Embed Size (px)
Citation preview
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
ii
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
iii
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.
iv
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
v
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
vi
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
vii
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
viii
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
1
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).
2
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
3
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
4
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.
The bioluminescent Green Fluorescent Protein (GFP) is found in the jelly fish
Aequorea victoria. The chromophore is formed by the cyclization of an internal Ser65-
Tyr66-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).
Figure 1.1: The chromophore of GFP consisting of Ser65-Tyr66-Gly67. 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.
5
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.
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
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.
6
Figure 1.3: The hydrogen bond network stabilizing the negative charge on the chromophore of photoactive yellow protein.
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
yellow protein (PYP) from Ectothiorhodopsira
halophila. On blue light absorption, PYP
undergoes a photocycle containing 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).
7
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: X-
ray 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,
8
some of the most important ones being PS-I (Fromme and Mathis 2004)and
bacteriorhodopsin (Grigorieff, Ceska et al. 1996).
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
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).
A
B
9
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).
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,
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)
A B C
10
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. 15N, 17O) 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 nuclei flip,
researchers can determine
molecular structure (McDonald
and Phillips 1967).
The first protein structure solved
using the NMR technique by the
Wuthrich group was of the alpha
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).
11
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
12
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
13
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
14
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
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 structure-
function studies of proteins: high structural sensitivity (proton transfer, hydrogen bonding
A B
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.
15
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.
16
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
Figure 1.8: The layout of time-resolved rapid-scan spectroscopy
17
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
18
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
19
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 time-
resolved infrared difference spectroscopy for structural characterization of functionally
important intermediate states of proteins, and consequently for understanding the
functional mechanism of proteins.
20
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
21
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
22
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°
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.
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
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°.
Figure 2.2: One water molecule can form up to 4 hydrogen bonds with the neighboring waters having an average life time of 10-
12 s, which makes the structure of water so dynamic(Voet and Voet 2004)
23
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.
24
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.
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
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].
25
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).
26
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.
Ion Ionic radius
(Å)
Hydration number
Li+ 0.6 4-6
Na+ 0.95 4-6
K+ 1.33 2-5 Cation
Cs+ 1.69 2-3
F- 1.27 -
Cl- 1.81 6-9
Br- 1.95 6-7 Anion
SO42- - 8
References: (Caminiti,
Licheri et al. 1978, Caminiti,
Paschina et al. 1979)
(Domene and Sansom 2003)
(Maslen 1967, Ramos,
Barnes et al. 2000)
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.
27
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.
28
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
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).
29
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
30
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,
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
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.
(c) (b) (a)
31
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
32
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.
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 hydrogen-
bonding 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).
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.
pCoumaric acid
Wavelength [nm]250 300 350 400 450 500
Abs
orpt
ion
[OD
]
0.0
0.1
0.2
0.3
0.4
0.5
0.6
33
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.
34
The chromophore configuration changes from C7=C8-trans C9-S-cis to C7=C8-cis C9-S-
trans.
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 pR-
pB, 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.
35
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-s-cis 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).
36
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
37
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 custom-
made 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.
38
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.
39
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).
40
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.
41
Wavenumber [cm-1
]
30003200340036003800
Abs
orpt
ion
[OD
]
0.0
0.2
0.4
0.6
0.8
1.0
Pure water
4.0 M NH 4Cl
4.0 M NaCl
Wavenumber [cm-1]
30003200340036003800
Abs
orpt
ion
[OD
]
0.0
0.2
0.4
0.6
0.8
1.0
2.0 M (NH4)2SO4
4.0 M NH 4Cl
Pure water
Wavenumber [cm-1
]30003200340036003800
Abs
orpt
ion
[OD
]
0.0
0.2
0.4
0.6
0.8
1.0
Pure water
2.0 M(NH 4)2SO4
2.0 M MgSO 4
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 SO4
2- 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, SO4
2-.
A B
C
42
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
43
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
44
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
45
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.
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
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.
Wavelength [nm]250 300 350 400 450 500
Abs
orpt
ion
[OD
]
0.0
0.1
0.2
0.3
0.4
0.5
4.0 M NaCl
wtPYP in
46
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).
47
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).
Abs
orpt
ion
[OD
]
0.2
0.4
0.6
0.8
wtPYP with 4 M NaCl
wtPYP in 0.5 M NaCl
wtPYP with 2 M NaCl
wtPYP with 1 M NaCl
wtPYP with 0 M NaCl
1726
1644
1688
1584
1635
1515
1537
1551
1455
1471
1498
1402
1440
Wavenumber [cm-1]
130014001500160017001800
2nd
Der
ivat
ive
of A
bsor
banc
e
-0.006
-0.004
-0.002
0.000
0.002
0.0
48
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
49
~500 water molecules per PYP. This partial dehydration can reduce the extent of
structural changes.
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
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
1726
1740
1689 16
09
1515
150015501600165017001750
Wavenumber [cm-1]
-4.0
0.0
4.0
Diff
Abs
orpt
ion
[mO
D]
1571
1497
wt PYP in solution
wt PYP in 4.0 M NaClwt PYP in P63 crystals
50
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
51
so far unassigned bands change with the salt concentration: the 1610 cm-1, 1550 cm-1 and
1440 cm-1 negative and the 1565 cm-1 and 1660 cm-1 positive bands.
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
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.
Wavenumber [cm-1]
1200130014001500160017001800
Diff
Abs
orpt
ion
[mill
i OD
]
-5
0
517
40
1689
1624
1497
1669
1515
1726
1302
0.0 M NaCl
0.5 M NaCl
1.0 M NaCl
2.0 M NaCl
4.0 M NaCl
1162
1641
1253
1571
1609
52
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).
53
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
WT PYP in 1.0 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
WT PYP in 1.0 M NaCl
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.
Diff
. A
bso
rban
ce [m
illiO
D] WT PYP in 0.0 M salt
Time [s]Wavenumber [cm-1]
Diff
. A
bso
rban
ce [m
illiO
D] WT PYP in 0.0 M salt
Time [s]Wavenumber [cm-1]
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
WT PYP in 0.5 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
WT PYP in 0.5 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
WT PYP in 2.0 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
WT PYP in 2.0 M NaCl
WT PYP in 4.0 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 4.0 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
A
D
B
C
E
54
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
Glu46 group remains protonated but its
frequency shifts in the presence of salt. The
position of these two carboxylic bands is
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.
55
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.
56
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+.
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.
1740
1689
1624
1497
1669
1515
1726
1302
0.0 M salt4.0 M CsCl
4.0 M NaCl
1162
1641
1253
1571
1609
4.0 M LiCl
Wavenumber [cm-1]
1200130014001500160017001800
Diff
Abs
orpt
ion
[mill
i OD
]
-5
0
5 4.0 M KCl
57
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.
