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RAMAN SPECTROSCOPY IN BIOLOGY: Principles and Applications
ANTHONY T. TU Department of Biochemistry Colorado State University
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:I807m1982 U8LIStn~
A Wlley-Interscience Publication
JOHN WILEY & SONS New York
Chichester
Brisbane
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Toronto
Singapore
Copyright © 1982 by John Wiley & Sons, Inc. All rights reserved. Published simultaneously in Canada. Reproduction or translation of any part of this work beyond that permitted by Section 107 or 108 of the 1976 United States Copyright Act without the permission of the copyright owner is unlawful. Requests for permission or further information should be addressed to the Permissions Department, John Wiley & Sons, Inc. Library of Congress Cataloging in Publication Data:
Tu, Anthony T., 1930Raman spectroscopy in biology. "A Wiley-Interscience publication." Includes bibliographical references and index. I. Raman spectroscopy. 2. Biology- Methodology. I. Title.
82-6901 QH324.9.S6T8 574.19'285 ISBN 0-471-07984-7 AACR2 Printed in the United States of America 10 9 8
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Preface
Although the Raman effect was discovered in 1928, the application of Raman spectroscopy to biological compounds is quite recent. Its progress, however, has been phenomenal. There are many books on Raman spectroscopy, but most are written for physicists and chemists. Because of its potential applicability to biological problems, Raman spectroscopy will become a more common tool in biological research once biologists become familiar with this powerful research technique. This book is written for those interested primarily in biological application rather than those doing research for Raman spectroscopy. Therefore, the description is more qualitative, without using complex mathematics and physical chemistry. Many scientists who do not have strong backgrounds in higher mathematics and advanced physical chemistry have been successfully using sophisticated spectroscopic techniques such as NMR, IR, and ESR. I had been using infrared techniques to study biological problems for a number of years and was impressed by the availability of many excellent books on infrared techniques that were written qualitatively for organic and biological chemists. I felt that such a book on Raman spectroscopy was needed. Even though this book emphasizes biological application, a brief description of the fundamental theory of Raman spectroscopy and Raman spectrometers is included. These are described in Chapters I and 2, which are expansions of my Colorado State University lectures in Physical Biochemistry and Biochemical Instrumentation. Since the application of Raman spectroscopy to biological systems is recent, many results remain highly controversial, even among Raman spectroscopists. Instead of being a judge, I took a neutral stand, presenting both views. I feel that we need more time to assess such publications, and through the process of natural selection, papers of poor quality will eventually be left to obscurity. Most biologists are still unfamiliar with how Raman spectroscopy can be .
...
vIII
Preface
applied to biological systems, so many examples of these applications are cited in this book. For the scientist, information from original papers is always important; therefore, many references are cited. Some highly mathematical papers that are not too relevant to the practical application of Raman spectroscopy to biological systems are omitted. I am grateful to the following colleagues who read parts of the manuscript: Drs. P. Azari, G. S. Bailey, R. Bansil, R. Callender, B. Gaber, V. J. Hruby, M. Lutz, C. R. Middaugh, F. P. Milanovich, H. Mitteldorf, M. D. Morris, M. Nicholas, M. Pezolet, R. L. Redington, I. Salmeen, J. A. Shelnutt, D. M. Thomas, E. W. Thomas, S. J. Webb, and S. S. York. I also give my thanks to those who supplied me with valuable photographs and figures, without which the book could not have been published. They are Drs. J. Abraham, M. Delhaye, E. S. Etz, W. Kiefer, P. S. Narayanan, J. Oswalt (Jovin-Yvon Instruments), S. Ramaseshan, J. A. Shelnutt, M. Yamada, and Spex Industries. Many comments provided by Drs. W. Caughey and R. Woody were also very helpful. I also thank my laboratory personnel, Drs. R. A. Hendon and H. Ishizaki, Mr. K. Liddle, and Ms. D. Stedman, for their help. Professional typing by Ms. L. Woeber is also appreciated. Thanks are extended to the three referees who reviewed the entire manuscript and provided useful critiques and suggestions. Finally, I appreciate the guidance of Dr. Stanley Kudzin, without whom this book may not have been written. ANTHONY Fort Collins, Colorado July 1982
T. Tu
Contents
PART I. CHAPTER 1.
GENERAL BACKGROUND BASIC CONCEPT AND ELEMENTARY THEORY
3
Absorption and Scattering of Light 3 1.1. AbsorptionSpectroscopy 4 1.2. Scattering 6 2. Raman Scattering 7 2.1. Stokes and Anti-Stokes Lines 8 2.2. Independence of the Incident Light 10 2.3. Intensity of Stokes and Anti-Stokes Lines 10 3. Classical Mechanics of Raman Scattering 12 4. Raman Spectroscopy and IR Absorption 15 5. Resonance Raman Spectroscopy 16 6. Basic Background for Vibrational Spectroscopy 19 6.1. Vibration 19 6.2. Dipole Moment 20 7. Vibrational Modes of Simple Molecules 20 7.1. Diatomic Molecules 21 7.2. Linear Triatomic Molecules 21 7.3. Nonlinear Triatomic Molecules 25 7.4. Vibrational Modes of Methylene Group 27 7.5. Vibrational Modes of Methyl Group 31 7.6. Cyclic Compounds 33 8. Depolarization Ratio 33 9. Group Frequency 36 I.
Ix
X
Contents
10.
Use of Isotopes 38 10.1. Isotopic Substitution 38 10.2. Natural Isotopes 40 11. Problem of Fluorescence 40 12. General References 41 References 43 CHAPTER 2. THE LASER RAMAN SPECTROMETER AND SPECIAL TECHNIQUES
44
1.
Laser 45 1.1. Basic Theory 45 1.2. Laser Tubes 47 2. Basic Units of Raman Instruments 48 3. Special Techniques 53 3.1. Raman Difference Spectroscopy (RDS) 53 3.2. Coherent Anti-Stokes Raman Spectroscopy (CARS) 54 3.3. Raman Optical Activity or Raman Circular Intensity Differential 58 3.4. Others 59 References 59
PART II. CHAPTER 3.
1.
2.
3.
BIOLOGICAL MOLECULES 65
PROTEINS
Peptide-Bond Vibrations 66 1.1. Amide A and B Bands 67 1.2. Amide I, II, and III Bands 68 1.3. Amide V, VI, and VII Bands 71 1.4. Amide IV Band 71 Secondary Structure (Peptide-Backbone Structure) 72 2.1. Conformational Analysis from Amide I and III Bands 2.2. Comparison of Amide I and III Bands 78 2.3. Other Structurally Sensitive Lines 80 2.4. The 0- and L-Amino Acid Copolymers 81 2.5. Right and Left-Handed a-Helices 82 2.6. Degree of Polymerization 82 2.7. Solid and Aqueous Phases 83 2.8. Effect of Protein on Water 84 2.9. Quantitative Estimation of Secondary Structure 84 Side Chains 86 3.1. Tyrosine 87 3.2. Tryptophan 89 3.3. Phenylalanine 90 3.4. Histidine 91
73
Contents
3.5. Disulfide Bond 91 3.6. C-S 94 3.7. Sulfhydryl Group (-SH) 96 4. Preresonance Raman 97 5. IR and Raman 97 6. Low-Frequency Vibrations 99 6.1. Internal Vibrations 100 6.2. Intermolecular Vibrations 101 7. Raman-Spectra Background 103 8. Application 103 8. I. Denaturation 103 8.2. Chemical Modification 106 8.3. Comparison of Related Proteins 8.4. Glycoproteins 108 8.5. Blood Coagulation 108 References 109 CHAPTER 4.
xl
107
ENZYMES AND IMMUNOGLOBULINS
117
Enzyme Action 117 I. I. Papain 117 1.2. Chymotrypsin 120 1.3. Carboxypeptidase A 121 1.4. Peroxidase 123 1.5. Thymidylate Synthetase 123 1.6. Others 124 2. Enzyme-Inhibitor Complexes 125 2.1. Chymotrypsin 125 2.2. Carbonic Anhydrase 125 2.3. Trypsin 127 2.4. Other Enzymes 127 2.5. Enzyme-Drug Interaction 128 3. Isozymes 128 4. Immunology 129 4.1. Antigen-Antibody Reactions 129 4.2. Hapten-Antibody Interactions 129 4.3. Cryoglobulin 130 References 131 I.
CHAPTER 5. I.
2.
NUCLEIC ACIDS
Background 134 1.1. Tautomerism 135 1.2. Hydrogen-Deuterium Exchange 1.3. Origin of Base Vibrations 135 Principal Raman Lines 137 2.1. Aqueous Solution 138 2.2. In D 2 0 141
134
135
xli
Contents
3.
Structurally Sensitive Lines 141 3.1. Heat Treatment 141 3.2. pH Treatment 141 3.3. Phosphodiester-Bond Vibrations 141 4. Conformational Analysis 145 4.1. Double-Stranded Polynucleotides, DNA, and RNA 4.2. Use of Phosphodiester Bands 145 4.3. Melting of Nucleic Acids 147 4.4. Base Stacking and Hydrogen Bonding 150 4.5. Single-Stranded Polynucleotides 151 4.6. Gel Formation 152 5. Native RNA 153 5.1. Transfer RNA 153 5.2. Ribosomal RNA 155 6. Reactions of Nucleic Acids 156 6.1. Alkylation 156 6.2. Polypeptides 157 6.3. Metal Ions 158 6.4. Drugs 163 Special Techniques 165 7.1. Preresonance and Resonance Raman Spectroscopies 7.2. Others 167 References 167
145
7.
CHAPTER 6.
NUCLEOPROTEINS-VIRUS AND CHROMOSOME
1. Viruses 174 1.1. Plant Viruses 175 1.2. Bacteriophages 180 2. Chromosome 182 2.1. Histones 182 2.2. DNA-Histone Interactions in Nucleosomes References 185 CHAPTER 7.
1. 2.
3.
165
174
183
LIPIDS AND BIOLOGICAL MEMBRANES
Brief Review of Lipids and Membranes 187 Vibrations of Fatty Acids and Phospholipids 190 2.1. Low-Frequency Vibrations 191 2.2. Structurally Sensitive Raman Bands 191 2.3. Unsaturated Fatty Acids 199 2.4. Phase Transition (Melting Behavior) 201 2.5. Quantitative Estimation of Conformation 206 Interaction of Lipids 207 3.1. Lipid-Lipid Interactions 207 3.2. Lipid-Protein Interactions 209 3.3. Lipid-Ion Interactions 213
187
Contents
xIII
Lipid-Antibiotic Interactions 213 Interactions with Other Compounds 218 4. Biological Membranes 219 4.1. Erythrocyte Membranes 219 4.2. Sarcoplasmic Reticulum Membranes 223 4.3. Streptococcus faecalis Membrane 224 4.4. Nerves 224 4.5. Other Plasma Membranes 225 References 226 3.4. 3.5.
CHAPTER 8.
CARBOHYDRATES
234
Assignment of Raman Bands 235 Glycosidic Linkage 235 2.1. IR Absorption Spectroscopy 235 2.2. Raman Spectroscopy 241 3. Functional Groups 242 3.1. 0- H Vibrations 242 3.2. C- H Vibrations 244 3.3. N-H Vibrations 245 3.4. Carboxyl and Acetyl Groups 245 3.5. Sulfates 245 4. Comparison of Solid and Aqueous Phases 248 5. Conformation 249 6. Lipid-Carbohydrate Interactions 251 7. Other Studies 252 References 253 1.
2.
CHAPTER 9.
CAROTENOIDS AND FLAVINS
256
I.
Carotenoids 256 1.1. Carotenoids from Plants, Microorganisms, and Blood 257 1.2. Lobster-Shell Carotenoproteins 261 1.3. Excitation Profiles 262 2. Flavins 262 2.1. Flavins and flavoenzymes 262 2.2. Flayocytochrome cm 265 2.3. Proflavin and Other Flavin Compounds 266 References 266 CHAPTER 10. VISUAL PIGMENTS AND BACTERIAL RHODOPSIN
1. Visual Pigments 270 1.1. Vibrational Assignments 272 1.2. Linkage Between Protein and Prosthetic Group 277 1.3. Experimental Techniques 279 1.4. Conversion of cis to trans 280
270
xiv
Contents
1.5. Opsin 282 1.6. Summary 283 2. Bacteriorhodopsin 283 2.1. Vibrational Assignments 285 2.2. Linkage Between Protein and Prosthetic Group References 293 CHAPTER 11.
NONHEME IRON COMPOUNDS
288
298
Hemerythrin 299 1.1. Bound Oxygen 299 1.2. Other Study 300 2. Transferrin and Uteroferrin 302 2.1. Transferrin 302 2.2. Uteroferrin 303 3. Oxygenases 303 3.1. Protocatechuate 3, 4-Dioxygenase 303 3.2. Pyrocatechase 305 4. Ferredoxin and Related Compounds 306 4.1. Ferredoxin 308 4.2. Rubredoxin 308 4.3. Adrenodoxin 309 5. Others 312 References 313 I.
CHAPTER 12.
HEMES AND PORPHYRINS
Vibrational Modes 317 1.1. Exci tation Profiles 318 1.2. Origins of Vibrations 319 I.3 Effect of Metals 321 2. Side Chains 322 2.1. Vinyl Groups 322 2.2. Formyl Group 324 2.3. Identification of Isomers 326 3. Spin State 326 3.1. Spin 327 3.2. Electrons in the Coordinated Iron 327 3.3. Spin States of Heme Compounds 330 3.4. Spin State and Raman Bands 331 4. Effect of Ligands 332 4.1. Ligand-Sensitive Raman Bands 333 4.2. Carbon Monoxide Ligand 334 4.3. Oxygenation 335 4.4. Fe-Axial-Ligand Vibrations 337 4.5. Pentacoordination and Hexacoordination 342 5. Oxidation States 343 5.1. Cytochrome c 343 I.
316
Contents
5.2. Cytochrome Oxidase (Cytochrome c Oxidase) 5.3. Others 346 6. Polarization 346 6.1. Inverse Depolarization 347 6.2. Heme Compounds 347 6.3. Metalloporphyrins 348 7. Quaternary Structure 349 8. Protein Environment 352 9. Geometry of the Heme Ring 352 10. Mitochondria 354 10.1. Intact Mitochondria 355 10.2. Cytochromes b and c 355 10.3. Cytochrome Oxidase 356 II. Other Studies 356 11.1. Effect of Redox Potential 356 11.2. Temperature 357 References 357 CHAPTER 13. SYSTEMS
XV
345
COPPER AND OTHER METALS IN BIOLOGICAL 369
Copper in Biological Systems 369 1.1. Hemocyanin 370 1.2. Other Blue Copper Proteins 372 1.3. Copper-Peptide Complex 375 2. Other Metals 376 References 377 1.
CHAPTER 14.
PHOTOSYNTHETIC PIGMENTS AND VITAMIN B12
381
Photosynthetic Pigments 382 1.1. Raman-Band Assignment 382 1.2. Plant Photosynthetic System 384 1.3. Bacterial Photosynthetic System 385 1.4. Carotenoids in Photosynthesis 386 2. Vitamin B I2 387 References 389 I.
PART III SPECIAL TISSUES AND THEIR COMPONENTS CHAPTER 15.
1.
EYES, TEETH, AND MUSCLES
Ocular Lenses 395 1. I. Peptide Backbone 397 1.2. Sulfhydryl Group and Disulfide Bond 1.3. Others 398
395
397
xvi
Contents
2. Cornea 398 3. Teeth 399 4. Muscle 401 4.1. Myosin 402 4.2. Myosin Subfragments 403 4.3. Tropomyosin and Troponin 404 4.4. Whole Muscle 404 4.5. Contractile State and Protein Structure 405 References 405
PART IV
ANALYTICAL TOOLS
CHAPTER 16.
RAMAN MICROPROBE, MOLE
411
I. Theory 412 2. Clinical and Histological Applications 414 3. Environmental Applications 419 4. Other Applications 420 References 420
CHAPTER 17.
CLINICAL AND ENVIRONMENTAL APPLICATIONS
422
I. Clinical Diagnosis: Tissues and Cells 422 2. Clinical Analysis 424 3. Environmental Problems 425 3.1. Pesticides 426 3.2. Food Additives 426 4. Others 427 References 427 APPENDIX. INDEX
ABOUT PROFESSOR C. V. RAMAN
430
437
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CHAPTER
Basic Concept and Elementary Theory
The purpose of this book is to describe how Raman spectroscopy can be applied to biological systems. The researcher should know some basic theory and general background on the Raman scattering effect, vibrational modes, lasers, and the basic units of the instrument before applying this technique to examine biological systems. This chapter gives this background; however, it is written in a qualitative fashion without an in-depth analysis. Readers who would like to know more about the concept and theory of Raman spectroscopy will find that there is no shortage of books on these topics. Some of these books are listed at the end of this chapter. The laser and basic components of the Raman spectrometer are described in Chapter 2.
1.
ABSORPTION AND SCATTERING OF LIGHT
Light is an electromagnetic wave composed of an electric vector E and a magnetic vector H. These vectors are perpendicular to each other· and to the 3
4
Basic Concept and Elementary Theory
I. Amplitude
>.. Wavelength
Electric Field
.1
Magnetic Field
iC:{j/?\ '----_..- Direction of light propogotion
I·
·1
Wavelenath
>..
FIGURE 1.1. Two components of light (electromagnetic wave). The vectors of the electric and magnetic components are perpendicular to each other and to the direction of light propagation.
direction of propagation (Figure 1.1). When the electric field of this wave interacts with matter, light can be scattered or absorbed. 1.1.
ABSORPTION SPECTROSCOPY
When incident light interacts with a compound, some light may be absorbed. Spectroscopy utilizing this property is called absorption spectroscopy. Readers are probably familiar with ultraviolet (UV) absorption spectroscopy, visible absorption spectroscopy, and infrared (IR) absorption spectroscopy, as these techniques are commonly used in modern biological laboratories. These are indeed part of absorption spectroscopy. There are many different kinds of electromagnetic waves, such as y rays, X rays, UV light, visible light (white light), IR light, microwaves, and radiowaves. The difference lies in their wavelength >.. One can also say the difference is due to the frequency P. Both expressions are correct because wavelength and frequency are inversely proportional:
c
}; =
P
(1.1)
where C is the velocity of the electromagnetic wave. Ultraviolet light has a shorter wavelength than IR light; UV light has a higher frequency then IR. In spectroscopy the term wave number ii is frequently used. The wave number is defined as the number of waves contained in a I-em length and can be expressed as ii = 1/>.. Wave number, wavelength, and frequency have the following relationship: I P ii=X= C
Different types of electromagnetic waves are diagrammed in Figure 1.2.
(1.2)
5
Absorption and Scattering of Light Wavelength in em
JOO
Hl
10
5
10
8
Radio waves
FIGURE 1.2. Different types of electromagnetic waves and their wavelengths. Note that visible light is only a small portion of the overall electromagnetic spectrum.
All electromagnetic waves contain inherent energy, which is expressed by Planck's famous equation:
(1.3)
E = hv
where h is Planck's constant. Since UV light has a higher frequency (or shorter wavelength) than IR light, the former contains more energy than the latter. Ultraviolet and visible absorption spectroscopies involve electronic transitions of a molecule. In other words, a molecule receives the energy of the incident light (UV or visible light) and is excited to a higher electronic level (Figure 1.3). Infrared light has low energy. In IR absorption a molecule absorbs the incident light (lR light) and is excited to a higher vibrational or rotational quantum level, but not to a higher electronic quantum level (Figure 1.4). Frequently IR absorption is expressed by percentage transmission T (the ratio of the intensities of the transmitted I and incident light 10 ) or as absorbance A, which is defined as A = 10g(Io/1).
Higher Electronic {. Level .
CD
o c: .c (; U> .c
vvv--
"
«
f-- -
Electronic Ground Level (Lower Electronic Level)
Wavelength ( Ultraviolet. Visible Region)
FIGURE 1.3. The diagram at the left shows the absorption of light by a molecule, raising it to a higher energy level. This results in the absorbtion spectrum diagramed at the right.
6
Basic Concept and Elementary Theory
J1
Higher Electronic Level
Wavelength (A) (Infrared Light Region)
J
/\ /\ ~--------~:~l V
V-
t
V=O
100,"",
>
.:;l5 Lower
Electronic Level
°1
.=
;f. O,~I
------
A
FIGURE 1.4. The IR absorption spectrum is due to the elevation of vibrational energy of a molecule (left). It does not involve electronic energy transition. IR absorption is customarily expressed either by absorption (upper right) or by transmission (lower right).
1.2.
SCATTERING
Scattering refers to light deflected from the direction of incident-light propagation. The interaction of the electric vector of an electromagnetic wave with the electrons of a compound results in the scattering of the incident light. Such interactions induce periodic vibrations in the electrons of a compound, thereby producing oscillating electric moments. Such oscillating electrons become new sources for emitting radiation, that is, the scattered light. There are three basic types of scattering: 1.
Elastic. Same frequency (wavelength) as the incident light-Rayleigh scattering. 2. Inelastic. Lower frequency (longer wavelength) than that of incident light-Stokes Raman scattering. 3. Inelastic. Higher frequency (shorter wavelength) than that of incident light-anti-Stokes Raman scattering.
In other words, there are two types of Raman scattering. In one type the scattered light has lower energy than the incident light, hence it has lower frequency, and the effect is called Stokes Raman scattering. In the other, the scattered light has higher energy than the incident light, hence it has higher frequency than that of the incident light (anti-Stokes effect of Raman scattering). Rayleigh scattering (same frequency) does not involve a change in the energy content of the incident and scattered lights (Figure 1.5). Rayleigh scattering is most familiar to us. We are able to see objects as a result of light scattering. Experiments show that scattering efficiency is in-
Raman Scattering
,/"~
Incidenl Light
,, ",
"
7
A""-S"'" R,m'" S",,,,,".
R"",." S""",",
A Scattering Cenler J ' ~ -llV
Slokes Raman Scattering
FIGURE 1.5. Diagrammatic presentation of the three types of light scattering: anti-Stokes Raman scattering, Rayleigh scattering and Stokes Raman scattering.
versely proportional to the fourth power of wavelength. Sunlight includes light of many wavelengths. But because blue light has a shorter wavelength than red light, it is scattered more than red light. When we see the blue sky, we are seeing the scattered-primarily blue-portion of sunlight. At sunset and sunrise, sunlight passes through a greater thickness of atmosphere than when the sun shines from above. Since most of the blue short wavelength light has been scattered by molecules in the air, most of the remaining red to orange light passes through. This is the reason why sunset and sunrise appear red to our eyes. Compared with Rayleigh scattering, Raman scattering is less common in our daily lives; nevertheless, it is important for scientists who are interested in vibrational and rotational states of molecules. The Raman process involves two photons with different energies, as the incident photon and scattered photon differ in energy. This energy difference is due to a change in the vibrational or rotational state of a molecule caused by interaction with incident photons. For this reason, analysis of the Raman spectra provide information about molecular properties such as the manner and type of vibrations. The intensity of scattered light is influenced by many factors: 1. The size of the particle or molecule illuminated. 2. The location of observation. The scattering intensity is a function of the angle with respect to the incident beam. 3. The frequency of the incident light. 4. The intensity of the incident light.
2. RAMAN SCATTERING The Raman scattering effect arises from the interaction of the incident light with the electrons in the illuminated molecule. In nonresonance Raman scattering, the energy of the incident light is not sufficient to excite the molecule to a higher electronic level. Instead, Raman scattering results in changing the molecule from its initial vibrational state to a different vibrational state (Figure 1.6).
8
Basic Concept and Elementary Theory Electronic - - - - - - - - - - - - - - - - - ] Excited State -- --- --
-/"""<;---0-
Virtual State
~---?"""--
h/l
\/\r--.-,-.-,--,-,-,-1 ~I Stakes Line
U
I
] Vibrational Quantum Number Within The Electronic Ground State
Anti-Stokes Line
Raman Scattering FIGURE 1.6. The energy diagram of a molecule showing the onglO of Raman scattering (nonresonance Raman effect). Note the different mechanisms of the Stokes and anti-Stokes effects; yet their energy differences (Raman shifts) are the same. The net effect is either an increase or decrease in the vibrational-energy level of a molecule. The molecule is momentarily elevated to a higher energy level (virtual state) but it never reaches an electronic excited state.
2.1.
STOKES AND ANTI-STOKES LINES
In order for a molecule to exhibit the Raman effect, the incident light must induce a dipole-moment change or a change in molecular polarizability. Consider carbon dioxide as an example. The change in polarizability can be qualitatively visualized as a change in the shape of the electron cloud. As shown in Figure 1. 7, the electron cloud around the CO2 molecule is alternately stretched or shrunk in phase upon interacting with the oscillating wave of the
~
fi
II' ; !) JV\rhll
===>
ilfT'i~~ ::::{
~
~
FIGURE 1.7. An example of change in polarizability. Qualitatively one can visualize it as a change in the electron cloud of a molecule by interaction with light. For a molecule to show a Raman effect, a polarizability change must occur in the molecule during a vibrational normal mode.
Raman Scattering
9
electric component of the electromagnetic wave. An electron with certain vibrational modes can couple with the incident-light photon and lead to scattered photons with altered frequency. The scattered light contains a small portion of light due to Raman scattering in addition to that due to normal Rayleigh scattering (same frequency as the incident light). The Raman scattering contains both a Stokes line and an anti-Stokes line; their frequencies correspond to the sum and the difference of frequency of the incident light and the allowed molecular vibrational frequencies. When photons interact with a molecule, some of their energy can be converted into various modes of vibrations of the molecule. As seen in Figure 1.6, the scattered light loses energy equivalent to the energy given to molecular vibrations (Stokes Raman effect). If the energy is transferred to the scattering light from a molecule, then the scattered light has more energy than the original incident light (anti-Stokes Raman effect). This requires that the scattering molecule already be in an excited state. For typical applications this is seldom the case; therefore, Stokes Raman scattering is much stronger than anti-Stokes scattering. Only a small fraction of photons are scattered by Raman scattering, so Raman lines are usually very weak (only 10- 6 of the intensity of the Rayleigh line). The majority of the scattered light is the same as the original incident light in terms of photon energy. This is why a laser is used as a light source: it is an intense, monochromatic source of light and produces more scattered photons. Because both the Stokes line and anti-Stokes line involve the same vibrational-energy difference (Figure 1.6), the difference between the incident-light frequency and the scattered frequency in Stokes and anti-Stokes scattering is identical. That is, the difference in frequency for Stokes and anti-Stokes scattering is symmetrical.
11 ~
500
°
19,992
20,492
T
6Cm' Cm'
•
I
500 20,992
FIGURE 1.8. In Raman spectroscopy it is important to find the Raman frequency difference, ~cm - I, rather than the absolute wave number (or frequency) of scattered light (cm -I). One notices that the Raman frequency difference of Stokes and anti-Stokes lines is always symmetrical.
10
Basic Concept and Elementary Theory
For instance, a compound illuminated with the blue line of an argon ion laser at 488 om (20,492 cm -I wave number) produces two Raman lines at 19,992 cm -I and 20,992 cm -I. Although one cannot see the significant correlation between these lines merely by inspecting the two values, one can readily see an interesting result by taking the frequency difference between the scattered and the incident light (Figure 1.8). Normally, Raman scattering is expressed by this wave number difference (~ wavenumber) rather than the absolute wave number of Raman-scattered light. A modern Raman spectrometer automatically gives the wave number shifts, which is what contains the molecular information. 2.2.
INDEPENDENCE OF THE INCIDENT LIGHT
The frequency of the Raman-scattered light is independent of the wavelength of the incident light. One should not confuse this with the fact that the intensity of Raman scattering depends on the wavelength of the incident light and, actually, is inversely proportional to '11.4 . Thus whether one excites a molecule with green light (514.5 nm or 19,436 cm- I ) or blue light (488.0 nm or 20,492 cm -I), one will obtain a Raman ~ line at exactly the same wave number difference. This can be readily understood from Figure 1.9. 2.3.
INTENSITY OF STOKES AND ANTI-STOKES LINES
From Figure 1.5, one sees that both Stokes and anti-Stokes lines have identical frequency values. However, the Stokes line has a higher intensity than the anti-Stokes line at room temperature. The Stokes line originates when a
..---.,
-f---- "" .....- -Energy of
G,ee~r~~
·01
1
Energy of Blue Light
]![
FIGURE 1.9. In Raman scattering, the frequency of the Raman-scattered light is independent of the excitation wavelength. One should not confuse this with the fact that the intensity of Raman scattering depends on the excitation wavelength.
....
Raman Scattering
11
molecule at a low vibrational energy (see Section 6) is elevated to higher energy hv, (b..v in Figure 1.10) by interacting with the incident light, whose energy is equal to hvo' On the other hand, the molecule at a higher vibrational-energy level gives away the energy hv,; therefore, the molecule becomes lower in vibrational energy, and the scattered light increases in energy by hv,. At low temperatures (or at room temperature), more of the molecules are at lower vibrational-energy levels than at higher vibrational-energy levels (Figure 1.10). Thus a larger fraction of molecules will have Stokes-type transitions than anti-Stokes transitions, and the Stokes line will have higher intensity than the anti-Stokes line. This can also be seen by examining the Boltzman distribution law, which states that the relative population of molecules with higher energy
.:
,-- r---- - --,--, --
-r--~------'----I
I
,
I1~ /\.
. V
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I
I
I
I I
I I
I :
--t,
:
; .0. V
V-Z /\. V-I V
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more
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level
:
':
:
than
V-Z V-I
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Anti-Stokes Effect
that
of
1\
Higher
temperature - The
molecules
at
energy
of V= I.
ttensit y
)
higher
I
I
.-..-1- -
temperature - The population
level V=O is
~
I
.
Stokes Effect ~
II (~~.o.II)
.0.11'
.......-.L--L------V·O
~
I
I
11(16-611)
-..-:- -
:
I
,
/
population
of
I
Lower
~ntensity
molecules
at
energy
V= I increases relative to that of energy V=O.
1\
~
"Lower ~ntensity
JA
Higher
~ntensity
FIGURE 1.10. The origin of Stokes and anti-Stokes effects and their intensity differences. The relative intensity of Stokes and anti-Stokes lines varies depending on temperature.
12
Basic Concept and Elementary Theory
increases as the temperature is increased. N =e -AE/kT -' No
where N; is the number of molecules at energy state Ei and No is the number of molecules at energy state Eo; NJNo is the fraction of molecules at the energy state E i ; k is the Boltzman constant; T is temperature in the Kelvin scale; and tiE is the energy difference between energy states Ei and Eo. Therefore the anti-Stokes-line intensity increases (or the Stokes-line intensity decreases) as the temperature is elevated. The ratio of the anti-Stokes to the Stokes line is directly related to the fraction of molecules at higher vibrational-energy levels. This intensity ratio may have interesting physical-chemical implications, but it is generally not used by biochemists and biologists. 3.
CLASSICAL MECHANICS OF RAMAN SCATTERING
So far Stokes and anti-Stokes Raman scattering have been explained qualitatively. The scattering of light by a molecule can be expressed easily in terms of classical mechanics. Since the mathematics involved is relatively easy and all three effects (Rayleigh scattering, Stokes, and anti-Stokes Raman scattering) can be explained by one equation, the classical-mechanical treatment of scattering is presented here. If a molecule interacts with light, the electric field of photons will exert oppositely directed forces on the electrons and the nuclei. As a result, the electrons will be displaced relative to the nuclei, and the polarized molecule will have an induced dipole moment caused by the external field. The induced dipole moment P (see Section 6 for the definition of dipole moment) is proportional to the electric field and to a property of the molecule called the polarizability a P= aE
(3.1)
The electric field is an oscillating function dependent upon the frequency of the light Po E = Eocos 27TPot
(3.2)
Substituting this into Eq. (3.1), we obtain P = aEocos 27TPot
(3.3)
The polarizability a is dependent upon the position of the nuclei in the molecule. For a molecule containing N atoms, there are 3N degrees of freedom
......
Classical Mechanics of Raman Scattering
13
available to the nuclei. Of these, 3N - 6 (3N - 5 for a linear molecule) result in vibrations of the molecule. In general, the vibrational motion of all but the simplest molecules (e.g., diatomic) is quite complicated (see Section 7). However, by using group theory it is possible to reduce this complicated vibrational motion to a set of independent normal modes of vibration. Instantaneous positions of the nuclei can therefore be expressed relative to their equilibrium positions in terms of the normal coordinates Qi' where i == 1,2, ... , 3N - 6. Considering a diatomic molecule with the single normal coordinate Ql' the dependency of a on Q) is expressed as a series expansion
==
a
ao +
( aa~1
L
QI + ...
(3.4)
where a o is the equilibrium value of the polarizability. The position of the nuclei is time dependent because the molecule is vibrating with frequency VI' Information on the frequency of vibration can be obtained from knowledge of the forces between the vibrating nuclei, and the application of the classical mechanics of small vibrations. This motion can be expressed as
QI ==
(3.5)
Q?COS27TV)t
where Q? is the maximum vibrational amplitude. It is seen, therefore, that a also oscillates at the frequency VI' Substituting Eqs. (3.4) and (3.5) into Eq. (3.1), we have
p == [ ao +
( aa~1 )0QI] Eocos 27Tvot
== [a o + (
aa~1 L(Q?COS27TV1t)] Eo cos 27Tvot
From cos (J X cos
== t[cos( (J
+ <1» + cos( (J -
P == aoEocos 27Tvot
<1»],
+ II EoQIo(~) aQI 0
X [cos 27Tt(VO + VI)
+ cos 27Tt(VO - VI)]
(3.6)
This classical derivation for a diatomic molecule predicts three basic lightscattering modes due to the induced dipole moment P oscillating at frequency VI:
1.
The a o term produces scattered light unshifted in frequency (Rayleigh scattering).
14
Basic Concept and Elementary Theory
2.
If aa/aQ =1= 0, Raman scattering occurs, and incident light of frequency Po is shifted to scattered light with higher frequency Po + PI (anti-Stokes) and lower frequency Po - PI (Stokes).
For a polyatornic molecule containing N atoms, there are 3N - 6 vibrational modes; therefore, Eq. (3.6) becomes 1 3N-6 ( P=Eoaocos27TPot+2" .~ EoQ?
aa ) aQ
1=1
X [cos 27T(PO + pJt
1
+ cos 27T(PO -
0
pi)t]
(3.7)
The first term refers to Rayleigh scattering (elastic), the second term to anti-Stokes lines, and the third term to Stokes lines. The fundamental difference between Raman and infrared spectroscopy is that aa/aQ does not equal zero for Raman scattering. This means that a molecule must have a change in its polarizability as it vibrates in order to be Raman active. Equation (3.1) is valid for molecules with isotropic polarizability and states that the dipole-moment vector P induced by the electric-field vector E is parallel to the electric-field vector. This is not true in general, as the polarizability of most molecules is anisotropic, and the induced-dipole-moment vector points in a different direction than the electric-field vector. A vector is a quantity that has a magnitude as well as a direction. The polarizability should therefore properly be expressed as the symmetric matrix axx a = a yx ( a zx
a xy ayy a zy
xz aa yz )
(3.8)
a zz
rather than as a constant. Using this matrix expression for a, the components of P become Px = axxEx + axyEy + axzEz Py
= ayxEx + ayyEy + ayzEz
Pz = azxEx + azyEy + azzEz
(3.9)
Thus the induced dipole moment should be expressed as
P= aE
(3.10)
or
P=
axx a yx (a zx
a xy a yy a zy
xz aa yz ) E a zz
(3.11)
~
15
Raman Spectroscopy and IR Absorption
4.
RAMAN SPECTROSCOPY AND IR ABSORPTION
Raman spectroscopy and IR absorption spectroscopy are complementary. If one wants to know the complete set of vibrational modes of a molecule, one needs to examine both spectra. There are some similarities between them, but there is also a fundamental difference. Look at the vibrational-energy diagram of Figure 1.11. The energy gained by a molecule is identical whether it is obtained through the Raman effect (IlERaman) or through IR absorption (IlE 1R ). This means that the Raman frequency (Ilv) and the IR frequency are the same. The IR spectrum and Raman spectrum of toluene are shown in Figure 1.12. Some spectral lines appear at exactly identical wave numbers, although their intensities may not be identical. However, there is a basic difference in the principles involved in the two spectroscopies. The selection rules for Raman and IR spectroscopy are not identical. A molecule absorbs IR light only when the dipole moment changes during the molecular vibration. The Raman effect is caused by an oscillating induced dipole moment, which means that the molecular interaction with light is through the polarizability of the molecule. The electromagnetic radiation must change the molecular vibration and the shape of the electron cloud of the molecule. Therefore, sometimes a molecule shows an IR line for a particular vibration, but not a Raman line, or vice versa. The difference depends on molecular symmetry. If a molecule has a center of symmetry, for example, the benzene molecule, the Raman and IR spectra will be mutually exclusive with no overlap. This is because the molecule interacts with light differently in the two processes; one is a scattering process and the other is an absorption process. Raman and IR are both vibrational spectroscopies, yet there are certain advantages with Raman spectroscopy. The Raman bands are usually sharper than IR bands for biological macromolecules. Raman spectra can be obtained for compounds in crystal, powder, aqueous solution, and gel forms. Raman frequencies arise from changes in the electronic polarizability associated with nuclear vibrational displacements. By analyzing the Raman intensity changes and frequency shifts, one can examine the molecular properties of a compound. As in UV and visible absorption spectroscopies, there are both Raman ------r-;------
~~1M.h' L '"\/'v-3
\/V--
Raman
----3 ------2
t
6E jr =hv
Infrared
~=o
FIGURE 1.11. Both Raman scattering (Stokes effect) and IR absorption increase the vibrational-energy level of a molecule, but their mechanisms are totally different. The former is a scattering effect and the latter is an absorption effect.
16
Basic Concept and Elementary Theory
i r Spectrum
2800
2600
2400
2200
2000
1800
1600
1400
1200
1000
BOO
600
em-'
FIGURE 1.12. Comparison of IR (top) and Raman (bottom) spectra of toluene. Some lines appear at the same frequency in both the IR and the Raman spectrum. However, some lines show in the IR but not in the Raman spectrum. The intensities of the IR lines are different from those of the Raman lines, although many of them appear at the same frequency. This renects the difference in selectivity of two fundamentally different processes.
hypochromism and Raman hyperchromism, which can be used to detect molecular order-disorder parameters or the progress of a particular reaction. Unlike IR or any other spectroscopic methods, Raman spectroscopy can measure depolarization ratios (discussed below); these data are useful to determine the symmetry of vibrational centers.
5.
