our study. We have examined and discussed the mechanism in which different salt solutions from the classic Hofmeister series inhibit protein structural dynamics. We use time-resolved FTIR spectroscopic technique to probe the structural dynamics of proteins, including the proton transfer process and global conformational motions. Infrared absorption spectroscopy is an important technique for structural and kinetic studies of protein functional mechanism and vibrational band assignment is a crucial step in order to obtain structural information from infrared absorption spectra. We report computational methods for developing vibrational structural markers of tyrosine. The time-resolved infrared spectroscopy combined with vibrational structural markers allows time-resolved structural characterization of proteins, making it possible to “watch” proteins in action over a broad time scale from picoseconds to seconds. Findings and Conclusions: We found that high concentration of salt (4.0 M NaCl, KCl, CsCl and LiCl) solution strongly inhibits the structural dynamics of PYP upon blue light excitation. The results provide insights to fundamental understanding of protein crystallization of water-soluble proteins. We observe that the large Amide I difference signals for PYP are suppressed by the salts, in a pattern that follows the Hofmeister series. The degree to which the amide I signals are suppressed provide a direct indication of the extent to which these salts suppress the functional structural changes. Time-resolved infrared data can be used to probe the dynamic nature of proteins. Using density function theory (DFT) based first principle computational studies combined with experimental data; we found that it is possible to unambiguously determine the participation of the functional hydroxyl group in Tyrosine in hydrogen bonding. In addition, we show that it possible to determine the number and nature of hydrogen bonding interactions of a phenolic group in proteins using a combination of C-O stretching and O-H stretching frequencies (2D vibrational spectroscopy). ADVISER'S APPROVAL: Dr. Aihua Xie