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 0.0 M salt
Time [s]Wavenumber [cm-1]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 0.0 M salt
Time [s]Wavenumber [cm-1]
WT PYP in 4.0 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 4.0 M NaCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
]
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 4.0 M KCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 4.0 M KCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 4.0 M CsCl
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 4.0 M CsCl
A
D C
B
58
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.
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
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).
59
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
60
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
61
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.
62
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
63
Figure 3.1: Tyrosine molecule with phenol functional group
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
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
64
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 Tyr-
FADH 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
65
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 Ser65-
Tyr66-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).
66
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.
67
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
+
Y92
E222
H148
Y145S65
G67Y66
+++
Y92
E222
H148
Y145S65
G67Y66
B
RET
D212
K126
Y185
RET
D212
K126
Y185
RET
D212
K126
Y185
C
A
68
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-
69
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
70
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
A B C
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.
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
71
Figure 3.4: Model compound for calculation of vibrational frequencies of tyrosine
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
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
72
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
73
Figure 3.5: (A) Initial structure from ChemDraw; (B) Geometry optimized using empirical method (PM3) from Chem3D (C) Geometry optimized using DFT in Gaussian03
A B C
employed to view the vibrational motions of any chosen mode from the output file of
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:
2
2)(exp(*)()(
d
xxIY ννν −
−= )
where I(ν) is the intensity of the vibrational frequency Xν in Gaussian03 output and
FWHM (full width at half maximum) is d2ln2 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
74
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:
75
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.
76
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 hydrogen-
bonding 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
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) .
A
B
C
1255
1327
1163
1236
1327
1171
1272
1350
1222
1211
125513
53
1234
Wavenumber [cm-1
]
110012001300140015001600
Inte
nsity
A
B
C
D
3612
3606
3361
3408
Wavenumber [cm-1
]
2900310033003500
Inte
nsity
D
77
10001100120013001400150016001700
Sec
ond
Der
ivat
ive
1258
0.0
0.2
0.4
0.6
0.8
Abs
orba
nce
Wavenumber [cm-1]
1615
1589
1599
1517
1454
1429
1264
1178 10
55 1048
1246
A
B
frequencies that are sensitive to the hydrogen- bonding interaction in the fingerprint
region.
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 hydrogen-
bonding 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
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.
78
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
79
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 hydrogen-
bond 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 hydrogen-
bond dissociation energy of these hydrogen-bonding interactions is approximately half of
80
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
81
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).
82
TABLE 3.1 Calculated hydrogen-bonding properties of a phenol group interacting with neutral groups
Structure Amino Acid* No. of
H-bond†
H-bond
Length†
(Å)
H-bond
Angle†
(deg)
H-bond
Energy‡
(ε = 1 /2 /4)
(kJ/mol)
νC-O
(cm-1)
νO-H Bending
(cm-1)
νO-H Stretching
(cm-1)
TYR 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
ASN/GLN −
A 1 3.06 172.1 10.7/12.0/6.6 1236.4 1164.1§ 3609.6
THR − A 1 2.9 159.9 11.9/9.7/3.7 1234.3 1161.1 3614.4
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
ASP/GLU −
A 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
84
ASN/GLN −
B 1 2.77 165.4 26.9/18.2/6.6 1273.1 1227.3 3312.0
H2O – B 1 2.82 179.0 23.0/17.9/12.8 1271.8 1221.5 3408.4
ASP/GLU –
B 1 2.81 158.4 27.7/11.3/-3.8 1268.9§
1221.3
1210.6
3427.8
3383.8
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
ASP/GLU −
B 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
85
CYS − B ~0.5 3.35 149.1 17.1/8.5/1.1 1258.7 1194.6 3452.0
ARG 1.5
2.72
3.03
157.4
144.8 48.4/35.8/25.1 1265.8 1249.6 3028.2§
ASN/GLN 1.5
2.72
2.94
157.5
136.6 33.0/24.0/11.4 1257.5§ 1235.9 3221.9
ASP/GLU 1.5
2.72
2.77
145.0
154.0 37.5/19.4/3.56 1245.9 1221.3
3338.9
3252.9
2 H2O 2 2.77
2.89
173.1
164.7 45.8/35.7/27.5 1254.7 1233.8 3361.0
2 SER 2
2.76
2.87
173.1
166.0 39.3/30.5/22.0 1251.6 1237.1 3332.2
* 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.
85
86
†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
87
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 hydrogen-
bonding 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 COO-
stretching frequency at 1672 cm-1 (Nie, Stutzman et al. 2005).
TABLE 3.2 The hydrogen bonding interactions of neutral Tyrosine with positively charged amino acid side chains
Structure Amino Acid* No. of
H-bond†
H-bond
Length†
(Å)
H-bond
Angle†
(deg)
H-bond
Energy‡
(ε = 1 /2 /4)
(kJ/mol)
νC-O
(cm-1)
νO-H Bending
(cm-1)
νO-H Stretching
(cm-1)
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
ARG+ 2
2.90
2.92
149.3
147.6 58.7/31.9/15.7 1194.1 1177.8 3585.6
88
TABLE 3.3 The hydrogen bonding interactions of neutral Tyrosine with negatively charged amino acid side chains
* 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.
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
90
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 hydrogen-
bonding 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 hydrogen-
bonding interactions (1252–1255 cm−1), it is difficult to use the C–O stretching frequency
91
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 hydrogen-
bonding 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.
92
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 1215-
1230 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.
93
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 (�).
C-O Stretching νCO [cm-1]118012001220124012601280
C-O
-H B
endi
ng δ C
OH [c
m-1
]
1160
1180
1200
1220
1240
C-O Stretching νCO [cm-1]
12001220124012601280
O-H
Str
etch
ing
ν OH
[cm
-1]
3200
3300
3400
3500
3600
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).
94
C-O Stretching νCO [cm-1]1265127012751280
C-O
-H B
endi
ng δ
CO
H [
cm-1
]
1230
1235
1240
1245
1250
1255PS II
PS II
TePixD
Human MnSOD
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 (▲).
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 p-
cresol in CCl4, i.e., zero hydrogen-bonding
interaction. In proton-acceptor solvents, the
frequency of C−O stretching mode of p-
cresol is up−shifted from 1255 cm−1 to
1263 − 1272 cm−1 and the frequency of
O−H bending mode of p-cresol is
up−shifted from 1176 cm−1 to 1199 − 1251
cm-1. In proton-donor solvents (strong
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
95
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.