RESONANCE RAMAN SPECTROSCOPY
When a compound is excited with incident light whose frequency is within an electronic absorption band, the intensity of some Raman spectral lines is greatly enhanced. The effect is called resonance Raman scattering, and is due to a coupling of electronic and vibrational transitions. From the quantum theory, Placzek (1934) predicted that the intensity of a Raman line would depend on the exciting frequency and that the intensity would be enhanced as the exciting frequency came close to an electronic absorption frequency. The experimental result of the resonance Raman effect was first reported by a Russian, Professor P. P. Shorygin, in 1953, at a meeting in Gmunden, Austria. He showed the advantage of using an excitation wavelength within the region of electronic absorption (Brandmuller and Kiefer, 1978). In resonance Raman scattering, the energy of the exciting laser beam coincides with that of an electronic transition. When the energy of the laser is close to, but not higher than an electronic excitation level of a molecule, the process is called preresonance Raman scattering. In normal resonance Raman scattering, the energy level of the virtual state falls far below that of the
Resonance Raman Spectroscopy
17
---=-rr=rrl EXCit~~at~lectraniC ---------~------
--
--~ -----.--~-----
--'-'--I~
---
- - } Vibrational Levels Within Ground Electronic Level
-11--'-'-----'1-1--
~LStokes
Anti-Stokes
Stokes
NON-RESONANCE RAMAN EFFECT
_'_~_
Anti-Stokes
RESONANCE RAMAN EFFECT
FIGURE 1.13. Diagrams of the difference between nonresonance and resonance Raman effects. In the resonance Raman effect, the energy of the excitation wavelength reaches the energy level of the excited electronic state.
electronic transition (Figure 1.13). As a photon interacts with a molecule, the energy level of a molecule rises momentarily to a higher energy level. This unstable energy level is called the virtual state. The components a qr of the polarizability tensor can be calculated by a semiclassical treatment of the interaction of the molecule and the incident-radiation field. The reason for the enhancement can be seen in a theoretical quantum-mechanical expression for the Raman-scattering tensor elements a qr , which are proportional to the scattered-light intensity. This calculation yields the following expression: (a ) qr mn
=! ~ [(Mr)me(M:)en + (Mq)me(M~)en 1 h e
Pe - Po
+ Ire
Pe
+ Ps + Ire
(5.1)
where q and r refer to the coordinate directions x, y, or z, Po is the frequency of incident radiation, Pe is the frequency of the transition between the ground and the excited electronic state, Ps is the scattered (Raman) frequency, and ire is a damping term related to the width of the excited state Pe (the bandwidth of the electronic transition). The initial and final states of a molecule are represented by m and n, respectively, and the (Mr)me and (Mq)en are electric dipole transition moments along the directions of rand q from the initial state of m to an excited state e and from e to the final state n. The h is Planck's constant. The first term of the equation describes a transition involving an absorbed photon and subsequently an emitted photon, whereas the second term describes the emission of a scattered photon followed by absorption of a photon from the laser light field. The first term is the resonant term. The denominator
18
Basic Concept and Elementary Theory
of this term (Pe - Po + ife) becomes very small, so that the polarizability element becomes very large, when the incident laser frequency Po is chosen near the excited electronic transition Pe • This explains the resonance Raman effect, which can effectively increase the Raman cross section by many orders of magnitude. In resonance Raman spectroscopy, the lines due to the vibrational modes of the chromophore or adjacent groups of atoms are selectively enhanced, and the number of resonance Raman lines is less than the number of lines in the nonresonance Raman spectrum (Strommen and Nakamoto, 1977). In this way detailed selective information about the chromophore can be obtained. Obtaining more-detailed information by an increase in sensitivity using resonance Raman spectroscopy outweighs the lost information. In other words, in most cases, the gain in selectivity surpasses the loss of broad information. Furthermore, the significant increase in intensity allows one to work at a much lower sample concentration. The exact theory of resonance Raman spectroscopy is complex, but it can be visualized qualitatively from Figure 1.13. Resonance Raman spectroscopy is tremendously useful in the study of structure-function relationships of biological compounds. Normally the site of biological activity is close to the positions of biological chromophores. For example, hemoglobin contains a chromophore, a heme group; an oxygen molecule attaches to the central iron atom of the heme group. Because resonance Raman spectroscopy requires an excitation frequency near that of an absorption band, sometimes this becomes the limiting factor. Resonance Raman spectroscopy requires laser light of a particular wavelength for excitation. Sometimes this is achieved by using either a tunable laser, specific lines from different laser tubes, or a frequency doubler. As the excitation wavelength comes close to the absorption maximum, the intensity of the resonance Raman line increases. However, one should keep in mind that sometimes there are no clear dividing lines between nonresonance, preresonance, and resonance Raman lines. The intensity of a Raman band is a function of the frequency of the incident light. It also depends on the shape of the absorption band. If the absorption band is narrow, a much clearer distinction among the three types of Raman scattering can be obtained. Many biological compounds contain chromophore groups that give rise to resonance Raman spectra. In this book, resonance Raman spectra of many compounds are discussed. For instance, carotenoids and flavins (Chapter 9), rhodopsin and bacteriorhodopsin (Chapter 10), various iron- and copper-containing compounds (Chapters 11, 12, 13), and chlorophylls (Chapter 14) are discussed from the viewpoint of resonance Raman spectroscopy. Simple proteins normally give no resonance Raman spectra; however, they become resonance Raman active after conjugation with chromophores. These are discussed in Chapter 4. In this section, resonance Raman spectroscopy has been described qualitatively. Readers who wish to know more about the theoretical aspects of resonance Raman spectroscopy are directed to the articles written by Lewis
....
19
Basic Background for Vibrational Spectroscopy
and Spoonhower (1974), Johnson and Peticolas (1976), Spiro and Stein (1977), and Warshel (1977).
6.
6.1.
BASIC BACKGROUND FOR VIBRATIONAL SPECTROSCOPY VIBRATION
A vibration (oscillation) is a type of motion in which a particle (or atom) changes its position periodically in time. Vibration can best be visualized as two particles connected by a spring. When the spring is stretched and released, the two particles (which represent atomic nuclei) make a stretching vibration (Figure 1.14). The mode of elastic vibration described by Hooke's law depends on the masses of the particles and the force constant of the spring. When two atoms A and B are vibrating, the mode of a vibration is related to the masses or reduced mass }J- of the atoms and the force constant k by v = _1
27T
[k (_1 + _1 ) ] mA
mB
1/2
= _1
27T
[k ( m
A
+ m B )] 1/2 =
mAmB
_1
27T
(~ ) 1/2 }J-
(6.1 )
The potential energy (V) for the harmonic oscillation of two particles is V = ~kX2 where X refers to the displacement of two particles from their equilibrium positions. A molecule has discrete quantum energy levels instead of continuous energy states as in the case of two particles connected by a spring. A quantum-mechanical treatment of the harmonic-oscillator problem gives a set of states of motion of the atoms with energies given by the formula E v = hvo(v +~)
(6.2)
where v is the vibrational quantum number (0, 1, 2 ... ) specifying the energy level, h is Planck's constant, Vo is the frequency of a diatomic molecule, and m A and m B are the masses of the atoms. The levels will be equally spaced with a separation of 1 quantum of vibrational energy hv. The harmonic-oscillator problem is applicable to polyatomic molecules as well, since the vibrations of a larger molecule can be separated into normal modes of vibration, each of which can be treated independently as simple harmonic motion.
~
FIGURE 1.14. Vibration of two atoms can be visualized as the vibration of two balls connected by a spring.
20
Basic Concept and Elementary Theory
Wi
~
FIGURE 1.15. Diagram of different types of O-H vibrations. I, O-H stretching vibration, 3200-3700 em -I; 2, O-H in-plane bending vibration, 1200-1500 em-I; 3, O-H out-of-plane bending vibration, 250-650 em-I.
A normal mode of vibration is a special combination of motions of the nuclei of the molecule that can be treated in an independent fashion. Although the motion of many atoms is involved in each of the normal modes, a simplification of the expressions for many properties of the molecule results. For example, the total vibrational energy of the molecule is then just the sum of the vibrational energy from Eq. (6.2) for each normal mode. The normal coordinate is a set of N vector displacements defining the motion of each of the atoms of the molecule for the normal mode. Vibration is not restricted to bond stretchings; bending vibrations are also common. Take for example the -OR functional group in Figure U5 attached to atom C; it has two bending vibrations.
6.2.
DIPOLE MOMENT
When two charges (positive and negative charge of q) are separated by a distance I the dipole moment P, which is a vector, is defined as
P == Iq The dipole moment consists of two independent quantities, the charge and the distance between the two charges. In the dipole-moment concept, two oppositely charges particles separated by distance I are considered as a single unit. A dipole produces its own electric field, thus influencing other charged particles, and it is also influenced by an external electric field.
7.
VIBRATIONAL MODES OF SIMPLE MOLECULES
The total number of vibrations for a nonlinear molecule is 3N - 6 where N is the number of atoms. Many biological molecules are macromolecules, which contain a large number of atoms. This means that they have a tremendously large number of fundamental vibrations. It is impossible to identify all vibrations for a biological molecule. Nevertheless, the understanding of basic vibrational modes in a relatively simple molecule is helpful. In this section different types of vibrations in diatomic molecules, triatomic molecules, the methyl group, and cyclic compounds are briefly reviewed.
....
Vibrational Modes of Simple Molecules
7.1. 7.1.1.
21
DIATOMIC MOLECULES Identical Atoms(A-A)
Bond stretching is the simplest example of vibration (Figure 1.16). Since there is a change in polarizability (aajaQ), such a vibration should appear in the Raman spectrum (Raman active). In order to have a Raman effect, a molecule must have a change in the induced dipole moment. The induced dipole moment can qualitatively be visualized as a distortion of the charge distribution within the molecule upon interacting with the electric vector of light. Or we may simply say that the shape of the electron cloud of the molecule changes with light. There is no separation of charge in a molecule consisting of identical atoms; thus there is no dipole moment. In such a case, there is also no change in dipole moment when the vibration is excited, so it is IR inactive. Therefore, nitrogen molecule, N2 , has a Raman spectrum but no IR vibrational absorption spectrum. 7.1.2.
Nonidentical Atoms (A-B)
In a diatomic molecule with nonidentical atoms, the charges are not symmetrically arranged; thus the molecule has a permanent electric moment. HCl is used as an example in Figure 1.17. As the hydrogen and chlorine atoms vibrate, there is a change in the polarizability ellipsoid (electron cloud); therefore, there is a change in the polarizability derivative, aajaQ. Likewise, there is a change in the dipole moment derivative, ap jaQ. From the example of HCl, it is clear that the HCl stretching vibration will show up in both Raman and IR spectra. 7.2.
LINEAR TRIATOMIC MOLECULES
For a linear molecule with N atoms, the total number of degrees of freedom is 3N, as each atom requires three coordinates to specify its position. Of 3N total degrees of freedom, two are for rotational motion of the molecule, and three degrees of freedom are for translations. Thus the maximum number of vibrations (maximum degrees of freedom for vibration) is 3N - 5. Carbon dioxide, CO2 , is a good example of a linear triatomic molecule. In CO2 the charges are symmetrically arranged, so this is a nonpolar molecule with no permanent electric dipole moment. However, there is an induced dipole moment when an electric-field vector of light interacts with a molecule, causing a shift in charges. The CO2 molecule has symmetrical-stretching, asymmetrical-stretching, and bending vibrations. As can be seen from Figure 1.18, a symmetrical-stretching vibration of CO2 is Raman active but IR inactive. For the asymmetrical-stretching vibration, CO2 is Raman inactive but IR active (Figure 1.19).
Vi bration Mode
®-------@
0--------0
Polarizability (Ellipsoid shape of Electron Cloud)
fa fa
Polarizability Variation With Normal Coordinates
Polarizability Derivative
(~~)al
0=0
Ramon activity
Dipole Moment
Dipole Moment Variation With Normal Coordinates
Dipole Moment Derivative
(~~Lo=o IR Activity
oa) (oQ
>"0 at 0=0
Active
o
o
+a +a
oP) (SQ
=
0
at Q=O
No
FIGURE 1.16. Diagram showing the stretching vibration of a diatomic molecule consisting of identical atoms.
.....
Vibrational Modes of Simple Molecules Vibrational Mode
23
@--@
@-----@
Polarizability Ellipsoid
Polarizability Variation With Normal Coordinates
Polarizability Derivative
(~)at Q~O
+0 +0 Active
Raman Activity
Dipole Moment
+,
Dipole Moment Variation With Normal Coordinates
Dipole Moment Derivative
(~~)at Q~O I R Activity
oa) ~o (oQ at Q~O
>
+ I
+0 +0
>
*0 oP\ ( oQl Q~O at
Active
FIGURE 1.17. Stretching vibration of a heterogeneous diatomic molecule. With Hel, such a vibration appears at 2885.9 cm- I in the gas phase, 2785.0 cm- I in the liquid phase, and 2768.0 cm - I in the solid phase.
For the bending vibration, there is no change in polarizability; therefore, it is Raman inactive. But it is IR active (Figure 1.20). In the CO2 , molecule, there is no overlap in fundamental vibrational frequencies between Raman and IR spectra. This is because the CO2 molecule possesses a center of symmetry. This is called the principle of mutual exclusion. There are many other compounds possessing a center of symmetry such as benzene, pyrole, pyrazine, and ethylene (Figure 1.21). The CO2 molecule consists of three atoms. The number of normal vibrations for a linear triatomic molecule should be 3 X 3 - 5 = 4. But actually only
24
Basic Concept and Elementary Theory Vibrational Mode
o-c-o
o-c-o
~
¢:
PoIarizability Ellipsoid
Polarizability
+ +0
Variation
With Normal Coordinates
Polarizability Derivative
(~QL
Q-O
Dipole Moment Variation With Normal Coordinates
Dipole Moment Derivative
(~L
0-0
IR Activity
Sa' ;0/0 SOlat 0 0 0
Active
Raman Activity
Dipole Moment
(
0
0
+0 +-0
(~6)
=0 at 0 0 0
No
FIGURE 1.18. Symmetrical-stretching vibration of a linear triatomic molecule such as CO 2 . It appears at 1285 cm- I in the Raman spectrum but is absent from the IR spectrum because there is no change in dipole moment.
three fundamental vibrational modes appear in Raman and IR spectra. This is due to the degenerate bending of CO2 , One bending takes place in the plane (Figure 1.22A) and the other out of plane (Figure 1.22B). Such "in-plane vibration" and "out-of-plane vibration" have the same energy, hence show the same frequency and are degenerate. So far we have discussed fundamental vibrations only. Actually CO2 produces more than four bands due to other mechanisms such as overtones and
....
Vibrational Modes of Simple Molecules Vibrational Mode
o-c-o
¢=
25
o-c-o
::;>¢=
Polarizability Ellipsoid
Polarizability Variation With Normol Coordinates
Polorizability Derivative
(~~L
0=0
Raman Activity
Dipole Moment
- ++ - (p=o) ¢:=-===:>
With Normal Coordinates
Dipole Moment Derivative 0=0
IR Activity
sa) (SO
=0 ot 0=0
No
Dipole Moment Variation
(~6t
-+a -f-a -
+ ====> + - (P ~ 0
¢:=. -
I
+a
fa
~o (M:.\ S °lat 0=0
Active
FIGURE 1.19. Asymmetrical-stretching vibration of a linear triatomic molecule such as CO 2 , The asymmetrical-stretching vibration of CO 2 is IR active (2349 cm -I) but Raman inactive because there is no change in polarizability, but there is a change in the dipole moment.
Fermi resonance (see Chapter 3, section 1.1). Usually fundamental vibration bands have a higher intensity than other bands. 7.3.
NONLINEAR TRIATOMIC MOLECULES
For nonlinear molecules the total degrees of freedom for vibrations is 3N - 6. Three coordinates are required to specify the position of each atom; thus 3N coordinates are needed for N atoms. Of all these degrees of freedom, three are
<}>
o/c" 0
o-c-o
Vibrational Mode
\1
{J Polarizability Ellipsoid
Polarizability Variation With Normal Coordinates
Polarizability Derivative
(~~at
0:0
-+0 fo
Ramon Activity
=0 (ba) bQ at 0:0
No
c 0 / "0
(p=o)
O-C-O
Dipole Moment
Dipole Moment Variation With Normal Coordinates
Dipole Moment Derivative
(~:t I R Activity
0=0
~+
~_
0" /O~C ~+
-+0 +0
(~at 0=0~O
Active
FIGURE 1.20. Bending vibration of a linear triatomic molecule such as CO 2 , This is IR active (667 cm -I for CO 2 ) but Raman inactive.
...
Vibrational Modes of Simple Molecules
o
00
II
C II
o CO 2
Benzene
27
,C=C H ,
H
/
/
H
H
Ethyl ene
Pyrazi ne
FIGURE 1.21. Examples of molecules with a center of symmetry. The symmetry of a molecule has an important relation to IR and Raman activities.
rotational motions and three are translational motions; thus the degrees of freedom remaining for vibrations are 3N - 6. Sulfur dioxide, S02' will be used as an example. The molecule is nonlinear; therefore, a possible maximum number of fundamental vibrations can be calculated from the equation 3N - 6. Accordingly, S02 should have 3 X 3 - 6 = 3 vibrational modes, which are symmetrical-stretching ("I' 1151 cm -I), asymmetrical-stretching ("3' 1361 cm -\), and bending vibrations ("2' 519 cm- 1). Usually the assignment is made for the highest frequency symmetric vibration as "\' and the next highest "2. After all symmetrical vibrations are assigned, then the asymmetric vibrations are counted starting at the highest frequency. In the symmetrical-stretching vibration there are changes in the electron-cloud size (polarizability) and dipole moment; hence this vibration is active in both Raman and IR spectra (Figure 1.23). For the asymmetrical-stretching vibration there is no change in the shape of the polarizability ellipsoid, but the orientation changes as it rocks. Such a vibration is Raman active. This vibration is also IR active, because there is a change in the dipole moment (Figure 1.24). For the bending vibration, both the ellipsoid and dipole moments change; therefore, it is both Raman and IR active (Figure 1.25). The vibrational bands described here are due to fundamental vibrations. Actually S02 shows more than three bands due to overtones (see Chapter 3, section 1.1) and combinations. 7.4.
VIBRATIONAL MODES OF THE METHYLENE GROUP
The C- H in the methylene group has complex vibrations, especially for bending (deformation) vibrations. For stretching vibrations, there are symmetrical and asymmetrical types (Figure 1.26). _o~
·A
P:':'o
,/
I
,/
C
\
o~o
B
[p
O-C
ft /1
d
FIGURE 1.22. Two types of bending vibrations of the CO 2 molecule producing degenerate bending. (A) Bending in the plane. (B) Bending out of plane.
s
{}
CO/s"cf
Vibrational Mode
0/""0
Polarizability Ellipsoid
+a
Polarizability Variation With Normal Coordinates
fa
Palarizability Derivative
(~t
0=0
Ramon Activity
Dipole Moment
n+
\1- 0
/5"
With Normal Coordinates
Dipole Moment Derivative
IR Activity
#0
SQ 010=0
Active
Dipole Moment Variation
(~l 0'
(k)
0=0
0
n+
~-o
/5~
a
+a fa
(~t O=~O
Active
FIGURE 1.23. Symmetrical-stretching vibration of a nonlinear triatomic molecule using S02 as an example (1151 em-I).
"a
lit...
s
S VI brat iona I Mode
0/ '0
0/ "0
Polarizability Ellipsoid
+0 fo
Polarizability Variation With Normal Coordinates
Polarizability Derivative
(~dt
0'0
Roman Activity
Dipole Moment
n:
Dipole Moment Derivative
FIGURE 1.24.
Activity
0=0
n+ s JJ_O/ "0
s 0/ "0
With Normal Coordinates
IR
f'!O SO at 0=0
Active
Dipole Moment Variation
(~~L
(so)
-f-
0
~o
(SP) SO at
¢O 0=0
Active
Asymmetrical-stretching vibration of a nonlinear triatomic molecule using S02 as
an example (1361 em-I).
~Q
Vibrational Mode
<}
5
/5\
/\ o 0
O~
00
Polarizability Ellipsoid
+0
Polarizability Variation With Normal Coordinotes
Polori zabil ity Derivative
~o
(h.....) SQ at 0 = 0
Raman Activity
/5"
o
Dipole Moment Variation With Normal Coordinates
Dipole Moment Derivative
I R Activity
FIGURE 1.25. em-I).
~O
atO=O
Active
Dipole Moment
(~6L
(k) ~O
0=0
n+ 6b~-
n+ 0{7-
-f-
5
0
-fa
~o ( ~P) ~o atO=O
Active
Bending vibration of a nonlinear triatomic molecule using 50 2 as an example (519
~
Vibrational Modes of Simple Molecules
H",~H
H~yH
C
C
31
I ASYfJIlIetrical 2926 cm- l
Symmetrical 2853 cm- l FIGURE 1.26. -CH 2 -·
Symmetrical- and asymmetrical-stretching vibrations of the methylene group,
For bending vibrations, there are in-plane as well as out-of-plane types (Figure 1.27). 7.5.
VIBRATIONAL MODES OF THE METHYL GROUP
The C- H in the methyl group has stretching as well as bending vibrations, as can be seen in Figure 1.28. In- Plane Type
H~
~H
\C / Rocking 724 -1I74cm- 1
Symmetrica I Bending (scissors deformation) 1463 em-I
Out-of-Plane Type
-p"
H
\,
H
H """,,;
J
,"1/{1
"c ......::...ll'HP"V-..c:w.I rr~j!
TwistinO 1063 -1295cm-1
Wagging
1170-1382cm- 1
FIGURE 1.27. In-plane and out-of-plane methylene bending vibrations.
Stretching Vi brat ion
~ '\J
Symmetric
~ ~
Asymmetric
Bending (Deformation) Vibration
~~
~ ~ C
H
c:;:::-
Asymmetric
H
Symmetric (Umbrella)
FIGURE 1.28. Stretching (symmetrical and asymmetrical) and bending (symmetrical and asymmetrical) vibrations of the methyl group, -CH 3 .
A
~o~ ~
4-
Y
Extreme of compressed position in stretching vibration FIGURE 1.29.
Equilibrium position
Extreme of stretching vibration
Breathing vibration of a cyclic compound.
Depolarization Ratio
~n~ s X
1?
Ii
H
33
x or expressed as'
(±)HOHe (±)H H@
'Ii
H
e The (±J indicates that the H atom vi brates out of the plane.
,M '.'
-
X expressed or as·
~(±) e e @
Ring Pucker Vibration
FIGURE 1.30.
7.6.
Vibrational modes of monosubstituted benzene giving IR lines.
CYCLIC COMPOUNDS
The vibrational modes of a cyclic compound can be extremely complex. Not only do they possess many different bond-stretching vibrations, but the ring compounds also show a special type of vibration called a breathing vibration. Breathing vibration refers to the in-phase stretching vibration; all bonds involved stretch by the same length at the same time. Breathing vibration can best be visualized from diagrams shown in Figure 1.29. Normally the breathing vibration and other ring vibrations give prominent Raman bands. Even for a relatively simple aromatic compound such as monosubstituted benzene, a complete assignment of all vibrational modes is not an easy task. A total of 30, near-normal modes of vibration of the monosubstituted benzene ring are listed by Colthup et al. (1975). Two types of in-phase and out-of-phase C- H wagging vibrations and the ring-pucker vibration are shown in Figure 1.30. These examples serve to show the complexity of even the simplest cyclic compounds. 8.
DEPOLARIZATION RATIO
When a parallel-oriented polarizer is placed in the path of polarized light, the light will pass through (Figure 1.3IA). However, when the polarizer is rotated 90°, no light will pass through (Figure l.3IB). Consider a molecule at 0 that scatters the incident light. Figure 1.32 show only the light scattered toward the direction of the Y axis, toward the observer.
34
Basic Concept and Elementary Theory
z
(aJ
Y--Evv+ftv0J-PO:~;:;'d Z
~90. (b)
y-fvv+l' FIGURE 1.31.
No polarized light passes through
Effect of a polarizer on polarized light.
For highly symmetrical molecules, such as CH 4 or SF6 , the polarizability is isotropic. When totally symmetrical vibrational modes of such molecules, for example, the in-phase stretching of the four C-H bonds of CH 4 , interact with incident laser light that is polarized in the XZ plane, the scattered light is polarized in the YZ plane. Thus when a polarizer is placed parallel to the YZ plane, only scattered light in this plane is allowed to pass. The intensity of this light is called III' The light in the YZ plane will not pass through when the polarizer is rotated 90°. The scattered intensity measured for this position of the polarizer is called I.l' It is seen that the depolarization ratio, defined as p = I.l/ III' is zero for the symmetrical C-H stretching mode of CH 4 . Most molecules are less symmetrical than CH 4 or SF6 ; therefore, the polarizability is usually anisotropic, and, in general, the molecule scatters light polarized in both the XZ and XY planes. The intensity of the scattered light observed to the polarized parallel (Z direction, Figure 1.32A) and perpendicular (X direction, Figure l.32B) to the incident light is different, but the depolarization ratio is not usually zero. When incident light is plane polarized, as is the case of a laser beam the depolarization p is expressed by the equation I.l
p= -
III
=
3(aaf 45(a j )2
+ 4(aa)2
where Isotropic part of the polarizability (the polarizability for a spherical molecule). au: Anisotropic part of the polarizability (the polarizability for an asymmetrical molecule). I.l: Intensity of the scattered radiation polarized perpendicular to the incident light. III: Intensity of the scattered radiation polarized parallel to the incident light. aj :
11....
Depolarization Ratio
35
z A.
I!
I
Y_10LJL_JJh~
..
.I
/\ /\
I~I"
Parallel Oriented Polo' nz~
X
Incident Light
The intensity of light observed III
Z
B.
ru I n ._ .lI-W--Vr ! l XJjVVY-' p
Y _. P
II _ _
.:
,
,
X
1
P~rpendicularl
Oriented Polo r1zer .y
The intensity of light observed 1.1.
FIGURE 1.32. The intensity of scattered light can be measured two different ways. (A) Light comes through a parallel-oriented polarizer and is parallel to the incident light (III)' (B) Light comes through a perpendicularly oriented polarizer and is perpendicular to the incident light (1,L)' The ratio of 1,L to III is called the depolarization ratio, and it is related to symmetry of vibrational modes.
For plane-polarized light, the depolarization ratio has a value from 0 to i, depending on the degree of antisymmetry of that vibration. For non-totally symmetrical vibrations of any molecule, the value is t (a; = 0). For a totally symmetrical vibration, p ::5 i. It was zero for the example of CH 4 mentioned above, but has values between 0 and i for" totally symmetrical" vibrations of molecules in general. A totally symmetrical vibration is unchanged on performing any symmetry operation (rotation axis, reflection plane, etc.) that simply permutes the equilibrium positions of equivalent atoms in the molecule.
36
Basic Concept and Elementary Theory
Because of this, measurement of the depolarization ratio gives useful information and is frequently used to support the assignment of a Raman line to a particular vibrational mode.
9.
GROUP FREQUENCY
As the molecular weight increases, so does the number of atoms present in a molecule. This means the number of vibrational modes also increases. For a nonlinear molecule, the maximum number of fundamental vibrations is 3N - 6. Usually vibrations of a particular submolecular group of atoms appear with characteristic group frequencies. For instance, for compounds containing the -OH group, the stretching vibration of the -OH group usually appears in the region of 3100-3600 em-I. This is because the force constants for O-H bonds are roughly the same for different molecules. Glucose contains five -OH groups, and the stretching vibrations appear at 3200-3500 em -I. In ethanol, the -OH frequency appears at 3360 em- '. The C- H stretching vibration normaUy appears at 2800-3000 em-I. For glucose, the C-H bands appear at 2875, 2890, 2910, 2940, and 2960 em -I (She et a1., 1974). For erythrocyte ghosts, the C-H vibrations appear at 2842, 2890, 2935, 2975, and 3017 em-I (Milanovich et a1., 1976). The C=O stretching vibration normally appears as a prominent band in the region of 1600-1800 cm- I in IR and Raman spectra. There are many functional groups that contain C=O; the frequency of the C=O stretching vibration depends on the functional group. Thus from the C=O stretchingvibration band, one can sometimes postulate the type of functional group. A few examples of functional groups and their C=O frequencies are shown here: Peptide Carboxylic acid Carboxylate Ketone Aldehyde Ester Acid anhydride
1640-1670 em-I 1750-1780 em-I 1500-1610 em-I 1690-1800 em-I 1720-1730 em-I 1720-1760 cm- 1 1760-1820 em-I
These are just a few examples of group frequencies. It is helpful to know the frequency ranges of several groups so that one can get some ideas about the origins of vibrations of a complicated Raman spectrum of a biological compound. The group-frequency assignment is made strictly from empirical fact; therefore, it is not unusual that the vibration of some functional groups appears at an atypical frequency. For instance, for a strongly hydrogen bonded C=O group, the stretching frequency tends to shift to lower frequency.
I-- -
CH
II NH
.-H C=C jH CH 3 CH 2
,..
I··
-
• -..
• I C-5
IS-H
• II Cli) J!C ~C
1-
--. --C;C• mon
~~
,..N-R II
III
I
N·N-, II CH \-.
811l
-.
I
3600
FIGURE 1.33.
3200
2800
2400
2000
1600
1,;o:~
,0$,.,
P'I'
- .... .... 8 1,4.
...
~
"
I'"lIr
-Ne 2 -
--,...,,.... II
1200
~1,~
mor 10.
R-.
-.NC bl
4000
"8~
mor 10.,
TI ICOOt
800
,
~.,
CI·t IIk8 nl l-811U1
J'es
8.1<8. 1l'8
400
Examples of group frequencies. The diagram was reproduced from a brochure of Varian Instruments.
o
38
Basic Concept and Elementary Theory
Some group frequencies are shown in Figure 1.33, but the figure is by no means complete. Many Raman frequencies of functional groups coincide with IR frequencies. However, there are usually large intensity differences between the bands for Raman spectra and IR absorption spectra. Some functional groups such as C=C, C=N, C=C, N=N, S-H, C-S, and S-S give intense Raman bands but show relatively weak IR absorption bands.
10. USE OF ISOTOPES 10.1.
ISOTOPIC SUBSTITUTION
Isotopic substitution is very useful in identifying particular vibrational modes. For a diatomic molecule, vibration frequency is given by the equation
P
= _1 [k ( _1 + 2'TT
mA
= _1
(~) 1/2
2'TT
_1 )] 1/2
= _1 [k ( m A + m B )] 1/2 2'TT
mB
mAm B
J.L
where m A and m B refer to the mass of A and B, respectively; k is the force constant; and J.L is the reduced mass equal to mAmB/(m A + m B ) for the diatomic molecule. When one of the atoms is replaced with an isotope, the mass is changed, thus causing a shift in vibration frequency. Let's assume the hydrogen atom in N-H is replaced with deuterium, giving N-D. The atomic weights are N, 14; H, 1; and D, 2. Thus the ratio is 1 [ (14
PN-H P
N
=
- D
I; k _1 [ 2'TT k
+
1 )] 1/2
l4Xl (14 + 2 )]1/2 14 X 2
The ratio is roughly equal to :N-H N-D
= (~) 1/2 _ 1 - 1.4
Assuming the force constant (or bond strength) does not change for N-H and N-D, the N-H stretching vibration will shift to lower frequency by the factor of approximately /2 when N-H is isotopically exchanged to N-D.
r
~
S!
b 459.3
~1
Carbon Tetrachloride
g 462.4
a
oCD
o .....
2----'
456.1 c
o
'" g
452.7
~
d
~ FIGURE 1.34. Raman spectrum of carbon tetracholoride, CCI 4 . There are so many Raman lines because of various naturally occurring isotopic chlorides. Only the symmetrical-stretching vibration (v)) is shown in this figure. Carbon tetrachloride also has other vibrations such as V2 around 210 em -), V4 around 308 em -I, V3 at 790 em -), and a v) + V4 combination at 760 em -I, but these are not shown in the figure.
39
40
Basic Concept and Elementary Theory
This method is actually used to assign the amide III band (see Chapter 3). In lipids there are many C- H vibrations. In order to determine the origin of a particular C-H vibrational band, deuterium can be introduced to a particular position by an organic synthesis, and the shifted C- D band can be identified. From this, one can precisely identify the origin of a particular C- H vibration.
10.2. NATURAL ISOTOPES If a compound contains sufficient amounts of isotopes with significant natural
abundance, this will be reflected in the Raman spectrum. The best example is CCI 4. Carbon tetrachloride contains 75.53% 35Cl and 24.47% 47Cl. Thus the naturally occurring CCI 4 is a mixture of C 35 Cl 4, C35C1337CI, C 35 Cl/ 7Cl 2 , C 35 CI 37CI 3, and C 37 Cl 4 (Carter, 1976). As can be seen in Figure 1.34, these are indeed shown in the Raman spectrum. Resolving such closely spaced Raman bands requires a high-resolution Raman spectrometer. Therefore, CC1 4 is frequently used to check whether a Raman spectrometer is in good condition or not. If the resolution is not good, the a and b bands tend to overlap (Figure 1.34). The author measures the intensity at positions 1 and 2 occasionally to check the condition of the instrument.
11.
PROBLEM OF FLUORESCENCE
Only a small fraction of incident-light energy is manifested as Raman scattering; most of the input energy is expressed as Rayleigh scattering. So in terms of energy, Raman spectroscopy is highly inefficient scattering spectroscopy. If a sample is fluorescent, Raman spectra are often masked by fluorescence spectra. Often fluorescence comes from impurities present in the sample. There are several ways to eliminate or suppress the fluorescence. 1.
2.
3.
Purification of sample. The sample to be analyzed has to be biochemically as well as optically pure, which means that the sample should have no fluorescence. Lyophilization of the sample sometimes improves the quality of the spectra. Of course, this will improve the spectra only when the impurity is relatively volatile. Prolongation of exposure of a sample to the laser beam. Nobody knows exactly why prolonged exposure to illumination suppresses the fluorescence. One possible explanation is that this bleaches the fluorescent moiety. Choice of excitation wavelength. There are basically two competing processes at work when one illuminates a biological sample with visible or ultraviolet laser radiation: (i) the sample (or impurities) may absorb the radiation and emit it as fluorescence, and (ii) the sample scatters the
General References
41
light. The Raman intensity is inversely proportional to the fourth power of incident light wavelength (I a: 1M). Consequently, there exists an optimum choice of incident wavelength to achieve a maximum signalto-noise ratio for any given sample. Thus by lowering the wavelength of the excitation light, one can increase the intensity of the scattered light. Sometimes there is an advantage to using different excitation lines. Fluorescence depends on the wavelength of excitation. At some wavelengths fluorescence is produced, whereas at other wavelengths there is no fluorescence. For instance, by using the 333.6-nm excitation line, Raman scattering shifts away from the fluorescence band and a goodquality Raman spectrum can be obtained (Bowman and Spiro, 1980). 4. Signal averaging. If the degree of fluorescence is small, signal averaging will improve the signal-to-noise ratio. After SCAMP, a computer from Spex Industries, Inc., was installed in the author's laboratory, the quality of spectra markedly improved with repetitive scanning and averaging of their signals. 5. Use of pulsed laser. Instead of using a continuous-wave source, using a pulsed laser may improve the quality of the spectra. The purpose is to record the Raman-scattered light before fluorescence takes place. The lifetime of Raman scattering is on the order of 10- 13 _10- 11 s, whereas that of fluorescence is around 10- 9 _10- 7 s. In order to achieve temporal resolution of the Raman scattering and fluorescence signals, it is possible to create a short-lived Raman signal by using a pulsed laser. An electronic time gate is made on the photomultiplier-tube detection circuit and is synchronized with the pulsed-laser source so that short-time events, or primarily Raman photons, are preferentially recorded. The article by Van Duyne et at. (1974) provides more information about this device. 6. CARS. As discussed in Chapter 2, CARS is a good technique to obtain fluorescence-free Raman spectra. If present technical difficulties are overcome, CARS will become very common in biological fields in the future.
12.
GENERAL REFERENCES
Since the object of this book is not to review the fundamental theory of Raman spectroscopy, readers are encouraged to look at general reference books as necessary. The following are a few: Wilson, E. B., Decius, J. C. and Cross, P. C. (1955) Molecular Vihration, The Theory of Infrared and Raman Vihrational Spectra, McGraw-HiU Book Co., New York, 388 pp. Gilson, T. R., and Hendra, P. J. (1970) Laser Raman Spectroscopy, Wiley, New York, 266, pp. ~ TObin, M. C. (1971) Laser Raman Spectroscopy, Wiley, New York, 171 pp.
42
Basic Concept and Elementary Theory
Dollish, F. R., Fateley, W. G., and Bentley, F. F. (1974) Characteristic Raman Frequencies of Organic Compounds, Wiley, New York, 443 pp. Freeman, S. K. (1974) Application of Laser Raman Spectroscopy, Wiley, New York, 336 pp. Colthup, N. B., Daly, L. H., and Wiberley, S. E. (1975) Introduction to Infrared and Raman Spectroscopy, 2nd ed., Academic, New York, 523 pp. Long, D. A. (1977) Raman Spectroscopy, McGraw-Hill, New York, 276 pp.
The book by Dollish et al. (1974) does not describe basic theory, but the author found this book very useful in determining the particular frequency of a functional group. For theoretical aspects of Raman spectroscopy one can use: Sushchinskii, M. M. (1972) Raman Spectra of Molecules and Crystals. Israel Program for Scientific Translation, New York, 446 pp.
The book by K. Nakamoto (1978) Infrared and Raman Spectra of Inorganic and Coordination Compounds, 3rd ed., Wiley, New York, 448, pp., is an excellent reference book, especially for someone working on inorganic or metallo compounds. A small booklet published by Beckman Instruments, Inc., Introduction to Raman Spectroscopy, by R. J. Obremski (1971), is written in such easy language that it will be useful for someone without much background in physical chemistry. Another very useful study aid is the American Chemical Society Audio Course called Application of Raman Spectroscopy by Stanley K. Freeman. It consists of six cassette tapes and a book with concise descriptions. J. Huley Associates (P. O. Box 910, Boca Raton, FL 33432) issues an audiovisual course entitled Raman Spectral Interpretation arranged by William G. Fateley. This consists of tapes and slides. The author found this very useful for understanding Raman characteristics of organic compounds. Both are very helpful for beginners who want to get into Raman spectroscopy for the first time. Sometimes Raman spectroscopy is described as a part of other spectroscopy books. For instance, the following books contain information about Raman spectroscopy and are relatively easy to read: Brittain, E. F. H., George, W.O., and Wells, C. H. J. (1970). Introduction to Molecular Spectroscopy, Theory and Experiment, Academic, New York, 387 pp. Jones, D. W. (1976) Introduction to the Spectroscopy of Biological Polymers, Academic, New York, 328 pp. Nakanishi, K., and Solomon, P. H. (1977). Infrared Absorption Spectroscopy, Holden-Day, San Francisco, 287 pp. Theophanides, T. M. (1979). Infrared and Raman Spectroscopy of Biological Molecules, Reidel, Dordrecht, Holland, 372 pp. Walton, A. G., and Blackwell, J. (1973). Biopolymers, Academic, New York, 604 pp.