96
Table 3.5 Vibrational modes of C−O stretching and O−H bending of p-cresol in
different solvents
Solvent* Solvent structure No. of
H-Bond
Exp. νC−O
(cm−1)
Exp. νO−H
(cm−1) Ref.#
CCl4 CCl4 0 1255 1177
1175
Acetamide NH2
O
1274 1240 3
Triethylamine N
1272 N/A 1
4-MeImH NH
NH3C
1271 1251 2
1-MeIm N
N
H3C
1269 1251 2
Pyridine N
1268 1246 2
Ethyl acetate OH3C
O
CH3 1267 N/A 1
Diethyl ether OH3C CH3 1267 N/A 1
N,N’-DMF H
O
N
CH3
CH3
1267 1230 2
1,4-Dioxane O
O
1265 1224 2
1-MeImH+ +HN
N
H3C
1
1263 1229 2
97
Trifluoroacetic acid HO
O
CF3 1240 N/A 1
TCL-acetic N/A
1
1235 N/A 2
Water H2O 1260 1240 1−2
2-propanol H3C
OH
CH3
2 1261 1244 2
Acetamide NH2
O
1241* 1241* 3
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)
Table 3.6 Correlations between the C−O stretching frequency and the hydrogen-bonding status of tyrosine side−chain
groups in proteins
No. of
H-bonds
IR/Raman
Freq.
C−O (cm-1)
IR/Raman
Freq.
O−H (cm-1)
Protein* Phenol
Group of
H-bond
Partner
Role of
Tyrosine
H-bond Length
(Å)
PDB
code Resolution
1279 1255 PSII TyrZ His190 Acceptor 2.78 2AXT 3.00 Å
1275 1250 PSII TyrD His189(0) Acceptor 2.59 2AXT 3.00 Å
1274 N/A CcO (P.D.,
oxidized) Tyr35 SER134 Acceptor 3.09 1QLE 3.00 Å
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
1
1240 N/A bR (K) Tyr185 Asp212 Donor 2.75 1M0K 1.43 Å
2 1265 1232 Human
MnSOD Tyr34
H2O
Gln143
Donor
Acceptor
2.8
3.0 1LUV 1.85 Å
98
99
3.7 CONCLUSION
Our density function theory based ab initio computational studies on the hydrogen-
bonding 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 two-
dimensional 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.
100
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
101
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
102
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.
103
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.
104
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.
105
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).
There are other polar side−chain groups at the active sites other than protonated
Asp/Glu and Tyr. We will proceed to further to calculate the hydrogen−bonding
properties of Asn/Gln, Arg+, neutral His and His+, negatively charged Asp/Glu
(Asp/Glu−), negatively charged Tyr (Tyr−), Lys+, and Ser/Thr. Our preliminary results
show that some of the vibrational frequencies of these polar groups are sensitive to
hydrogen−bonding interactions. With Gaussian data analysis and experimental support
from the literatures, a series of vibrational structural markers can be established. Thus
these vibrational structural markers are powerful tools to get insights into the hydrogen-
bonding status and local environments of important amino acids during functionally
important intermediate states of proteins.
106
References
Adam, G., and J. H. Gibbs. 1965. On temperature dependence of cooperative relaxation properties in glass forming liquids. J. Chem. Phys. 43:139.
Ames, J. B., M. Ros, J. Raap, J. Lugtenburg, and R. A. Mathies. 1992. Time-Resolved Ultraviolet Resonance Raman Studies Of Protein-Structure - Application To Bacteriorhodopsin. Biochemistry 31:5328-5334.
Arakawa, T., and S. N. Timasheff. 1984. Mechanism of protein salting in and salting out by divalent-cation salts - balance between hydration and salt binding. Biochemistry 23:5912-5923.
Ayala, I., J. J. P. Perry, J. Szczepanski, J. A. Tainer, M. T. Vala, H. S. Nick, and D. N. Silverman. 2005. Hydrogen bonding in human manganese superoxide dismutase containing 3-fluorotyrosine. Biophys. J. 89:4171-4179.
Baker, D. 2000. A surprising simplicity to protein folding. Nature 405:39-42.
Bakker, H. J., M. F. Kropman, and A. W. Omta. 2005. Effect of ions on the structure and dynamics of liquid water. J. Phys.-Condens. Mat. 17:S3215-S3224.
Berthomieu, C., R. Hienerwadel, A. Boussac, J. Breton, and B. A. Diner. 1998. Hydrogen bonding of redox-active tyrosine Z of photosystem II probed by FTIR difference spectroscopy. Biochemistry 37:10547-10554.
Billeter, M., T. Schaumann, W. Braun, and K. Wuthrich. 1990. Restrained Energy Refinement With 2 Different Algorithms And Force-Fields Of The Structure Of The Alpha-Amylase Inhibitor Tendamistat Determined By NMR In Solution. Biopolymers 29:695-706.
Blundell, T. L., and S. Patel. 2004. High-throughput X-ray crystallography for drug discovery. Curr. Opin. Pharmacol. 4:490-496.
Borgstahl, G. E., D. R. Williams, and E. D. Getzoff. 1995a. 1.4 A structure of photoactive yellow protein, a cytosolic photoreceptor: unusual fold, active site, and chromophore. Biochemistry 34:6278−6287.
Borgstahl, G. E. O., D. R. Williams, and E. D. Getzoff. 1995b. 1.4 Å Structure Of Photoactive Yellow Protein, A Cytosolic Photoreceptor - Unusual Fold, Active-Site, And Chromophore
Biochemistry 34:6278-6287.
107
Braatsch, S., M. Gomelsky, S. Kuphal, and G. Klug. 2002. A single flavoprotein, AppA, integrates both redox and light signals in Rhodobacter sphaeroides. Mol. Microbiol. 45:827-836.
Braiman, M. S., T. Mogi, T. Marti, L. J. Stern, H. G. Khorana, and K. J. Rothschild. 1988a. Vibrational spectroscopy of Bacteriorhodopsin mutants - light -driven proton transport involves protonation changes of Aspartic acid residue-85, residue-96 and residue-212. Biochemistry 27:8516-8520.
Braiman, M. S., T. Mogi, L. J. Stern, N. R. Hackett, B. H. Chao, H. G. Khorana, and K. J. Rothschild. 1988b. Vibrational spectroscopy of bacteriorhodopsin mutants: I. Tyrosine-185 protonates and deprotonates during the photocycle. Proteins 3:219-229.
Braiman, M. S., T. Mogi, L. J. Stern, N. R. Hackett, B. H. Chao, H. G. Khorana, and K. J. Rothschild. 1988c. Vibrational spectroscopy of Bacteriorodopsin mutants. 1. Tyrosine-185 protonates and deprotonates during the photocycle. Proteins-Struct. Funct. Genet. 3:219-229.
Braiman, M. S., and K. J. Rothschild. 1988. Fourier-Transform Infrared Techniques For Probing Membrane-Protein Structure. Annu. Rev. Biophy. Biochem. 17:541-570.
Broering, J. M., and A. S. Bommarius. 2005. Evaluation of Hofmeister effects on the kinetic stability of proteins. J. Phys. Chem. B 109:20612-20619.
Brown, I. D. 1976. Geometry Of O-H...O Hydrogen-Bonds. Acta Crystallogr. A 32:24-31.
Brudler, R., T. Meyer, U. K. Genick, S. Devanathan, T. Woo, D. Millar, K. Gerwert, M. Cusanovich, G. Tollin, and E. Getzoff. 2000a. Coupling of hydrogen bonding to chromophore conformation and function in photoactive yellow protein. Biochemistry 39:13478−13486.
Brudler, R., T. E. Meyer, U. K. Genick, S. Devanathan, T. T. Woo, D. P. Millar, K. Gerwert, M. A. Cusanovich, G. Tollin, and E. D. Getzoff. 2000b. Coupling of hydrogen bonding to chromophore conformation and function in photoactive yellow protein. Biochemistry 39:13478-13486.
Brudler, R., R. Rammelsberg, T. T. Woo, E. D. Getzoff, and K. Gerwert. 2001. Strucutre of the I1 early intermediate of photoactive yellow protein by FTIR spectroscopy. Nat. Struct. Biol. 8:265−270.
Cacace, M. G., E. M. Landau, and J. J. Ramsden. 1997a. The Hofmeister series: salt and solvent effects on interfacial phenomena. Q. Rev. of Biophys. 30:241-277.
Cacace, M. G., E. M. Landau, and J. J. Ramsden. 1997b. The Hofmeister series: salt and solvent effects on interfacial phenomena. Q. Rev. Biophys. 30:241−277.
108
Caminiti, R., G. Licheri, G. Piccaluga, and G. Pinna. 1978. NO-3-H2O interactions in aqueous solutions. J. Chem. Phys. 68:1967-1970.
Caminiti, R., G. Paschina, G. Pinna, and M. Magini. 1979. Experimental evidence of interactions SO4(2-)-H2O in an aqueous solution. Chem. Phys. Lett. 64:391-395.
Carugo, O. 2007. Recent progress in measuring structural similarity between proteins. Curr. Prot. Pep. Sci. 8:219-241.
Caughey, B., and P. T. Lansbury. 2003. Protofibrils, pores, fibrils, and neurodegeneration: Separating the responsible protein aggregates from the innocent bystanders. Annu. Rev. Neurosci. 26:267-298.
Chandra, A. K., T. Uchimaru, M. Sugie, and A. Sekiya. 2000. Correlation between hardness and activation energies for reactions of OH radical with halomethanes. Chem. Phys. Lett. 318:69-74.
Chayen, N. E., T. J. Boggon, A. Cassetta, A. Deacon, T. Gleichmann, J. Habash, S. J. Harrop, J. R. Helliwell, Y. P. Nieh, M. R. Peterson, J. Raftery, E. H. Snell, A. Hadener, A. C. Niemann, D. P. Siddons, V. Stojanoff, A. W. Thompson, T. Ursby, and M. Wulff. 1996. Trends and challenges in experimental macromolecular crystallography. Q. Rev. of Biophys. 29:227-278.
Collins, K. D. 1995. Sticky Ions in Biological-Systems. Proc. Natl. Acad. Sci. USA 92:5553-5557.
Collins, K. D., and M. W. Washabaugh. 1985. The Hofmeister Effect And the Behavior of Water at Interfaces. Q. Rev. of Biophys. 18:323-422.
Creighton, T. E. 1993. Proteins: Structure and Molecular Properties. W. H. Freeman and Company, New York.
Creighton, T. E., and R. B. Freedman. 1993. Protein-Structure - A Model Catalyst Of Protein Disulfide Bond Formation. Curr. Biol. 3:790-793.
Crosson, S., and K. Moffat. 2001. Structure of a flavin-binding plant photoreceptor domain: insights into light-mediated signal transduction. Proc. Natl. Acad. Sci. USA 98:2995−3000.
Cubitt, A. B., R. Heim, S. R. Adams, A. E. Boyd, L. A. Gross, and R. Y. Tsien. 1995. Understanding, Improving and using Green Fluorescent Proteins. Trends Biochem. Sci. 20:448-455.
Desiraju, G. R., and T. Steiner. 1999. The Weak Hydrogen Bond in Structural Chemistry and Biology. Oxford University Press, Oxford.
Devanathan, S., R. Brudler, B. Hessling, T. T. Woo, K. Gerwert, E. D. Getzoff, M. A. Cusanovich, and G. Tollin. 1999. Dual photoactive species in Glu46Asp and
109
Glu46Ala mutants of photoactive yellow protein: a pH-driven color transition. Biochemistry 38:13766−13772.
Dobson, C. M. 1999. Protein misfolding, evolution and disease. Trends Biochem. Sci. 24:329-332.
Dobson, C. M. 2002. Protein-misfolding diseases: Getting out of shape. Nature 418:729-730.
Dohlman, H. G., J. Thorner, M. G. Caron, and R. J. Lefkowitz. 1991. Model systems for the study of 7-transmembrane segmant receptors. Annu. Rev. Biochem. 60:653-688.
Domene, C., and M. S. P. Sansom. 2003. Potassium channel, ions, and water: Simulation studies based on the high resolution X-ray structure of KcsA. Biophys. J. 85:2787-2800.
Ducruix, A., J. P. Guilloteau, M. RiesKautt, and A. Tardieu. 1996. Protein interactions as seen by solution X-ray scattering prior to crystallogenesis. J. Cryst. Growth 168:28-39.
Durbin, S. D., and G. Feher. 1996. Protein crystallization. Annu. Rev. Phys. Chem. 47:171-204.
Dux, P., G. Rubinstenn, G. W. Vuister, R. Boelens, F. A. Mulder, K. Hard, W. D. Hoff, A. R. Kroon, W. Crielaard, K. J. Hellingwerf, and R. Kaptein. 1998. Solution structure and backbone dynamics of the photoactive yellow protein. Biochemistry 37:12689−12699.
Endom, L., H. G. Hertz, B. Thul, and M. D. Zeidler. 1967. A Microdynamic Model Of Electrolyte Solutions As Derived From Nuclear Magnetic Relaxation And Self-Diffusion Data. Ber. Bunsen. Gesell. Phys. Chem. 71:1008-1031.
Faller, P., R. J. Debus, K. Brettel, M. Sugiura, A. W. Rutherford, and A. Boussac. 2001. Rapid formation of the stable tyrosyl radical in photosystem II. Proc. Natl. Acad. Sci. USA 98:14368-14373.