There are several monographs dealing with specialized topics of spectroscopy. Infrared and Raman Spectroscopy published by Marcel Dekker of New
f References
43
York so far has three volumes. Advances in Infrared and Raman Spectroscopy by Heyden, London, has eight volumes published so far. Elsevier Scientific Publishing Company has a series of spectroscopy monographs, Vibrational Spectra and Structure, which includes Raman spectroscopy.
REFERENCES Bowman, W. D., and Spiro, T. G. (1980). Fluorescence-free resonance Raman spectra of reduced nicotinamide adenine dinucleotide via ultraviolet excitation. J. Raman Speclrosc., 9, 369. Brandmiiller, J., and Kiefer, W. (1978). Physicist's view, Fifty years of Raman spectroscopy. Spex Speaker, 23, 310. Carter, R. L. (1976). Inorganic materials. In Infrared and Raman Spectroscopy, E. G. Brame, 1r., and 1. Grasselli, Eds.. Dekker, New York, pp. 71-206. Colthup, N. B., Daly, L. H., and Wiberley, S. E. (1975). Introduction to Infrared and Raman Spectroscopy, 2nd cd., Academic, New York. 1ohnson, B. B., and Peticolas, W. L. (1976). The resonant Raman effect. Ann. Rev. Phys. Chern. 27, 465.
Lewis, A, and Spoonhower, 1. (1974). Tunable laser resonance Raman spectroscopy in biology. In Spectroscopy in Biology and Chemistry, S. H. Chen, Ed., Academic, New York. Milanovich, F. P., Shore, B., Harney, R. c., and Tu, AT. (1976). Raman spectroscopic analysis of Dutch Belt Rabbit erythrocyte ghosts. Chern. Phys. Lipids 17, 79. Placzek, G. (1934). Rayleigh-Streung und Raman-Effekt, Handbuch der Radiologic, E. Marx, Ed. Leipzig. She, C. Y., Dinh, N. D., and Tu, A T. (1974). Laser Raman scattering of glucosamine, N-acetylglucosamine, and glucuronic acid. Biochim. Biophys. Acta 372, 345. Spiro, T. G., and Stein, P. (1977). Resonance effect in vibrational scattering from complex molecules. Ann. Rev. Phys. Chern. 28, 501. Strommen, D. P., and Nakamoto, K. (1977). Resonance Raman spectroscopy. J. Chern. Educ. 54, 474.
Van Duyne, R. P., 1eanmaire, D. L., and Shriver, D. F. (1974). Mode-locked laser Raman spectroscopy-a new technique for the rejection of interfering background luminescence signals. Anal. Chern. 46, 213. Warshel, A (1977). Interpretation of resonance Raman spectra of biological molecules. Ann. Rev. Biophys. Bioeng., 6, 273.
....
CHAPTER
2
The Laser Raman Spectrometer and Special Techniques Raman spectroscopy is occasionally called laser Raman spectroscopy. There is a good reason for this. Practically all modern Raman spectrometers employ lasers as the light source. Only a small fraction of scattered light is emitted as Raman scattering, whereas the majority of the scattered light has the same energy as the incident light (Rayleigh scattering). The laser is not only monochromatic and coherent light, but it is also a powerfully intense source of light. Therefore, even the small fraction of scattered light that shows the Raman effect is sufficiently strong in intensity to be detected by conventional detection methods such as the photomultiplier. In this chapter the basic theory of laser-light production, the basic units of Raman spectrometry, and some special techniques are briefly described.
r
r
Laser
1.
45
LASER
The laser was invented by C. H. Townes and A. L. Schawlow in 1958, and lasing action was first observed by T. H. Maiman in 1960. The word laser stands for Light Amplification by the Stimulated Emission of Radiation. In earlier days lasers were called optical masers. The maser generates coherent and amplified microwaves, and it was invented before the laser was. Maiman observed lasing using a single crystal of ruby with the ends ground flat and silvered (Hellman, 1969). Prior to the invention of the laser, Raman spectroscopy was performed using mercury lamps as the illuminating source. Due to the small cross section for Raman scattering, only the strongest vibrations were observed and then only on long exposure of photographic plates to the scattered light. The laser compacts intense radiation into a narrow frequency range (i.e., it is nearly monochromatic). The intensity is such that all but the weakest Raman-active vibrations can be observed photoelectrically. In laser Raman spectroscopy, the laser is used only as a source for the incident light; yet it is worthwhile to have some understanding of basic laser theory. Lasers are also used for many other purposes, such as photochemistry, laser photophysics, remote sensing of atmospheric pollutants, communication, and different types of spectroscopy (Brown, 1968; Bloembergen, 1975; Kimel and Speiser, 1977; Hudson, 1977). Edward Teller made an interesting comment about the laser; "Lasers have been discovered by the wrong people, namely by physicists. The result is that the chemists have not grabbed it to the extent they should. . . . The only people who can make real progress are the chemists" (Teller, 1973). Many different varieties of lasers have been developed. We will discuss the basic principles briefly. 1.1.
BASIC THEORY
Light is emitted from an atom or molecule due to a transition from a higher energy level of the atom or molecule to a lower energy level (Figure 2.1). Depending on the energy difference, different wavelengths of light are emitted. When the energy difference is large, the atom releases radiation as ultraviolet light. If the energy difference is less, then visible light is emitted. If the energy difference is even smaller, then the emitted light is in the IR region. Even-smaller energy differences lead to microwave emission, which is why the maser was invented before the laser. It is easier to obtain population inversion (see below) for microwave transitions, and harder for ultraviolet-visual transitions. An excited atom or molecule can lose energy by either stimulated or . spontaneous emission of radiation. Examine the situation when an atom a~l.readY "pumped" (generally "pumping" refers to exciting the molecule, not ~ the stimulated-emission process) to a higher energy level (Figure 2.2A)
k..
46
The Laser Raman Spectrometer and Special Techniques
• • "h'
-
'vv-l"
h
B.
A. An atom absorbs light
:r l ~II_-M-
energy
An atom
C. An
at
atom
at
lower energy due
higher energy due
to
~orPtion
spontoneous emission
This process is called "PumpinlJ"
FIGURE 2.1.
Absorption and emission of light by an atom or a molecule.
receives radiation energy of exactly the same amount that the atom would emit to make a transition (A E) to a lower energy level. The atom under these conditions is stimulated by the incident photon to lose its energy AE, and the energy lost is converted to light (Figure 2.2B). The emitted light possesses the same phase as the stimulation light, thus resulting in the coherent property of the laser. Since the two waves have the same energy, they possess the same frequency, thus resulting in the monochromatic nature of laser radiation. The mechanism of laser-beam production is diagramed in Figure 2.3. The first step (Figure 2.3A) is to pump energy into the atoms so that a higher proportion of atoms or molecules will be in higher energy states (Figure 2.3B). In other words, one artificially changes the natural thermal balance and creates a situation with an abnormally large number of high-energy atoms or molecules. This is said to establish a population inversion. When one of the atoms or molecules in the higher energy state _falls back to the lower state, it emits a photon of energy AE. This photon stimulates other high-energy atoms to emit a similar photon immediately, as shown in Figure 2.2, and to fall to the lower energy state. Each atom emits light with energy of AE (Figure 2.3C). The emitted light moves back and fourth between two ends. The light becomes very intense (light is amplified) between the mirrors. Some of it is taken out through one end mirror, which is not completely silvered and therefore transmits part of the light instead of reflecting it all (Figure 2.3D). A molecule higher energy
Photon 2
1I='~'\/'v h
• I 6E
Radiation
1 A.
FIGURE 2.2.
at level
P~II=~}
-...
Photon 2
AJV-
h II = 6 E h
Stimulated emission
B.
Production of enhanced coherent light by stimulated emission.
r Laser
47
~ +M! ---.--.-.-.---.-.-.-0-.-.-.-.-
--.-0-.-0-0-0-.-.-
A
B
11
~
.
~ c
l~lvlv lv~jv ~ ---.-.-.-.-.-e-.-o-
o
~
--0-0-0-0-0-0-0-.-
-
t----
~--=--r -~--r---
~~
--~
~~~~r-
FIGURE 2.3. Production of a laser beam. (A) Pumping process (either by light or other means) raises many atoms (or molecules) to excited states. (B) Many atoms or molecules in excited states. The "population inversion" is established. This means that more atoms (or molecules) are excited than are at lower energy levels. (C) Lasing begins when photons are spontaneously emitted, as metastable excited atoms or molecules fall back to the lower energy level. (D) The spontaneously emitted photon starts a cascade of many photons (light waves). When photons of identical energy and phase are reflected back and forth, they can leave the laser tube through a partly transparent end mirror as coherent light.
Ordinary light consists of beams propagated in different directions with different frequencies and phases (Figure 2.4). We may say that such light is optically not pure or not coherent. 1.2.
LASER TUBES
Many types of laser tubes are commercially available. Frequently 514.5 nm is used for excitation light for Raman spectroscopy because this is the strongest line for an argon ion laser. There are several other lines from argon ion lasers that can be used (Table 2.1). The choice of excitation wavelength is particularly important in resonance Raman spectroscopy, since a particular chromophore is resonance enhanced when the excitation wavelength is within the absorption spectrum of a compound. Sometimes Raman intensity of a particular band is ~ plotted against excitation of different wavelengths (excitation profile, Raman
48
The Laser Raman Spectrometer and Special Techniques
FIGURE 2.4. Ordinary light, such as that emitted from a tungsten light bulb, is a mixture of different frequencies, directions, and phases. Therefore, it is an example of noncoherent light.
dispersion spectroscopy). For such purposes, one needs laser lights of different wavelengths. Some broadly tunable dye lasers are commercially available. The dye laser consists of two main components: an organic dye solution and a pumping source. The nitrogen laser has become the accepted pumping source for tunable dye lasers because the short wavelength (337.1 nm) permits dye tuning over a broad range (360-750 nm), and peak powers are high enough to double frequency to the range 260-360 nm. The dye laser absorbs energy from the pump laser, and then it can lase over a broad band that is characteristic of each dye. In other words, the lasing action occurs through the fluorescence transition of an electronically excited organic molecule. Molecules suitable for dye-laser action have virtually a continuum of energy levels in the ground electronic state. This permits tuning of the laser output by proper cavity adjustments. For more detail readers are advised to see other books or articles (e.g., Cunningham, 1974).
2.
BASIC UNITS OF RAMAN INSTRUMENTS
A Raman spectrometer essentially consists of a light source, sample compartment, monochromator, photomultiplier system, and electronic-signal-processing unit (Figure 2.5). The overall view of a Raman spectrometer currently used in the author's laboratory is shown in Figures 2.6A and 2.6B. The principle of the Raman spectrometer shown in Figure 2.5 is briefly described here. As the incident light hits a sample, light is scattered in all directions. The scattered light includes both Rayleigh (elastic) and Raman (inelastic) scattered light. The intensity of elastically scattered light is much greater than that of inelastically scattered light. In a Raman spectrometer, an
TABLE 2.1.
Wavelength of Varfous Laser Tubes
Gas Fill Options Output Power Wavelength (nm) (All Output Power TEM oo ) Far red
Red Yellow
Green
Blue
Violet Ultraviolet
799.3 793.1 752.5 676.4 647.1 568.2 530.9 520.8 514.5 501.7 496.5 488.0 482.5 476.5 476.2 472.7 465.8 457.9 454.5 341.1 + 363.8 350.7 + 356.4
Argon Model 164-00 (mW)
Krypton Model 164-01 (mW) 30 10 100 120 500 150 200 70
800 140 300 700 30 300
Argon/Krypton Model 164-02 (mW)
20 200 80 80 20 200 20 50 200 10 60
Argon Model 164-03 (mW)
1400 250 400 1300 500
50 60 50 150
20 20
150 100 250 100 20
40
Source: Reproduced from Spectra-physics manual for Model 164/166 Ion Laser with Model 265 Exciter, Spectra-Physics, Mountain View, California (1976).
50
The Laser Raman Spectrometer and Special Techniques
C
LASER TUBE
~ fr-~-_._~----ll ~.._.:_.:>{] __.-4'~
!!-=: - -. _ -]II._~.-
~=/~~ DOUBLE
MONOCHROMATOR
)--(])_.y I
"'''' I DJ"'""",," """~~ II'\'
""" .
"'~~""~ ~ N'JJ\g;\ '. I R=D~R' "1 ELECTRONIC
•
... .
,
," ,,
"
.
~ITill"" QJ .. 1~ .. ~ ~~~ ~-
I
A
.
"
','
1\ COMPUTER
FIGURE 2.5. examined.
Essential parts of a Raman spectrometer. Normally the scattered light at 90° is
instrument is often designed to collect scattered light at 90 0 with respect to the incident light. The collected scattered light is aligned and focused to the slit of a double monochromator. In a sophisticated Raman spectrometer, usually two to three monochromators are used. As the ratio of incident light to Ramanscattered light sometimes exceeds 10 9 , high spectral purity is needed to unveil weak Raman spectra. In a double monochromator, light is dispersed in the first monochromator, and it is again dispersed in the second. For special experiments, such as observing Raman lines at extremely low frequency, a third monochromator may be needed. The light coming out the exit slit of the final monochromator is collected and focused on a photomultiplier tube, which converts photons of light into an electrical signal after amplification. For special purposes a pulsed laser is also used as a light source. Had the laser been invented 20-30 years earlier, the popularity of Raman and IR spectroscopy might have been reversed. In a modern Raman spectrometer, the Raman wave number (wave number difference of the incident and the scattered light) and the absolute value of the wave number of the scattered light are automatically shown on the instrument panel. For a liquid sample, a capillary tube is the most commonly used container. There are different devices for holding samples depending on the need. For instance, for deuterated samples, the author used a specially constructed sealed
r
~
FIGURE 2.6. A Raman spectrometer and computer attachment used at the author's laboratory at Colorado State University. (A) Ramalog 5 manufactured by Spex Industries. The laser tube is placed in the back and is not shown in the photograph. (B) Spex SCAMP computer. The computer can average spectra after repetitive scanning, calculate intensity ratios or the ratio of areas of two lines, and measure the depolarization ratio.
1;;1
\\11 I \ I \'.-1\C
> l-
e;; Z
\
'
~ z « :E «c::
~1!1j
I
W
I
~
II~
1700
\
l!l!t,'1
~ '~1 j
;,,1
1600
B
\ ~ All
! \ I
III
I
~ I l'll~ ~
I~I~ ~!lI!I.~ ~ I
~
'I
1\
"
~
J
A
I
i
1500
1400 -1
1300
1200
eM
FIGURE 2.7. (A) Raman spectrum of lysozyme obtained from one scan. (B) Additive spectrum derived from 10 scans registered in the SCAMP computer. (C) Averaged Raman spectrum of lysozyme derived from 10 scans with smoothing action. These spectra were obtained using a SCAMP computer in the author's laboratory.
r Special Techniques
53
glass chamber to prevent reexchange with moisture in the air. For Raman-intensity measurement at elevated temperatures, the author constructed a copper heating block that is connected to a rheostat to control the heat generation. The temperature of the block is read on a thermocouple (Fox and Tu, 1979). For colored samples or any that are ordinarily destroyed by absorption of heat from laser light, Raman spectra can be obtained by rotating the sample. A sample rotator attachment can be obtained from Raman-spectrometer manufacturers. Usually Raman signals are very weak, so background noise occasionally becomes high. Signal averaging with the aid of a computer can improve the quality of the signal-to-noise ratio. The author uses computer averaging routinely (Figure 2.7). Using conventional Raman spectroscopy, a relatively long time is required to obtain a complete spectrum. This limits the use of Raman spectroscopy to relatively stable molecules. Recently, because of technological advances, multiwavelength detectors have become available for Raman spectroscopy. With the combined use of a picosecond laser and multiwavelength detector, one can obtain Raman spectra of short-lived compounds. For instance, a 7-ns time-resolved resonance Raman spectrum of cytochrome c with good resolution was obtained by Woodruff and Farquharson (1978). Recently an optical multichannel analyzer with sufficient sensitivity has become available to Raman spectroscopists. It consists of a large number of detectors, with each detector "channel" measuring a different wavelength of scattered light. Since normal scanning method is time consuming, it is not suitable for a fast reaction. A multichannel analyzer records the frequencies and intensities of all scattered Raman bands simultaneously. Rotating choppers with variable-size slits are also frequently used for time-resolved resonance Raman spectroscopy. The choppers and a continuous-wave laser function as a pulsed laser with variable pulse width. 3.
SPECIAL TECHNIQUES
As Raman spectroscopy is advancing, many special techniques are becoming more common. In this section, a few of them are briefly mentioned. 3.1.
RAMAN 01 FFERENCE SPECTROSCOPY (RDS)
Double-beam spectrometers are extensively used in IR, visible, and UV absorption spectroscopy in order to cancel the absorption by water or solvent. Such a two-channel technique has just been introduced for Raman spectroscopy. Raman difference spectra are obtained by the use of a cylindrical cell with a .~.partition along a di~meter' so th~t two sample~ are alternately illuminated by . the laser as the ceills rotated (Kiefer, 1977) (Figure 2.8). The computer-generated difference between the Raman spectra of the two samples is plotted. By
54
The Laser Raman Spectrometer and Special Techniques RAMAN
DIFFERENCE
SPECTROSCOPY
CD
CD-® CYLINDRICAL ROTATING RAMAN SPUT CELL
q q
Q)® FIGURE 2.8. Simplified diagram of the arrangement for Raman difference spectroscopy. As the cell is rotated, a synchronous signal is sent to gating and counting electronic instruments to allow for the independent accumulation of data from each sample with subsequent independent storage in a computer. To obtain a difference spectrum, the intensity of a preselected Raman line in one spectrum is adjusted to equal the intensity of that line in the other spectrum. The two spectra are then subtracted digitally.
the RDS technique one can measure frequency differences as small as 0.02 cm- 1 between the maxima of very similar Raman bands of differen t compounds. An actual example of RDS on cytochrome c from tuna and horse is shown in Figure 2.9 (Shelnutt et al., 1979; Rousseau, 1981). Because of further technical improvements, four-channel Raman difference spectroscopy has been developed (Laane and Kiefer, 1981). 3.2. 3.2.1.
COHERENT ANTI-STOKES RAMAN SPECTROSCOPY (CARS) Technique
The CARS phenomenon was discovered by Terhune in 1965. Recently CARS has attracted great attention in biochemical research, because one can get Raman spectra free from fluorescence background.
....
Special Techniques
Tuna -
55
1.24 X Horse
~
i
'iii
c:
.•c:.. c:
II
E II
a:
780
720
800
FIGURE 2.9. Raman difference spectrum of tuna cytochrome c versus horse cytochrome c. The 750 cm - I line of tuna cytochrome c is 0.1 cm - I higher in frequency than that of horse cytochrome c. The figure was reproduced from Shelnutt et al. Proc. Natl. Acad. Sci. USA 76 (1979) by permission of the copyright owner, National Academy of Science.
CARS is based on the nonlinear conversion of two laser beams into a coherent laserlike beam of high intensity in the anti-Stokes region. The essence of this technique is that the sample is illuminated by two nearly colinear laser pulses with wave number "'I and "'2' The laser with wave number "'I (pump laser) is stationary in wave number, whereas the second laser, with wave number "'2 (probe laser), is scanned. When the value (2",. - "'2) becomes coincident with the wave number of anti-Stokes Raman scattering ("'3 = "'. + Il",), the light is emitted in the forward direction at wave number "'3' The emission is often many orders of magnitude greater than with normal scattering. This relationship is explained in more detail in Figure 2.10, using an energy diagram of illuminated light, emitted light, scattered light, and the energy level of the sample. The light "'3 emitted by the CARS process is identical to the normal anti-Stokes scattering effect "'3' These relations are summarized as follows: I.
The condition of illuminating by two lasers to produce the emitted light (Figures 2.1OA-F):
'" I + ("'. - "'2) = 2'" I - "'2
= "'I + Il", = "'3 2.
The condition to produce anti-Stokes Raman line (Figure 2.1OG):
"'. + Il",
= "'3
56
The Laser Raman Spectrometer and Special Techniques
CARS Effect:
6~E~~-=-~_~:
../\j. 6W-..,.t--
•
-~r- ~T:~
±
.
--
B.
A.
C.
6~lT=== ,,
===·1-===
J\/.
WI
WI
i,
-e D.
iI
w?,= WI
+{),.W
_ F
E.
Romon Effect (Anti-Stokes Line):
:~!6W'~ •
The final result is F effect = G effect or CARS effect (W 3 )= Anti-Stokes effect (w 3 )
G.
FIGURE 2.10. Diagram illustrating the CARS effect and anti-Stokes Raman effect. By coincidence, the final results of the two processes have the same energy.
where ~W
=
WI -
W
z
Therefore, our main interest is not just to produce new emitted light - wz ), but to detect the light whose wave number is coincidentally identical to the anti-Stokes Raman line with wave number W 3 • The CARS spectrum is obtained by plotting the intensity of the emitted light against the frequency (or wave number) of scanned light W z or against ~w. In normal Raman scattering, anti-Stokes lines are weaker than Stokes lines because at normal temperatures the lower vibrational states of a molecule are more populated than the higher vibrational states. Under these conditions, more of the molecules in the population show the Stokes effect (for details, see Section 2.3, Chapter I). In the CARS process, the emission of light of
(2w I
~
Special Techniques
57
frequency identical to that of the anti-Stokes line is not subject to this rule and therefore has a higher intensity at normal temperatures. For biochemists the biggest attraction of this method is to obtain clearly resolved Raman spectra of biological samples without fluorescence interference. Readers are advised to see more-comprehensive reviews on this subject (Begley et aI., 1974; Hudson et aI., 1976; Nibler et aI., 1977; Long, 1977; Harvey, 1978; Morris and Wallan, 1979). CARS is not without technical difficulty (Tolles et aI., 1977). It is very critical to adjust the angles of the pump and probe laser beams properly. It also suffers from the presence of a large, broadbanded, nonresonant background signal (Wallan et aI., 1977; Morris et aI., 1978). 3.2.2.
Biochemical Application
Several compounds have been investigated by CARS. In resonance CARS, the spectrum is obtained in resonance with the electronic excitation. The intensity and shape of the CARS line depends on the frequency of excitation. For instance, the C= 0 stretching band of FAD (flavin adenine dinucleotide) appears at 1635 cm -I. As the excitation wI is changed from 485 to 525 nm, the shape of the 1635 cm- I line changes from positive Lorentzian through dispersion to negative Lorentzian (Dutta and Spiro, 1978). It has been difficult to obtain good spectra of certain compounds by conventional resonance Raman spectroscopy because of high levels of fluorescence. It is therefore quite an achievement that Raman spectra were obtained for FAD and glucose oxidase using the CARS technique (Dutta et aI., 1977; Dutta et aI., 1978). By this technique, even the spectrum of FMN (flavin mononucleotide) semiquinone was obtained, and it was shown that it is quite different and readily distinguishable from that of oxidized flavin (Dutta and Spiro, 1980). When coherent anti-Stokes Raman spectra of lumazine protein in the presence and absence of excess luciferase are compared, there are considerable differences in the 1264-cm -1 pyrimidine breathing mode, suggesting an interaction of lumazine protein and luciferase. Lumazine protein is a blue fluorescent protein isolated from the bioluminescent bacterium Photobacterium phosphoreum (Lee et aI., 1981). CARS spectra of cytochrome c and CO hemoglobin also show resonance Raman bands that are mainly of porphyrin vibration modes (Nestor et aI., 1976; Dallinger et aI., 1978). As the wavelength moves away from the absorption maximum, the inverse Raman effect takes place for cytochrome c and Vitamin B12 , and the dispersion curve becomes negative. This effect takes place when the solution is transparent at that wavelength (Nestor et aI., 1976). Figure 2.11 shows the CARS spectra of two compounds, light- and darkadapted bacteriorhodopsin (Tretzel and Schneider, 1979; 1980). CARS has also been applied to the interaction of acridine orange with DNA. The addition of the dye to DNA changes the CARS spectra, suggesting that the acridine orange intercalates into the spaces between the base pairs of DNA (Tretzel and Schneider, 1978).
58
The Laser Raman Spectrometer and Special Techniques
'"g.
CD N
1.0
!!!
!! ~
~Q.5 e
~ o o
2500
2000
1700 1500
Frequency (cm- I )
o
~!!! ~
1000
1100
1200
1300
1400
1500
1600
CARS Frequency(cm- I )
FIGURE 2.11. Examples of resonance CARS spectra. (A) Light-adapted bacteriorhodopsin (BRLA ); (B) dark-adapted bacteriorhodopsin (BRDA ). The figure was reproduced from Tretzel and Schneider (1979).
3.3. RAMAN OPTICAL ACTIVITY OR RAMAN CIRCULAR INTENSITY DIFFERENTIAL
Natural optical activity has long been a valuable tool for stereochemical analysis of molecular structure. In its basic application, circular dichroism measures electronic optical activity. With limited success, circular dichroism has been extended to vibrational transitions in a technique called vibrational circular dichroism (VCD). Since optical activity increases with the frequency of incident or exciting light, the vibrational (lR) optical activity is vanishingly small. Raman spectroscopy, however, retrieves vibrational information by using visible rather than IR incident or exciting light. Raman optical activity, therefore, is a promising stereochemical tool. Raman optical activity (ROA) or Raman circular intensity differential (CID) was first predicted by Barron and Buckingham in 1971. It was first observed by Barron et al. (1973a, 1973b) and confirmed by Hug et al. in 1975. In its basic application, alternating right and left circularly polarized laser radiation is incident on a sample. The Raman-scattered light from right-circular-polarized incident radiation (/R or 1+) is recorded independently from
~
References
59
that scattered from left-circular-polarized incident radiation (IL or J _). The resulting experimental parameter is A = (IR - JL)/(IR + JL). The value of A varies from positive values through zero to negative values, depending on the stereochemical features of the vibrating moiety that give rise to the Ramanscattered radiation. Because of the small value of A, signals must be accumulated until the statistical noise is reduced to approximately 10- 4 . This requires complicated equipment and long exposure times. For these reasons Raman CID has found limited use and even then in samples possessing high optical activity. The reader is directed to two recent reviews (Barron and Buckingham, 1975; and Nafie and Diem, 1979) for details on Raman CID and to the paper by Hug and Surbeck (1979) for a recent advance in instrumentation.
3.4. OTHERS Recently a new technique called surface-enhanced Raman spectroscopy (SERS) has been developed. For instance, the use of silver and copper electrodes enhances the Raman scattering of a compound. Using this technique, Raman spectra of amino acids (Nabiev et al., 1981), cytochrome c and cd! (Cotton et al., 1980; 1981), bile pigments (Lippitsch, 1980; 1981), adenine mononucleotides (Koglin et al., 1980a, b; 1981) and nucleic acids (Sequaris et al., 1981) have been obtained.
REFERENCES Barron, L. D., and Buckingham, A. D. (1971). Rayleigh and Raman scattering from optically active molecules. Mol. Phys. 20, 1111. Barron. L. D., and Buckingham, A. D. (1975). Rayleigh and Raman optical activity. Ann. Rev. Phys. Chern. 26, 381. Barron, L. D., Bogaard, M. P., and Buckingham, A. D. (l973a). Differential Raman scattering of right and left circularly polarized light by asymmetric molecules. Nature 241, 113. Barron, L. D., Bogaard, M. P., and Buckingham, A. D. (1973b). Raman scattering of circularly polarized light by optically active molecules. J. Arn. Chern. Soc. 95, 603. Begley, R. F., Harvey, A. B., Byer, R. L., and Hudson, B. S. (1974). Raman spectroscopy with intense, coherent, anti-Stokes beams. J. Chern. Phys., 61, 2466. Bloembergen, N. (1975). Lasers: A renaissance in optics research. Am. Sci. 63, 16. Brown, R. (1968). Lasers, Tools of Modern Technology, Doubleday, Garden City, N.Y. Cotton, T. M., Schultz, S. G., and Van Duyne, R. P. (1980). Surface-enhanced resonance Raman scattering from cytochrome c and myoglobin adsorbed on a silver electrode. J. Am. Chern. Soc. 102, 7960. Cotton, T. M., Timkovich, R. and Cork, M. S. (1981). Resonance Raman and surfaced-enhanced resonance Raman studies of cytochrome cdi'> FEBS Lell., 133, 39. Cunningham, R. C. (1974). Dye lasers today-and tomorrow? Electrical-Optical Systems Design, pp. 13-18.
60
The Laser Raman Spectrometer and Special Techniques
Dallinger, R. E, Nestor, 1. R., and Spiro, T. G. (1978). Resonance coherent anti-Stokes Raman scattering evidence for out-of-plane heme iron displacement within 6 ns of CO dissociation in CO hemoglobin. J. Am. Chem. Soc. 100,6251. Dutta, P. K., and Spiro, T. G. (1978). Resonance CARS line shapes: Excited state parameters for flavin adenine dinucleotide. J. Chem. Phys., 69, 3119. Dutta, P. K., and Spiro, T. G. (1980). Resonance coherent anti-Stokes Raman scattering spectra of oxidized and semiquinone forms of Clostridium MP flavodoxin. Biochemistry 19, 1590. Dutta, P. K., Nestor, J. R., and Spiro, T. G. (1977). Resonance coherent anti-Stokes Raman scattering spectra of fluorescent biological chromophores: Vibrational evidence for hydrogen bonding of flavin to glucose oxidase and for rapid solvent exchange. Proc. Nat. Acad. Sci., 74, 4146. Dutta, P. K., Nestor, J., and Spiro, T. (1978). Resonance coherent anti-Stokes Raman scattering (CARS) spectra of flavin adenine dinucleotide, riboflavin binding protein and glucose oxidase. Biochem. Biophys. Res. Comm. 83, 209. Fox, J. W., and Tu, A T. (1979). Sample heating apparatus for Raman spectroscopy. Appl. Spectrosc. 33, 646. Harvey, A B. (1978). Coherent anti-Stokes Raman spectroscopy. Anal. Chem. SO, 905A Hellman, H. (1969). Lasers, U.S. Atomic Energy Commission, Washington, D.C. Hudson, B. S. (1977). New laser techniques for biophysical studies. Ann. Rev. Biophys. Bioeng 6,
135. Hudson, B., Hetherington, III, W., Cramer, S., Chabay, I., and Klauminzer, G. K. (1976). Resonance enhanced coherent anti-Stokes Raman scattering. Proc. Nat. Acad. Sci. 73, 3798. Hug, W., and Surbeck, H. (1979). Vibrational Raman optical activity spectra recorded in perpendicular polarization. Chem. Phys. Lett. 60, 186. Hug, W., Kint, S., Bailey, G. F., and Scherer, J. R. (1975). Raman circular intensity differential spectroscopy. The spectra of (- )-a-phenylethylamine. J. Am. Chem. Soc. 97, 5589. Kiefer, W. (1977). Recent techniques in Raman spectroscopy. In Advances in Infrared and Raman Spectroscopy, Vol. 3, R. H. 1. Clark and R. E. Hester, Eds., Heyden, Bellmawr, N.1. Kimel, S., and Speiser, S. (1977). Lasers and chemistry. Chem. Rev. 77, 437. Koglin, E., Sequaris, 1. M., and Valenta, P. (1980a). Surface Raman spectra of nucleic acid components adsorbed at a silver electrode. J. Mol. Struc. 60,421. Koglin, E., Sequaris, J. M., and Valenta, P. (1980b). Surface enhanced Raman spectra of adenine mononucleotides adsorbed at a silver electrode. In Proc. VlIth Int. Conf. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam, p. 628. Koglin, E., Sequaris, 1. M., and Valenta, P.(l981). Hydrogen-deuterium exchange in adenosine 5'-monophosphate detected by surface enhanced Raman scattering (SERS). Z. Naturforsch., C: Biosci. 36, 809. Laane, J., and Kiefer, W. (1981). Applications of four-channel Raman difference spectroscopy. Appl. Spectrose., 35, 428. Lee, J., Carreira, L. A, Gast, R., Irwin, R. M., Koka, P., Small, E. D., and Visser, A 1. W. G. (1981). Bioluminescence and Chemiluminescence, Academic, New York. Lippitsch, M. E. (1980). Surface enhanced Raman spectra of bile pigments. In Proc. VIIth Int. Conf. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 630-633. Lippitsch, M. E. (1981). Surface enhanced Raman spectra of biliverdin and pyrromethemone adsorbed to silver colloids. Chern. Phys. Lett. 79, 224. Long, D. A (1977). Raman Spectroscopy, McGraw-Hili, London. Morris, M. D., and Wallan, D. J. (1979). Resonance Raman spectroscopy. Anal. Chem. 51, 182A
110..
References
61
Morris, M. D., Wallan, D. 1., Ritz, G. P., and Haushalter, J. P. (1978). AC-coupled inverse Raman spectroscopy. Anal. Chern. SO, 1796. Nabiev, I. R., Trakhanov, S. D., Efremov, E. S., Marinyuk, V. V., and Lazorenko-Manevich, R. V. (1981). Anomalously intense Raman spectra of some biological molecules adsorbed on silver electrodes. Bioorg. Khirn. 7, 941. Nafie, I.. A and Diem, M. (1979). Optical activity in vibrational transitions: Vibrational circular dichroism and Raman optical activity. Accts. Chern. Res. 12,296. Nestor, 1., Spiro, T. G., and Klauminzer, G. (1976). Coherent anti-Stokes Raman scattering (CARS) spectra, with resonance enhancement of cytochrome c and vitamin B I2 in dilute aqueous solution. Proc. Nat. Acad. Sci. 73, 3329. Nibler,1. W., Shaub, W. M., McDonald, 1. R., and Harvey, A B. (1977). Coherent anti-Stokes Raman spectroscopy. In Vibrational Spectra and Structure, Vol. 6, J. R. During, Ed., Elsevier, Amsterdam, Oxford, and New York, pp. 173-225. Rousseau, D. I.. (1981). Raman difference spectroscopy as a probe of biological molecules. J. Rarnan Spectrosc. 10, 94. Sequaris, J. M., Koglin, E., Valenta, P., and Nuernberg, H. W. (1981). Surface-enhanced Raman scattering (SERS) spectroscopy of nucleic acids. Ber. Bunsenges. Phys. Chern. 85, 512. Shelnutt,1. A, Rousseau, D. 1.., Dethmers, J. K., and Margoliash, E. (1979). Protein influence on the heme in cytochrome c: Evidence from Raman difference spectroscopy. Proc. Nat. Acad. Sci. 76, 3865. Teller. E. (1973). Lasers in chemistry. In Proc. Robert A. Welch Found. Conf. Chern. Res., XVI, Theoretical Chemistry, W. O. Milligan, Ed., Houston, Tex., p. 205. Tolles, W. M., Nibler, 1. W., McDonald, J. R., and Harvey, A B. (1977). A review of the theory and application of coherent anti-Stokes Raman spectroscopy (CARS). Appl. Spectrosc., 31, 253. Tretzel, 1., and Schneider, F. W. (1978). Resonance CARS spectroscopy of biological systems intercalation of DNA by acridine molecules. Proc. Sixth Int. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 478-479. Tretzel, 1., and Schneider, F. W. (1979). Resonance CARS spectroscopy of bacteriorhodopsin. Chern. Phys. Lett. 66, 475. Tretzel, J., and Schneider, F. W. (1980). Resonance CARS spectroscopy of bacteriorhodopsin. In Proc. Vllth Int. Conf. Rarnan Spectrosc., w. F. Murphy, Ed., North-Holland, Amsterdam and New York. Wallan, D. 1., Ritz, G. P., and Morris, M. D. (1977). Adjustable angle beam crossing system for coherent anti-Stokes Raman spectroscopy. Appl. Spectrosc. 31, 475. Woodruff, W. H., and Farquharson, S. (1978). Seven-nanosecond time-resolved resonance Raman spectrometry of cytochrome c. Anal. Chern. SO, 1389.
(0'
en -
-0 CD D.>
e --
o
CDo
o~(l] ___ 0--
CHAPTER
Proteins
Proteins and synthetic polypeptides consist of amino acids joined together by peptide bonds. In the study of the conformation of proteins, one must look at two aspects. One is the peptide-backbone structure or secondary structure. The :other is the arrangement of the many side chains in a protein, or tertiary :~tructure. Of course, at times there is no clear dividing line between secondary .·and tertiary structures. There are several typical peptide-backbone arrange·ments such as the a-helix, ,B-sheet, ,B-turn, y-turn, w-turn, and random coil. Although no one physical method can elucidate all the structural aspects, X-ray diffraction is the most powerful. From it one can unequivocally deduce the absolute conformation of protein molecules in crystals. However, X-ray diffraction is only applicable to crystallized substances, and practically all protein molecules in biological systems are in aqueous solution. Thus a question always arises whether the conformational structure of a protein in the natural state is identical to the structure deduced by the X-ray diffraction method. Fluorescence spectroscopy is another analytical tool for studying proteins. Yet this method is incapable of determining peptide-backbone structure. Fluorescence spectroscopy is rather limited to the study of the environments of tyrosine and tryptophan side chains. Circular dichroism (CD) is a powerful physical methods for the study of protein structure, especially the
66
Proteins
peptide backbone. CD, however, does not provide much structural information concerning side chains or disulfide bonds. To their disadvantage, both fluorescence and CD spectroscopies require aqueous solutions. Raman spectroscopy overcomes most of these objections and recently has become a powerful tool for studying proteins. Raman spectroscopy can be carried out with crystals, powders, gels or-unlike IR-aqueous solutions. Recently, using a highly sophisticated IR spectrometer equipped with a computer, it has been demonstrated that the spectra can be obtained in aqueous solution by subtracting the intensity of the water blank. Raman spectroscopy provides information on the peptide backbone, geometry of disulfide bonds, and the environment of some side chains such as those of tyrosine, tryptophan, and methionine. Raman spectroscopy can detect the presence of disulfide bonds, methionine residues, and sulfhydryl groups (-SH); such detection involves tedious procedures when done by conventional chemical methods.
1.
PEPTIDE-BOND VIBRATIONS
How can Raman spectroscopy be used to analyze the secondary structures of proteins? What is the theoretical basis for these procedures? The peptide bond -CONH- is nearly planar because of resonance stabilization. The bonds'" and ep on both sides of a peptide bond are fixed for
.<
FIGURE 3.1. The rotation of a peptide backbone is restricted to the C(a)-C and N-C(a) bonds. Spatial orientation of the peptide backbone (conformation) is determined by the angles of rotation of '" and
j ':i\
Peptide-Bond Vibrations
67
a particular conformation and ultimately affect the vibrational modes of a peptide bond (Figure 3.1). A peptide bond gives rise to many different types of vibrational modes such as amide A and B bands, amide I, II, III, IV, V, VI, and VII bands. Among these, the amide I and III bands yield very prominent Raman bands that are correlated with structural properties of protein molecules (Peticolas et al., 1981).
1.1.