Fenimore, P. W., H. Frauenfelder, B. H. McMahon, and R. D. Young. 2004. Bulk-solvent and hydration-shell fluctuations, similar to alpha- and beta-fluctuations in glasses, control protein motions and functions. Proc. Natl. Acad. Sci. USA 101:14408-14413.
Force, D. A., D. W. Randall, R. D. Britt, X. S. Tang, and B. A. Diner. 1995. H-2 ESE-ENDOR study of hydrogen bonding to the tyrosine radicals Y-D center dot and Y-Z center dot of photosystem II. J. Am. Chem. Soc. 117:12643-12644.
Foresman, J. B., and Æ. Frisch. 1996. Exploring chemistry with electronic structure methods. Gaussian Inc., Pittsburgh.
110
Frauenfelder, H., P. W. Fenimore, G. Chen, and B. H. McMahon. 2006. Protein folding is slaved to solvent motions. Proc. Natl. Acad. Sci. USA 103:15469-15472.
Frauenfelder, H., S. G. Sligar, and P. G. Wolynes. 1991. The energy landscapes and motions of proteins. Science 254:1598−1603.
Frauenfelder, H., and P. G. Wolynes. 1985. Rate theories and puzzles of hemeprotein kinetics. Science 229:337−345.
Frisch, Æ., M. J. Frisch, and G. W. Trucks. 2003. Gaussian 03 user's reference. Gaussian, Inc., Carnegie.
Fromme, P., and P. Mathis. 2004. Unraveling the Photosystem I reaction center: a history, or the sum of many efforts. Photosynthesis Research 80:109-124.
Gao, F. P., and T. A. Cross. 2005. Recent developments in membrane-protein structural genomics. Genome Biol. 6:244.
Gauden, M., J. S. Grinstead, W. Laan, H. M. van Stokkum, M. Avila-Perez, K. C. Toh, R. Boelens, R. Kaptein, R. van Grondelle, K. J. Hellingwerf, and J. T. M. Kennis. 2007. On the role of aromatic side chains in the photoactivation of BLUF domains. Biochemistry 46:7405-7415.
Gauden, M., I. H. M. van Stokkum, J. M. Key, D. C. Luhrs, R. Van Grondelle, P. Hegemann, and J. T. M. Kennis. 2006. Hydrogen-bond switching through a radical pair mechanism in a flavin-binding photoreceptor. Proc. Natl. Acad. Sci. USA 103:10895-10900.
Genick, U. K., G. E. Borgstahl, K. Ng, Z. Ren, C. Pradervand, P. M. Burke, V. Srajer, T. Y. Teng, W. Schildkamp, D. E. McRee, K. Moffat, and E. D. Getzoff. 1997. Structure of a protein photocycle intermediate by millisecond time-resolved crystallography. Science 275:1471−1475.
Genick, U. K., S. M. Soltis, P. Kuhn, I. L. Canestrelli, and E. D. Getzoff. 1998. Structure at 0.85 Å resolution of an early protein photocycle intermediate. Nature 392:206−209.
Getzoff, E. D., K. N. Gutwin, and U. K. Genick. 2003. Anticipatory active-site motions and chromophore distortion prime photoreceptor PYP for light activation. Nat. Struct. Biol. 10:663−668.
Gilchrist, M. L., J. A. Ball, D. W. Randall, and R. D. Britt. 1995. Proximity of the Manganese cluster of Photosystem-II to the Redox-active Tyrosine Y-Z. Proc. Natl. Acad. Sci. USA 92:9545-9549.
Gilliland, G. L., M. Tung, and J. E. Ladner. 2002. The biological macromolecule crystallization database: crystallization procedures and strategies. Acta Cryst. D. 58:916-920.
111
Goormaghtigh, E., V. Cabiaux, and J. M. Ruysschaert. 1990. Secondary Structure And Dosage Of Soluble And Membrane-Proteins By Attenuated Total Reflection Fourier-Transform Infrared-Spectroscopy On Hydrated Films
Eur. J. Biochem. 193:409-420.
Grigorieff, N., T. A. Ceska, K. H. Downing, J. M. Baldwin, and R. Henderson. 1996. Electron-crystallographic refinement of the structure of bacteriorhodopsin. J. Mol. Biol. 259:393-421.
Hajdu, J., and I. Andersson. 1993. Fast Crystallography And Time-Resolved Structures. Annu. Rev. Biophys. Biomol. Struct. 22:467-498.
Heberle, J. 2000. Proton transfer reactions across bacteriorhodopsin and along the membrane. Biochim. Biophys. Acta. 1458:135−147.
Hendrickson, W. A., and C. M. Ogata. 1997. Phase determination from multiwavelength anomalous diffraction measurements. Methods Enzymol. 494-523.
Hienerwadel, R., A. Boussac, J. Breton, B. A. Diner, and C. Berthomieu. 1997. Fourier transform infrared difference spectroscopy of photosystem II tyrosine D using site-directed mutagenesis and specific isotope labeling. Biochemistry 36:14712-14723.
Hoff, W. D., A. Xie, I. H. Van Stokkum, X. J. Tang, J. Gural, A. R. Kroon, and K. J. Hellingwerf. 1999. Global conformational changes upon receptor stimulation in photoactive yellow protein. Biochemistry 38:1009−1017.
Hofmeister, F. 1888. Zur Lehre von der Wirkung der Salze. II. Arch. Exp. Pathol. Pharmakol. 24:247-260.
Illman, D. L. 1994. Chem. Engg. News Ap. 25.
Imamoto, Y., M. Kataoka, and F. Tokunaga. 1996. Photoreaction cycle of photoactive yellow protein from Ectothiorhodospira halophila studied by low-temperature spectroscopy. Biochemistry 35:14047−14053.
Imhoff, J. F. 1984. Reassignment Of The Genus Ectothiorhodospira Pelsh 1936 To A New Family, Ectothiorhodospiraceae Fam Nov, And Emended Description Of The Chromatiaceae Bavendamm 1924. Int. J. Sys. Bacter. 34:338-339.
Isaacs, E. D., A. Shukla, P. M. Platzman, D. R. Hamann, B. Barbiellini, and C. A. Tulk. 2000. Compton scattering evidence for covalency of the hydrogen bond in ice. J. Phys. Chem. Sol. 61:403-406.
Kameda, Y., H. Saitoh, and O. Uemura. 1993. The Hydration Structure Of NO3- In Concentrated Aqueous Sodium-Nitrate Solutions. Bull. Chem. Soc. Japan 66:1919-1923.
112
Kendrew, J. C., G. Bodo, H. M. Dintzis, R. G. Parrish, H. Wyckoff, and D. C. Phillips. 1958. 3-Dimensional Model Of The Myoglobin Molecule Obtained By X-Ray Analysis. Nature 181:662-666.
Kita, A., K. Okajima, Y. Morimoto, M. Ikeuchi, and K. Miki. 2005. Structure of a cyanobacterial BLUF protein, Tll0078, containing a novel FAD-binding blue light sensor domain. J. Mol. Biol. 349:1-9.
Kowalsky, A. 1962. Nuclear Magnetic Resonance studies of Proteins. J. Biol. Chem. 237:1807-1819.
Krestov, G. A. 1962. Standard state of ions in solution. Zhur. Fizi. Khim. 36:1549-1552.
Krimm, S., and J. Bandekar. 1986. Vibrational Spectroscopy And Conformation Of Peptides, Polypeptides, And Proteins. Adv. Protein Chem. 38:181-364.
Kulhanek, P., E. W. Schlag, and J. Koca. 2003. A novel mechanism of proton transfer in protonated peptides. J. Am. Chem. Soc. 125:13678-13679.
Laan, W., M. A. van der Horst, I. H. van Stokkum, and K. J. Hellingwerf. 2003. Initial characterization of the primary photochemistry of AppA, a blue-light-using flavin adenine dinucleotide-domain containing transcriptional antirepressor protein from Rhodobacter sphaeroides: A key role for reversible intramolecular proton transfer from the flavin adenine dinucleotide chromophore to a conserved tyrosine? Photochem. and Photobiol. 78:290-297.
Lanyi, J. K. 2004. X-ray diffraction of bacteriorhodopsin photocycle intermediates. Mol. Membr. Biol. 21:143−150.
Lu, J. R., and R. K. Thomas. 1998. Neutron reflection from wet interfaces. J. Chem. Soc.-Faraday Transactions 94:995-1018.
Lundstrom, K. 2006. Structural genomics for membrane proteins. Cell. Mol. Life Sci. 63:2597-2607.
Magini, M., G. Paschina, and G. Piccaluga. 1982. NI-Cl Bonding In Concentrated NI(II) Aqueous-Solutions At High CI-NII2+ Ratios - An X-Ray-Diffraction Investigation. J. Chem. Phys. 76:1116-1121.
Makinen, M. W., and W. A. Eaton. 1973. Polarized single-crystal absorption spectra of carboxyhemoglobin and oxyhemoglobin. Annu. N. Y. Acad. Sci. 206:210−222.
Makinen, M. W., and A. L. Fink. 1977. Reactivity and cryoenzymology of enzymes in the crystalline state. Annu. Rev. Biophys. Bioeng. 6:301−343.
Maslen, V. W. 1967. Crystal Ionic radii. Proc. Phys. Soc. 91:259.
113
McDonald, C. C., and W. D. Phillips. 1967. Manifestations Of Tertiary Structures Of Proteins In High-Frequency Nuclear Magnetic Resonance. J. Am. Chem. Soc. 89:6332-6341.
Membrane proteins of known structure. [http://blanco.biomol.uci.edu/Membrane_Proteins_xtal.html]. 2005.
Meyer, T. E. 1985. Isolation and characterization of soluble cytochromes, ferredoxins and other chromophoric proteins from the halophilic phototrophic bacterium Ectothiorhodospira halophila. Biochim. Biophys. Acta. 806:175−183.
Meyer, T. E., J. C. Fitch, R. G. Bartsch, G. Tollin, and M. A. Cusanovich. 1990. Soluble cytochromes and a photoactive yellow protein isolated from the moderately halophilic purple phototrophic bacterium, Rhodospirillum salexigens. Biochim. Biophys. Acta. 1016:364−370.
Meyer, T. E., E. Yakali, M. A. Cusanovich, and G. Tollin. 1987. Properties of a water-soluble, yellow protein isolated from a halophilic phototrophic bacterium that has photochemical activity analogous to sensory rhodopsin. Biochemistry 26:418−423.
Michel, H. 1983. Crystallization of Membrane Proteins. Trends Biochem. Sci. 8:56-59.
Morais, F., J. Barber, and P. J. Nixon. 1998. The chloroplast-encoded alpha subunit of cytochrome b-559 is required for assembly of the photosystem two complex in both the light and the dark in Chlamydomonas reinhardtii. J. Biol. Chem. 273:29315-29320.
Murphy, R. M. 2002. Peptide aggregation in neurodegenerative disease. Annu. Rev. Biomed. Engg. 4:155-174.
Musinu, A., G. Paschina, G. Piccaluga, and M. Magini. 1982. The Sulphate Ion in aqueous solution - An X-ray diffraction study of a ZnSO4 solution. J. App. Crystallogr. 15:621-625.
Nagle, J. F., and S. Tristram-Nagle. 1983. Hydrogen bonded chain mechanisms for proton conduction and proton pumping. J. Membr. Biol. 74:1−14.
Nambu, J. R., J. O. Lewis, K. A. Wharton, and S. T. Crews. 1991. The Drosophila Single-Minded Gene Encodes A Helix-Loop-Helix Protein That Acts As A Master Regulator Of CNS Midline Development
Cell 67:1157-1167.
Neilson, G. W., R. D. Broadbent, I. Howell, and R. H. Tromp. 1993. Structural and Dynamical aspects of Aqueous Ionic Solutions. J. Chem. Soc. Faraday Trans. 89:2927-2936.
114
Nelson, D. L., and M. M. Cox. 2000. Lehninger principle of biochemistry. Worth Publishers, New York.
Ng, K., E. D. Getzoff, and K. Moffat. 1995. Optical studies of a Bacterial Photoreceptor Protein, Photoactive Yellow Protein, in single crystals. Biochemistry 34:879-890.
Nie, B. 2006. Probing hydrogen bonding interactions and proton transfer in proteins. Oklahoma State University. 243.
Nie, B., J. Stutzman, and A. Xie. 2005. A vibrational spectral marker for probing the hydrogen-bonding status of protonated Asp and Glu residues. Biophys. J. 88:2833-2847.
Ohnishi, S., and K. Takano. 2004. Amyloid fibrils from the viewpoint of protein folding. Cell. Mol. Lif Sci. 61:511-524.
Parak, F. G. 2003. Proteins in action: the physics of structural fluctuations and conformational changes. Curr. Opin. Struct. Biol. 13:552-557.
Parikh, S., D. P. Moynihan, G. P. Xiao, and P. J. Tonge. 1999. Roles of tyrosine 158 and lysine 165 in the catalytic mechanism of InhA, the enoyl-ACP reductase from Mycobacterium tuberculosis. Biochemistry 38:13623-13634.
Parsegian, V. A. 2002. Protein-water interactions. Int. Rev. Cytol. 215:1-31.