AMIDE A AND B BANDS
An overtone is a multiple of a fundamental vibrational frequency. If anharmonicity is present, new bands involving overtone and combination transitions can appear in the spectrum but usually are weak. Sometimes, however, the frequency of an overtone or combination band may have nearly the same value as the frequency of another fundamental. In such a case it may happen that two relatively strong bands may be observed where only one strong band for the fundamental was expected. These are observed at somewhat higher and lower frequencies than the expected unperturbed positions of the fundamental and overtone. This effect is called Fermi resonance. The apparent intensification of the overtone results from the fact that the fundamental is involved in both bands. The amide A and B bands arise from the Fermi resonance between
3236 3134
em' em'
.
N-H s tretchi ng (fundamenta 1 vibrational frequency)
t
2xamlde II
-----.......
overtone
1567
amide A
,.,
amide B
3280
em'
3090
em1
em1 - - - - - - - - - amide II
ground state FIGURE 3.2. Origins of the amide A and B bands, The 3134 cm - I band is twice the frequency of the amide II fundamental vibrational band (overtone). The N-H stretching vibration appears at 3236 cm -I, which is very close to the value of the amide II overtone band at 3134 cm -I, The result is the interaction of two vibrational levels; one appears at a somewhat higher (3280 cm - 1 amide A band) and the other at a lower frequency (3090 cm - I amide B band). This effect is called Fermi resonance.
68
Proteins
the first excited state of the N-H stretching vibration and the second excited state (overtone) of the amide II vibration (Figure 3.2) 1.2.
AMIDE, I, II, AND III BANDS
Amide I, II, and III bands arise from in-plane vibration of the peptide bond. Different modes of vibration (Figure 3.3) are responsible for each of these bands. 1.2.1.
Amide I Band
The main contribution to the amide I band (Table 3.1) is the peptide carbonyl stretching vibration, although there is a small contribution from the N - H in-plane bending vibration. There are many functional groups that give C=O stretching vibrations, The amide I vibrational mode is the C=O of -CONH-, and therefore it is different from the C=O vibration of -COOH (carboxylic acid) or -COOR (ester). For instance, free proline has a C=O stretching vibration of -COOH that appears at 1615 em-I. In poly (L-Pro) the C=O stretching vibration of -CONH- (amide I band) appears at 1650 em - I (Smith et aI., 1969). The ester C=O vibration usually appears at a
R
--8
\f
/
/
\
C-N
o
;
R
Amide I
R
'8
\ / +-C-N / ~R o
1
, ..
j
\R
~c-N-
Amide II
Amide III
FIGURE 3.3. The in-plane vibrational modes of the peptide bond. Among the three modes shown in the figure, the amide I and III bands in the Raman spectra are indices of the peptidebackbone conformation of a protein. The amide II mode is either Raman inactive or very weak.
Peptide-Bond Vibrations
TABLE 3.1.
69
Amide I Band for Synthetic Polypeptides
Frequency
Compound
Reference
a-Helix Right Handed 1632 1645 1652 1652 1653 1654 1656 1657 1657 1660 Left Handed 1663
T.-I. Yu et aI. (1973) T.-I. Yu et aI. (1973)
PolY(L-Lys) (in D 2 O) PolY(L-Lys) PolY(L-Glu) Poly( y-benzy1-L-G1u) Poly(L-Leu, Glu) random copolymer Poly(L-G1y)II Po1y(L-AJa) Poly(L-Ala) Poly(L-Lys) (H 2 O) PolY(L-Ala) (fibers)
Frushour and Koenig (1974) SmaIl et aI. (1970) Frushour and Koenig (1974) Fanconi et aI. (1969) Painter and Koenig (1976a) Fanconi et aI. (1969)
Poly( p-benzyl-L-Asp)
Frushour and Koenig (1975c)
F asman et aI. (1978b) Fasman et aI. (1978b)
p-Sheet Right Handed 1658 1669 1670 1671 1672 1674 Left Handed 1679 1660 1665 1665
PolY(L-Lys) Poly(L-Ala) PolY(L-Lys) PolY(L-Lys) PolY(L-Lys) Poly(L-His) Poly(Gly)I
(in D 2 O)
T.-I. Yu et aI. (1973) Frushour and Koenig (1974)
T.-I. Yu et aI. (1973) (H 2 O)
Po1y(p-benzyl-L-Asp)
Random Coil PolY(L-Lys) (in D 2 O) PolY(L-Ala) PolY(L-Glu)
Painter and Koenig (1976a) WaIlach et aI. (1970) Ashikawa and Itoh (1979) SmaIl et aI. (1970) Frushour and Koenig (1975c)
T.-I. Yu et aI. (1973) Frushour and Koenig (1974) Lord and Yu (1970a)
higher frequency, in the vicinity of 1700 cm -I. Because the amide I band originates from the peptide bond C=O stretching vibration, it should not reach a frequency of 1700 cm- I. For instance, an antibiotic peptide, valinomycin, contains ester as well as amide bonds. It shows the ester C=O stretch at 1767 and 1742 cm- I and the amide c=o stretch at 1675 and 1649 cm- J (Asher et al., 1974). Similarly, the cyclic peptide antibiotics nonactin, monactin, and dinactin contain ester groups and have C=O stretching vibrations at 1726, 1725, and 1726 cm - 1, respectively (Phillies et aI., 1975).
70
Proteins
Because of the major contribution of C=O, deuterium exchange does not shift the amide I band significantly. ""
/H
~C-N
O~
D 20
"
/D
~C-N
O~
"
"-
Actual examples of amide I before and and after D 20 treatment are shown here: Original Amide I (em-I)
Protein
After Frequency Deuteration Shift (em-I) (em-I)
Reference
Myotoxin a Lapemis hardwickii toxin Oxytocin a-Chymotrypsin
1674
1668
6
Bailey et aL (1979)
1675 1666 (solid) 1669
1668 1663 1664
7 3 5
Histone 3
1663
1657
6
Histone 4
1663
1655
8
Fox et aL (1979) Tu et aL (1978) Lord and Yu (1970a) Pezolet et aL (1980) Pezolet et aL (1980)
By comparing the rates of deuteration of various amides, it was found that the a-helical regions of lysozyme are more accessible to solvent than the j3-sheet regions (Peticolas et aI., 1979). 1.2.2.
Amide III Band
The torsional angles (Ramachandran angles) of tf; and ep determine the conformation of peptide-backbone structure. To establish the correlation between these angles and the Raman frequency p theoretically is not easy. However, from the experience of X-ray crystallographic research, it is known that the tf; angle is the parameter most indicative of a- and j3-conformations by amide III frequency (Richards and Wyckoff, 1971; Lord, 1977; Chen and Lord, 1980). The deuterium-exchange reaction shows significant isotopic effect on the amide III band because the major contribution originates from the N - H vibration. Usually the amide III band shifts to a lower frequency approximately by the factor of {f = 104. This method is important in ascertaining the amide III band which is located in a region of highly mixed vibrational modes. For determining the peptide backbone conformation of proteins, it is essential to ascertain the amide III band by deuteration.
Peptide-Bond Vibrations
71
Examples of the isotopic shift of the amide III band are shown here:
Protein
Original After Amide III Deuteration (em-I) (em-I) Ratio
Reference
Lapemis hardwickii toxin Enhydrina schistosa
1240
980
1.27
Yu et al. (1975)
toxin Oxytocin Chymotrypsinogen
1242 1260 1241
980 994 961
1.27 1.27 1.29
Yu et al. (1975) Tu et al. (1978) Koenig and Frushour (1972b)
Recently it was proposed that the amide III' of D 2 0-exchanged protein obtained by subtracting the spectrum from the corresponding spectrum in water is better resolved than the amide I band (Williams et al., 1980).
1.2.3.
Amide II Band
The amide II band is usually weak in Raman spectra or is not shown at all. The amide II band is very strong in IR spectra, and contains useful structural information complementary to that obtained from Raman spectroscopy. However, by the use of a laser line of low wavelength, namely 257.3 nm, the amide II band at 1555-1560 cm- I can be observed in Raman spectra of N-ethylacetamide and N-methylpropionamide (Harada et al., 1975).
1.3.
AMIDE V, VI, AND VII BANDS
These vibrational bands are associated with out-of-plane vibrations that are antisymmetrical with respect to the CONH plane (Figure 3.4). These bands appear in IR and Raman spectra, but their Raman intensities are weak.
1.4.
AMIDE IV BAND
This is basically due to the bending mode of the CONH group. The amide IV band is not restricted to the peptide group, and it appears in IR spectra at 647 cm-I for N-methylacetamide. It is rarely identifiable in either the IR or Raman spectra of polypeptides.
72
Proteins
-0.02M
H+0.56 / +0.16 C - N -0.15 '-...
/l
-0.04 0 +0.05M
'-...
-0.20· C ~
+0.07 0 -0.11 M
'-...
+0.06 C +0.04 0
~
'-...
Amide V
M +0.04
H+0.61 / N -0.03
Amide VI
'-...
M +0.02 H+0.46 / N +0.13
Amide VII
'-...
M-0.14
FIGURE 3.4. Three out-of-plane vibrations that are antisymmetrical with respect to the CONH plane. The + indicates that the atom vibrates upward, and - indicates downward vibration. The uni ts are in angstroms.
2.
SECONDARY STRUCTURE (PEPTIDE-BACKBONE STRUCTURE)
Proteins can form different types of helices such as the a-helix, 310 helix, and fJ-sheet. In general these helices are formed through intramolecular hydrogen bonding between the carbonyl oxygen of one peptide bond and the hydrogen atom of another. A hydrogen bond is formed with every fourth peptide bond for the a-helix, and every third peptide bond for the 310 helix (Figure 3.5). Synthetic polypeptides are much simpler in structure than natural proteins. Proteins usually consist of nearly all 20 amino acids in varying proportions. Moreover, many synthetic polymers can form different conformations precisely by varying the conditions of pH, ionic strength, type of salts, temperature, or degree of polymerization. Much of the basic information on the Raman spectroscopy of proteins was actually obtained by studying synthetic peptides first.
o
o
0
0
0
II II II II II -C-N-C HR1-C N-CHR2-C-N-CHR3-C-N-CHR4-C-NI I I I I H H H H H
y-turn
~-turn
a-helix
rr-helix
(J,o helix)
FIGURE 3.5.
Hydrogen bonding in the y-turn, {l-turn, a-helix, and 'IT-helix.
Secondary Structure (Peptide-Backbone Structure)
2.1.
73
CONFORMATIONAL ANALYSIS FROM AMIDE I AND III BANDS
2.1.1 . a-Helix
A protein with an a-helical structure has a low amide I band at 1645-1657 cm- 1 (Table 3.1). Values as high as 1660 cm- 1 were reported for poly (L-Ala), but it is rare for the band from an a-helix to occur that high. The amide III band is extremely high (1264-- 1300 cm- I ) (Table 3.2) and it is relatively easy to detect a protein with high a-helical content. The range of the amide I and III bands for different conformations is summarized in Figure 3.6. The intensities of amide I and III bands are sometimes related to protein conformation, although this is not as clear as the relationship to differences in frequency. A hyperchromic shift in the intensity of the amide III line accompanying the helix-to-coil transition has been reported (Koenig and Frushour, 1972a). Zinc insulin was well studied by X-ray crystallographic analysis. In this molecule, a-helical structure exists in the central part of the B chain and between residues A 2-6 and A 13-19. Antiparallel-pleated-sheet interactions are involved in the formation of the insulin dimer. The rest of the peptide groups have unordered structure. Therefore, it is of interest to correlate Raman data with X-ray data. The amide I Raman frequencies of the zinc-insulin crystal are at 1662 and 1680 cm - I. The 1662-cm- I band is assigned to a-helical structure and the 1680-cm- 1 band to the random-coil form (Yu et al., 1972a, b). The 1662-cm ~ I band is rather high for a-helix of a synthetic polypeptide, but the shift to high frequency can be explained in terms of TABLE 3.2.
Amide III Band Frequency for Synthetic Polypeptides
Frequency (cm - I)
Compound
Reference
a-Helix (crystal)
1264 1274
POlY(L-Ala) Poly(L-Ala)
1296 1296
POlY(L-Glu) Poly( y-benzyl-L-Glu)
1230 1234 1236 1240 1243
f12- Poly(L-Glu) Poly(Gly)I f11- Poly(L-Glu) POlY(L-Lys) POlY(L-Ala)
1242 1243
POlY(L-Ala) POlY(L-Lys)
Fanconi et al. (1969) Frushour and Koenig (1974) Fanconi et aI. (1969) Fasman et aI. (l978b) Fasman et aI. (1978b)
f1-Sheet Fasman et al., (I 978b) Small et al. (1970) Fasman et aI., (I 978b) T.-I. Yu et aI. (1973) Frushour and Koenig (1974)
Random Coil Frushour and Koenig (1974) T.-J. Yu et aI. (1973)
74
Proteins
a-helix
.a-sheet
l3-tu r n
random coli
1680
1660
AMIDE
16"40
I
I
I
1300
1260
AMIDE
1220
III
FIGURE 3.6. Amide I and III frequencies and different types of protein conformations. There are always exceptions in band frequencies, but the ranges shown in the figure usually cover the majority of bands for a given conformation.
weakness of hydrogen bonding to the carbonyl group. Oriented-fiber poly (L-Val) has a high content of a-helix, as can be seen from the amide I band at 1655 cm -I. Low- and high-molecular-weight poly (L-Val) treated with trifluoroacetic acid are in the ,B-sheet conformation, and the amide I shifts to 1676 cm- I with a low amide III band at 1233 cm- I (Yamashita and Yamashita, 1975). 2.1.2.
,B-Sheet
Vibrational modes of antiparallel-,B-sheet structure are of four different types (Figure 3.7). Calculated amide I and III frequencies of ,B-sheet structures of polY(L-Ala), polY(L-Ala-Gly), and poly(Gly)1 are extremely close to the values observed experimentally (Moore and Krimm, 1976a, b) (Table 3.3). PolY(L-Lys) shows Raman amide III band hypochromism while undergoing a transition from the disordered conformation to a-helix and shows Raman hyperchromism when the antiparallel-,B-sheet conformation is formed (Painter and Koenig, 1976b). Detergents, usually having both polar and nonpolar groups within a molecule, bind to proteins and change their conformation. Dodecylpyridinium bromide is a detergent that alters the conformation of ,B-1actoglobulin. The Raman spectrum of ,B-lactoglobulin has three bands or shoulders corresponding to the amide III region at 1266, 1242, and 1235 cm- I . When the protein binds to the detergent, the 1266 cm- I band disappears, whereas the 1242 and
Secondary Structure (Peptide-Backbone Structure)
75
Antiparallel II-sheet Chains
6II 0
i
H ,
a
~I
",
"N,C....C'N....C... C" I o"'r.L\ Io±I H: : :
"'0 1
a
o -
:
6
~r.L\
a
H
lI I
11o±I,,,
a
6II",0
a
"N'C"C"'N"C,C"
1o±I,,,0
H r.L\
a
,
"c c.... N...C"C... N"
"C"'C"N"'C"C"'N"
0-
,........ "
"0
a
,
H i
:
~
:
:
v11l",01
~
6II.....0
a
"N,C C...N....C C"
o" I I
H
: :
v[O,OI
~,
I
a
0
,... I
"'C\ ~0 b
a
~H
6 o
, I
':'0 b
OHI
a
' ..... " "Ca C.... N...C"C'N"
"" "Ca C.... N...C"C...N"
,...
0-
~
"
a
0
:
,
':I
"0
I
a
H :
i
;
0
I a "'" "N"'C"C"'N"C...C"
vla,rrl
vl 7T ,:rr1
FIGURE 3.7. Vibrational modes of a peptide bond of antiparal1e1-,B-sheet structure. The figure was redrawn based on Miyazawa (1960).
1235 bands are shifted slightly to 1247 and 1230. This indicates that the cationic detergent causes a decrease in a-helical content and probably stabilizes the ,a-sheet conformation (Wasylewski, 1979). When a gel is formed, there is a tendency for the ,a-sheet content to increase with a simultaneous decrease in a-helical content. This change can be detected by Raman spectroscopy (Clark et aI., 1981). From X-ray diffraction and CD evidence, it is known that pancreas chymotrypsinogen A, pancreas RNase, and egg white ovalbumin have a low a-helical content but are high in unordered and ,a-sheet conformations. The Raman frequencies of amide I and III bands are consistent with this evidence TABLE 3.3. Observed and Calculated Frequencies of ~-Sheet Synthetic Polypeptides
PolY(L-Ala) Vibrational obsd. Type
11(0,0) 11(0, 'IT) II( 'IT, 0) II( 'IT, 'IT)
1669 1695 1630 -
PolY(L-Ala-Gly)
ealed.
obsd.
(em~l)
ealcd.
Poly(Gly)I obsd.
1669 1695 1630 1701
1665 1702 1630 1693
ealed. (em-I)
(em-I)
1665 1702 1630 1694
1674 1685 1636 -
1674 1685 1637 1690
76
Proteins
(Koenig and Frushour, 1972a). These frequencies are shown here: Amide I (cm- 1)
Amide III (cm- I )
Chymotrypsinogen (solid) RNase (H 2O)
1668 1664
RNase (solid)
1665
Ovalbumin
1665
1241 1239 1261 1235 1258 1245
Protein
If the proteins were a-helical in configuration, they would be expected to have low amide I (less than 1650 cm- I) and high amide III bands. 2.1.3.
,8-Turn (,8-Bend, ,8-Reverse Turn)
,8-Turn is also called 310 helix because it represents three amino acid residues and 10 atoms involved per turn in which a hydrogen bond closes a ring (Figure 3.8). Compared to a-helical and ,8-sheet structures, ,8-turn structure has not been studied extensively. Any structure not belonging to a-helical and ,8-sheet is frequently considered to be random coil. Recently, however, it has been recognized that reverse-turn structures such as ,8-turns and y-turns constitute important conformational components of protein structure and play an important role in protein-structure determination. This is particularly the case in regions where the peptide backbone folds back upon itself. There is relatively little Raman data on this subject, but some compounds known to have ,8-turns have been studied and are summarized in Table 3.4.
J ~Y ~
C'
/ N
1
Gf-" @:
ref? i
\
~~~ N
~, e,)H
.
I
:I
~
cif~~~ I
]I
FIGURE 3.8. Two basic types of p-turn conformation. The figure was reproduced from Dickerson et al. (1971) by permission of the copyright owner, American Society of Biological Chemists.
- . - - - - - - - . - - - - - - - - - •• -
•• - _ •• _ - -
•••• - - _ . -
Amide I (em-I)
Type of {1-Tum
Oxytocin
1663-1666
-
Amide III (em-I)
:;:11
r- . _ ••• _
Reference for Raman
Type II 1260-1260 Tu et aI. (1978)
Pro-Leu-GlyNH 2
1649 1688
1238 1266 1283
s- Benzyl-Cys-
1644
1235 1261 1294
Pro-Leu-GlyNH 2
- - - - - - _ •• - -
Hseu and Chang (1980), Fox et aI. (1981) Fox et aI. (1981)
A Mixture of Type 1I {1-Turn and {1-Sheet 1668 1240 Fox et aI. ({1-Sheet) (1981) 1258 1273 1293
Gramicidin S
__ .._--
Evidence of {1-Tum
NMR
Xray
~. " . . .' ." ~. .,:",:~ :,: :;.~(;.:.·.:.7,.,. ,.·i.;.:~.'••:,>.:.~,.~.3.~.;r.~:.~~.~,;~ - ...~.~.·~~~,,~'t.~**.::,~.~./!:.:/~.tt'.;\··,.:.;·.;.;,>(R ef erence for {1-Turn
Brewster et aI. (1973), Ballardin et aI. (1978) Reed and Johnson (1973)
Xray
Rudkoand Low (1975)
X ray
Hodgkin and Oughton (1975), Stem et aI. (1968)
NMR
VenkatachaIapathi and BaIaram (1982) ShamaIa et aI. (1977) Prasad et aI. (1979)
TypeIlr BOC-Cys-Pro-VaI-Cys-CONHCH 3 I
I
S
S
Z-Aib-Pro-AibAla-COOCH 3 Z-Aib-Pro-CONHCH 3
aROC_
terti~rv hlltvlnxvr.~rhnnvl prnnn°
7.
1668
1267
1665
1265
1667
1286
hf"n7vlnYVr~rhnnvl arnl1n
Ishizaki et aI. (1982) Ishizaki et aI. (1982) Ishizaki et aI. (1982)
Xray Xray
78
Proteins
Experimental results are often within the range of calculated frequencies, most consistently for the amide I band (Bandekar and Krimm, 1979a, b; Krimm and Bandekar, 1980). For the amide III band, some frequencies fall within the calculated range, although none of the experimental data appear as high as 1300 cm -1 (Han et aI., 1980). Many amide III bands are actually lower than the calculated values. For gramicidin S, t,here are four amide III bands at 1240, 1258, 1273, and 1293 cm -1. The 1240-cm -1 band is associated with ,8-sheet structure, as gramicidin S is known to contain both ,8-sheet and ,8-turn structure. Pro-Leu-Gly-NH 2 is a carboxamide C-terminal tripeptide from oxytocin; the tripeptide is known from X-ray analysis to possess a type II ,8-turn (Reed and Johnson, 1973). With the exception of a band at 1238 cm- I , all band frequencies are close to the calculated values (Hseu and Chang, 1980; Fox et aI., 1981). It seems that there is a fairly wide range in values of the amide III band, but it is definitely higher than those of ,8-sheet and random coil. Thus Raman spectroscopy can provide diagnostic evidence for ,8-turn structures by careful analysis of both amide I and III bands. As more model compounds having ,8-turn are examined by Raman spectroscopy, more-precise information on the amide I and III band frequencies will be established. Although the Raman data on the ,8-turn are relatively few, the ranges of the amide I and III bands so far investigated are summarized in Figure 3.6. 2.1 .4.
Omega-Helix
Poly(,8-benzyl-L-Asp) can form the w-helix (left handed) from the left-handed a-helix by heating at 160°C under vacuum. The amide I band appears at 1675 cm- 1 in Raman and at 1676 cm- I in IR spectroscopy (Frushour and Koenig, 1975b). This is the only investigation on omega-helix; therefore, the correlation between Raman data and the omega-helix is not definite at this stage. 2.1.5.
y- Turn
The only y-turn-containing peptide investigated by Raman spectroscopy is [l-penicillamine]-oxytocin. This compound has been shown by nuclear magnetic resonance (NMR) to contain y-turn (Meraldi et aI., 1977). Therefore, it is premature to draw any definitive conclusions on its relationship to Raman data. For solid samples of [I-penicillamine] oxytocin, amide I and III bands appear at 1668 and 1255 cm- I , respectively. For aqueous samples, the amide I band appears at 1656 and 1666 cm ~ 1, whereas the amide III band appears at 1255 cm -1 (Hruby et aI., 1978). 2.2.
COMPARISON OF AMIDE I AND III BANDS
The amide III band is more sensitive to structural changes than the amide I band. Lysozyme has a-helix in residues 5-15, 24-34, and 88-96; antiparallel ,8-sheet in residues 41-45 and 50-54; and random coiled structure. These three different conformations are reflected in the amide III bands at 1272, 1238, and
Secondary Structure (Peptide-Backbone Structure)
79
1258 cm- I , respectively. However, only one distinct amide I band at 1660 cm- I can be observed (Yu and Jo, 1973a). The best and most accurate way to determine protein conformation is by the combined use of both amide I and amide III bands. It is risky to determine conformation based solely on the amide III band, since this region is a highly mixed vibration zone. The amide III band can be accurately determined by dissolving protein in D2 0 to allow isotopic exchange. The deuterated protein should give a new amide III' band in the vicinity of 980 cm -I, and any bands not shifted in the region of 1200-1300 cm- I are not amide III bands. The amide I band usually has less interference, but even so, the amide I band is an average over the different conformational distributions. For instance, the amide I band of native feather keratin can be resolved into two components (Hsu et al., 1976). With the simultaneous use of both amide I and III bands, it is possible to differentiate a-helix, ,B-sheet, ,B-turn, and random coil, where any structure that does not belong to the first three structures is grouped together as random coil, or disordered structure. These relationships are shown in Figure 3.6. a-Helix, ,B-sheet, and random-coil structures have been extensively studied by many investigators; thus good correlations exist between the Raman spectroscopic data and their respective structures. Only a few peptides with ,B-turn have been investigated by Raman spectroscopy. The investigation of more compounds is necessary to make more precise correlations between ,B-turn structure and the amide I and III band frequencies. Hydrogen bonding involving C=O exerts a great effect on the frequency of the amide I and III bands. When a peptide-bond C=O is involved in hydrogen bonding, it decreases the amide I band frequency, whereas it increases the amide III band frequency. Cis and trans amide bonds have different frequencies. In proteins all peptide bonds are in the trans form. Occasionally a glutamic acid residue at the N-terminal forms an amide bond to become a pyroglutamyl residue. In such a case, the amide bond is in the cis form. Thyroid releasing factor, a hormone, is a very interesting peptide from the viewpoint of the amide bond (Bellocq et al., 1973). The sequence is pGlu-His-ProNH 2 • As can be seen from the chemical structure, the hormone contains four amide bonds (Figure 3.9). One is the pyroglutamyl bond, which is in the cis form; the glutamyl-histidine bond is in the trans form; the histidyl-proline bond is an example of a tertiary amide bond, and the carboxy-terminal amide is a primary amide bond. The frequencies for these amide bonds are different and are shown here. Amide Bond cis-form amide trans-form amide tertiary amide primary amide
In H 2 O (cm- I )
In D 2 0 (cm- I )
1677 1660 1640 1677
1663 1640 1640 1640
80
Proteins pGlu-His-Pro NH
r--u---;
~2
0,... " ....C .{ I
Z
N~N
I
~2
I
" CH 2 CH2,.----' CI H2C/ II H, CH2 I I' I ,.---, ,.-----, 1 I ' N ;C 'c I 'C NH2: :______."' H...N....--CH-\Cr- C .11, H,,II - N I H..,II I
I
I'
'L 0 .J' cis form
trans form
I
'0
1.
I
",I
tertiary form
L .0
.JI
primary form
FIGURE 3.9. Structure of pGlu·His·Pro·NH 2 .
The cis-form and primary-amide-bond vibrational modes appear at the same frequency-1677 cm - 1 - by coincidence, and these are separated in DzO. The assignment was made from N-acetylprolinamide, which has bands at 1677 and 1615 cm- I • These bands shift to 1650 and 1610 cm- I in DzO. The 1677-cm- 1 band is assigned to the vibrational mode of the primary amide, and the band at 1615 cm- I to the tertiary amide in N -acetylprolinamide. 2.3. 2.3.1.
OTHER STRUCTURALLY SENSITIVE LINES
a-Helix
The amide I and amide III bands are sensitive to conformational changes in the peptide backbone. Besides these two bands, the 890-945-cm- I band is also sensitive to structural change. There is an a-helix line at 890-945 cm -I that disappears or displays weak intensity upon conversion to jJ-sheet or random coil (Frushour and Koenig, 1974, 1975b). Thus the 890-945-cm - I line is a characteristic Raman line for a-helical conformation. The band arises from the skeletal C-C stretching vibration. There is a wide range in the position of this line. For instance, for a-helical polypeptides of polY(L-Ala), it appears at 931 cm- I ; for polY(L-Lys), at 945 cm- I ; for polY(L-Leu), at 931 cm- I ; for poly(L-Glu), at 931 cm- I ; for polY(L-Met), at 907 cm- I ; and for poly(jJ-benzyl-L-Asp), at 890 cm- I . For a-helix of polY(L-Glu), the characteristic line appears at 924 cm- I (Fasman et a1., 1978a, b). Muscle is known to be rich in a-helical proteins that show a strong band at 939 cm -I with the amide I band at 1648 cm - I, which is a characteristic of the a-conformation (Pezolet et aI., 1978). One of the muscle components, tropomyosin, is known to have mainly a-helical structure. It too shows the characteristic a-helical C-C stretching vibration at 940 cm - I (Frushour and Koenig, 1974).
Secondary Structure (Peptide-Backbone Structure)
2.3.2.
81
,8-Sheet
, For the ,8-sheet, characteristic lines lie in the 1020-1060 cm ~ 1 region. For ,81-poly(L-Glu), the line appears at 1042 cm-I, and for ,82' lines appear at 1021 and 1059 cm - I (Fasman et a1., 1978b). The difference between ,81- and ,82-type conformations lies in the spacing distance of the hydrogen-bonded pleated sheets. The ,82-form is much shorter than the ,81-form as studied by X-ray diffraction, IR, and CD spectroscopy (Hoh et a1., 1976). 2.3.3.
C-H Stretching Vibrations and Conformation
The C-H stretching vibrations appear in the region of 2800-3000 cm- I . Structural implications of C-H bands to protein structure have not been extensively studied. However, there is an indication that some bands may indeed have structural implications. The 2930-cm - I band arises from the C-H stretching vibration of the CH 3 group. Unfolding of RNase produces a large increase in Raman intensity at 2930 cm ~ I. This is interpreted as the exposure of previously buried aliphatic amino acid residues to the surrounding water (Verma and Wallach, 1977). The ratio of integrated Raman intensities of C-Hand 0- H stretching bands of aqueous lysozyme solutions is related to hydrogen bonding or hydration between protein and water (Cavotorta et a1., 1976; Samanta and Walrafen, 1978).
2.4.
THE D- AND L-AMINO ACID COPOLYMERS
PolY(L-amino acid) has a tendency to form the right-handed a-helix. When L and D forms of amino acids are mixed to form a random copolymer, the a-helix is disrupted. As a matter of fact, polY(DL-Ala) in the solid state is more like the disordered-chain conformation, as indicated by its Raman data, shown here (Frushour and Koenig, 1975a):
PolY(L-Ala) PolY(DL-Ala)(powder)
Conformation
Amide I (cm ~ I )
Amide III (cm ~ I)
a-Helix Random coil
1656 1665 1674
1274 1242 1247
X-ray diffraction data indicate that polY(DI:-Ala) does not have significant a-helical content (Brown and Trotter, 1956; Arnott and Wonacott, 1966).
82
Proteins
2.5.
RIGHT AND LEFT-HANDED a-HELICES
A polypeptide that consists of nothing but L-amino acid residues forms a right-handed a-helix, and a polypeptide with only D-amino acid residues produces a left-handed a-helix. Raman spectra for the pure L- or D-poly( y-benzyl-Glu) are identical. However, the racemic (50:50 mixture) poly( y-benzyl-Glu) has small but definite spectral changes. It is known that side-chain interactions in the racemic mixture lead to small conformational changes. Therefore, the observed change in the Raman spectra for the racemic mixture is attributed to changes in side-chain conformation and very small changes in the backbone. The amide I peak is at 1650.5 cm- I, a shift of about - 5 cm -1, and the amide III peak is at 1291 cm- I , a shift of +2.5 cm- t (Wilser and Fitchen, 1974). 2.6. 2.6.1.
DEGREE OF POLYMERIZATION Synthetic Oligopeptides
The secondary structures of proteins and polypeptides have been extensively studied, but relatively little study has been done on linear oligopeptides. It is generally agreed that their structures vary with different side chains. Baron et al. (1979) investigated this problem using homooligopeptides of L-valine, L-isoleucine, and L-phenylalanine by IR and Raman spectroscopy. For the peptides with n > 4, the amide I band appears at 1655-1665 cm- I and the amide III band at 1215-1228 cm- I; these spectra are more like those of the ,8-conformation. X-ray data show that some dipeptides and tripeptides such as Gly-Gly, L-Ala-L-Ala, and (L-Alah have an antiparallel ,8-structure and AC-L-Ala-NHMe has a twisted antiparallel structure (Rao and Parthasarathy, 1973; Tokuma et aI., 1969; Fawcett et aI., 1975; Harada and litaka, 1974). All this evidence suggests that even small peptides can have ordered structure as solids. Raman spectra of some simple peptides such as Gly-Gly, (GlY)4' Ala-Gly, and Gly-Ala in water and deuterium oxide are catalogued in the paper of Lord and Yu (1970b). Those who wish to see these lines are advised to consult the original reference. Even small peptides may have their own specific conformation. For instance, N-acetylserine methylamide, N-acetyltyrosine methylamide, and Nacetylhistidine methylamide have different rotational isomers as shown by Raman spectroscopy (Koyama et aI., 1977). 2.6.2.
Synthetic Polypeptides
The conformation of certain synthetic peptides depends on the degree of polymerization. The conformation can be readily determined by the combined use of the amide I and III bands from Raman spectra. Poly(L-Val) has ,8-sheet
secondary Structure (Peptide-Backbone Structure)
83
structure, but as the degree of polymerization increases above 500, a-helical structure begins to form (Fasman et al., 1978a). This a-helical structure of polY(L-Val) is rather unstable. The a-helix can be stabilized by introducing a strong helix-former such as L-alanine into a copolymer. 2.7.
SOLID AND AQUEOUS PHASES
Because of the low intensity of the 0- H vibration, Raman spectroscopy is ideal to study conformation in aqueous solution. This is a particular advantage because it is in this medium that the majority of biological molecules exist. When one obtains a conformation from X-ray diffraction, one always wonders whether it is identical to that in aqueous solution. Since Raman spectra can be obtained from either solid or aqueous phases, Raman spectroscopy is a good tool with which to investigate this problem. The conformations of snake neurotoxins (Yu et al., 1975; Tu et al., 1976a), Mojave toxin from Mojave rattlesnake venoms (Tu et al., 1976b), oxytocin agonists and antagonists (Hruby et al., 1978), and apamin (Nurkhametov et al., 1981) are identical in solid and aqueous phases, as there are no significant changes in amide I and III bands. Glucagon, however, exists predominantly as a-helix in crystal form, whereas it exists as l3-sheet in gel, since it shows different amide I and III bands in the two phases (Yu and Liu, 1972b). With lysozyme, the amide I and III bands are identical for crystals and aqueous solutions, but there are many spectral changes at other frequencies. This is interpreted to mean that the main peptide conformation is the same between the two phases; however, there are some side-chain conformational changes (Yu and Jo, 1973a, b). The disulfide stretching vibration (500-550 cm -I) is frequently examined for clues of conformation change, since the disulfide bridge is very important in maintaining a particular tertiary structure of a protein. When rat-liver acid phosphatases I and III are dissolved in water, the S-S stretching vibration frequency changes from 515 to 506 and 505 cm -I, respectively. This reflects the fact that the geometry of disulfide bridges changes' from a solid-phase protein to an aqueous phase. This effect is a consequence of changes in the secondary and tertiary structures of the enzyme (Twardowski, 1978). Lyophilization is frequently used by biochemists in order to store protein in a powder form instead of keeping it in solution. From experience, biochemists know that some proteins subjected to this process retain full biological activities, whereas others lose activity completely. In order to see the effect of lyophilization, RNase was examined by Raman spectroscopy. There is a marked difference in the tyrosine doublet at 830 and 850 cm-) for the powder and aqueous solution. Such a difference is attributed to changes in the local environments of the tyrosine side chains (Yu et a1., 1972a; Yu and Jo, 1973a, b). This is clear evidence that lyophilization may change the protein conformation slightly.
84
Proteins
Raman spectra obtained from a single insulin crystal at two orientations give essentially identical spectra. Insulin fibrils of "wet" and "dried" samples, however, give different spectra, especially at 1227 cm -1, indicating that the loss of bound water or loss of the hydrogen bonding does perturb the protein conformation (Yu et aI., 1974). Amide I and III bands of a-lactalbumin crystals are essentially identical to those in solution, indicating that crystallization produces no detectable effect on backbone conformation. Lyophilizedpowder samples of a-lactalbumin show significant change in the amide III band, a clear indication of alteration in protein conformation upon lyophilization (Yu, 1974). In pancreatic trypsin inhibitor, the amide I frequency increases by 3 cm- I , and the amide III bands at 1238, 1263, and 1284 cm- 1 shift to slightly lower frequencies when lyophilized powder is dissolved in water. This is explained by a weakening of hydrogen bonds in the inhibitor protein when it dissolves in water. Since the relative intensities of the three amide III peaks are the same in the aqueous as in the powder form, the relative distribution of secondary structures remains the same for the powder and the aqueous-solution forms. The greatest changes are observed in Raman lines due to the aromatic-side-chain vibrations of tyrosine and phenylalanine, indicating that the microenvironments of aromatic groups are greatly altered by converting the solid phase to an aqueous phase (Brunner and Holz, 1975). 2.8.
EFFECT OF PROTEIN ON WATER
There are many studies of proteins in aqueous solution, but most studies are focused on the effect of water on the protein molecule itself. It is equally important to understand how a water molecule is affected by introducing a protein into water. At low protein concentrations there is no competition among protein molecules for their own water shell; however, at certain concentration values, competition will make probable a superposition of the hydration shells of different molecules (Cavotorta et aI., 1976; Aliotta et aI., 1981). One way to measure the effect of proteins on water molecules is to plot the I CHI I OH ratio (intensity ratio) against the protein concentration. The deviation from a straight line indicates interaction between protein and water molecules (Samanta and Walrafen, 1978). 2.9.
QUANTITATIVE ESTIMATION OF SECONDARY STRUCTURE.
So far we have discussed different types of protein conformations separately, but natural proteins are usually a mixture of the different conformations. Thus a method for determining the relative amounts of different secondary structures becomes important. The prediction of secondary structure from the amino acid sequence and CD is fairly common and has been applied to many proteins. Structure prediction based on Raman spectroscopy has not been extensively used since it
Secondary Structure (Peptide-Backbone Structure)
85
is a comparatively new technique. Pezolet et aI. (1976) proposed methods based on the position and intensity of the amide III band and CH 2 bending .vibration of proteins to quantitate secondary-structure forms from Raman . spectra. The validity of these two methods has not been widely established to date; however, successful results were obtained for human carbonic anhydrase B (Craig and Gaber, 1977), human immunoglobulin (Pezolet et aI., 1976), and sea snake neurotoxin (Fox and Tu, 1979). In Lippert's method (Lippert et al., 1976), a set of four simultaneous equations is used. They are obtained from Raman spectra taken in both H 20 and D20 and use the relative intensities of the amide I and III bands. The purpose of using 0 2 0 is to avoid the small influence of the water-molecule vibration on the amide I band. The amounts of various types of secondary structures can be obtained by solving the following simultaneous equations:
C proteinIProtein 1240
-
/, a 1240
R + J{3(I{31240 + J:R I 1240
C proteinIprotein 1632
-
/, I a a 1632
R + J{3( I{31632 + J:R I 1632
C proteinIProtein 1660
-
/, I a a 1660
R + J{3( I{31660 + J:R I 1660
fa
+ /p + fR
r
= 1
where fa' /p, and f R refer to the fractions of a~ helix, ,a-sheet, and random coil, respectively. The experimental values of I protem (intensity) are related to the I;, If, and I: intensities previously determined for poly (L-Lys) in its various secondary-structural forms. cprotein is a scaling constant that represents the relative intensity of the methylene band in the protein. As can be seen from these equations, they do not include a term for ,a-turn (,a-reverse turn) structure; therefore, the method gives the content of a-helix, ,a-sheet, and unordered structure only, without giving any information on ,a-turn content. The ,a-turn content is probably distributed to the unordered-structure (random coil) term in this method (Fox and Tu, 1979; Bailey et aI., 1979). Lippert's equation could become a better method for predicting protein structure if it were modified to include the ,a-turn structure. The secondary structure of human carbonic anhydrase B was estimated using the methods of Lippert et aI. (1976), and the result was compared with results from X-ray diffraction (Craig and Gaber, 1977). Remarkably close results were obtained: Raman
X-Ray*
Secondary Structure
(%)
(%)
a-Helix ,a-Sheet Disordered
19
17
39 42
40
'X-ray data was based on the work of Kannan et aL (1975).