Pellequer, J. L., K. A. Wager-Smith, S. A. Kay, and E. D. Getzoff. 1998. Photoactive yellow protein: a structural prototype for the three-dimensional fold of the PAS domain superfamily. Proc. Natl. Acad. Sci. USA 95:5884−5890.
Pinna, M. C., P. Bauduin, D. Touraud, M. Monduzzi, B. W. Ninham, and W. Kunz. 2005. Hofmeister effects in biology: Effect of choline addition on the salt-induced super activity of horseradish peroxidase and its implication for salt resistance of plants. J. Phys. Chem. B 109:16511-16514.
Procopio, M., A. Lahm, A. Tramontano, L. Bonati, and D. Pitea. 2002. A model for recognition of polychlorinated dibenzo-p-dioxins by the aryl hydrocarbon receptor. Eur. J. Biochem. 269:13−18.
Ramos, S., A. C. Barnes, G. W. Neilson, D. Thiaudiere, and S. Lequien. 2000. The hydration structure of Br- from anomalous x-ray diffraction. J. Phys.-Condens. Mat. 12:A203-A208.
The RCSB Protein Data Bank [http://www.rcsb.org/pdb]. 2007.
Rothschild, K. J. 1992. FTIR difference spectroscopy of bacteriorhodopsin: toward a molecular model. J. Bioenerg. Biomembr. 24:147−167.
115
Rothschild, K. J., M. S. Braiman, Y. W. He, T. Marti, and H. G. Khorana. 1990. Vibrational spectroscopy of bacteriorhodopsin mutants. Evidence for the interaction of aspartic acid 212 with tyrosine 185 and possible role in the proton pump mechanism. J. Biol. Chem. 265:16985-16991.
Rothschild, K. J., and H. Marrero. 1982. Infrared evidence that the Schiff base of bacteriorhodopsin is protonated: bR570 and K intermediates. Proc. Natl. Acad. Sci. USA 79:4045−4049.
Rozwarski, D. A., C. Vilcheze, M. Sugantino, R. Bittman, and J. C. Sacchettini. 1999. Crystal structure of the Mycobacterium tuberculosis enoyl-ACP reductase, InhA, in complex with NAD(+) and a C16 fatty acyl substrate. J. Biol. Chem. 274:15582-15589.
Rupley, J. A., and G. Careri. 1991. Protein hydration and function. Adv. Protein Chem. 41:37−172.
Rutherford, A. W., A. Boussac, and P. Faller. 2004. The stable tyrosyl radical in Photosystem II: why D? Biochim. Biophys. Acta 1655:222-230.
Samoilov, O. Y., Yashkich.Vi, V. G. Markova, Alekseev.Ls, and Goncharo.Vv. 1972. Ionic State In Aqueous-Solutions According To Self-Diffusion Data .1. Self-Diffusion Of Cl(-)-36 And I(-)-131 In Aqueous-Solutions Of Alkali-Metal And Ammonium Chlorides
Zhur. Strukt. Khim. 13:384.
Sanchez, R., and A. Sali. 1997. Advances in comparative protein-structure modelling. Curr. Opin. Struct. Biol. 7:206-214.
Saunders, M., and A. Wishnia. 1958. Nuclear Magnetic Resonance spectra of Proteins. Annu. N.Y. Acad. Sci. 70:870-874.
Scapin, G. 2006. Structural biology and drug discovery. Curr. Pharm. Des. 12:2087-2097.
Schotte, F., J. Soman, J. S. Olson, M. Wulff, and P. A. Anfinrud. 2004. Picosecond time-resolved X-ray crystallography: probing protein function in real time. J. Struct. Biol. 147:235-246.
Skipper, N. T., and G. W. Neilson. 1989. X-Ray And Neutron-Diffraction Studies On Concentrated Aqueous-Solutions Of Sodium-Nitrate And Silver-Nitrate. J. Phys. Chem.- Condens. Mat. 1:4141-4154.
Sprenger, W. W., W. D. Hoff, J. P. Armitage, and K. J. Hellingwerf. 1993. The eubacterium Ectothiorhodospira halophila is negatively phototactic, with a wavelength dependence that fits the absorption spectrum of the photoactive yellow protein. J. Bacteriol. 175:3096−3104.
116
Takahashi, R., K. Okajima, H. Suzuki, H. Nakamura, M. Ikeuchi, and T. Noguchi. 2007. FTIR study on the hydrogen bond structure of a key tyrosine residue in the flavin-binding blue light sensor TePixD from Thermosynechococcus elongatus. Biochemistry 46:6459-6467.
Takeuchi, H., N. Watanabe, Y. Satoh, and I. Harada. 1989. Effects of hydrogen bonding on the tyrosine Raman bands in the 1300-1150 cm-1 region. J. Raman Spectrosc. 20:233-237.
Tamm, L. K., and S. A. Tatulian. 1997. Infrared spectroscopy of proteins and peptides in lipid bilayers. Q. Rev. of Biophys. 30:365-429.
Tanford, C. 1961. Ionization linked changes in protein conformation theory. J. Am. Chem. Soc. 83:1628.
Taylor, B. L., and I. B. Zhulin. 1999. PAS domains: Internal sensors of oxygen, redox potential, and light. Microbiol. Mol. Biol. Rev. 63:479−506.
Thompson, W. E., and G. C. Pimentel. 1960. The 1st Overtone Of The Hydrogen Bending Mode Of Chloroform - Enhancement By Hydrogen Bonding. Zeitschrift Fur Elektrochemie 64:748-753.
Ujj, L., S. Devanathan, T. E. Meyer, M. A. Cusanovich, G. Tollin, and G. H. Atkinson. 1998. New photocycle intermediates in the photoactive yellow protein from Ectothiorhodospira halophila: picosecond transient absorption spectroscopy. Biophys. J. 75:406−412.
Un, S., X. S. Tang, and B. A. Diner. 1996. 245 GHz high-field EPR study of tyrosine-D degrees and tyrosine-Z degrees in mutants of photosystem II. Biochemistry 35:679-684.
van Brederode, M. E., T. Gensch, W. D. Hoff, K. J. Hellingwerf, and S. E. Braslavsky. 1995. Photoinduced volume change and energy storage associated with the early transformations of the photoactive yellow protein from Ectothiorhodospira halophila. Biophys. J. 68:1101−1109.
van Thor, J. J., A. J. Pierik, I. Nugteren-Roodzant, A. Xie, and K. J. Hellingwerf. 1998a. Characterization of the photoconversion of green fluorescent protein with FTIR spectroscopy. Biochemistry 37:16915−16921.
van Thor, J. J., A. J. Pierik, I. Nugteren-Roodzant, A. H. Xie, and K. J. Hellingwerf. 1998b. Characterization of the photoconversion of green fluorescent protein with FTIR spectroscopy. Biochemistry 37:16915-16921.