43
86
Proteins
TABLE 3.5. Relative Intensity8 of the 1240-cm- 1 Amide III Raman Band and Fraction of ~-Structure for Different Natural or Synthetic Polypeptides
Protein or polypeptide Poly(L-Lys) Ribonuclease A Basic Pancreatic Trypsin Inhibitor a-Chymotrypsin Lysozyme IgG
Frequency (em-I)
Average number of CH z per residue
Relative intensity
Fraction of IJ-Structure
1240 1239
4.0 1.3
4.8 1.6
1.0 0.38
1242 1243 1238 1240
1.4
1.9
1.1 1.2 1.3
1.1
0.45 0.34 0.16 0.37
0.3 1.5
Source: The table was reproduced from pezolet et aI. (1976). "The relative intensity is expressed as the ratio of the peak height of the 1240-cm- 1 band to that of the 1450-cm- 1 band, times the average number of CH z per residue. The fraction of ,a-structure was obtained from the X-ray data.
The method of Pezolet and co-workers (1976) is to calculate the amount of l3-sheet structure using the relative intensity, which is the ratio of intensity at 1240 cm- I to that at 1450 cm- I multiplied by the average number of CH 2 group per residue, instead of using straight-intensities. They normalized intensities of the amide III band against the methylene-bending-vibration band at 1450 cm - I because of its insensitivity to structural change (Table 3.5). As they pointed out, this method can be applied only when there is a well-defined amide III band characteristic of the l3-sheet structure at 1240 ± 3 cm- I. Moreover, this method can be used only for the content of l3-sheet structure. The method is based on the fact that the fraction of l3-sheet structure has a linear relation to the relative amide III intensities of proteins with known l3-sheet content. By plotting a standard curve, one can estimate the l3-sheet content of a protein one wishes to examine. The proteins used to construct straight-line curves are POlY(L-Lys), RNase, trypsin inhibitor, a-chymotrypsin, lysozyme, and IgG, which have l3-contents of 1.0, 0.38, 0.45, 0.34, 0.16, and 0.73 fraction, respectively.
3.
SIDE CHAINS
Unlike CD, Raman spectroscopy can be used to study the microenvironment of certain side chains. Often different biological properties of proteins are determined by the presence of certain side chains; therefore, it is important to understand the microenvironment of the side chains.
Side Chains
3.1.
87
TYROSINE
Yu, N.-T. et aI. (1973) found that the relative intensities of the Raman bands at 830 and 850 cm -1 are related to the environment of the tyrosine side chain. On some occasion~, the frequency of 860 cm - I has been reported instead of 850 cm-] (Bicknell-Brown et aI., 1981). Working with the dipeptide Gly-Tyr as an example of an exposed tyrosine, they found that the intensity of the 850/830-cm- 1 bands is 1/0.71 (Figure 3.10). An enzyme, RNase, is known to have a "buried tyrosine residue," and the intensity ratio is 0.8/1.0. On heating, the protein unfolds, thus exposing the tyrosine residue, and the 850-cm- 1 band becomes more intense than the 830-cm -] band. According to the investigation of Siamwiza et aI. (1975), the 850 and 830 cm-] doublet bands are due to Fermi resonance between the ring-breathing vibration and the overtone of an out-of-plane ring-bending vibration of the parasubstituted benzenes (Figure 3.11). If the tyrosine is buried, it acts as a strong hydrogen-bond donor; the ratio of 850/830-cm -] bands is about 0.5, resulting from the higher intensity of the 830-cm -I band (Figure 3.12 right). When the tyrosine is on the surface of a protein in aqueous solution (exposed), the ratio is higher because of the higher intensity of the 850-cm- 1 band (Figure 3.12 left). Using this property, one can quantitatively determine the number of buried and exposed tyrosine residues by using the following equation (Craig and Gaber, 1977). One should be
853cm- 1
82~ Gly-Tyr
)
l 853
830 830
RNase
3
Sea Snake Neurotoxin
)
85'C
~5/6 to"A~ v
25'C
IOCC
FIGURE 3.10. Raman spectra of (top) GlyTyr as an example of exposed tyrosine; (middle) RNase before and after heat denaturation; (bottom) sea snake neurotoxin at 25 and 100°C.
+0.257 + T 0.122 P16 u
+
-·0.122
+
-->O.lamu- I /
2
0
OH
-0.257
•
CH 3
FIGURE 3.11. Vibrational modes of p-cresol as a model for the tyrosine residue. The figure was reproduced from Siamwiza et al. (1975) by permission of the copyright owner, American Chemical Company.
~~
(EJ llAJ em
em
H 0. 2
C'' "'©.:P'"."0·" 0 .. ·.,
"2°
",
IIp-osed Tyrosine Hydrogen Bond Acceptor and Donor
I
"
"Buried" Tyrosine Hydrogen Bond Donor
FIGURE 3.12. Raman doublet for the tyrosine ring. The intensity ratio of 830 to 850 cm- I is frequently used to determine whether the tyrosine residue is buried or exposed. The ratio depends on the manner of hydrogen bonding of the tyrosine hydroxyl group.
DD
Side Chains
89
careful with the equation originally shown in their paper because it had errors which are corrected here:
N buried O.5Nburied
+ Nexposcd = 1
+ 1.25Ncxposed = 1850/1830
For instance, human carbonic anhydrase B has a total of 8 tyrosine residues and the Raman intensity ratio of 850/830-cm - I bands is 0.9. Therefore, the above equation becomes N buried
0.5Nburicd
+ Nexposed =
+
1.25Nexposcd
1
= 0.9
where Nburied and Nexposcd refer to the mole fractions of buried and exposed tyrosine residues. Solving these equations, the values are Nburied = 0.46 and Nexposcd = 0.54. The values correspond to 3.7 moles of buried tyrosine and 4.3 moles of exposed tyrosine residues. X-ray diffraction studies indicate that the number of buried tyrosine residues is 4, and 4 are exposed (Kannan et aI., 1975). The two methods give excellent agreement. Sea snake neurotoxins have one tyrosine residue (Tu, 1973). Iodination of the neurotoxin gives only an 84% reaction even with an excess of 12 reagent, and nitration is only 50% complete (Raymond and Tu, 1972). This leads to the conclusion that the tyrosine residue in the neurotoxin is "buried" or "masked." Later studies of various sea snake neurotoxins by Raman spectroscopy revealed that the single tyrosine residue is, indeed, buried (Yu et aI., 1975; Tu et aI., 1976a). There is a change in the tyrosine vibrational bands when a tyrosine residue is involved in binding. A protein called LIV-protein specifically binds to leucine, isoleucine, or valine, causing a spectral change in the Raman frequencies for the aromatic residues, tyrosine and tryptophan (Vorotyntseva et aI., 1981). This illustrates the usefulness of the Raman technique in observing the microenvironment of tyrosine residues in a protein. 3.2.
TRYPTOPHAN
The characteristic bands of the tryptophan indole-ring vibrations appear at 544, 577, 761, 879, 1014, 1338, 1363, 1553, and 1582 cm- I (Lord and Yu, 1970b). Among these lines, the 1363-cm- 1 band can be correlated with the environment of the indole side chain. A sharp, intense Raman line at 1360 cm- 1 is diagnostic for a buried tryptophan (Yu, 1974). In carbonic anhydrase B, three residues are buried, and two residues are partially exposed, as studied by X-ray diffraction (Kannan et al., 1975). Raman analysis indicates that some
90
Proteins
of the residues are buried. The application of this method is also shown with sea snake neurotoxins, which have one tryptophan residue (Tu, 1977). The tryptophan residue in the neurotoxins can readily be modified by different chemical reagents, indicating that the tryptophan is exposed (Tu and Toom, 1971; Tu et aI., 1971; Tu and Hong, 1971). The Raman spectra of different sea snake neurotoxins indicate that the tryptophan is indeed exposed, because there are no 1360-cm- 1 lines (Yu et aI., 1975; Tu et aI., 1976a).
3.3.
PHENYLALANINE
The breathing vibration of the benzene ring of phenylalanine can be detected at 1005 cm -1 when the concentration of phenylalanine is above 1%. Although there is no particular band that can be correlated to the environment of the phenylalanine side chain in a protein, the ratio of phenylalanine to tyrosine can be determined. Yu and East (1975) estimated the phenylalanine-to-tyrosine ratio in lens proteins by comparing the intensities at 624 cm -1 (phenylalanine) and 644 cm- 1 ( tyrosine). When this method is applied to carbonic anhydrase B, the ratio is 1.4, which is in good agreement with the ratio (1.38) determined from the amino acid composition (Craig and Gaber, 1977). This method is based on the Raman intensity ratio multiplied by the factor 1.25: /624 /644
X
Phe 1.25 = Tyr
In order to check the validity of this method, 64 Raman spectra of 25 proteins were analyzed (Liddle and Tu, 1981). Of these, 28 out of 64 (44%) showed a positive correlation within a range of ±25% error. The results of this investigation indicate that the intensity-ratio method is of limited use for all proteins. Ribonuclease has three phenylalanines out of a total of 124 amino acids. Vibrational bands that originate from the phenylalanine ring can be seen in the Raman spectrum of RNase (Lord and Yu, 1970a). The aromatic-vibration bands are relatively insensitive to the state of aggregation. The lines due to the phenylalanine ring are summarized here: 622 Weak 1006 Strong- "breathing" vibration of the monosubstituted ring 1033 Weak 1183 } 1207 1585 These lines are overlapped when there are tyrosine residues 1605
Side Chains
3.4.
91
HISTIDINE
The imidazole group of histidine has two tautomeric forms (Figure 3.13). Tautomer I represents the I-N-protonated form, and II is for the 3-N-protonated form. The two forms show different breathing vibrations, which appear at 1282 cm- 1 for tautomer I and 1260 cm- I for tautomer II. The Raman spectrum obtained at neutral pH indicates that the imidazole group exists in an equilibrium between the two tautomeric forms (Ashikawa and Itoh, 1979). 3.5.
DISULFIDE BOND
Disulfide bonds (disulfide linkages, disulfide bridges) help to provide additional stability to a protein molecule. Clearly, understanding disulfide bonds in a protein is quite important to the structural study of proteins. Raman spectroscopy can be used for this purpose, furnishing information that other physical methods cannot. Raman spectroscopy shows a strong S-S stretching vibrational band in the region of 500-550 cm- I . The origin of the S-S stretching vibrations has been extensively studied. It has been found that the S-S stretching vibration depends on the internal rotation about the C-S and C-C bonds of C-C-S-S-C-C. A disulfide bond is normally expressed as -S-S(Figure 3.14) but more strictly speaking, it is C(a)-C(jJ)Hz-S-Sc(,B)Hz-C(a). The C(a) refers to the a-carbon atom to which peptide bonds attach. As elucidated from studies of many disulfide model compounds of known conformation, the symmetrical-stretching vibration of the disulfide bond is influenced by the conformation of the carbon atoms in the disulfide bridge Ca-Cp-S-S-Ca,-Cf/" The vibration at 510 cm- I can be assigned to the gauche-gauche-gauche (g-g-g) conformation (Figure 3.15). The gauche-gauchetrans (g-g-t) rotamer gives the band at 525 cm -1, and the trans-gauche-trans (t-g-t) at 540 cm- 1 (Sugeta et aI., 1972, 1973). Many naturally occurring proteins give the disulfide stretching vibration at or near 510 cm -(, suggesting that the gauche-gauche-gauche form is the most preferred conformation. Even when proteins are denatured by heat treatment, very little change in the H
H
I
I
~~'N;,H a
N
\4
1,
5
C=C'H I R
Tautomer J
--+ +--
C
H'N .... 2~N
a
1
\4
5
1
C=C'H I R
Tautomer II
FIGURE 3.13. Tautomerism of the imidazole ring of histidine.
92
Proteins
o
II
~
-N-CH- C - - H I CH 2
I
}
S
I
s
o II
I
CH 2
I
- C - CH -
FIGURE 3.14.
NH
-
Disulfide bond in protein.
intensity of the 510 cm- I band is observed. Occasionally new shoulders appear at 525 or 540 cm - I upon heat denaturation, suggesting that trans-gauche-gauche and trans-gauche-trans forms are formed in addition to the original gauchegauche-gauche conformation (Tu et aI, 1976a). However, other conformations are common among synthetic cyclic compounds such as [I-penicillamine]oxytocin and [I-penicillamine, 2-leucine]oxytocin (Hruby et al., 1978). Naturally occurring cyclic peptides such as mesotocin and
FIGURE 3.15. Dependence of the S-S stretching frequency on the internal rotation. (A) Gauche-gauche-gauche form, (B) gauche-gauche-trans form, (C) trans-gauche-trans form. Trans and gauche refer to the atoms from the left sides.
Side Chains
93
: argininylvasopressin have trans-gauche-trans conformation in addition to the .gauche-gauche-gauche form (Tu et aI., 1979). Occasionally the S-S stretching vibrational frequency appears somewhere other than at 510 cm- 1 for natural proteins. For instance, the Bence-Jones protein has one S-S vibrational band at 524 cm -1, indicating the transgauche-gauche type (Kitagawa et aI., 1979), and a sulfide band appears at 521 cm - 1 for pepsin (Tobin, 1968). In lysozyme and snake neurotoxins, only one intense disulfide stretching vibration was observed at 510 cm -I, suggesting that all the disulfide linkages have a similar local geometry. Insulin is an interesting example, because it has different types of disulfide bonds. It has two interchain and one intrachain disulfide bonds; there are two different disulfide Raman bands, one at 505 cm -I and the other at 517 cm- I. Moreover, the intensity ratio of these two bands is about 2 to 1. The same ratio is obtained from the C-S stretching vibrations at 668 and 680 cm -I. Since insulin does not contain methionine, the observed C-S bands are all from C-S-S-C of the disulfide linkages (Yu et aI., I972b). The S-S stretching vibrational band is rather prominent in Raman scattering, and often the appearance and disappearance of this band are correlated with a structural change in proteins. RNase has four disulfide bonds and upon reduction is converted to random coil. The disulfide-bond vibration at 516 cm- I reappears in the correctly refolded RNase (Galat et aI., 1981). The coagulation of egg white by heating is often thought to involve the formation of a disulfide bond. However, no band appeared in the 510-cm- 1 region when egg white protein was aggregated (Painter and Koenig, 1976c). Therefore, it can be concluded that no disulfide bonds are formed by the coagulation of egg white. 3.5.1 . Theory Proposed by Sugeta
Sugeta and co-workers (Sugeta et aI., 1972, 1973) proposed that the S-S stretching frequency does not depend on the C-S-S-C dihedral angle, but depends on the torsional angles in C-C-S-S-C-c. According to this theory, Raman spectroscopy can distinguish three types of disulfide-bond geometry, namely, gauche-gauche-gauche, trans-gauche-gauche, and transgauche-trans conformations (Figure 3.15). The Raman band appears at 510 cm - I for the g-g-g, 525 cm - I for the t-g-g, and 540 cm - I for the t-g-t conformation of C-C-S-S-C-c. There is some experimental evidence to support this theory, as shown in Streptomyces subtilisin inhibitor and lysozyme (Satow et aI., 1980). Lysozyme has three disulfide linkages at the positions Cys(6)-Cys(l27), Cys(30)-Cys(115), and Cys(64)-Cys(80), all with the g-g-g conformation, and one disulfide at Cys(76)-Cys(94) with the t-g-g conformation, as deduced from X-ray crystallographic analysis (Blake et aI., 1967). Raman spectra of lysozyme have one prominent peak at 507 cm- I for g-g-g and a smaller band at 528 cm- I for t-g-g (Nakanishi et aI., 1974).
94 3.5.2.
Proteins
Theory Proposed by Van Wart
The essence of this theory is that the S-S stretching frequency has a linear correlation with the C-S-S-C dihedral angle (Van Wart et al., 1973; Maxfield and Scheraga, 1977). Martin (1974) reported that there is no such correlation. After reevaluation, both Van Wart and Martin (Van Wart et al., 1976) agreed that the S-S stretching frequency is invariant to the C-S-S-C dihedral angle in the 65-85° range, but the frequency of the S-S vibration does vary with the C-S-S-C dihedral angle between 0 and 65° as originally proposed by Van Wart et al. In proteins, the S-S stretching frequencies quite often appear at 510, 525, and 540 cm- I. Van Wart and Scheraga ( I 976a, b) summarized the relationship between the S-S stretching frequencies and the C-C-S-S and C-S-S -C dihedral angles. These results are shown here:
"s-s (em-I) 510 ± 5 525 ± 5 540 ± 5
Dihedral Angle of C-C-S-S (deg)
Dihedral Angle of C-S-S-C (deg)
50-180 50-180 0-50 50-180 0-50 0-50
85 ± 20 85 ± 20 85 ± 20
According to the study of Van Wart et al. (1976), the band at 525 cm- 1 is not due to the trans conformation of C-C-S-S in C-C-S-S-C-C, but rather to a low value of the C-C-S-S dihedral angle. Similarly, the band at 540 em -I is considered to be associated with the trans-gauche-trans conformation of the C-C-S-S-C-C unit by Sugeta et al. However, according to Van Wart and Scheraga (1976) a value for "s-s of 540 cm- I arises from the presence of conformations with small C-C-S-S dihedral angles of about 30°. Thus there are two opposing theories concerning the origin of the S-S stretching vibration. 3.6.
C-S
Methionine and cystine residues have a C-S stretching vibration. There is a correlation between the C-S stretching frequencies and the internal rotation about the C-S bond in methionine and isobutyl methylsulfide (Nogami et al., 1975). This relationship is illustrated in Figure 3.16. The C-S stretching
'.'
\";
/CMETHIONINE RESIDUE
'Vk'
}t~
·.tl~'
,<
~~'
!~:i
~
~
655 em- 1 724 em-1
H-C-CH2-CH2f'S-CH3
'~
~
'-~
f"
~
700em- 1
,",-C',-'
FIGURE 3.16. Trans and gauche forms of methionine and their Raman frequencies.
-EN
OCI-~,ACH2-
~
S - s-
~
PH
Pc
PN CYSTINE RESIDUE
FIGURE 3.17. Internal rotation of cystine residue along the C-C axis. This produces three conformational forms that give different C-S stretching vibrations.
9S
96
Proteins
frequencies of a methionine residue depend on its conformation. The 655- and 724 cm - I bands are for the trans form of the methionine side chain, whereas the 700 ± 5 cm- I band is for the gauche form (Lord and Yu, 1970a). Similarly, in cysteine and cystine, the C-S stretching vibration is related to the internal rotation along the C-C axis of X-C-CH 2 -S-S. It depends upon the atom X at the trans site with respect to the sulfur atom adjacent to the C-C bond. When X is a hydrogen atom (expressed by the symbol, PH)' the C-S stretching vibration lies between 630 and 670 cm -I. It is at 720 cm - I for the conformation when X = carbon (PC> and at 700 cm- I when X = nitrogen (PN ) as shown in Figure 3.17. It has been proposed that there is a correlation between the intensity ratio of the C-S stretch near 660 cm- I and the S-S stretch around 510 cm- I and the CSS angle (Lord and Yu, 1970a, b). Using a large number of model compounds, however, it has been concluded that there is no correlation between the Raman intensity ratio for C-S and S-S stretching bands and the CSS angle (Bastian and Martin, 1973). 3.7.
SULFHYDRYL GROUP (-SH)
The S- H stretching vibrations appear in the 2570-2580 cm- 1 region for many alkyl thiols (Dollish et a1., 1974). The Raman spectra of proteins normally originate from C-C, C=C, C-N, C=N, C-H, C-S, C-O, C=O, N-H, O-H, and S-H vibrations. Among these vibrational modes, only the S-H vibration appears in the 2500-2600 cm- I region. Therefore, little interference or ambiguity is involved in the detection of a sulfhydryl group. A disulfide bond of pancreatic trypsin inhibitor can be reduced to form
...> III
...... Z
z z c
~
c
'" 2580
2620
eM -1
2560
2500
FIGURE 3.18. S-H stretching vibration of sea snake (Lapemis hardwickii) neurotoxin. The figure was reproduced from Fox et al. (1977) by permission of the copyright owner, North-Holland Publishing Company.
IR and Raman
97
sulfhydryl groups. The reduced protein shows a new Raman peak at 2580 cm- I , which is the -SH vibrational mode (Brunner and Holz, 1974). A neurotoxin of Lapemis hardwickii sea snake venom contains four disulfide bonds and one sulfhydryl group. However, the sulfhydryl group is masked, and its presence cannot be detected by conventional chemical analysis. Yet the sulfhydryl group of the toxin can readily be detected by Raman spectroscopic methods since the typical S- H stretching vibration at 2585 cm - I can be seen (Fox et aI., 1977) (Figure 3.18). Similarly, the coat protein (A-protein) of tobacco mosaic virus contains one masked sulfhydryl group that cannot be detected by normal chemical analysis. Again, this can readily be detected in a Raman spectrum at 2567 cm- I (Fox et aI., 1979; Figure 6.5). These examples further demonstrate the usefulness of Raman spectroscopy as an analytical tool in protein studies. Another way to determine sulfhydryl groups is to attach a Raman-active compound. For instance, by adding 5,5'-dithiobis (2-nitrobenzoic acid) to sulfhydryl groups, one can detect strong Raman bands due to the nitro group vibrations at 1325 and 850 cm - I (Banford et aI., 1981).
4.
PRERESONANCE RAMAN
Practically all Raman spectra of proteins and synthetic peptides have been obtained by using excitation light in the visible region (nonresonance Raman). It is interesting to see the Raman spectra of polypeptides with excitation light in the ultraviolet region. Sugawara et aI. (1978), using the 257.3-nm laser line as the light source, examined poly(L-Lys) and polY(L-Glu)(preresonance Raman)(Table 3.6). The vibrations associated with the C=O and C- N stretching modes are preresonantly enhanced. Especially the amide I band becomes clearer in D 2 0 because of the lack of water band interference. This result is highly encouraging because some day we will be able to excite polypeptides with the laser line corresponding to the peptide-bond absorption band, and we expect to see resonantly enhanced amide I and III bands. When lysozyme is excited by the near-ultraviolet laser line at 363.8 cm -I, the tryptophan Raman bands at 757, 1010, 1330, 1360, 1552, and 1582 cm- I are greatly enhanced. Preresonance Raman spectroscopy may be very useful in studying enzyme mechanism involving tryptophan residues (Brown et aI., 1977).
5.
IR AND RAMAN
Infrared and Raman spectroscopies are complementary to each other. In order to make detailed vibrational assignments, it is necessary to have both IR and Raman data. There are variations of intensity between IR and Raman spectra, especially when the molecule has a high degree of symmetry. In this book we
TABLE 3.6.
Nonresonance and Preresonance Raman Spectra of POly(L-Glu) and POly(L-Lys)
Amide I
Amide III
N onresonance 488.0 nm
Preresonance 257.3 nm
PolY(L-Glu) (H 2 O) PolY(L-Glu) (D 2 O) Poly(L-Lys) (H 2 O) PolY(L-Lys) (D 2O) PolY(L-Lys) (H 2O)
1668 1662 1665 broad 1663 m 1647 s
1674 1664 1664 s 1662 m 1648 s
a-Helix ,B-Sheet
Poly(L-Lys) (D 2O) Poly(L-Lys)(H 2 O)
1641 s 1665 s
,B-Sheet
PolY(L-Lys)
1638 m 1665 s broad 1659 s
Conformation
Compound
Random coil Random coil Random coil Random coil a-Helix
Source:
(D 2O)
The data was reconstructed from Sugawara et aI. (1978).
1657 s
N onresonance 488.8 nm
Preresonance 257.3 nm
1251
1250
1247 m
1248 s 1200 1300 (broad)
1244 s
1243
Low-Frequency Vibrations
99
TABLE 3.7. Peptide Bond Vibrational Bands of Poly(Gly) I (IJ-Sheet) and a-Helical Poly(L-Val) POlY(L-Gly)
POlY(L- Val)
Raman (em-I)
IR (em-I)
3301
3308 3088 1685 1636
1674 1564 1515 1234 1220
-
1517 1236 1214 708 628
601
-
207
589 217
Vibrational Assignments Amide A Amide B Amide I Amide I Amide II Amide II Amide III Amide III Amide V Amide VI Amide VI Amide IV Amide VII
Raman (em-I)
IR (em-I)
3300
3293
1655
1655
-
1253 1246 1237
-
1535
1238 612
Source: The table was constructed based on the data of Small et aI. (1970) for Poly(Gly) and from Yamashita and Yamashita (1975) for Poly(L-Val).
are mainly concerned with Raman data, but it is of interest to compare IR and Raman spectra directly for the same compound. The example given here is polyglycine I, which has extended-,B-sheet form (Small et al., 1970). As can be seen in Table 3.7, some IR-active bands such as amide B, amide IV, V, and VII bands are Raman inactive. Structurally important amide I and III bands are relatively strong in a Raman compared with IR spectra. Amide I assignments for different conformations in Raman and IR spectra are different. For instance, in Raman spectra the bands from a-helix, antiparallel ,B-sheet, and unordered structures usually appear at 1650-1657 cm- I , 1665-1680 cm- I , and 1666-1668 cm -1, respectively. However, for IR spectra the bands from a-helix, ,B-sheet, and unordered structures appear at 1650, 1633, and 1655 cm - I, respectively (Doyle et al., 1975).
6.
LOW-FREQUENCY VIBRATIONS
The origin of low-frequency vibrations of biological compounds is complicated. They arise from any different modes. For example, the breathing vibration is a molecule as a whole undergoing symmetrical vibration. Or the vibration may be due to two lobes of a protein molecule moving relative to one another, interaction between a neighboring molecule and a subunit, interaction
100
Proteins
with water molecules, lattice vibrations, metal-ligand-bond vibrations, and so forth. The study of the low-frequency region, especially below 100 em - I, is hampered by the presence of strong Rayleigh scattering. This is because as the monochromator approaches the Rayleigh line in frequency, it is incapable of discriminating between the Rayleigh line and the low-frequency Raman bands. Consequently, the Raman bands are obscured. Brown et aI. (1972) used an iodine filter to block the Rayleigh light and observed Raman bands as low as 10-12 em-I. 6.1.
INTERNAL VIBRATIONS
6.1.1.
Breathing Vibrations
The exact assignment of low-frequency bands is difficult. Chymotrypsin shows a broad peak at 35 em -1, which may be a breathing or swelling motion of the entire molecule (Brown et aI., 1972). The low-frequency band at 264 em - I is assigned to the a-helical-coil breathing vibration (Itoh and Shimanouchi, 1970). The intensity of this band decreases markedly by introducing D residues into a-helical polY(L-Ala) (Itoh et aI., 1974). This suggests that long sequences of L-alanine residues in the a-helical conformation are required to have a good breathing vibration (Figure 3.19). Low-frequency vibrations can arise from the internal vibrations of the protein molecule. They are dependent upon the conformation of the protein molecule, but are relatively independent of the form of the sample, that is, (a)
--- ---ססoo
. ~
~
4
0lf00
(b)
FIGURE 3.19. The breathing vibration of poly( L-Ala) a-helix, and the effect of D residues on the a-helical conformation and on vibrational modes. (A) Breathing vibration. (B) No breathing vibration.
Low-Frequency Vibrations
101
. whether it is a film or a crystal. a-Chymotrypsin shows a peak at 29 em- I that arises from vibrations that involve all, or a very large portion, of the protein molecule (Brown et al., 1972). The vibrational frequencies of the longitudinal acoustical modes depend on chain length (Fanconi and Peticolas, 1971; Fanconi et al., 1971). Therefore, the longer the peptide chain, the lower the frequency. 6.1.2.
Torsional Vibrations
Low-frequency vibrations can also arise from the torsional mode of CH 3 . Skeletal-deformation bands of alanine appear in the range of 260-410 em - I. The CH 3 torsional bands of alanine also appear within this range. As a result, it is difficult to make specific band assignments without observing isotopic effects. By obtaining deuterated CD3 in DL-alanine-a, f3-d 4 , it has been shown that CD3 torsional-mode bands appear at 200 and 290 em -I (Machida et al., 1979). The calculated CH 3 torsional frequencies for L-alanine crystals are 394 em - I for the a and 285 em - I for the b species. The a and b refer to the crystal axis. The observed CH 3 torsional-vibration bands are at 398 em-I for the a species and at 282 em - J for the b J species (Machida et al., 1978). 6.2.
INTERMOLECULAR VIBRATIONS
In the solid state, low-frequency vibrations can also arise from the linking forces between neighboring macromolecules. Solid lysozyme has one band at 75 em -I that is absent in aqueous samples. This suggests that this band cannot be attributed to an internal molecular vibration. In concentrated lysozyme solutions, a band at 170 em - I appears. In concentrated protein solutions, water-water interactions are weakened, thus the protein-water interactions become important, and the low-frequency band appears (Samanta and Walrafen, 1978). There is a band at 25 em - I in the Raman spectrum of solid lysozyme. This vibrational mode is probably associated with an intermolecular vibration, since it disappears in the spectrum of an aqueous solution (Genzel et al., 1976). Disappearance of this band in aqueous solution may also by explained in terms of "dampening" by solvent. From theoretical calculations, the intramolecular vibration can be as low as 30 em -I. Therefore, the possibility is not ruled out that such a vibration might exist within the molecule (Ataka and Tanaka, 1979). 6.2.1.
Subunit Interactions
Two globular lobes of lysozyme move relative to one another in a vibration. This mode can be dampened by solvent. In immunoglobulin IgG the same phenomenon can be observed (Painter and Mosher, 1979). In low humidity, IgG gives low-frequency bands at 28, 35, and 60 em -I. When IgG is saturated
102
Proteins
FIGURE 3.20. Highly simplified view of immunoglobulin. H, heavy chain; L, light chain. Chains are held together by many disulfide bridges.
with water, these bands almost disappear. An immunoglobulin monomer is a Y-shaped molecule (Figure 3.20). These vibrations may be due to relative displacements of the subunits, or to "elbow-bending" or "hinge-bending" modes. The loss of resolution in 100% humidity conditions probably means that these vibrational modes are being dampened in the presence of water molecules. 6.2.2.
Lattice Vibrations
In crystalline samples, the lattice vibration also gives low-frequency bands (Figure 3.21). For instance, several lattice-vibrational bands can be observed in the region of 25-200 cm-\ (Machida et aI., 1978; Medina et aI., 1980; Andrews et aI., 1980).
FIGURE 3.21.
Lattice vibration of a crystal.
Application
103
RAMAN-SPECTRA BACKGROUND
One frequently encounters a large, broad, and persistent "background" band. origin of this mysterious background is not yet fully understood. Normally this background is considered to be a fluorescent band of small impurities present in the sample (Smith et aI., 1969) or the lack of optical homogeneity of the sample (Lord and Yu, 1970a). The background band often decays in intensity with time (Lord and Yu, 1970b; Careri et aI., 1973). Thus it is recommended that a sample be exposed to a laser beam for several hours in order to decrease this background and obtain better-quality spectra. Some investigators do not consider this band to be due to fluorescence but believe it to be a true Raman spectrum (Careri et aI., 1970a, b; Biscar et aI., 1972a, b; Biscar and Kollias, 1973). They think this background relates to the conformation of proteins. Sometimes this broad Raman background is called pseudo-Raman in order to differentiate it from an ordinary Raman spectrum. The pseudo-Raman spectrum is claimed to be correlated with the chain length of proteins and nucleic acids (Biscar et aI., 1974). An attempt was made to correlate the frequency to a collective molecular electronic effect on chymotrypsin (Biscar and Malone, 1975; Biscar, 1976). This subject remains controversial, and many investigators consider the pseudo-Raman effect to be an artifact without any scientific significance. Readers who wish to know more about this subject are advised to see the original references on the theory, called electromagnetic molecular electronic resonance.
8.
APPLICATION
8.1. 8.1.1 .
DENATURATION Peptide Backbone
Denaturation is a process that converts ordered structure to disordered structure. This process can readily be followed by Raman spectroscopy. As is shown in Figure 3.22, a lysozyme molecule undergoes a conformational change from ordered structure to random coil as the temperature is increased. The amide I band shifts to a higher frequency, and the amide III band moves to a lower frequency (Porubcan et aI., 1978; Fox and Tu, 1979). Snake venoms of Elapidae and sea snakes contain potent neurotoxins that are very stable molecules that contain relatively large numbers of disulfide bonds for their small size. Heat denaturation of a sea snake neurotoxin does not result in any change in the amide I and III bands, suggesting that there is no significant change in the peptide-backbone conformation (Tu et aI., 1976a). This is consistent with the fact that the neurotoxins are relatively stable to heat treatment; neurotoxins from Laticauda semifasciata venoms retained their toxicities even when boiled at 100°C for 30 min (Tu et al., 1971).
104
Proteins
~
AMIDE I 1700
~
AMIDE III
16001300
CM-1 8.1.2.
1200
FIGURE 3.22. Denaturation of lysozyme. As temperature increases, the amide I band shifts to higher frequency, and the amide III band shifts to lower frequency.
Side Chains
The change in the microenvironment of side chains of certain amino acid residues can be detected by Raman spectral changes. In the case of Pelamis platurus sea snake neurotoxin, there is no significant change in the disulfide band at 512 em- 1, but a new shoulder appears at 546 cm -) (Figure 3.23). This suggests that a small fraction of the disulfides change conformation to the trans-gauche-trans configuration of C-C-S-S-C-C. On the other hand, the structurally sensitive tyrosine bands at 832 and 850 cm - I change in relative intensity, suggesting that on heat denaturation, the tyrosine side chain becomes exposed. In the case of insulin, there is no change in the S-S stretching vibration at 516 cm-) on heat treatment, suggesting that there is no change in the local geometry of CSSC. However, the C-S stretching vibration at 670 cm-) changes to 668 and 680 cm-), suggesting that the CSS angles of the disulfide linkages have changed appreciably (Yu and Liu, I 972a).
Pelamis platurus major toxin (solid) Amide III
Amide I
(a) Native
1245
1672
o
N
':!
~
"in c: Ql
C ..... c: o
(b) Heat - treated
E
1241
o
'\
a::
1600
1400
1200
1000
800
600
400
Wove Number (C M- 1 ) FIGURE 3.23. Raman spectra of Pelamis platurus (yellow-bellied sea snake) neurotoxin before and after heat treatment. The figure was reproduced from Tu et al. (1976) by permission of the copyright owner, Munksgaard Company"
.. ft.,
106
Proteins
The effect of heat denaturation on side chains is mainly to disrupt those side chains involved in hydrogen bonding and hydrophilic and hydrophobic interactions. The covalently linked side-chain disulfide bridge is not broken by heat treatment. The disulfide stretching band at 500-550 cm - I is normally very strong in Raman spectra, and in no case does this band disappear because of heat denaturation of proteins. A specific example of sea snake toxin is given above. In lysozyme the S-S stretching vibration band at 509 cm- I does not change even when heated to 76°C (Chen et al., 1973). Heat denaturation studies of RNase using the amide I and III lines indicates that the process proceeds via a stepwise unfolding rather than a transition between two states. It also reveals that substantial amounts of the helical and pleated-sheet conformations remain even at 70°C (Chen and Lord, 1976). So far we have discussed denaturation by heat treatment, but Raman spectroscopy is equally useful in following protein denaturation by chemical agents (Lord and Mendelsohn, 1972), that is, acid or base treatment. The amide I and III bands are most frequently used to determine the conformation of proteins. There are relatively few studies of protein polarization in relation to conformation. Gunde et al. (1977) reported that all bands are more polarized in the denatured state than in the native state. The amide III bands of the protein in the denatured state are completely polarized. This is explained by the fact that denaturation changes local environment and affects local symmetry. When the disulfide bond is cleaved by chemical modification, there is a drastic change in the Raman spectra. For instance, when the S-S bonds of lysozyme are cleaved by reduction and subsequent cyanoethylation of the sulfhydryl group, the original 507 cm- 1 S-S stretching vibration band disappears, and there is a large shift in the amide I band from 1672 to 1660 cm- I and in the amide III bands from 1254 to 1243 cm- I and from 1271 to 1263 cm- I (Chen et al., 1974). This is logical since the disulfide bond is important in maintaining the ordered structure of a protein. Cleavage of this bond readily causes the denaturation of the protein. 8.2.
CHEMICAL MODIFICATION
Chemical modification is a useful technique to elucidate structure-function relationships. One important aspect is to ascertain that the specific group is modified as originally intended, and that other functional groups are not affected by the modifying agent. Raman spectroscopy can be used to detect some chemical modification of proteins. 8.2.1.
Tryptophan
N-bromosuccinimide is commonly used for the chemical modification of tryptophan residues. It oxidizes the indole ring to oxyindole.
~J,
Application
107
:p;
~2
When lysozyme is treated with this reagent, the tryptophan peak at 1010 :tl'cm- I disappears, and a new strong band of oxyindole appears at 10 17 cm- I :\:-(Schmidt and Bieker, 1979). Shifts are also observed in the amide I and III 7'bands, indicating conformational changes resulting from tryptophan modifica?',tion. In an excess of N-bromosuccinimide, tyrosine vibrational bands can no . longer be detected, indicating that the tyrosine residue is also modified. Tryptophan residues can also be modified with ozone to form N-formylkynurenine. When lysozyme is treated with ozone, the tryptophan residue is completely destroyed. This is shown by the disappearance of the indole-ring-vibration bands at 760 and 1011 cm - I and the appearance of a new band at 1604 cm- I , which is assigned to the benzene system of N-formylkynurenine (Bieker and Schmidt, 1979). 8.2.2.
Tyrosine
The coupling of certain agents to proteins yields strongly resonance-enhanced Raman spectra because of the introduction of a chromophoric group. When carbonic anhydrase is modified with the diazonium salt of sulfanilamide, it causes the loss of sulfanilamide-binding capability and esterase activity (Li et al., 1979). Resonance Raman spectra of the chemically modified enzyme indicate that the reagent is coupled to a tyrosine residue. 8.2.3.
Histidine
Histidine side groups give rise to a characteristic ring frequency at 1491 cm -1. In RNase this line is masked by other lines. In D20 the strong and sharp lines of the N-deuterated imidazolium ring of histidine appear at 1409 cm- I . This line is sensitive to change from imidazolium to imidazole (Lord and Yu, 1970a).
8.3. 8.3.1.
COMPARISON OF RELATED PROTEINS S-Ovalbumin and Ovalbumin
S-ovalbumin is a more-heat-stable form of ovalbumin. A Raman difference spectrum (ovalbumin minus S-ovalbumin) shows differences in the intensities of the amide I and III regions. These intensity differences are due to the conversion of ovalbumin to S-ovalbumin, involving a conformational change of the protein from a-helix to antiparallel ,B-sheet. Although only 3-4% of the total structure is involved in the change, the small difference may account for the large difference in the thermodynamic stability of ovalbumin and Sovalbumin (Kint and Tomimatsu, 1979).
108
Proteins
8.3.2.
Proinsulin and Insulin
Proinsulin is a precursor of insulin. A segment is removed from the middle of proinsulin by hydrolysis, and the active hormone insulin, C-peptide, and amino acids are liberated. X-ray study indicates that there are two segments of a-helix in the C-peptide portion of proinsulin (Fullerton et al., 1970). The amide I and III bands for proinsulin and insulin are indeed very similar, suggesting similarities between the conformation of insulin and proinsulin. Quantitative estimation indicates that proinsulin has more a-helix contribution. Thus Raman data correlate remarkably well with the data from X-ray crystallographic studies (Yu et al., 1972b, c). 8.4.
GLYCOPROTEINS
Antifreeze glycoprotein from the sera of an Antarctic fish has been relatively well studied (Tomimatsu et al., 1976). Because two components, proteins and carbohydrates, are present, the overall Raman spectrum is the sum of the contribution from both components; thus the overall spectrum is more complicated. However, the amide I band is clearly shown for antifreeze glycoproteins, components 4 and 8. Except for a band near 1632 cm- I due to the N-acetyl chromophore, the carbohydrate moiety has no bands in this region (She et al., 1974). Several bands occur in the amide III regions; however, amide III assignments are not certain because of the lack of deuterium-exchange data. The Raman spectra of water present in a 1% antifreeze glycoprotein solution and of ice frozen from this solution are indistinguishable from the spectra of pure water and ice, respectively. These results indicate that the bulk properties of water and ice are unaffected by the presence of the antifreeze glycoprotein. Examination of active and inactive glycoproteins reveals that they differ in conformation and possibly in the environment of their carbohydrate hydroxyls. These observations suggest that hydrogen bonding of the carbohydrate hydroxyls of the active glycoprotein at the ice-solution interface may physically prevent growth of the ice lattice. 8.5.
BLOOD COAGULATION
Conversion of fibrinogen to fibrin increases the fJ-sheet form. Fibrinogen has a predominant amide III band at 1254 cm-I, whereas fibrin has a major band at 1236 cm- I and a minor band at 1256 cm- I . The interpretation of the amide-I-band change associated with the conversion is less clear-cut (Marx et al., 1978; 1979). There is a considerable change in the region of 1000 cm-I, which is a mixed-vibration region of C-C and C-N vibrations. This suggests that there is considerable change in the C-C and C- N skeletal conformation when the reorganization of fibrinogen monomer to fibrin polymer takes place.
References
109
REFERENCES . Aliotta, F., Fontana, M. P., Giordano, R., Migliardo, P., and Wanderlingh, F. (1981). Raman scattering in lysozyme solutions. J. Chem. Phys. 75, 4307. Andrews, B., Torrie, B. H., and Powell, B. M. (1980). Lattice modes of deuterated a-glycine. In Proc. Vllth Int. ConI Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 110-111. Arnott, S., and Wonacott, A J. (1966). Atomic co-ordinates for an a-helix: Refinement of the crystal structure of a-poly-L-alanine. J. Mol. BioI. 21, 371. Asher, 1. M., Rothschild, K. J., and Stanley, H. E. (1974). Raman spectroscopic study of the valinomycin-KSCN complex. 1. Mol. Bioi. 89, 205. Ashikawa, 1., and Itoh, K. (1979). Raman spectra of polypeptides containing L-histidine residues and tautomerism of imidazole side chain. Biopolymers 18, 1859. Ataka, M., and Tanaka, S. (1979). Far infrared spectrum of crystalline lysozyme. Biopolymers 18, 507. Bailey, G. S., Lee, J., and Tu, A T (1979). Conformational analysis of myotoxin a (muscle degenerating toxin) of prairie rattlesnake venom. J. Bioi. Chem. 254, 8922. Ballardin, A, Fischman, A J., Gibbons, W. A, Roy, J., Schwartz, 1. L., Smith, C. W., Walter, R., and Wyssbrod, H. R. (1978). Conformational studies of [Pro3 ,Gly 4]-oxytocin in dimethyl sulfoxide by I H nuclear magnetic resonance spectroscopy: Evidence for a type II ,B-tum in cyclic moiety. Biochemistry 17, 4443. Bandekar, J., and Krimm, S. (1979a). Vibrational analysis of pep tides, polypeptides, and proteins: Characteristic amide bands of ,B-tums. Proc. Nat. Acad. Sci. 76, 774. Bandekar,1., and Krimm, S. (I 979b). Vibrational analysis of peptides, polypeptides, and proteins, VII. Normal modes and vibrational spectra of a type I ,B-tum tetrapeptide. In Pept. Struct. Bioi. Funct., Proc. Am. Pept. Symp., 6th, E. Gross and J. Meienhofer, Eds., Pierce Chemical Co., Rockford, IU., pp. 241-244. Banford, J. c., Brown, D. H., McConnell, A A, and Smith, W. E. (1981). Resonance Raman spectra produced by the reaction of 5, 5'-dithiobis (2-nitrobenzoic acid) with glutathione and with sulfhydryl groups of biological fluids. Inorg. Chim. Acta 56, Ll5. Baron, M. H., DeLoze, c., Toniolo, c., and Fasman, G. D. (1979). Infrared and Raman study in the solid state of fully protected, monodispersed homooligopeptide of L-valine, L-isoleucine, and L-phenylalanine. Biopolymers 18, 411. Bastian, J I., E. J., and Martin, R. B. (1973). Disulfide vibrational spectra in the sulfur-sulfur and carbon-sulfur stretching region. J. Phys. Chem. 77, 1129. Bellocq, AM., Boilot, J. c., Dupart, E., and Dubien, M. (1973). Analyse conformationnelle de I'hormone hypothalamique TRF par spectroscopie Raman. C. R. A cad. Sci. Paris 276, 423. Bicknell-Brown, E., Lim, B. T, and Kimura, T (1981). Laser Raman spectroscopy of adrenal iron-sulfur apoprotein: The anomalous tyrosine residue at position 82. Biochem. Biophys. Res. Commun. 101, 298. Bieker, L., and Schmidt, H. (1979). Raman spectra of N-formylkynurenine derivatives of lysozyme produced by ozone oxidation. FEBS Lett. 106, 268. Biscar, J. P. (1976). Photon enzyme activation. Bull. Math. Bioi. 38, 29. Biscar, J. P., and Kollias, N. (1973). Pseudo-Raman behavior of the scattering broad band of BSA Polymer Letters Edition 11, 725. Biscar, J. P., and Malone, R. (1975). Laser-enzyme investigations. In Proc. Laser 75, Munich, West Germany. Biscar, J. P., Dhall, P., Pennison, J. (I 972a). Raman behavior of bovine serum albumin. Chem. Phys. Lett. 14, 569.
110
Proteins
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·,~ ;"
~f
5"1\::
:}~
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j~tTobin, M.
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Ii
}i,)Tomimatsu, Y., Kint, S., and Scherer, 1. R. (1976). Resonance Raman spectra of iron(I1I)-, ,i,;: copper(II)-, cobalt(I1I)-, and manganese(I1I)-transferrins of bis(2,4, 6-trichlorophenolato)di,:" imidazolecopper(II) monohydrate, a possible model for copper(II) binding to transferrins. '1' Biochemistry 15, 4918. y.Tu, A. T (1973). Neurotoxins of animal venoms: Snakes. Ann. Rev. Biochem. 42, 235. Tu, A. T (1977). Venoms: Chemistry and Molecular Biology, Wiley, New York. .Tu, A. T, and Hong, B. S. (1971). Purification and chemical studies of a toxin from the venom of Lapemis hardwickii (Hardwick's sea snake). J. Bioi. Chem. 246, 2772. Tu, A. T, and Toom, P. M. (1971). Isolation and characterization of the toxic component of Enhydrina schistosa (common sea snake) venom. J. Bioi. Chem. 246, 1012. Tu, A. T., Hong, B. S., and Solie, T N. (1971). Characterization and chemical modifications of . toxins isolated from the venoms of the sea snake Laticauda semifasciata, from Philippines. Biochemistry 10, 1295. Tu, A. T., Jo, B. H., and Yu, N.-T (1976a). Laser Raman spectroscopy of snake venom neurotoxins: Conformation. 1m. J. Pept. Prot. Res. 8, 337. Tu, A. T, Prescott, B., Chou, C. H., and Thomas, Jr., G. J. (1976b). Structural properties of Mojave toxin of Crotalus scutulatus (Mojave rattlesnake) determined by laser Raman spectroscopy. Biochem. Biophys. Res. Comrnun. 68, 1139. Tu, A. T, Bjarnason, J. B., and Hruby, V. 1. (1978). Conformation of oxytocin studied by laser Raman spectroscopy. Biochim. Biophys. Acta 533, 530. Tu, A. T, Lee, 1., Deb, K. K., and Hruby, V. J. (1979). Laser Raman spectroscopy and circular dichroism studies of the peptide hormones mesotocin, vasotocin, lysine vasopressin, and arginine vasopressin: Conformational analysis. J. Bioi. Chem. 254, 3272. Twardowski, J. (1978). Laser Raman spectroscopy of acid phosphatase from rat liver. Biopolymers 17, 181. Van Wart, H. E., and Scheraga, H. A. (1976a). Raman spectra of cystine-related disulfides. Effect of rotational isomerism about carbon-sulfur bonds on sulfur-sulfur stretching frequencies. J. Phys. Chern. SO, 1812. Van Wart, H. E., and Scheraga, H. A. (1976b). Raman spectra of strained disulfides. Effect of rotation about sulfur-sulfur bonds on sulfur-sulfur stretching frequencies. J. Phys. Chern. SO, 1823. Van Wart, H. E., Lewis, A., Scheraga, H. A., and Saeva, F. D. (1973). Disulfide bond dihedral angles from Raman spectroscopy. Proc. Nat. Acad. Sci. 70, 2619. Van Wart, H. E., Scheraga, H. A., and Martin, R. B. (1976). Agreement concerning the nature of the variation of disulfide stretching frequencies with disulfide dihedral angles. J. Phys. Chern. 80, 1832. Venkatachalapathi, Y. V., and Balaram, P. (1982). Personal communication. Verma, S. P., and Wallach, D. F. H. (1977). Changes of Raman scattering in the CH-stretching region during thermally induced unfolding of ribonuclease. Biochern. Biophys. Res. Cornmun. 74,473. Vorotyntseva, T 1., Surin, A. M., Trakhanov, S. D., Nabiev, I., and Antonov, V. K. (1981). Spectral properties of the leucine-isoleucine-valine-binding protein and its complexes with substrates. Biorg. Khim. 7, 45. Wallach, D. F. H., Graham, J. M., and Oserhoff, A. R. (1970). Application of laser Raman spectroscopy to the structural analysis of polypeptides in dilute aqueous solution. FEBS Lett. 7,330.
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CHAPTER
A --rmr
Enzymes and Immunoglobulins An enzyme-catalyzed reaction is a dynamic process involving the interaction of an enzyme E and substrate S to form an intermediate complex of enzyme-substrate ES, which yields the product P and regenerated free enzyme. By introducing an active chromophore group to a substrate, S
+E
->
ES
->
P
+E
one can apply the resonance-Raman-labeling technique to clarify the enzymereaction mechanism. There is a good review article by Carey (1981), which the reader is advised to see. 1.
1.1.
ENZYME ACTION PAPAIN
Hydrolysis of a protein with the enzyme papain involves a cysteine residue of papain. For both chymotrypsin and papain, ester hydrolysis proceeds through 117
118
Enzymes and Immunoglobulins
a covalently linked acyl-enzyme intermediate:
papain-SH
+-
Rm
Rn
Rm
I
I
I
CH-CONH- CH- ---+
-
Rn
CH-CO-S-papain
Rm
+ H2 N-
I CH-
Rm
I - CH-CO-S-papain
+ H 20
I ---+
-CH-COOH
+ papain-SH
Several synthetic compounds can be used as substrates for papain action. For resonance Raman spectroscopy, the substrate should contain an ester bond that can be hydrolyzed by papain, and also a chromophore that can be detected by resonance Raman. An example of such a substrate is 4-dimethylarnino-3-nitro-( a-benzarnidocinnamate) (Figure 4.1). The resonance Raman spectrum of the intermediate ES is totally distinct from that of the substrate or product. The ES complex gives an intense peak at H
IC~12NOb=T-COOCH3 I NHC04>
N
~ Papain
H
ICH3 12N Q t N O 2
=~- COS-Papain I
+
NHC04>
CH 0H 3
H 0 2
H
ICH31:ro-~=~-COOH N
+
Papain-SH
NHC04>
O2
FIGURE 4.1. A synthetic substrate, 4-dimethylarnino-3-nitro-(a-benzarnidocinnarnate), for papain action. The nitrobenzene group can be resonance enhanced and monitored during hydrolysis.
....
Enzyme Action
119
~ III
I
ENZYME-SUBSTRATE COMPLEX
i"'
III
~
~
i
FIGURE 4.2. Resonance Raman spectra of the catalytic intermediate 4-dimethylamino-3-nitro(a-benzamido cinnamoyl)-papain (top) and the substrate methyl 4-dimethylamino-3-nitro-( abenzamido cinnamate) (bottom). S indicates solvent bands. The figure was reproduced from Carey and Schneider (1978) by permission of the copyright owner, American Chemical Society.
1570 cm- I that is not observed in either free S or P (Figure 4.2). The interaction with the active site produces drastic changes in the properties of the acyl residue. This change is a direct expression of the catalytically important forces that are present in the active site (Carey et al., 1978; Carey and Schneider, 1978). Resonance Raman spectra indicate that there is a rearrangement of the
o II
a-benzamido group in the bound substrate, NH- C-Phe
becoming
ox I
- N = C - Phe. Therefore, it is concluded that deacylation proceeds from an acyl-residue structure differing from that of the substrate in solution (Carey et al., 1976). S
II If the substrate contains thioester -C-OR instead of ordinary ester o S
II
II
- C-OR, the intermediate should have a thioacyl group, - C-S-, which is a new chromophore; thus it should give a resonance-enhanced spectrum (Figure 4.3). Therefore, using a single atom replacement, during enzymolysis the catalytically crucial bonds in the ester moiety can be monitored with resonance Raman spectroscopy (Storer et al., 1979).
120
Enzymes and Immunoglobulins
5
OCONHCH2~-OCH3 H 5- papain
5
OCONHCH2~-S-
papai n
+
CH30H
H 0 2
o
II CONHCH C -SH 2
0>
+
. Papal n-SH
5 II
FIGURE 4.3. Dilhioacyl compounds can also be used as substrates for papain. The - C - 5 group is a cbromophore that can be resonance enhanced.
1.2.
CHYMOTRYPSIN
In proteolysis by chymotrypsin, a serine residue of the enzyme is involved in catalysis. The same reagents used for papain can be used with resonance Raman spectroscopy for elucidation of the mechanism of chymotrypsin catalysis (Carey and Schneider, 1974, 1976; Phelps et al., 1981). For this study, 4-amino-3-nitrocinnamoyl ester is used as a substrate, and the cinnamoyl group is covalently attached at serine-195 of a-chymotrypsin (Figure 4.4). The spectrum of the acyl enzyme ES shows absorption at 280, 340, and 430 nm. The 280-nm peak is due entirely to chymotrypsin, whereas the other two peaks are due to the enzyme-bound substrate. Therefore, resonance Raman spectra can be obtained using an excitation laser wavelength of either 340 or 430 nm. Similarly, 2-hydroxy-3-nitro-a-toluyl ester can also be used as a substrate to make the corresponding acylchymotrypsin. At pH 3, there is no change in the resonance Raman spectra of ES and S. Thus at pH 3 the conformation of the acyl group in the active site is not perturbed by forming an intermediate complex. At this pH, chymotrypsin is inactive, and the acyl enzyme ES is stable for days. However, on raising the pH to 7.0, the enzyme becomes active. The acyl enzyme at this pH has a
Enzyme Action
121
1 1625cm
H2N~CH=CH-COOR
+ HO-Ser-a-Chymotrypsln
N02
S
I
E -ROH
H2N.p-eH=CH-COO -Ser-a- Chymotrypsin N0 2
E-S H 20
H2N"9-CH=CH-COOH
+ HO-Ser-a-Chymotrypsln
N0 2
P
E
FIGURE 4.4. Enzymatic reaction scheme of hydrolysis of 4-amino-3-nitro-trans-cinnamic acid S catalized by chymotrypsin E. Using a resonance-Raman-active chromophore in the substrate, one can study the mechanism of an enzyme reaction by means of resonance Raman spectroscopy.
half-life of seconds. At an active pH, there is a drop in the intensity of the 1625 cm- I band due to the C=C stretching vibrational mode. Thus it can be concluded that twisting occurs about the C=C-C=O single bond in the catalytic process. In other words, distortion takes place in the bond near the linkage undergoing cleavage as shown here:
o II C=C-C- OR i i twist cleavage
1.3.
CARBOXYPEPTIDASE A
Many azo compounds are known to bind to proteins; these bindings can be studied by resonance Raman spectroscopy (Kim, 1975; Thomas and Merlin,
122
Enzymes and Immunoglobulins
1979). When azo compounds bind to enzymes, this allows the study of the enzymes' catalytic mechanisms. Carboxypeptidase A is a metalloenzyme containing zinc. Chemical modification of the enzyme with diazotized p-arsanilic acid (Figure 4.5) modifies tyrosine-248 with retention of catalytic activity. Since the reagent contains the azo group, -N=N-, the binding of the reagent to the enzyme generates a visible chromophore that can be studied by resonance Raman spectroscopy, since it shows a band at 1400-1500 cm- 1 (Lorriaux et al., 1979; Merlin and Thomas, 1979). On complexing, the bands responsible for C-O and N=N groups are shifted or changed in intensity. From these results, it is proposed that the microenvironment of the carboxypeptidase-arsanil complex in the active site is as shown in Figure 4.5. The same technique was applied to aldolase to make resonance-Raman-active azoaldolase (Masetti et al., 1976). The azo group is in the trans conformation, as this gives the - N = N stretching mode at 1440 cm-], whereas in the cis form the band appears at 1500 cm- I . The fact that complex formation decreases the N=N stretching vibration at 1440 by only 4 cm-] implies that the interaction of Zn(II) with the N=N group occurs through a lone pair of electrons rather than through a 'IT-orbital (Scheule et al., 1977, 1979). Crystal and aqueous samples of azocarboxypeptidase show different resonance Raman spectra at N=N (1380-1450 cm -]) and N -If> (1150-1210 cm-]) stretching vibrations, suggesting they have different azo Tyr-248 structure in the two phases. In aqueous solution, phenolic oxygen ionizes, depending on pH. In crystals, the phenolic proton of Tyr-248 is involved in an intermolecular hydrogen bond to a protein group. This interaction may be related to the marked reduction of the enzyme's activity brought about by crystallization (Scheule et al., 1980). Methyl orange binds to bovine serum albumin, and the azo group in this complex is also in a
9"'
N L--o-N
zn[:I[),
-
o
~
CH2
!J
~ ~ ~ .. ~~
FIGURE 4.5. Structure of the azotyrosyl(248) zinc complex of arsanilazotyrosyl(248)-carboxypeptidase A deduced from its resonance Raman spectra. The figure was redrawn from Scheule et aI. (1977).
Enzyme Action
123
trans conformation, as the N =N stretching vibration bands appear at 1415 and 1423 cm- 1 (Carey et al., 1972; Kim et al., 1975). It is, therefore, the trans conformation that is the likely structure in a complex involving the azo group. The distribution of the hydrogen-bonding system of azotyrosine-248 in water is perturbed by many inhibitors (Scheule et al., 1981). 1.4.
PEROXIDASE
The ES complex is the intermediate compound before a product is formed in an enzyme-catalyzed reaction. Yet the identification of this intermediate complex is usually very difficult because of the unstable nature of the complex. Peroxidase catalyzes the conversion of hydrogen peroxide to water in the presence of a hydrogen donor: H 20 2
+ AH 2 -> 2H 20 + A
The reaction proceeds through intermediate compounds I and II. It is known that horseradish peroxidase forms a stable intermediate complex in the absence of a hydrogen donor. The reddish brown color of peroxidase solution becomes olive green with the addition of H 20 2 (compound I or complex I), then it turns to pale red (compound II or complex II). Compound II accepts hydrogen from a donor, AH 2 , and eventually produces water. H 20 2 + 1ePeroxidase Compound I Compound II (Fe3+) 1AH 2 +le2H 20
(Fe3+ ) Resonance Raman spectroscopy can be used to study the intermediate compound I. The spectra of peroxidase and compound I are indeed different, indicating that the addition of hydrogen peroxide significantly alters the heme 7T-ring system (Woodruff and Spiro, 1974). The oxidation-state marker at 1382 cm-\ can be seen, but the Fe-O band observed in hemoglobin cannot be observed in compound II (Rakshit et al., 1976). Like peroxidase, myoglobin can form a stable complex with hydrogen peroxide. The resonance Raman spectrum indicates that the iron atom in the complex is formally in the Fe(IV) state with a low spin configuration (Campbell et al., 1980). 1.5.
THYMIDYLATE SYNTHETASE
Before a product is formed from a substrate, the intermediate complex ES is first produced. It is always curious to know whether there is any change in
124
Enzymes and Immunoglobulins
conformation of an enzyme when it forms a complex with a substrate. Raman spectroscopy can be used to study this as long as the amide I and III bands are not masked by the vibrational bands from the substrate moiety. The reaction catalyzed by thymidylate synthetase is as follows: thymidylate synthetase Deoxyuridate + 5, lO-methylenetetrahydrofolate - - - - - - - thymidylate + dihydrofolate 5, lO-Methylenetetrahydrofolate serves as a coenzyme in the reaction. In this case the intermediate is a ternary complex of the enzyme-substrate-coenzyme. The enzyme itself gives the amide I band at 1655 cm- I with a shoulder at 1680 cm- I . Upon forming a ternary complex, the amide I band remains at 1678 and 1660 cm -I. This is interpreted to mean that no change occurs in the overall secondary structure on formation of the ternary complex (Sharma et aI., 1975). Unfortunately, in these experiments the amide III band of the enzyme was not clear; thus they could not use the amide III band to verify this conclusion. 1.6.
OTHERS
Resonance Raman scattering can be observed when p-nitrophenylphosphate is mixed with rat-liver acid phosphatase, because they form a complex. It seems that histidine and aspartic acid residues are involved in the formation of the enzyme-substrate complex (Twardowski and Proniewicz, 1980). Alcohol dehydrogenase catalyzes the following reaction: -CHO
+ NADH + H+
--. -CH 2 0H
+ NAD+
During the formation of the ternary complex of enzyme-substrate-coenzyme, zinc is involved, as the Zn(II)-O stretching band at 363 cm -1 can be observed (Jagodzinski and Peticolas (1981). Because of recent technological advances in laser tubes, one can obtain a laser in the near-ultraviolet range (325-360 nm). Therefore, the furylacryloyl and the thienylacryloyl group can be attached to chymotropsin. From such resonance-Raman-active chromophores, the detailed structural aspect of enzyme-substrate intermediates can be studied (MacClement et aI., 1981). Glyceraldehyde-3-phosphate dehydrogenase catalyzes the oxidative phosphorylation of D-glyceraldehyde-3-phosphate to 1, 3-diphosphoglycerate in the presence of NAD+ via the formation of a covalent acyl-enzyme intermediate in which the acyl group is linked to the sulfur of an active-site cysteine. The compound ,B-(2-furyl)acryloylphosphate (FA) can be used as a substrate analogue, and this compound is resonance enhanced. The enzyme is composed of four identical subunits, and the resonance Raman spectra of FA-enzyme indicate that the complex exhibits heterogeneous binding in the active site (Storer et al., 1981).
Enzyme-Inhibitor Complexes
2.
125
ENZYME-INHIBITOR COMPLEXES
Inhibitors are frequently used to study the mechanism of enzyme reactions, as they sometimes bind to the active site of the enzyme. Unlike the enzyme-substrate complex, the enzyme-inhibitor complex does not undergo further reaction to produce a product and regenerated enzyme. Thus it offers a unique opportunity to study the active site of an enzyme or mechanism of an enzymatic reaction. Raman spectroscopy can be used for the mechanistic study of enzyme -inhibitor reactions. 2.1.
CHYMOTRYPSIN
2-Phenylethane-boronic acid-a-chymotrypsin complex will be used as an example. The inhibitor shows a 685-cm -I band at pH 11 and a 775-cm -I band at pH 5.0. The first band is characteristic of tetrahedral boron, whereas the second is characteristic of trigonal boron. The enzyme-inhibitor complex does not show the 685-cm -1 band, indicating that boron remains in the trigonal form. The inhibition is pH dependent, and this occurs in a subsequent step that leads to a tetrahedral adduct between enzyme and inhibitor, and also gives rise to a 684-cm- I band. Thus Raman spectroscopy shows that inhibition occurs in two steps (Hess and Seybert, 1975).
2.2.
CARBONIC ANHYDRASE
Carbonic anhydrase is a zinc metalloprotein that is strongly inhibited by aromatic sulfonamides. Zinc is essential for enzyme activity. X-ray studies have indicated that the zinc atom is located in a deep crevice that constitutes the active site, and it is coordinated to three histidyl side chains and a water molecule (Liljas et aI., 1972). The sulfonamide inhibitor binds to the enzyme very strongly and forms a very stable complex with a stability constant as high as 10 9 • A possible mechanism of the inhibition by sulfonamide is by blocking the active site of zinc, as shown in Figure 4.6. The binding site of the enzyme-inhibitor complex was studied by resonance Raman spectroscopy. Most unconjugated proteins are resonance Raman inactive. However, by introducing certain chromophore groups into a protein, one can convert resonance-Raman-inactive proteins into active ones. Figure 4.7 shows the compounds attached to enzymes by Kumar et ai. (1974, 1976).
0<Ei
;
-§-NH .....Znlnl, II
o
FIGURE 4.6. Mechanism of sulfonamide inhibition of carbonic anhydrase activity.
126
Enzymes and Immunoglobulins
The compounds I, II, and III all contain sulfonamide groups, as shown by the dotted lines. At the same time they contain an azo group that produces a resonance Raman effect. Resonance Raman study indicates that there is a change in geometry about the sulfonamide sulfur atom when forms enzymeinhibitor complex. From this finding it is postulated that inhibition may be due to the influence of the bound sulfonamide group, which closely mimics the transition state of the reactants in the reversible hydration of CO2, An increase in the frequency of the 1139 (I) and 1131 (II) cm - I bands upon binding and ionization strongly suggests that the bound sulfonamide group is present as S02 NH - . However, a different explanation was offered by Petersen et al. (1977). They concluded that the sulfonamide was bound to the enzyme as -S02NH2 rather than as -S02NH- . Moreover, they concluded that the sulfonamide is bound to the enzyme surface in the acidic (protonated)-S02NH2 form. The Raman spectrum of Neoprontosil (sulfonamide 4'-sulfamylphenyl-2-azo-7-acetamidoI-hydroxynaphthalene-3, 6-disulfonate) bound to the enzyme at pH 9.5 shows a shift in the intense -N=N stretching mode from 1414 (free) to 1407 cm- 1 (bound), suggesting that a slight conformational change about the -N=N single-bond linkage occurs upon binding (Carey and King, 1979). In order to clarify this discrepancy, cadmium-substituted carbonic anhydrase and 15N_ enriched sulfonamide were used by Evelhoch et al. (1981). The result indicates that aromatic sulfonamides, which inhibit carbonic anhydrase, are found with the ionized sulfonamide moiety. They concluded that the report of Peterson et al. (1977) is based on the misassignmen t of the band at ~ 900 cm- I. A
L1 ~~;~~1 -ow"'::::-'" ...:'."";
(C"l,N
II
r~"
"2
'y' --.~~??~
IN ·.. ···_·r··_ZtJ.:i ~
0
N
A
! : ' ; ' S()I~~: III
resona
active c~ce Raman romophore
:~~~c~~st~Oe ~~i~~ enZ)1TIe Sulfonar.l1de
FIGURE 4.7. Structures of three inhibitors of carbonic anhydrase action. Each compound contains a resonance-Rarnan-active chromophore and the enzyme attachment site. (I) 4sulfonamido-4'-dimethylarninoazobenzene; (II) 4-sulfonarnido-4'-hydroxyazobenzene; (Ill) 4sulfonarnido-4'-aminoazobenzene.
)· :{ I
-!- '/
;m>' Enzyme-Inhibitor Complexes
CI
-0-
~ + ,..~~N=N~ NICH 3)2
NH 2 ,
-
NH
FIGURE 4,8.
127
2
Structure of the trypsin inhibitor 4-amidino-4'-dimethylaminoazobeDzene.
comparison of the pH dependence of the resonance Raman spectra of sulfanilazocarbonic anhydrase with pH dependence of the spectra of sulfanilazotyrosine, sulfanilazohistidine, and sulfanilazotryptophan suggests that histidine is the site of modification of the carbonic anhydrase derivative (Li et aL, 1979). 2.3.
TRYPSIN
4-Arnidino-4'-dimethylarninoazobenzene is a competitive inhibitor of trypsin (Figure 4.8). The binding of the inhibitor to a specific pocket of the enzyme narrows the band widths compared with those found with free inhibitor. This can be explained by the freezing of the rotational motions of the inhibitor molecule (Dupaix et al., 1975). This interpretation is in agreement with crystallographic results (Krieger et aL, 1974). Studies of spin-labeled trypsin have shown that the mobility of the labels was decreased when bound to the protein (Berliner and Wong, 1974). 2.4.
OTHER ENZYMES
Zinc ion is required for alcohol dehydrogenase action. Zincon is a dye that forms a one-to-one complex with zinc, and is a coenzyme-competitive inhibitor (Figure 4.9). Since zincon contains an azo group, the binding of zincon to liver alcohol dehydrogenase can be studied by the resonance Raman technique. When the enzyme-inhibitor complex is formed, there are significant changes in the Raman bands at 1500, 1272, and 1100 cm -1, suggesting that zincon is
~
H03S'(jl
yOH
yCOOH
N':::::-N
,...NH ,
~N
C
6 Zineon
fIl
H03S~ y~, 09C~
Znltll
zn
N'::::-N N/NH . , ~
C
6 Zn-zineon
FIGURE 4.9. Structure of zincon, which is a competitive inhibitor for a coenzyme in the alcohol dehydrogenase reaction.
128
Enzymes and Immunoglobulins COOH I H2
I
:x
~ N~
NH2
H 2
N
N
JCHd~
II: rr
H2
:H-00:
~
N
pterldlne portlOn
FIGURE 4.10.
I
'I '\
:I
-
C,NH-CH :I I COOH
:: I'.- amInobenzoic acId portion
: :I
L-glutamate portion
Structure of methotrexate.
complexed with a zinc atom at the enzyme's active site (McFarland et aI., 1975). Protocatechuate 3, 4-dioxygenase [3, 4-dihydroxybenzoage: oxygen oxidoreductase (decycling)] is a nonheme enzyme catalyzing the cleavage of catechols to cis-cis-muconic acid (see Chapter 11, Section 3.1). Various compounds form enzyme-inhibitor complexes. Resonance Raman spectroscopy indicates that some compounds inhibit enzyme activity by attaching to the active-site iron, whereas others attach to the tyrosine side chain. Thus Raman spectroscopy can be used to identify the site of interactions in enzyme-inhibitor complexes (Que and Epstein, 1981). 2.5.
ENZYME-DRUG INTERACTION
Methotrexate (Figure 4.10) is a drug used for the treatment of childhood leukemia. Dihydrofolate reductase catalyzes the NADPH-linked reduction of dihydrofolate to tetrahydrofolate. Methotrexate inhibits this enzyme. Methotrexate is structurally similar to folate and contains p-aminobenzoyl and pteridine groups, which are both resonance-active chromophores. These groups can be excited at 324 and 350 nm, respectively. By using two different wavelengths for excitation, one can monitor the geometric conformations of both chromophores (Ozaki et aI., 1981). The 1685 cm -1 Raman band can be observed in the NADPH-enzyme binary complex but is absent from the NADPH-methotrexate-enzyme ternary complex. This is ascribed to stabilization of the polarized form of the carboxamide of NADPH by hydrogen bonding to the NH and CO groups of the peptide backbone of dihydrofolate reductase in the ternary complex (Dwivedi et al., 1982). Raman spectra reveal that protonation of the pteridine ring occurs when the drug is bound to the enzyme. At pH 7, there is a strong band at 659 cm- 1 that is present only in the drug-enzyme complex (Saperstein et aI., 1978).
3.
ISOZYMES
Some enzymes have different electrophoretic mobility and different physical properties, although they have the same enzyme activity (isozymes). Acid phosphatase from rat liver has two isozymes, I and II. The two isozymes have
Immunology
129
different Raman spectra in the range 10-1800 cm - I (Twardowsky, 1978), illustrating that some isozymes can be differentiated by Raman spectroscopic technique. The difference in the Raman spectra probably originates from different quantities of carbohydrate attached to the apoenzyme moieties. Sialic acid is a prosthetic group of acid phosphatases. After neuraminidase treatment to hydrolyze neuraminic acid, the low-frequency bands below 300 cm- I disappear, and the difference in Raman spectra for the two isozymes is eliminated (Twardowsky, 1979).
4.
IMMUNOLOGY
Antibodies, or immunoglobulins, are Y-shaped molecules consisting of two heavy and two light polypeptide chains. Several disulfide bonds connect these chains. There are several classes of immunoglobulins, such as IgG, IgA, IgM, IgD, and IgE. An immunoglobulin molecule has both "constant" and" variable" regions and the same class of immunoglobulins contains similar "constant regions." 4.1.
ANTIGEN-ANTIBODY REACTIONS
When antigen (Ag) combines with its antibody (Ab) to form an Ag-Ab complex, it frequently precipitates. Raman study indicates that immunoglobulins are composed predominantly of antiparallel-f3-sheet structure, and they change conformation on binding with an antigen so that the protein becomes more disordered (Painter and Koenig, 1975).
HumanIgG Rabbit IgG Human IgM Ovalbumin-rabbit antiovalbumin precipitate
4.2.
Amide I (cm- I )
Amide III (cm- I )
1673 1673 1673 1673
1238 cm1239 1238 1236 } 1248
I
}
Anti parallelf3-sheet A mixture of f3-sheet and random coil
HAPTEN-ANTIBODY INTERACTIONS
Antigenicity is usually a property of proteins and some polysaccharides. Small molecules are usually not antigenic. However, some small molecules, when attached to a protein, do cause production of antibody that can specifically bind to this small molecule. Such a small compound is called a hapten. The
130
Enzymes and Immunoglobulins
H~9;_", ,Q!NO/'----6\
O
-NiT
o SO; \...
,
- -.- -
I
21
I
,
I
\~02 //
I "l
Serve as a
chromophore for resonance Raman spec-
Serve as an active moiety for hapten
troscopy
FIGURE 4.11. Structure of l-hydroxy(2,4-dinitrophenylazo)-2, 5-napthalenedisulfonic acid. This compound has two active functional groups. One is a resonance-Raman-active chromophore and the other is a hapten-active group.
dinitrophenyl group is a classical example of a hapten. An azo compound is a good chromophore for resonance Raman scattering; therefore, the dinitrophenyl group conjugated with the azo group will serve as an antigen as well as a resonance-Raman-active compound. One example of such a conjugated compound is shown in Figure 4.11. The trans form of the N =N stretching vibration is in the region of 1380-1440 cm- I . When the hapten shown in Figure 4.11 is complexed with its antibody, the PN=N increases, suggesting that the C=M bond of the azo linkage is twisted. The barrier to rotation about the Phe-N bond is larger than 7 kcaljmol. Therefore, quite large amounts of distortion energy are involved (Carey et al., 1973). The IgA protein secreted by the mouse myeloma cell line MOPC 315 has a high affinity for e-DNP-L-lysine and related compounds. Since the DNP group is a chromophore for resonance Raman spectroscopy, the DNP-binding sites of MOPC 315 IgA and of MOPC 460 IgA can be studied. All ligands show significant shifts in their Raman bands, but the shifts are different depending on the source of the antibodies. From this it can be concluded the DNP sites in the two proteins are distinct (Kumar et al., 1978). 4.3.
CRYOGLOBULIN
Monoclonal cryoglobulins decrease in solubility as the temperature of sera is reduced from 37 to O°e. They belong to the immunoglobulin classes IgM, IgG, and IgA. It is important to discover whether a conformational change is involved when they undergo cryoprecipitation. Raman spectral analysis indicates that there is no peptide-backbone structural change over the temperature interval of 40 to - 8°C, during which cryoprecipitation of IgM-K McE occurs. Thus the temperature-dependent solubility may be due to changes in the environment of certain aromatic groups (Middaugh et al., 1977, 1978; Thomas et al., 1979; Scoville et al., 1980).