Voet, D., and J. Voet. 2004. Biochemistry. John Wiley & Sons, Inc., New Jersey.
Vollmar, P. M. 1963. Ionic interactions in aqueous solution - A Raman spectral study. J. Chem. Phys. 39:2236.
117
Vonhippe.Ph, and T. Schleich. 1969. Ion effects on solution structure of biological macromolecules. Acc. Chem. Res. 2:257.
Vorontsov, A. V., and Y. V. Novakovskaya. 2007. Ionization of Sodium in Water Clusters. Russ. J. Phys. Chem. 81:759-767.
Wang, M., T. E. Wales, and M. C. Fitzgerald. 2006. Conserved thermodynamic contributions of backbone hydrogen bonds in a protein fold. Proc. Natl. Acad. Sci. USA 103:2600-2604.
Whisstock, J. C., and A. M. Lesk. 2003. Prediction of protein function from protein sequence and structure. Q. Rev. of Biophys. 36:307-340.
Wishart, D. S., and B. D. Sykes. 1994. Chemical-Shifts as a tool for structure determination. Methods Enzymol. 363-392.
Woutersen, S., U. Emmerichs, and H. J. Bakker. 1997. Femtosecond mid-IR pump-probe spectroscopy of liquid water: Evidence for a two-component structure. Science 278:658-660.
Wuthrich, K. 2001. The way to NMR structures of proteins. Nat. Struct. Biol. 8:923-925.
Wuthrich, K. 2003. NMR studies of structure and function of biological macromolecules (Nobel Lecture). Angew. Chem., Int. Ed. 42:3340-3363.
Xie, A., W. D. Hoff, A. R. Kroon, and K. J. Hellingwerf. 1996a. Glu46 donates a proton to the 4-hydroxycinnamate anion chromophore during the photocycle of photoactive yellow protein. Biochemistry 35:14671−14678.
Xie, A., L. Kelemen, J. Hendriks, B. J. White, K. J. Hellingwerf, and W. D. Hoff. 2001a. Formation of a new buried charge drives a large-amplitude protein quake in photoreceptor activation. Biochemistry 40:1510−1517.
Xie, A. H., W. D. Hoff, A. R. Kroon, and K. J. Hellingwerf. 1996b. Glu46 donates a proton to the 4-hydroxycinnamate anion chromophore during the photocycle of photoactive yellow protein. Biochemistry 35:14671-14678.
Xie, A. H., L. Kelemen, J. Hendriks, B. J. White, K. J. Hellingwerf, and W. D. Hoff. 2001b. Formation of a new buried charge drives a large-amplitude protein quake in photoreceptor activation. Biochemistry 40:1510-1517.
Yamaguchi, T., M. Niihara, T. Takamuku, H. Wakita, and H. Kanno. 1997. Scandium(III) hydration in aqueous solution from X-ray diffraction and X-ray absorption fine structure measurements. Chem. Phys. Lett. 274:485-490.
Yoo, H., J. Boatz, V. Helms, J. McCammon, and P. Langhoff. 2001. Chromophore protonation states and the proton shuttle mechanism in green fluorescent protein:
118
inferences drawn from ab Initio theoretical studies of chemical structures and vibrational spectra. J. Phys. Chem. 105:2850−2857.
Zhang, L., Y. T. Kao, W. Qiu, Y. Yang, O. Okobiah, L. Wang, and D. Zhong. 2007. Mapping hydration dynamics around a protein surface. Proc. Natl. Acad. Sci. USA.
Zhang, Y. J., and P. S. Cremer. 2006. Interactions between macromolecules and ions: the Hofmeister series. Curr. Opin. Chem. Biol. 10:658-663.
Zhao, H. 2006. 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.
119
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.
120
• 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.
121
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
122
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 (saved at
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
123
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)
124
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
125
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.
126
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.
127
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.
128
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.
129
PART C: Selected Salts from Hofmeister Series
Salt Chemical formula Molecular
weight Solubility
(% by weight)
Solubility (by volume)
Sodium fluoride NaF 41.99 3.97% 0.95M
Sodium chloride NaCl 58.44 26.45% 5.13M
Sodium bromide NaBr 102.9 48.6% 9.2M
Sodium iodide NaI 149.89 64.8% 11.3M
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%
Ammonium chloride
NH4Cl 53.49 28.34% 5.5M
Ammonium sulphate
(NH4)2SO4 132.14 43.3% 5M
Magnesium sulphate
MgSO4 120.5 26.3% 2.56M
Sodium phosphate monobasic
NaH2PO4 119.99 48.68%
Sodium phosphate dibasic
Na2HPO4 141.96 10.55%
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
130
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.
131
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.
132
** 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.
133
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)
134
• 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.
135
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)
Time to centrifuge (min)
@ 9000 rpm (RCF=6,610)
@ 12000rpm (RCF=11,750)
NaCl 20 10 CsCl 20 5 LiCl 20 20 KCl 20 10
NaBr 20 20
136
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.
Wavelength [nm]250 300 350 400 450 500
Abs
orpt
ion
[OD
]
0.0
0.1
0.2
0.3
0.4
0.5
0.64.0 M NaCl
1.0 M NaCl2.0 M NaCl
0.5 M NaCl
wtPYP in
137
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
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
Wavenumber [cm -1
]
250 300 350 400 450 500
Abs
orpt
ion
[OD
]
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.84.0 M NaBr
4.0 M CsCl4.0 M KCl
4.0 M LiCl
4.0 M NaCl
138
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
Wavenumber [cm-1]
1200130014001500160017001800
Diff
Abs
orpt
ion
[mill
i OD
]
-4.0
0.0
4.0
8.0
1740
1689
1624
1497
1669
1515
1726
1302
0.0 M NaCl
4.0 M NaBr
4.0 M NaCl
1162
1641
1253
1571
1609
139
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.
1689
1624
1497
1669
1515
1726
Salt-free wt PYP2.0 M MgSO 4
1641
1571
Wavenumber [cm-1]
1200130014001500160017001800
Diff
Abs
orpt
ion
[mill
i OD
]
-4.0
0.0
4.0
0.67 M MgSO 4
1253
1162
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.
140
Wavenumber [cm-1]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 2.0 M MgSO4
Wavenumber [cm-1]
Diff
. Abs
orba
nce
[mill
iOD
] 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.
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 0.67 M MgSO4
Wavenumber [cm-1]Time [s]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 0.67 M MgSO4
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 0.0 M salt
Time [s]Wavenumber [cm-1]
Diff
. Abs
orba
nce
[mill
iOD
] WT PYP in 0.0 M salt
Time [s]Wavenumber [cm-1]
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