References
131
REFERENCES Berliner, L. J., and Wong, S. S. (1974). Spin-labeled sulfonyl fluorides as active site probes of protease structure. J. Bioi. Chem. 249, 1668. Campbell, 1. R., Clark, R. J. H., Clore, G. M., and Lane, A N. (1980). Characterization of the electronic properties and geometric environment of the iron atom in the "myoglobin hydrogen-peroxide" complex by Soret-excited resonance Raman spectroscopy. Inorganica Chimica Acta 46,77. Carey, P. R. (1981). Resonance Raman spectroscopic studies of transient enzyme-substrate complexes. Can. J. Spectros., 26, 134. Carey, P. R., and King, R. W. (1979). Neoprontosil binding to carbonic anhydrase. Resonance Raman and other studies on the ionization behavior of the sulfonamide. Biochemistry 18, 2834. Carey, P. R., and Schneider, H. (1974). Resonance Raman spectra of chymotrypsin acyl enzymes. Biochem. Biophys. Res. Commun. 57, 83!. Carey, P. R., and Schneider, H. (1976). Evidence for a structural change in the substrate preceding hydrolysis of a chymotrypsin acyl enzyme: Application of the resonance Raman labelling technique to a dynamic biochemical system. J. Mol. Bioi. 102, 679. Carey, P. R., and Schneider, H. (1978). Resonance Raman labels: A submolecular probe for interactions in biochemical and biological systems. Accts. Chem. Res. 11, 122. Carey, P. R., Schneider, H., and Bernstein, H. 1. (1972). Raman spectroscopic studies of ligand-protein interactions: The binding of methyl orange by bovine serum albumin. Biochem. Biophys. Res. Commun. 47, 588. Carey, P. R., Froese, A, and Schneider, H. (1973). Resonance Raman spectroscopic studies of 2,4-dinitrophenyl hapten-antibody interactions. Biochemistry 12, 2198. Carey, P. R., Carriere, R. G., Lynn, K. R., and Schneider, H. (1976). Resonance Raman evidence for substrate reorganization in the active site of papain. Biochemistry 15, 2387. Carey, P. R., Carriere, R. G., Phelps, D. 1., and Schneider, H. (1978). Charge effects in the active site of papain: Resonance Raman and absorption evidence for electron polarization occurring in the acyl group of some acylpapains. Biochemistry 17, 108!. Dupaix, A, Bechet, J.-J., Yon, 1., Merlin, J.-c., Delhaye, M., and Hill, M. (1975). Resonance Raman spectroscopic studies of the interactions between trypsin and a competitive inhibitor. Proc. Nat. Acad. Sci. 72, 4223. Dwivedi, C. M., Plante, L. T., Kisliuk, R. L., Pastore, E. J., Verma, S. P., and Wallach, D. F. H. (1982). Interaction of the carboxamide of NADPH with Lactobacillus casei dihydrofolate reductase. Arch. Biochem. Biohpys. 213, 338. Evelhoch, J. L., Bocian, D. F., and Sudmeier, J. L. (1981). Evidence for direct metal-nitrogen binding in aromatic sulfonamide complexes of cadmium(II)-substituted carbonic anhydrases by cadmium-I 13 nuclear magnetic resonance. Biochemistry 20, 495!. Hess, G. P., and Seybert, D. (1975). Tetrahedral intermediate in a specific a-chymotrypsin inhibitor complex detected by laser Raman spectroscopy. Science 189, 384. Jagodzinski, P. W., and Peticolas, W. L. (1981). Resonance enhanced Raman identification of the zinc-oxygen bond in a horse liver alcohol dehydrogenase-nicotinamide adenine dinucleotidealdehyde transient chemical intermediate. J. Am. Chem. Soc. 103, 234. Kim, B.-K (1975). Interaction of methylorange with some proteins and cationic surfactants. Seoul. Univ. Faculty Papers 4, 75. Kim, B.-K, Kagayama, A, Saito, Y, Machida, K, and Uno, T. (1975). Resonance Raman spectra of methylorange bound to proteins and cationic surfactants. Bull. Chem. Soc. Japan 48, 1394. Krieger, M., Kay, L. M., and Stroud, R. M. (1974). Structure and specific binding of trypsin: Comparison of inhibited derivatives and a model for substrate binding. J. Mol. Bioi. 83, 209.
132
Enzymes and Immunoglobulins
Kumar, K, King, R. W., and Carey, P. R. (1974). Carbonic anhydrase-aromatic sulfonamide complexes, a resonance Raman study. FEBS Lett. 48, 283. Kumar, K, King, R. W., and Carey, P. R. (1976). Resonance Raman studies on some carbonic anhydrase-aromatic sulfonamide complexes. Biochemistry 15, 2195. Kumar, K, Phelps, D. J., Carey, P. R., and Young, N. M. (1978). Resonance Raman spectroscopic studies of the hapten features involved in the binding of 2,4-dinitrophenyl haptens by the mouse myeloma proteins MOPC 315 and MOPC 460. Biochem. J. 175, 727. Li, T-Y., Chen, J. F., Watters, K 1., and McFarland, J. T (1979). Identification of enzyme coupling sites with aromatic diazonium salts-a resonance Raman study. Arch. Biochem. Biophys. 197, 477. Liljas, A, Kannan, K K, Bergsten, P.-C., Waara, I., larupp, 1., Lovgren, S., Petef, M. (1972). Crystal structure of human carbonic anhydrase C. Nature, New BioI. 235, 131. Lorriaux, J. 1., Merlin, 1. c., Dupaix, A, and Thomas, E. W. (1979). Spectres Raman de Resonance de derives parasubstitutes de I'Azobenzene. J. Raman Spectrosc. 8, 81. MacClement, B. A E., Carriere, R. G., Phelps, D. J., and Carey, P. R. (1981). Evidence for two acyl group conformations in some furylacryloyl and thienylacryloy1chymotrypsin: Resonance Raman studies of enzyme-substrate intermediates at pH 3.0. Biochemistry 20, 3438. Masetti, G., Dellepiane, G., and Zerbi, G. (1976). Resonance Raman spectra of azoaldolase and model molecules. In Proc. Int. Conf. Raman Spectrosc. 5th, E. D. Schmid, J. Brandmueller, and W. Kiefer, Eds. Hans Ferdinand Schulz Verlag: Freiburg/Br., Germany, pp. 230-1. McFarland, J. T, Watters, K 1., and Petersen, R. 1. (1975). Resonance Raman investigation of an enzyme-inhibitor complex. Biochemistry 14, 624. Merlin, 1. c., and Thomas, E. W. (1979). Resonance Raman spectroscopic studies of 2-(4'-hydroxyphenylazo)-benzoic acid and some substituted analogs-I. pH effect on spectra. Spectrochimica Acta 35A, 1243. Middaugh, C. R., Thomas, Jr., G. 1., Prescott, B., Aberlin, M. E., and Litman, G. W. (1977). Investigations of the molecular basis for the temperature-dependent insolubility of cryoglobulins. II. Spectroscopic studies of the IgM monoclonal cryoglobulin McE. Biochemistry 16, 2986. Middaugh, C. R., Gerber-Jensen, B., Hurvitz, A, Paluszek, A, Scheffel, c., and Litman, G. W. (1978). Physicochemical characterization of six monoclonal cryoimmunoglobulins: Possible basis for cold-dependent insolubility. Proc. Nat. Acad. Sci. 75, 3440. Ozaki, Y., King, R. W., and Carey, P. R. (1981). Methotrexate and folate binding to dihydrofolate reductase. Separate characterization of the pteridine and p-aminobenzoyl binding sites by resonance Raman spectroscopy. Biochemistry 20, 3219. Painter, P. C., and Koenig, J. 1. (1975). Raman spectroscopic study of the structure of antibodies. Biopolymers 14, 457. Petersen, R. 1., Li, T.-Y., McFarland, J. T., and Watters, K 1. (1977). Determination of ionization state by resonance Raman spectroscopy. Sulfonamide binding to carbonic anhydrase. Biochemistry 16, 726. Phelps, D. J., Schneider, H., and Carey, P. R. (1981). Correlations between reactivity and structure of some chromophoric acy1chymotrypsins by resonance Raman spectroscopy. Biochemistry 20, 3447. Que, 1., and Epstein, R. M. (1981). Resonance Raman studies on protocatechuate 3,4dioxygenaseinhibitor complexes. Biochemistry 20, 2545. Rakshit, G., Spiro, T G., and Uyeda, M. (1976). Resonance Raman evidence for Fe(IV) in compound II of horseradish peroxidase. Biochim. Biophys. Res. Commun. 71, 803. Saperstein, D. D., Rein, A J., Poe, M., and Leahy, M. F. (1978). Binding of methotrexate to Escherichia coli dihydrofolate reductase as measured by visible and ultraviolet resonance Raman spectroscopy. J. Am. Chem. Soc. 100, 4296.
References
133
Scheule, R. K., Van Wart, H. E., Vallee, B. L., and Scheraga, H. A. (1977). Resonance Raman spectroscopy of arsanilazocarboxypeptidase A: Determination of the nature of the azotyrosyl248, zinc complex. Proc. Nat. Acad. Sci. 74, 3273. Scheule, R. K., Van Wart, H. E., Zweifel, B. 0., Vallee, B. L., and Scheraga, H. A. (1979). Resonance Raman spectroscopy of arsanilazocarboxypeptidase A: Assignment of the vibrations of azotyrosyl-248. J. Inorgan. Biochem. 11,283. Scheule, R. K., Van Wart, H. E., Vallee, B. L., and Scheraga, H. A. (1980). Resonance Raman spectroscopy of arsanilazocarboxypeptidase A: Conformational equilibria in solution and crystal phases. Biochemistry 19, 759. Scheule, R. K., Han, S. L., Van Wart, H. E., Vallee, B. L., and Scheraga, H. A. (1981). Resonance Raman spectroscopy of arsanilazocarboxypeptidase A: Mode of inhibitor binding and activesite topography. Biochemistry 20, 1778. Scoville, C. D., Turner, D. H., Lippert, J. L., and Abraham, G. N. (1980). Study of the kinetic and structural properties of a monoclonal immunoglobulin G cryoglobulin. J. BioI. Chern. 255, 5847. Sharma, R. K., Kisliuk, R. L., Verma, S. P., and Wallach, D. F. H. (1975). Study of thymidylate synthetase-function by laser Raman spectroscopy. Biochim. Biophys. Acta 391, 19. Storer, A. c., Murphy, W. F., and Carey, P. R. (1979). The use of resonance Raman spectroscopy to monitor catalytically important bonds during enzymic catalysis. J. BioI. Chern. 254, 3163. Storer, A. c., Phelps, D. J., and Carey, P. R. (1981). Resonance Raman and electronic absorption spectral studies of some ,B-(2-furyl)acryloylglyceraldehyde-3-phosphate dehydrogenases. Biochemistry 20, 3454. Thomas, E. W., and Merlin, 1. C. (1979). Resonance Raman spectroscopic studies of 2-(4'hydroxyphenylazo)-benzoic acid and some substituted analogs - II. Binding to avidin and bovine serum albumin. Spectrochimica Acta 35A, 1251. Thomas, Jr., G. J., Prescott, B., Middaugh, C. R., and Litman, G. W. (1979). Raman spectra and conformational structures of Fabl' and (Fc)51' fragments of cryoglobulin IgM-k McE. Biochim. Biophys. Acta 577, 285. Twardowski, 1. (1978). Laser Raman spectroscopy of acid phosphatase from rat liver. Biopolymers 17, 181. Twardowski, 1. (1979). Laser Raman spectroscopy of acid phosphatase from rat liver after neuraminidase treatment. Biochim. Biophys. Acta 578, 116. Twardowski, 1., and Proniewicz, L. M. (1980). Chemical aspects of the study of the active site of acid phosphatase from rat liver by spectroscopic methods. Zesz. Nauk. Uniw. Jagiellon., Pro Chern. 25, 93. Woodruff, W. H., and Spiro, T. G. (1974). Resonance Raman spectroscopy of reaction intermediates by a rapid mixing, continuous flow technique. Appl. Spectrosc. 28, 576.
CHAPTER
:)
Nucleic Acids
Raman spectroscopy is extensively used for the study of nucleic acid conformation and mechanism of interaction with other compounds. Nucleic acids show many structurally sensitive Raman lines; by using these lines, one can follow the progress of conformational changes or interactions. In nature the majority of nucleic acids occur as nucleoproteins such as ribosomes, chromatins, and viruses. How does a nucleic acid interact with protein? How do some drugs combine with nucleic acids? To answer these questions, Raman spectroscopy is now being utilized more often. With the advances in computer technology, the intensity changes in Raman lines of nucleic acids can be measured more accurately than before. It can be expected that Raman spectroscopy will playa dominant role in such studies in the future. 1.
BACKGROUND
Nucleosides, nucleotides, and polynucleotides are all constituted of purine and pyrimidine bases. One really cannot discuss the Raman spectroscopic properties of bases alone. It is best to discuss some of the properties of purine and pyrimidine bases together with those of nucleosides, nucleotides, and nucleic acids. In this section a few selected topics are discussed.
Background
1.1.
135
TAUTOMERISM
The structures of purine and pyrimidine bases can be written in keto or enol form. An important question is which form guanine, inosine, and cytosine take. Raman spectroscopy solves this problem readily, as these compounds show an intense keto band at 1670 em- I and other characteristic bands associated with the keto group. Therefore, guanine, inosine, and cytosine in nucleosides and nucleotide derivatives exist in the keto form (Lord and Thomas, 1968; Medeiros and Thomas, 1971a). However, the free bases can show tautomerism in solution depending on the dielectric constant of the solvent, the temperature, and the ionic strength of the solution. Detailed Raman analyses of free uracil and thymine have been made to differentiate between tautomers (Lippert, 1979).
1.2.
HYDROGEN-DEUTERIUM EXCHANGE
The hydrogen atom at C-8 of purine in nucleic acid can be exchanged with deuterium. In poly(A), C(8)-H and C(8)-D deformation vibrations appear at 1485 and 1462 cm- I , respectively (Livramento and Thomas, 1974) (Figure 5.1). Using the measurement of intensity changes at 1485 and 1462 cm- 1 at different temperatures, it is possible to measure parameters of hydrogendeuterium-exchange kinetics such as the activation energy and the frequency factor for different nucleotides (Thomas and Livramento, 1975; Lane and Thomas, 1979). Similarly, the C(8)-H of GMP and cyclic GMP can be exchanged with deuterium, and this can also be followed by Raman spectroscopy (Thomas and Lane, 1980). Ringland et al. (1979) attempted to use the slow C-8 exchange to obtain information on the tertiary structure of calf-thymus chromatin. 1.3.
ORIGIN OF BASE VIBRATIONS
Exact assignment of every vibrational band for all nucleosides and nucleotides is not an easy matter. By using deuterium and 15N isotopic substitutions, some assignments can be made. For instance, the Raman spectrum of guanosine can
~N ~.,A>-H N N I
R FIGURE 5.1.
N0
°2° H0 2
6:>-0 2
I
R
Hydrogen-deuterium exchange in adenine.
136
Nucleic Acids Purine ring
'"
r·
~
1--<1,...............-
keto group
1--'' ilH'N~N 1....1- i1 -- ,.,..""" ~
0
I
II
I
..~ _.~
!..1
I
.I i + - -.' - ,.I
pyrlm,d,ne
_ .._..•.•...•__.•....••. ~~ I 0
CHpHi '
'
~?••••_.•.••_ OH .•••...3I
part
FIGURE 5.2.
!
>
ribose
Vibrational modes of different parts of guanosine.
be traced to C=O stretching, overall ring vibration, the pyrimidine part of the purine ring, the imidazole part of the purine ring, and ribose (Figure 5.2). Oelabar and Guschlbauer (1979) assigned the guanosine vibrations as follows: Pyrimidine-portion vibrations Purine-ring vibrations Ribose-ring vibrations
1605 and 1577 cm- I at 60°C and 1585 cm- I at 10°C 1540, 1323, 1180, and 650 cm -I at 60°C and 1540,1325,1182, and 650 cm- I at 10°C 1080, 1030, and 865 cm - I at 60°C and 1080, 1030, and 870 cm - I at 10°C
The molecular breathing vibration usually appears at lower frequency. The 651 cm- I band of guanine is assigned to the ring-breathing vibration (Majoube, 1978). Some bases such as uracil, guanine, cytosine, and hypoxanthine contain carbonyl groups that show bands in the region of 1500-1700 cm -I. The band from the carbonyl group in uracil is much more intense than that from the carbonyl group in guanine and cytosine; thus it dominates the spectra of RNA (Rice and Thomas, 1972). Thymine contains two C=O groups; the one at C-2 shows a band at the highest frequency, and the C(4)=0 shows a band at slightly lower frequency, in the region of 1671-1674 em-I. Similarly, uracil also contains two C=O groups. The C(2)=0 of poly(D) shows a band at the higher frequency of 1695 cm- I , and C(4)=0 shows a band at 1658 cm- I at 25°C in 020. At 5°C in 020' the C(2)=0 band appears at 1698 em-I, and the C(4)=0 gives a doublet at 1675 and 1641 em -I. In water at 25°C, poly (D) gives two C=O bands at 1686 and 1629 em-I (Schmid and Gramlich, 1979). Temperature definitely has an effect on the C=O band. Lower temperatures tend to give better resolution. Examples that show the temperature effect are GpA and ApG. They show the C=O stretching at 1689 cm- 1 at 45°C, whereas at 5°C
137
Principal Raman Lines
the C=O bond exhibits two distinct stretching vibrations at 1725 and 1680 cm - I for both GpA and ApG. This change probably reflects the different states of hydrogen bonding at different temperatures. There are several chemical groups that give C-O stretching vibrations; they often have slightly different frequencies, which can be used to detect different functional groups. The range in keto stretching vibrations of different functional groups is summarized here:
Carboxyl
°II
-C-O-H
°II Carboxylate ion
-C-O-
1660-1740 cm- I asym 1340-1440 cm-
1
asym 1550-1690 cm- I
°II Ester
-C-OR
Amide
-C-NH 2
°II °II
Ketone
C-C-C
°II Aldehyde
H-C-C
1700-1746 cm- I
~
1675 cm- I
1700-1720 cm- I
1710-1740 cm- I
The C=O stretching vibration usually gives a prominent band. The band is quite stable and does not fluctuate in different solvents. For instance, the bands from the NADH nicotinamide C=O stretching vibration in H 2 0, D 2 0, and dimethyl sulfoxide all appear at 1689 cm- I (Patrick et aI., 1974). The primary-amide C=O band is different for NADH and NAD. The latter gives a weaker C=O stretching band at 1698 cm- I (Forrest, 1976).
2.
PRINCIPAL RAMAN LINES
Before we study the properties of nucleic acids and their derivatives or their interaction with other compounds, it is essential to know what each Raman band means. The band assignment, therefore, is important. This section looks into the origin of nucleic acid Raman bands. The next section discusses the Raman bands that are most useful in the study of nucleic acid structures and properties.
138 2.1.
Nucleic Acids
AQUEOUS SOLUTION
There are many structurally sensitive Raman bands in spectra of nucleic acids. Because nucleic acids usually contain at least four different types of bases, the band assignments are more complicated than individual nucleotide. Synthetic polynucleotides are convenient to study because some contain only one base. Before going over the structurally sensitive bands of nucleic acids, it is best to summarize the Raman bands and their assignment for synthetic polynucleotides in aqueous solution. The detailed assignments of the bands in spectra of poly(G), poly(A), poly(l), poly(C), and poly(U) are shown in Table 5.1. In the detailed band assignment of dA and dT, it is shown that the bands in the region from 1240 to 1700 cm - I are exclusively due to base vibrations, and bands in the 1000-1240 cm - I region are due to the backbone con tribu tions. For instance, a PO; antisymmetrical stretching vibration appears at 1240 cm - I, and a PO; symmetrical stretching vibration appears at 1100 cm - '. Some deoxyribose vibrations also appear in this region (Baret et a1., 1979). TABLE 5.1.
Assignment of Raman Frequencies of Synthetic Polynucleotides
Poly(A)
Poly(G)
Poly(l)
Poly(C)
Poly(U)
(em-I)
(em-I)
(em-I)
(em-I)
(em-')
Vibrations of Bases
705 725 1228 1252 1303 1336 1337 1424 1483 1508 1576
675 691 722 782 1207 1242 1267 1326 1330 1360 1367 1390 1419 1483-1485 1535-1543 1583-1585
723 1269 1323 1352 1384 1422 1518 1556 1594
750 783-784 790 991 1194 1248-1256 1290-1292 1365 1384 1408-1410 1527 1547
783 . 1233 1399 1476
Ribose
1466
1135 1470
1464
1144 1466
810
800
1097-109R
1094
Phosphate Diester Symmetric Stretch
811
819-820
794 (disordered)
Ionic (O-P-O) Symmetric Stretch IOQ'\
IOR7-IOQ6
10QI
i
IlTABLE 5.1. ·h"
Poly(A) (em-I)
'IJ ,,','
,Ie
Continued Poly(G) (em-I)
Poly(I) (em-I)
Poly(C) (em-I)
Poly(U) (em-I)
/.'
..
1/
Ribose-Phosphate
533 635 868 912
408 429 505 588 640 975
529 559 822 912 969 1158
429 545 596 601 625
556 639 997
644
710 760 845 866 915-916 975 1008 1047 Base External C-N Stretch
il79
1183
1042
915 (C-O and C-N) 1025-1026 1046 1070 (C-O and C-N)
1180 C-O Stretch
1050
1042
C-C and C-N Stretch
1100 1175 1220 1245 C-H Deformation
1465
1462 C=O Stretch
1608 1680
1680
Source: The table was constructed based on the papers of Petico1as (1971), Rice et al. () 973) and Chou and Thomas (1977).
TABLE 5.2.
Poly(rA)
360(0) 530(0) 560(0) 585(0)
643(1) 700(S) 716(5) 760(0) 810(4) 855(1) 870(0) 912(0) 985(0) 1043(0) 1094(3) 1185(4)
Raman Frequencies of POly(A), Poly(C), and Poly(U) In 0 20
Poly(rC) 275(0) 310(0) 362(1) 425(0) 547(0) 600(1) 613(S) 650(0) 705(0) 750(2) 775(10) 810(4) 840(0) 870(0) 910(0) 982(0) 1035(1) 1088(S) 1098(2) 1132(0) 1175(4) I 190(?)
1218(S) 1260(0) 1303(7) 1341(10) 1380(2)
1253(8) 1293(7) 1325(0) 1375(0)
1425(0) I463(S) 1480(5) 1520(1) 1550(0) 1572(5) 1623(0)
1415(0) 1462(1) 1503(3) 1520(2)
1618(3) 1650(2)
Poly(rU)
415(0) 550(2) 570(S) 620(1) 637(S) 710(0)
778(10) 797(4) 865(0) 915(0) 975(0) 1045(0) 1094(2) 1136(3) 1190(?) 1217(7) 1237(S) 1252(10) 1300(5) I380(S) 1400(3) 1422(0) I462( I)
1620(2) 1662(10) 1695(6)
Assignment a Ring Ring Ring Ring Ring Ribose Ribose Ribose Ribose Ribose Ribose Ring Ribose Ring OPO sym. str. Ribose-phosphate Ribose Ribose Ribose Ribose Ring P0 2 sym. str. Ring Ring Ring Ring Ring Ring Ring Ring Ring Ring Ring CH def. Ring Ring Ring Ring Ring C=O str. C=O str.
Source: The table was reproduced from Prescott et al. (1974). "Abbreviations: sym., symmetrical; str., stretching; def., deformation; s, shoulder. Numbers in parenthesis give relative line intensities on 0 to 10 scale.
Structurally Sensitive Lines
141
-;
.' 3.
STRUCTURALLY SENSITIVE LINES
There are many structurally sensitive Raman lines for different nucleotides and nucleic acids. Some originate from the vibrations associated with particular bases; some are associated with the sugar-phosphate-backbone bond (815-cm - I line). We will discuss the Raman bands that are useful in nucleic acid research.
3.1.
HEAT TREATMENT
Heating is one good way to alter the structure of nucleic acids. Structural changes of polynucleotides can be followed by measuring the intensity changes of some base vibrational bands. Raman lines sensitive to heat treatment are summarized in Table 5.3. Structurally sensitive lines are not restricted to nucleic acids and their components, but can also be found in spectra of special nucleotides. In NAD and NADH, the 1400-cm- 1 band is assigned to the nicotinamide-ring vibration, which is also sensitive to conformational changes (Barrett, 1980).
3.2.
PH TREATMENT
Some Raman lines are sensitive to pH. These lines are usually associated with functional groups or atoms involved in hydrogen bonding, or the atoms may be protonated or deprotonated. These lines are summarized in Table 5.4.
3.3.
PHOSPHODIESTER-BOND VIBRATIONS
There are two types of phosphodiester-stretching vibrations. One is the phosphodiester-stretching vibration and another is the phosphoionic bond-stretch-
TABLE 5.3.
Compound Uracil Cytosine Guanine
CMP-5'
Inosine, IMP
Poly(C)
Poly(A)
Poly(I) Poly(U)
Structurally Sensitive Raman Lines (Heat Treatment) Raman Line (em-I)
Assignment
785 1235 780 1280 670 1485 1580 783 1243 1530 723 820 1350 1382 1518 1554 1594 790
Base stacking O-P-O Base stacking Base stacking Base stacking Base stacking Base stacking Ring mode
1256
Ring mode
1547
Double-bond stretch
720 725
Base stacking Ring mode
1303
Ring mode
1380 1508 1520 722 790 1236 1240 1660
Ring mode Double-bond stretch Double-bond stretch Ring mode
Reference Arie et aI. (1971) Arie et aI. (1971) Arie et aI. (1971) Arie et aI. (1971) Arie et aI. (1971) Arie et aI. (1971) Arie et aI. (1971) Lord and Thomas (I 967a, b) Lord and Thomas (1967a, b) Lord and Thomas (1967a, b) Medeiros and Thomas (l97Ib) Medeiros and Thomas (1971 b) Medeiros and Thomas (1971 b) Medeiros and Thomas (1971 b) Medeiros and Thomas (1971 b) Medeiros and Thomas (1971 b) Medeiros and Thomas (1 971 b) Small and Peticolas (1971 a) Peticolas (1975) SmaIl and Peticolas (1971 a) Peticolas (1975) Small and Peticolas (1971 a) Peticolas (1975) Tomlinson and Peticolas (1970) Small and Peticolas (1971 a) Peticolas (1975) SmaIl and Peticolas (1971 a) Peticolas (1975) Small and Peticolas (1971 a) Small and Peticolas (1971 a) Peticolas (1975) SmaIl and Peticolas (1971 a) Peticolas (1975) SmaIl and Peticolas (1971 a) Peticolas (1975) SmaIl and Peticolas (1971 a)
,
;..:~,;;
.~i:
I~'~":
;~:fTABLE
i,I\f_'
5.4.
~n\l'
••~ <;,', "'.
'i4
,7£'5, Compound
Structurally Sensitive Raman Lines (pH Change)
Frequency (cm - I)
Change
_
Reference
1,1-'- - - - - - - - - - - - - - - - - - - - - - - - - - - - - - -
;,~,~,,';';'Cytidine
1180-1330
:4,i fi}f
·i~i}
:!~i
i
',~\,Uridine
?~~1 \'~~:. ;.'J
1444 1543 591 1017 1230 1293 1397
AMP-5'
980 1253 1378 1407 1483 1581
GMP-5'
1412
Poly(A)
1488 1533 1251 1378 1407 1560 1581
Poly(C)
1170-131O 1527
Increases greatly in intensity and becomes one band on protonation Increases in intensity at lower pH Shifts to 1527 cm - I at low pH Disappears at lower pH Disappears at lower pH Increases greatly in intensity at lower pH Disappears at lower pH Increases in intensity at lower pH, disappears at higher pH Disappears on protonation Disappears on protonation Shifts to 1407 cm - I on protonation Shifts to 1378 cm - I by increasing pH Disappears on protonation Shifts to 1560 em - I on protonation Increases in intensity on protonation Disappears on protonation Disappears on deprotonation Decreases in intensity on protonation Decreases in intensity on protonation Increases in intensity on protonation Increases in intensity on protonation Decreases in intensity on protonation Increases with degree of protonation Hypochromic on protonation, due to cytosine ring vibration
O'Connor et al. (1976a)
O'Connor et al. (1976a)
O'Connor et aI. (1976b)
O'Connor et al. (1976b)
O'Connor et aI. (1976b)
O'Connor and Scovell (1981)
144
Nucleic Acids
ing vibration. This can best be seen from:
o -11- 0 - P-O-
I
Phosphodiester bondstretching vibration
0-
t o
0
I:
II
1
- 0 - P - 0 - or - 0 - P - 0 ~ I 0o-
j
Phosphoionic bondstretching vibration
j
The phosphodiester stretching vibration is sensitive to conformational changes and usually appears at around 815 cm- '. On the other hand, the phosphoionic bond stretching vibration which appears at around 1100 cm - I is relatively insensitive to structural changes; therefore, this band is used as a standard line with respect to other Raman lines. Raman spectroscopy is a useful tool to elucidate the conformation of double-helix polynucleotides (for synthetic as well as natural ones). Since the phosphate group is connected to ribose or deoxyribose, the phosphodiester vibration is also referred to as the phosphosugar stretching vibration. For instance, in poly(G) . poly(C), the ratio 1815 /1 1100 can be used as an index of the fraction of A-type conformation (Brown et aI., 1972). Because the 815-cm-, band is due to phosphodiester-bond vibrations in a double-stranded helix, and not base vibrations, the band indicates the amount of ordering of the helix, and it is not influenced by the types of bases involved. In a completely ordered ribonucleic acid, the ratio 1815 /11100 is 1.64 (Yo). From the ratio 1815 /11100 (y) observed for a given RNA divided by 1.64 (Yo), one can calculate the fraction of ordered structure (Thomas et aI., 1973b; Thomas and Hartman, 1973). In single-stranded polynucleotides, the 815-cm- 1 band is either absent or very weak (Aylward and Koenig, 1970). The presence of the 815-cm-, band may indicate stability in single-stranded polynucleotides. For instance, poly(U) in the presence of Mg(II) shows a band at 814 cm-' at oDe, but the band disappears at 20 0 e (Small and Peticolas, 1971 b). Poly(rG) forms a single strand, but has a relatively stable secondary structure and shows a weak band at 815 cm - '. This band becomes more prominent in the presence of high salt concentrations (Rice et aI, 1973). Further evidence that the 815-cm- 1 band indicates the amount of ordered structure is that the band disappears on heating (Morikawa et a1., 1973; Peticolas and Lippert, 1973). Phosphate vibrations of cyclic nucleotides have been extensively studied. Symmetrical- and asymmetrical-stretching vibrations of o-p-o appear at
.~~~"
Conformational Analysis
145
-it:;'::';
,!¥i,803-812 and 812-822 cm -I, respectively. For POz vibrations, symmetrical~:stretching vibrations appear at 1111-1113 cm -I, whereas asymmetrical-
;~stretching vibrations are at 1228-1230 cm - I (Forrest and Lord, 1977). ::;~~::,. ,,~{
if:~,.·
i~4.
:~i
CONFORMATIONAL ANALYSIS
;~lNucleic
acids have conformational flexibility in solution. Even double-stranded ;:DNA is not entirely rigid, as can be seen from the fact that some drug <)molecules can intercalate into the grooves of DNA helices. Thus it is important ;'to analyze the conformation of nucleic acids in solution. Raman spectroscopy contributes to some aspects of the conformational study of nucleic acids; these ,aspects are summarized in this section.
4.1.
DOUBLE-STRANDED POLYNUCLEOTIDES, DNA, AND RNA
According to X-ray diffraction study, double-stranded DNA has three forms. When it is in the A form, the bases are tilted at an angle of 20° from perpendicular to the helix axis (Figure 5.3). When DNA is in the B form the bases are perpendicular to the helix axis. When DNA is in the C form, the configuration is similar to that of the B form, but the base pairs are farther away from the helix axis and tilted by about 6° so that the narrow groove in the helix is made deeper.
4.2. 4.2.1.
USE OF PHOSPHODIESTER BANDS A and B Forms
The 815-cm -I band of the phosphodiester-stretching vibration has been well studied for synthetic double-stranded polynucleotides. This band is considered to be an index of the amount of ordered structure of double-stranded A-form helix. This band is also sensitive to the conformations of the DNA and RNA backbone-chain phosphodiester bonds (Erfurth et al., 1972). In the disordered form of poly(A)' poly(U), the O-P-O band appears at 800 cm- I , but it is shifted to 815 cm -Ion forming an A-type helix (see Section 4.1) (Small and Peticolas, 1971 b). When they are in the B form, an 835-cm -I band is characteristic. 4.2.2.
C Form
The Raman spectrum of the C form of DNA is characterized by a band at 865-870 cm - I (Goodwin and Brahms, 1978).
A-Form DNA
B-Form DNA
•....
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Conformational Analysis
147
Interconversion of A and B Forms
the DNA fiber, conformational changes (A relative humidity. A
807-815 cm- I
±::;
B) take place depending on
B 790 cm- I 835 cm- I
In the aqueous solution, DNA always possesses the B-type structure. At
temperature, RNA always possesses the A-type structure, as it shows a sharp band at 814 cm- I (Peticolas, 1975). At 100% A-type conformation, the ratio of 1810-8151/1100 assumes the value of 1.64 ± 0.05 (Brown and Peticolas, 1975). Type-A and type-B structures are not restricted to DNA, but also exist in other double-stranded polynuc1eotides. The A-form-to-B-form transition was examined in poly(A) . poly(U) using a special technique, Raman temperaturejump (Sturm et aI., 1978). By this technique, Raman spectra can be obtained in the microsecond or longer time range after a temperature jump. At lower temperature, poly(A) . poly(U) shows the characteristic A form, with the band at 814 cm -I. Two milliseconds after the temperature jump, from 54°C to 57°C, the intensity of the 8l4-cm- 1 band decreases, whereas that of the 785-cm - I band increases. By this method one can study the detailed process of relaxation kinetics for the melting of poly(A) . poly(U). The conformation of the Na-DNA fiber is sensitive to relative humidity and salt concentration. The two types of DNA can be differentiated by Raman spectroscopy. A band at 807 cm - I is always present when a fiber shows an A-type diffraction pattern; the 790- and 835-cm - I bands always appear in the B form (Figure 5.3) (Lu et aI., 1977). Therefore, Raman spectroscopy correlates well with X-ray diffraction data (Brown and Peticolas, 1975; Erfurth et aI., 1975). Since Raman spectroscopy is much easier and faster than X-ray diffraction, there is a certain advantage to using the former. DNA in 80% ethanol shows the A-form conformation, but the B form appears in 60% ethanol. Photochemical modification of pyrimidine bases can covalently link bases laterally. By monitoring 807 cm - I (0- P-O vibration) and 784 cm- I (cytosine vibration) bands, one can determine the secondary conformation of DNA. Pyrimidine-dimer cross-links induce the B form of DNA and lock it irreversibly into that conformation, preventing it from converting to the A form in 80% ethanol. The A-form DNA can tolerate a few cross-links, but it converts cooperatively to the B form if a larger number of cross-links are introduced (Herbeck et aI., 1976). 4.3.
MELTING OF NUCLEIC ACIDS
Since some Raman bands of DNA are temperature dependent, it is possible to follow the melting behavior of DNA using these bonds. The temperature-sensitive bands are summarized in Table 5.5.
148
Nucleic Acids
TABLE 5.5.
Raman Lines of DNA sensitive to Temperature
Frequency (em-I)
Intensity Change
672 683 730 750 798 835
Small increase Decrease Large increase Decrease Increase Shift to lower frequency
879 920 1051 1144 1186 1214 1225 1240 1259 1303 1378 1463 1491 1534 1579 1660 1674 Source:
Decrease Decrease Decrease and shift to higher frequency Disappears Moderate increase Moderate increase Moderate increase Large increase Small increase Moderate increase Moderate increase Decrease Moderate increase Moderate increase Large increase Large increase Decrease
Assignment T G A T C+T P0 2 Diester antisymmetrical stretch Deoxyribose-Phosphate Deoxyribose C-O stretch Deoxyribose-phosphate T T A T C,A A T,A Deoxyribose-Phosphate G,A C G,A C=OofT T
The data based on the works of Rimai et al. (1974) and Erfurth and Peticolas (1975).
Melting of calf-thymus DNA measured by following conventional UV absorption at 260 nm, and Raman hyperchromism at 1300 cm -] for the adenine band and at 1658 cm - I for the thymine band is shown in Figure 5.4. All three measurements give essentially the same Tm at 75°C. Tm refers to a transition temperature or a midpoint temperature. Usually there are several Raman bands that can be used for a melting study, but each polynucleotide has its own specific bands. In poly(Br U-A), 1627-, 1352-, and 1230-cm- 1 bands all show Raman hyperchromism, but the 1627cm -] band, which is attributed to the carbonyl stretch, shows the most pronounced effect upon melting (Chinsky et aI., 1977). Similarly, in poly(l-Me-I), there are several temperature-sensitive Raman lines; the C=O vibration at 1710 cm-] shifts to 1680 cm- ], causing Raman hypochromism upon melting (Scovell et at, 1979).
1.65
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30 40
50 60 70 80 Temperature. • C
a:
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Temperature,
-c
(0)
(b)
FIGURE 5.4. Melting curve of calf-thymus DNA using (A) UV-absorption method, (B) Raman hyperchromism at BOO cm -I, and (C) Raman hyperchromism at 1658 cm -I. The figure was reproduced from Erfurth and Peticolas (1975).
Temperature, ·C (e)
150 4.4.
Nucleic Acids
BASE STACKING AND HYDROGEN BONDING
The phosphodiester-stretching-vibration band at 815 cm- 1, used to monitor the ordered structure of the double-stranded helix, can also be used to monitor base stacking if it affects the phosphate-backbone configuration. Different conformations of DNA have different base-stacking modes. The thymine line at 750 cm - I is strong in type-B DNA, but its intensity is diminished in type-A and type-C DNA. The type-C DNA is also distinguished from both A and B types by a strikingly different pattern of Raman frequencies in the 1250-1400-cm-] region, again attributable to its altered base-stacking geometry. The stabilization of DNA conformation is maintained not only by base stacking, but also by horizontal base interactions, that is, base pairing through hydrogen bonds. Actually, base pairing is a bigger stabilizing factor than the base-stacking effect. Therefore, there are different 1670/1680 ratios -these lines are due to the in-plane ring vibrations of thymine and guanine, respectively-for types A, B, and C DNA (Lord and Thomas, 1967a, b). The sequence of bases in a double-helical-polynucleotide complex can influence the Raman frequencies and intensities, as can be shown by the example of poly(A) . poly(U) and poly(A-U) . poly(A-U). Both have the. O-P-O band at 814 cm-I, indicating an identical backbone structure. The poly(A) . poly(U) contains repetitive sequences of AU pairs, which show Raman bands at 1680, 1631, and 1613 cm- I . The other copolymer complex poly(A- U) . poly(A- U) contains AU and UA pairs in alternating sequence and shows the bands at 1677, 1650, and 1624 cm- I (Morikawa et al., 1973; Lafleur et al., 1972; Thomas, 1975). Because of the lack of chain length, oligonucleotides do not form regions of intramolecular helix like tRNA (transfer RNA). However, in some oligonucleotides "base stacking" cannot be ignored. The hypochromism effect in UV absorption is well known and occurs when planar molecules such as purines and pyrimidines become stacked. Similarly, Raman hypochromism takes place when the bases are stacked. In the ordered poly(A) in which each A is stacked, the 725-cm-] line is 50% less intense than in the random-chain configuration of poly(A) (Small and Peticolas, 1971a, b). The 1338-cm-] band of adenine shows strong Raman hypochromism upon base stacking or base ordering (Tomlinson and Peticolas, 1970). This band is resonance enhanced with 257.3-nm laser light. The l338-cm -I band derives its intensity from the UV-absorption band at 260 nm (Pezolet et al., 1975). Even the simple oligonucleotide isomers with different base sequences have different conformations, as they show different Raman spectra (Gramlich and Schmid, 1978). For instance, ApU and UpA have different Raman spectra. Judging from the spectra, ApU undergoes base stacking, whereas UpA does not. Similarly, GpC and CpG show different spectra (Prescott et al., 1974). The dinucleotide GpC has a band at 809 cm-] that is assigned to the phosphodiester vibration. When GpC is heated, this band disappears. This is explained by the fact that the dimers become increasingly self-stacked at lower temperatures
Conformational Analysis
151
and the phosphate group assumes the geometry of A-form DNA (Erfurth et aI., 1972). The Raman data indicate that dinucleoside phosphates favor the C;-endo ribose conformation in aqueous solution (Kiser, 1976). Some lines are sensitive to the formation or dissociation of hydrogen bonds; these Raman lines can be used to examine the state of base pairing. Cocrystalization of 9-methyladenine and I-methylthymine takes place with a unimolar ratio of the two bases. A Raman frequency change is associated with the modes most sensitive to hydrogen-bonding interactions, suggesting that a cyclic dimer is formed (Strobel and Scovell, 1980). The double strand of poly(A, G) . poly(U) is an interesting one, as A-U and G-U form hydrogen-bonded pairs. The o-p-o band at 815 cm- I is used to assess the degree of double-helical content. Both G and U have C=O groups, but the C=O from G and the C=O from U give bonds at different frequencies. Analysis of Raman spectra indicates that the structure of poly(A, G)-poly(U) is very similar to the structure of poly(A)-poly(U); however, there is some thermal instability in the former complex. This is explained in terms of the wobble associated with G-U base pairs (Ackerman et aI., 1979). Poly(l) shows a distinct Raman band at 1464 cm- I that is assigned to the amide II band of the cis-amide group of the hypoxanthine base. When poly(l) forms a pair with poly(C), the 1464-em -I band shifts to 1484 cm -I. Thus by measuring the 1484-em- 1 band, one can determine the fraction of I-C pairs formed (Brown et aI., 1972). 4.5. 4.5.1.
SINGLE-STRANDED POLYNUCLEOTIDES Poly(C)
At neutral pH, poly(C) forms a single-stranded structure that contains stacked bases. At pH 4.5, poly(C) becomes double stranded by taking one-half mol of protons per mole of cytosine: poly(C) + H+ +poly(C) ---> poly(C)-poly(C+) (Figure 5.5). The structurally sensitive line at 815 cm- I (actual frequency is H
H \
/
H-C
\
\
N-H---O \
/
C-C
\ /
/
Chain / C-N
\
N-- H--N
C-H
\
/
N-C C-C \ / \ Chain O---H-H H
/
\
H
FIGURE 5.5. Two strands of poly(rc) and poly(rc+). The figure was reproduced from Chou and Thomas (1977).
152
Nucleic Acids
811 cm -]) is better resolved in D 2 0. The ratio 1815 /1 1]00 = 1.65 indicates that the complex has a high degree of helical content (Chou and Thomas, 1977). Judging from the 1380-cm-] line as a function of pH, the complex is stable within the pH range of 3.7-5.5. That poly(C) is single stranded but stabilized by base stacking at neutral pH is supported by heat-denaturation experiments. On heating the Raman bands at 790, 1256, and 1547 cm- 1 show Raman hyperchromicity. The first two bands are base ring vibrations and the latter corresponds to the double-bond stretch (Peticolas et al., 1971). 4.5.2.
Poly(U)
When Raman spectra of poly(U) at 20 and 80°C are compared, there are practically no changes. This reflects the fact that there is little conformational change in poly(U) because there is no base stacking (Peticolas et al., 1971). 4.5.3.
Poly(A)
The Raman spectrum of AMP is relatively insensitive to temperature change, whereas that of poly(A) is greatly perturbed. This reflects the lack of a long chain in AMP; the spectrum of poly(A) changes because of changes in the chain structure. The most pronounced changes in the spectrum of poly(A) are in the bands at 725, 1252, 1303, 1377, 1424, and 1508 cm- I , which show Raman hyperchromicity. The l576-cm- 1 band shifts in frequency without changing in intensity (Peticolas et al., 1971). Poly(A) is a single-stranded polynucleotide, but base stacking stabilizes the strand. 4.5.4.
Poly(l)
At high ionic strength, poly(I) shows the ordered conformation of the A-type helix, as shown by the presence of an 8l5-cm-] band (A-type band) and the absence of an 835-cm-] band (B-type band). The conformation is stabilized by specific hydrogen bonding involving hypoxanthine C(6)=O groups (Chou et al., 1977).
4.6.
GEL FORMATION
Some nucleosides or nucleotides form a gel from a solution on cooling. The formation of gel from solution is a reversible process. Such a phase change does involve fine alteration in structures such as hydrogen-bonding and base-stacking parameters. Raman spectroscopy can be used to detect such fine changes. It is known that guanosine solution (0.1 M KCl) forms a gel when it is cooled to lODe. It is of interest to correlate the Raman spectral change to gel formation. The most pronounced change is in the keto-stretching-vibration bands. The keto band at 1671 cm-\ present at 60°C disappears at lOoC, and
...
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t,~
Native RNA
• '
153
: small bands at 1700 and 1665 cm - I appear. This reflects the involvement of :C=O in hydrogen bonding during gel formation. Another big change occurs :in the vibration due to the pyrimidine portion of the purine ring. This is explained by the C=O's forming a strong hydrogen bond with the N-H of the pyrimidine ring at either N-l or N-3 (Delabar and Guschlbauer, 1979). At pH 5, 5'-GMP also forms a viscous gel with altered optical properties. This is believed to be due to a helixlike formation between two nucleotides joined together horizontally through hydrogen bonding and joined hiterally through base stacking. In order to understand the gel ~ solution behavior of GMP-5', Raman spectra at 0 and 40°C were examined by Gramlich et al. (1976a, b). There are marked differences in the 1487- and 1670-cm- 1 bands. The 1487-cm -} band is attributed to the guanine-ring stretching mode, whereas the 1670-cm- 1 band is due to the C=O stretching vibrational mode. At 40°C, the 1487- and 1670-cm- 1 band intensities increase. This indicates that both the N-7 position and the carbonyl group are involved in hydrogen bonding in the gel ~ solution transformation. Guanosine derivatives tend to form molecular aggregates. For instance, GpG forms an aggregate whose melting behavior can be followed by the Raman intensity increase at 1480 em-I. The melting point is near 60°C. Melting is due to unstacking of the bases and breaking of base-pair hydrogen bonds (Savoie et al., 1978).
5.
NATIVE RNA
So far we have reviewed basic Raman studies of bases, nucleosides, nucleotides, and synthetic polynucleotides. The most exciting and challenging thing is to study naturally occurring nucleic acids and nucleoproteins by Raman spectroscopy, which permits analysis of their overall conformation or detailed small changes occurring within nucleic acids. This section discusses how Raman spectroscopy contributes to the understanding of transfer RNA (tRNA) and ribosomal RNA (rRNA) 5.1.
TRANSFER RNA
Transfer RNA has been extensively studied by Raman spectroscopy. After thermal treatment of tRNA, Small et al. (1972) found several lines to be sensitive to the order-disorder effect of yeast tRNA. These lines are useful in the study of tRNA structure; they are shown in Table 5.6. Table 5.6 was constructed from different papers; sometimes the frequencies and assignments are slightly different because of different investigators' observations and judgments. Since both tRNA and 5 S ribosomal RNA have very similar Raman spectra, the same table can be used for band frequencies and band assignments (see Section 5.2) of both types of RNA.
154
Nucleic Acids
TABLE 5.6. Raman Band Frequencies of tRNA and Ribosome 5 S RNA on Thermal Disordering
Raman Lines (em-I)
Degree of Change
670
Large decrease in intensity
725 758
Moderate increase in intensity Disappears
783
Moderate increase in intensity due to shift in 814-cm- I band
785" 814
Shift in frequency
975
Large decrease in intensity
1044
Moderate decrease in intensity
1100 1158
Disappears Disappears
1180
Moderate increase
1230 1299 BOO" 1321 1339 1373 1375 a 1484 1485" 1531 1575"
Large increase in intensity Small increase in intensity Small increase in intensity Small increase in intensity Moderate increase in intensity Moderate increase in intensity Moderate increase in intensity Small increase in intensity Small increase in intensity Moderate increase in intensity Moderate increase in intensity
Assignment ChangeinG-backbone interaction Unstacking of A Change in G-backbone interaction O-p-O
C and U ring vibrations O-P-O disordering of ribose-phosphate backbone Disordering of ribosephosphate backbone Disordering of ribosephosphate backbone o-p-o- symmetrical stretch Disordering of ribosephosphate backbone Base external stretch increase due to breaking of interbase H-bonds Unstacking of U Unstacking of A C and A ring vibrations Unstacking of G Unstacking of A Unstacking of A G and A ring vibrations Unstacking of G G and A ring vibration Unstacking of G G and A ring vibrations
"The data were obtained from the papers of Small et al. (1972), Thomas et al. (1972), Chen et al. (1975; 1978) and Luoma and Marshall (1978a).
All tRNAs are stabilized by the cloverleaf-style, intrachain-hydrogen-bond pattern achieved through base pairing. Such high stability is also reflected in the characteristic Raman bands. For instance, all 11 tRNAs have a very high /815/[1100 ratio (1.73), indicating a high degree of order (Chen et at, 1978). Melting of tRNA (670,812, 1578 cm- I ) indicates that G residues remain stacked even at 90°C, but C and U unstack at 70°C (Thomas et al., 1973a).
Native RNA
155
From the evidence of Raman spectra, tRNAfMet, tRNAVal , tRNAPhe , G1u f, tRNA , and tRNAArg are all similar (Thomas et al., 1973b). 1~ The degree of ordered structure can be measured from the intensity ratio ~. 18l5 /1 1l00 . Both tRNA and rRNA have the same Raman spectra in solid and 1~' aqueous solution. Thus unlike DNA, RNA has a backbone conformation that .~ is insensitive to the degree of hydration (Chen and Thomas, 1974). Yeast .~~; tRNAPhe in two crystal forms (orthorhombic and hexagonal) and aqueous i~.solution has been examined using the structurally sensitive Raman lines at 670 ~'cm-l (G), 725 cm- 1 (A), 785 cm- 1 (C and U), and 814 cm- 1 (O-P-O .~~•. stretching vibration). These lines were normalized against the 11 OO-cm- I line, (~iwhich is the structurally insensitive ionic PO; stretching vibrational mode. All if'three forms of tRNAPhe give identical Raman bands. So the molecular struc',~r< ture of this tRNA is not altered by the differences in molecular packing in two ;~::,different unit cells or aqueous solution (Chen et al., 1975). In the tRNAPhe the 'ir1G-C bases have more effective stacking in the solid form than in aqueous ~~!isolution (Pham and Giege, 1981). }~ Raman spectroscopy was used in a unique study on the effect of a 100-MHz ;~(tradio-frequency field on E. coli tRNA. It was found that the tRNA lost several ',nof the Raman lines associated with intramolecular hydrogen bonding that appear in the region 800-2000 cm- 1 (Klainer and Frazer, 1975). Raman spectra of native 5 S RNA and 5 S RNA incorporated with 5-fluorouracil (Fu) are very similar, suggesting that both native and Fu-5 S RNA have highly base stacked structures (Marshall and Smith, 1980). ..
5.2. 5.2.1 .
RIBOSOMAL RNA Isolated RNA
The ribosome plays a central role in protein biosynthesis together with other compounds such as tRNA and mRNA (messenger RNA). The ribosome is a nucleoprotein; both protein and RNA components are heterogeneous. The exact biological role of RNA is still not clear. When rRNA (16 Sand 23 S) is heated to 85°C, all base pairs are broken, but there is an increase in the single-stranded stacked region. The stabilization energy of the stacking is in the order U < C < A < G. This means that the single-stranded stacked regions contain more G than A and more A than U or C. Heat treatment changes some Raman line intensities (Thomas et al., 1971). 5 S RNA and 5.8 S RNA are structural components of the large ribosomal subunit of eucaryotic cells. When yeast 5 S RNA and 5.8 S RNA were compared with yeast tRNA with respect to the relative intensities of Raman lines at 670 (G), 725 (A), 785 (C, U), 814 (OPO), 1100 (PO; ), 1234 (U), 1251 (C, A), 1300 (C, A), 1321 (G), 1338 (A), 1375 (G, A), and 1485 (G, A) em-I, they were all found to be very similar. From these results, Luoma and Marshall (1978a, b) proposed that eucaryotic 5 S RNA and 5.8 S rRNA have a new cloverleaf secondary structure with G-C and A-U pairings.
156
Nucleic Acids
Similarly, E. coli 5 S ribosomal RNA (MW 40,000) also has a highly ordered structure judging from the ratio 1815 /[1100 and the structurally sensitive lines of G at 670 cm -, and A at 725 cm- '. Actually, the stacking efficiency of the guanine bases is much higher in 5 S ribosome RNA than in yeast tRNA (Chen et aI., 1978; Fabian et aI., 1981). It is well recognized that Mg(II) is essential for maintaining the integrity of ribosome particles. The importance of Mg(II) can be demonstrated with Raman spectroscopy. The effect of Mg(II) on the ordered structure of RNA is monitored by measuring [8\511785' The 785-cm- 1 peak is independent of Mg(II) concentration; therefore, it is used as a normalization standard. In the presence of EDTA (ethylenediaminetetraacetic acid), a powerful chelating agent, Raman spectra of ribosomes and rRNA differ very little. But as the concentration of the Mg(II) is increased, the ribosome becomes more stable than the rRNA to changes induced by elevated temperature. These studies show that the Raman spectra are detecting a contribution to the backbone ordering that requires the presence of both Mg(II) and ribosomal proteins (King et al., 1980). 5.2.2.
RNA Within Ribosomes
Since ribosome particles are nucleoproteins, the Raman spectra of ribosomes should be like those of RNA and proteins. However, the Raman scattering of nucleotide residues is more intense than that of amino acid residues, and RNA components contribute more to the spectra of ribosomes. Thus one can study the state of RNA within the intact ribosome particles by analyzing the Raman spectra of ribosomes. The RNAs within the ribosomes (80 S rat-liver ribosome and 70 S E. coli ribosome) retain highly ordered structure with the backbone conformation of the A-type helix. Even when the ribosomes are dissociated into subunits, RNA within the subunits remains highly ordered (Thomas et aI., 1980). The integrity of ribosome particles is highly dependent on the magnesium-ion concentration. At low magnesium-ion concentration, Raman intensities of the phosphodiesterase at 813 cm- I for 30 Sand 50 S ribosomes decreases compared with the intensities for intact ribosomes. This suggests that a fraction of the ribose moieties have shifted from 3'-endo (ordered) to the 3'-exo (disordered) conformation (King et aI., 1981).
6. 6.1.
REACTIONS OF NUCLEIC ACIDS ALKYLATION
Nucleic acids and polynucleotides can be alkylated by chemical agents and drugs. These reagents usually modify bases. Nucleic acid bases can be methylated, and 7-Me-guanosine has quite a different spectrum from that of GMP;
Reactions of Nucleic Acids
157
the methylated derivative lacks the strong band at 1488 cm- I. The replacement of C(8)-H with C(8)-D shifts the 1488 band to 1465 cm- I . From this one can conclude that the 1488-cm- 1 band is the N(7)=C(8) double bond stretch. Thus this band can be used to detect the alkylation site on the guanine base. Upon alkylation of DNA with methylnitrogen mustard, the 1492-cm - I band decreases and shifts to 1530 cm -I, an indication of alkylation at the N-7 of guanine. No changes in the Raman bands of any other bases are observed in alkylated DNA. Furthermore, alkylated DNA forms a stable double-stranded helical complex at neutral pH in which the alkylated guanine residues are in keto form (Mansy and Peticolas, 1976). 6.2. 6.2.1.
POLYPEPTIDES DNA- Polypeptide Interactions
Understanding DNA-protein interactions is important, as many biological compounds in living cells are nucleoproteins. When DNA is mixed with polylysine at neutral pH it forms a complex. This is readily understandable, as DNA is negatively charged, whereas poly(Lys) is positive at neutral pH. Raman spectroscopy can be used to look into the nature of such interactions. In this case, judging from the lack of change in the O-P-O stretching band at 815 cm-I, it is concluded that the complex formation does not change the backbone structure of DNA (Prescott et aI., 1976). Somewhat similar to the case with polylysine, other polyamines such as spermine and spermidine interact with DNA. The protonated groups of the polyamine interact with the phosphate groups of DNA (Bertoluzza et aI., 1978). The DNA in DNA-proteinase complex has a modified B-form conformation. The modified B form possesses a slightly different (by about 4° with respect to the helical axis) OPO bisector angle when compared with the B form of DNA (Herskovits and Brahms, 1976). There is a large increase in the adenine band at 1582 cm - I when polyd(A-T) forms a complex with RNase, suggesting that the adenine base is involved (Chinksy et aI., 1978b). Although most Raman studies have been accomplished at high frequency, there is a report that the low-frequency (20-30 cm~l) vibrational spectra of DNA and poly(Lys)-DNA complex are sensitive to the dynamics or conformation flexibility of these macromolecules (Painter et aI., 1981). 6.2.2.
RNA-Polypeptide Interactions
Many compounds and viruses are RNA-protein complexes. In order to understand them, it is essential to know more about the nature of binding between these two components. Unlike DNA, poly(A) changes its backbone geometry, and the base-stacking modes are altered on complexing with poly(Lys) (Hernandez et aI., 1980). The
158
Nucleic Acids
TABLE 5.7. Raman Lines Assignment for Poly(rA) and Their Changes After Complex Formation with Poly(Lys)
Frequency
Assignment
726 811
A ring O-P-O sym str
887 917
C-O, C-O str (RP) A ring
1007 1025-1075 1221 1306 1379 1483 1578
C-O, C-O str (RP) C-O, C-C, C-N str (PLL, RP) A ring A ring A ring A ring A ring
Change Intensity increase Shift to 804 cm - J and intensity increase Intensity increase Shift to 912 cm - I and intensi ty decrease Shift to 1014 cm- I Intensity increase Intensity increase Intensity increase Intensity increase Intensity decrease Intensity increase
Source: This table was reproduced from Prescott et al. (1976). RP, ribose phosphate; PLL, polylysine.
Raman band changes on complexing and their assignments are shown in Table 5.7 (Prescott et al., 1976). PolY(L-Arg) increases the order of the base stacking of poly(A) and poly(C), whereas polY(L-Lys) causes disordering in the base stacking. Thus different polypeptides exert different effects on nucleic acids.
6.3.
METAL IONS
Many metal ions interact with nucleic acids and their components. The sites of metal-ion interactions with nucleosides, nucleotides, and nucleic acids have been extensively studied by many investigators. The binding mechanism is not simple, and it depends on the type of metal ion, ligand, and the pH (Tu and Heller, 1974). Because Raman spectroscopy can detect many modes of vibration including metal-ligand bonds, the site of complexing can be further clarified by the use of Raman spectroscopy. Hard metal and soft metal are terms used in inorganic chemistry. In hard metals, the electrons are firmly held; if the electrons are easily removed, the species is a soft metal. For instance, Na(l), Mg(ll), Ca(ll), Mn(ll), and Cr(lll) are examples of hard-metal ions, whereas Cu(I), Ag(I), and Hg(ll) are soft-metal ions. Fe(ll), Co(ll), and Cu(ll) are considered borderline. The hard-metal ions such as Mg(ll) tend to attach to the phosphate group. The phosphate group exhibits characteristic vibrational bands such as the antisymmetrical stretching at 1230 cm- J, the ionic-phosphate vibration at 1100 cm- I, and the 0- P-O
).
'.'. '':;'.
....
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~~:,
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Reactions of Nucleic Acids
159
if;:: ester vibration at 815 cm - '. When the phosphate complexes with metal ions, ~::
it, these bands are perturbed
and show either a frequency shift or a change in ",: Raman intensity (Theophanides, 1979). Soft-metal ions tend to attach to ,~~~ purine and pyrimidine bases at a variety of sites. Usually the site of attachment can be deduced from Raman or IR spectra. For instance, if a metal attaches to '~C(6)=0 of the inosine base, it induces perturbation of the keto stretching I~vibration (Millard et al., 1975; Hadjiliadis and Theophanides, 1976). On some ~1; occasions, metal ions attach to the delocalized 'IT-orbital ring instead of specific 't~~ groups. For instance, ferric ion forms a 'IT-complex with the nicotinamide ring ltof NADH (Siiman and Callaghan, 1981).
I.
':p' :--6.3.1.
Adenosine
Hard-metal ions such as Ba(II) and Pr(III) do not interact with adenosine. When Hg(II) and Zn(II) ions bind to adenosine, there is a large change in the Raman spectrum in the 1300-1600 cm- 1 region. It is believed that they attach to both N-I and N-7 (Marzilli et al., 1980). AMP-5' gives a very intense band at 248 cm- 1 when the compound is absorbed at a silver electrode. This line is attributed to an Ag-POf- vibration (Koglin et al., 1980b). 6.3.2.
Cytidine
When cytidine is mixed with Hg(II), there are significant Raman spectral changes in the region of 1600-1700 and 1200-1300 cm - 1 that correspond to C=O and C=N stretching vibrational modes, respectively (Lord and Thomas, 1968). It is very likely that Hg(II) attaches to N-3 and C=O moieties. Other metal ions such as Zn(II), Cd(II), Cu(II), and Mg(II) do not change the spectrum of cytidine; thus the mercuric-ion interaction with cytidine is rather specific. Cytidine also interacts well with alkaline-earth-metal ions. There is a large perturbation in the 1200-1300 cm - I region, and this is interpreted to mean that the ions attach to C(2)=0 (Marzilli et al., 1980). 6.3.3.
Uridine
Uridine does not form a complex with Pb(II), Ba(II), Hg(II), or Zn(II), as they show no Raman spectral changes (Marzilli et aI., 1980). Uridine seems to be unreactive to many metallic ions. Silver ion interacts well with guanosine and inosine, does not interact with uridine (Tu and Reinosa, 1966). 6.3.4.
Guanosine Monophosphate
GMP-5' interacts with a variety of metal ions (Pt, Hg, Ni, Co, Mn, Mg, Ca). There are changes in the Raman intensities of the guanine-base bands and the phosphate-group bands. This indicates that metal ions attach to the guanine base as well as to the phosphate group (Makrigiannis et aI., 1980).
160
Nucleic Acids
6.3.5.
Interaction With Platinum Compounds
Because of their importance, the interactions of platinum compounds with nucleic acids and their components have been extensively studied by Raman spectroscopy and other techniques. The anticancer properties of cis-dichlorodiamine platinum(II) and related metal complexes are well known. Although the cis isomer has antitumor activity, the corresponding trans isomer has not. When DNA complexes of both cis and trans isomers are examined by Raman spectroscopy, there is a significant difference in the carbonyl stretching region. This indicates that a perturbation of the guanine carbonyl bond occurs when the antitumor drug (cis isomer) attaches to DNA (Alix et al., 1981). This is one example of the use of Raman spectroscopy: to see structure-function relationships in biological compounds. Cis (NH3hPt(II)CI2 attaches to the N-7 of GMP and to a lesser extent to AMP-5' (Theophanides, 1980). The trans isomer is more selective for GMP-5'. Neither the cis nor the trans complex reacts with the pyrimidine nucleotides. There are decreases in the intensities of 1486- and 727-cm- 1 bands on complexing. The former is the guanine-base vibration, and the latter is the adenine-ring vibration. When the complexes are formed, the l486-cm- I band shifts to 1500 cm- I , and the 727-cm- 1 band shifts to 718 cm- I (Mansy et al., 1978; Moller et al., 1980). No significant change occurs in the carbonyl mode at 1680 cm- I , suggesting that C(6)=O is not involved in complexing (Chu et al., 1978). The attachment to the guanine base agrees well with other observations that the rate of reaction and platinum-compound-binding ability increase with increasing G-C content of DNA.
6.3.6.
Alkylmercury Compounds
Unlike inorganic mercury, alkylmercury, such as methylmercury, is much more toxic and extremely neurotoxic. The interaction of monomethylmercuric ion with nucleotides has been extensively studied by Raman spectroscopic methods. As thymine interacts with CH 3HgI, the 505-cm- 1 CH 3 HgOH band decreases, and the 1292-cm- I complex band increases. Using these two bands it is possible to measure the stability constant of the monomethylmercurythymine complex (Chrisman et al., 1977; Tobias et al., 1978). The equilibrium constant is 0.6, which is in reasonable agreement with an earlier value based on uv spectrophotometric data. The binding site is at N-3, with substitution of CH 3Mg+ for the proton. In contrast, native calf-thymus DNA does not bind to CH 3Mg(II). It is reported that methylmercury binds to the N-3 of cytidine and UMP, . causing perturbations of their Raman spectra (Tobias et al., 1976; Moller et al., 1980).
Reactions of Nucleic Acids
6.3.7.
161
ATP
The potential atoms or groups in ATP that metal ions can bind to are 0, N, and the phosphate group. Before analyzing the spectra of the metal-ATP complex, it is essential to know the major bands of the various vibrational modes of PO; ,C-N, C=N, C-C, and CH in ATP (Table 5.8). The metal ions Ca(II), Mg(II), Co(II), Cu(II), and Hg(II) attach to the triphosphate group in the pH range 3-12. Especially Ca(II) and Mg(II) interact strongly with the phosphate moiety at neutral pH. At neutral pH, Cu(II) also attaches simultaneously to N-7 and with the amino group of the adenine ring (Figure 5.6). At pH 10-11, Cu(II) binds to N-3, as indicated by the enhancement of the 760- and I360-cm- I vibrations. At neutral pH, Hg(II) shows a direct coordination at N-I, whereas at low pH, with N-l blocked by protonation, Hg(II) does not interact with the adenine moiety (Lanir and Yu, 1979). 6.3.8.
Mg(II)-ATP
Magnesium ion plays an important role in biological systems, especially when the reactions involve certain nucleotides. It is also known to stabilize the TABLE 5.8.
Raman Line Assignment of ATP at pH 7.0
Frequency
Assignmenta, b
320 731
8(N,C6 -NH 2) > 8(CS C6 -NH 2) All 12 bonds stretch in phase (adenine moiety) p ...:-. 0)
1118
p(O~'
1223 1255
8(C s H) > p(CSN?) p(CSN?) > p(CSNg) > 1', 8(N,C 2) p(N,C 2 ) > 8(C 2H) > p(NgC s ) > p(CgN?) p(NgC g) > p(N3C 2 ) > 8(C gH) > 8(C 2 H)
1309 1340
p~SN9»p~2~»8~gH)
p(N?C s ) > ,,(CgN?)
1380 1424 1485 1510 1583
b
p(N 1C6 ) > p(C6 -NH 2 ) p(C4 N 9 ) > 8(C g H)
Five-membered ring p(C4 C S ) p(C4 C S ) > p(N3 C4 )
Source: This table was reproduced from Lanir and Yu (1979). denotes stretching; 8, bending. bAssignment not available.
ap
~
162
Nucleic Acids
N H:Jc2
I
-
0-
0-
N~
IN N
-0-P-0-P-0-P-0- CH 2 III III II 000
?
r
U N;:/
0
OH
FIGURE S.6. The site of Cu(H) attachment to ATP.
OH
double-helical structure of nucleic acids. The stabilization is attributed to a reduction in the electrostatic repulsion between phosphate groups of the helical backbone in the presence of Mg(II). When monomethylphosphate is mixed with magnesium ion, the poisymmetrical vibrational band at 985 cm - I broadens and shifts to 990 cm - \; the C-O stretching vibration at 1057 cm-\ shifts to 1050 cm- I (Lord and Thomas, 1968). The most common and important reaction involving magnesium ion and nucleotide is the phosphate-transfer action of ATP: ATP
->
ADP + P
Mg(II)
It is important to know how the Mg(II) is involved. Free ATP has a band at 1117 em-I due to the PO; vibration. When magnesium ion is added to ATP, another band at 1123 cm-\ appears in addition to a 1116-cm- 1 band. The peak height of the 1I 23-em - I band is about twice that of the 1116-cm- I band. This suggests that Mg(II) binds to two phosphate groups, and one phosphate must be free. The ATP-Mg(II) intermediate can be stabilized by the presence of dimethysulfoxide and a dicarboxylie acid. The intermediate as well as the ADP-Mg(II) complex show a 1091-cm-\ band. The Raman spectrum of A-O-P-O-P-O-P + MgllIl
ATP
1
A-O-~-O-;P-O-P \
,
intermediate
ADP-MgIIII
complex
Nlg++
1
A-O-~-O-;P \
ATP-MgIIII
,
+ P
W++
FIGURES.7. Structure of ATP-Mg ion complex.
Reactions of Nucleic Acids
163
AMP-Mg(II) is depolarized while those of ATP-Mg(II) and ADP-Mg(II) are polarized. From this evidence it can be concluded that the magnesium ion attaches to the a- and ,8-phosphate of ATP, and the y-phosphate becomes free for the transfer reaction (Figure 5.7) (Lewis et al., 1975). 6.4. 6.4.1.
DRUGS
Actinomycin
Actinomycin D is an antibiotic effective against many gram-positive and gram-negative organisms. It consists of a phenoxazine linked to two different peptide chains (Figure 5.8). It is highly toxic to humans and is known to inhibit the transcription process. The actinomycin chromophore gives resonance-enhanced Raman bands at 1505,1489,1405,1385, and 1265 cm- I by laser excitation at 458 nm. Of these, the 1385-,1489-, and 1505-cm- 1 bands are sensitive to interaction with DNA. In particular, large intensity changes are observed for the 1489- and 1505-cm- 1 lines. This is explained by the insertion of actinomycin into the DNA groove, probably through the interaction of the phenoxazine ring with guanine bases (Chinsky et aI., 1975). Although the use of 458-nm laser excitation enables analysis of the fate of the actinomycin molecule, it does not reveal what happens to the DNA bases. By uv excitation at wavelengths of 300 and 280 nm, the l582-cm - I line of adenine and the 1492-cm - I line of guanine can be seen. Thus the use of uv incident light allows one to look at DNA bases free of any actinomycin contribution. In the actinomycin-DNA interaction, a large intensity decrease in the l492-cm- 1 line occurs, indicating that guanine bases are the ones disturbed by actinomycin (Chinsky and Turpin, 1978). Further analysis of the Raman spectra indicates that actinomycin D forms a 'IT-complex with a 2-amino group of guanine, and this complex is stabilized by the hydrogen bond between the carbonyl oxygen of the L-threonine residue and the guanine base (Turpin and Chinsky, 1978). In order to further understand the vibronic states of the actinomycin chromophore, Smulevich et al. (1980) investigated the Raman excitation profile.
L-N-Maval I Sar
L-N-Meval I Sar
L-Pro
L- Pro
I I
I
I
D-Val
D-Val
L-Thr
L-Thr
I
I
I
¢r I
I
~ NX)~
Qa CH
0
~ ~ CHa
. FIGURE 5.8.
Structure of actinomyCin D.
164
Nucleic Acids
6.4.2.
Bleomycin
Bleomycin is a glycopeptide drug used in the treatment of squamous-cell carcinoma and lymphoma. The drug is known to inhibit chromosomal DNA biosynthesis by binding to the bases. The 1543-cm -1 band is enhanced when bleomycin binds to calf-thymus DNA. This is interpreted to be preresonance enhancement of a ring vibration of the bithiazole ring present in bleomycin. The intensity of the band due to thymine vibrations decreases on complexing, suggesting that the site of the interaction is at the thymine bases (Fairclough et aI., 1978). 6.4.3.
Others
Netropsin and distamycin A (Figure 5.9) are oligopeptide antibiotics as well as antiviral agents that bind tightly to A-T-rich regions of DNA. For biochemists, these drugs are of particular interest because of their ability to recognize and bind to a specific site, that is, to a region of high A-T composition. After the drug has complexed with DNA, all methyl vibrations (1412, 1113, 1440 cm - I) with the exception of 1066 cm - I remain unchanged. The 1412-cm- I band is assigned to methyl symmetrical bending, the ll13-cm - I band to methyl rocking, the 1440-cm- 1 band to methyl asymmetrical bending, and the 1066-cm- 1 band to rocking of the methyl group and pyrrole-ring N-C vibration. The big changes occur for the ring N-C stretching at 1398 and 1485 cm -1. This indicates that the pyrrole ring is the portion that interacts with DNA. The amide III N -H vibration shifts to higher frequency, indicating that the peptide N - H is strongly hydrogen bonded with DNA. The A
Distamycin H
o
I
~C-NO I
H/
~
H I C-N
II CH3 0
I
+
H
UJL~- 0-C-~-CH2-CH2-C'NH2 NH2
I
N ION
CH3
1
I
H
//
H
+ NH
II 0
CH3
B
Netropsin
+
H2N "
H 1
H I
~C-N-CH2-C-NOI
H N 2
.
II 0
FIGURE 5.9.
H I C N II - N O I 0 N CH3 II I 0 CH3
I I C-~-CH2-CH2-d'/'NH22
Structures of distamycin A and netropsin.
Special Techniques
165
:~OH:'6 II
.~ T ".
3'
NH 2 QH
FIGURE 5.10.
Structure of adriamycin.
drug-DNA binding can be visualized so that the methyl groups on the pyrroles project away from the DNA, and the peptide N-H groups form hydrogen bonds with the DNA (Martin et aI., 1978; Martin, 1979). Ethidium bromide is a trypanocidal drug known to form complexes with nucleic acids. Ethidium bromide-bound DNA shows a different Raman spectrum from that of naturally occurring DNA, thus indicating an interaction (Chinsky et aI., 1976). Adriamycin is an antitumor and antibiotic drug isolated from Streptomyces peucetius that binds to DNA. The structure is shown in Figure 5.10. Raman spectroscopic analysis indicates that phenolic C-OH vibrational bands are strongly affected, suggesting that the phenol interacts with the A-T plane (Manfait et aI., 1980). Hexaziridinocyclotriphosphazene has antitumor activity against LI210 and P388 leukemias and B16 melanoma. A Raman investigation of the drug-DNA interaction suggests that the drug attaches to the N-7 and NH 2 positions of the adenine base (Manfait et aI., 1981). 7.
SPECIAL TECHNIQUES
7.1. PRE RESONANCE AND RESONANCE RAMAN SPECTROSCOPI ES
The Raman intensity of nucleic acids and their components depends on the excitation wavelength, especially when it is near the uv and near-UV regions (Blazej and Peticolas, 1980; Chinsky and Turpin, 1980). This is best illustrated by a three-dimensional profile of Raman spectra plotted against intensity, wave number and excitation wavelength of f3-uridine-5'-phosphoric acid (Figure 5.11) (Nishimura et aI., 1978). Preresonance or resonance Raman spectra can be obtained by the use of uv radiation of different wavelengths. There are several advantages to having resonance-enhanced Raman intensities. First, one can work with a lower concentration. Second, the solvent background spectrum can be eliminated. When Penicillium crysogenum mycophage RNA is excited at 514.5 nm, many bands are masked by the solvent background. However, the preresonanceenhanced spectrum of the same sample shows very little water-scattering
166
Nucleic Acids
Molar decadic 7 absorbance, «/M-1cm- 1
5000 3000 10001 ...\
20kl
250
~ ()"-
.,,,,,
~ 400 ~...
~
.,~
r 1500
1000 Wavenumber / cm-'
FIGURE 5.11. Three-dimensional view of Raman spectra of ,B-uridine-5'-phosphoric acid. The figure was reproduced (rom Nishimura et al. (1978).
background, and Raman bands due to RNA are shown clearly. When AMP, UTP, GMP, and CMP are illuminated with 300-nm laser light, the preresonance effect is observed. Their bromo derivatives, however, show the resonance Raman effect, and Raman intensity is greatly enhanced even in greatly diluted samples. This suggests that once the bromo-substituted nucleotide is incorporated into DNA, one can study a small part of the macromolecule with resonance Raman spectroscopy (Chinsky et aI., 1978a; Bushaw et aI., 1978).
....
References
167
By exciting AMP-5' with uv light, the scattering intensities of some Raman bands can be enhanced by about 10 5 . The Raman bands at 1484 and 1583 cm - I obtain their intensity from weak electronic transitions at 276 nm; the bands in the region of 1300-1400 cm - I derive at least part of their intensity from an electronic band whose 0-0 transition is in the 269-259 nm region (Blazej and Peticolas, 1977). The main role of tRNA is to transfer specific amino acids in the" translation" process of protein biosynthesis. tRNA is the smallest RNA; it contains unusual bases such as 4-thiouridine and ribothymidine. Among the bases present in tRNA, only 4-thiouridine has a strong absorption band at 330 nm. Therefore, one can obtain resonance-enhanced Raman spectra for tRNA because of the thiouridine vibration. The C=S stretching vibration is identified at 708 cm -1. This technique can be used for the analysis of thiouridine in tRNA, as some tRNAs contain this modified base, whereas others do not (Nishimura et al., 1976). However, the resonance Raman bands of thiouridine are short lived, as photochemical reactions take place because of the laser beam. Nicotinamide adenine dinucleotide has two different chromophores: the adenine base and nicotinamide. By selecting specific uv excitation lines, one obtains two different resonance Raman spectra of NADH (Rodgers and Peticolas, 1980). 7.2.
OTHERS
Excitation profiles-Raman intensities as a function of excitation frequencyfor AMP, ApA, and poly(A) have been obtained, but no appreciable differences between these compounds have been observed (Bushaw et al., 1980). It is reported that an order-disorder transition of poly(U) can be reflected in the excitation profile (Chinsky and Turpin, 1980).
REFERENCES Ackermann, Th., Gramlich, V., Klump, H., Knable, Th., Schmid, E. D., Seliger, H., and Stulz, J. (1979). Demonstration of G . U wobble base pairs by Raman and IR spectroscopy. Biophys. Chem. 10,231. Alix, A. 1. P., Bernard, L., Manfait, M., Ganguli, P. K., and Theophanides, T. (1981). Binding of cis- and trans-dichlorodiamine platinum(II) to nucleic acid studied by Raman spectroscopy. 1. Salmon sperm DNA. Inorg. Chim. Acta 55, 147. Arie, G., Da Silva, 12., Dumas, G., Rozannsza, H., and Scbenne, C. (1971). Etude des molecules biologiques par spectrometric Raman. Exemple d'application: Guanine, Uracyl et Cytosine. Biochimie 53, 1041.
l
Aylward, N. N., and Koenig, J. L. (1970). Raman spectrum of poly(adenylic) acid. Macromolecules 3,590. Baret, 1. F., Carbone, G. P., and Sturm, J. (1979). High frequency vibrational modes of poly(dA) . poly(dT) and poly(dAT) . poly(dAT). J. Raman Spectrosc. 8, 291. Barrett, T. W. (1980). pH-Induced modification of NAD and NADH solutions detected by Raman spectroscopy. J. Raman Spectrosc. 9, 130.
168
Nucleic Acids
Bertoluzza, A, Bertoluzza Morelli, M. A, Finelli, P., and Tosi, R. (1978). Raman and IR spectra of spermine and spermidine phosphate hexahydrate in connection with interactions between polyamines and nucleic acids. ltal. J. Biochem. 27, 323. Blazej, D. c., and Peticolas, W. L. (1977). Ultraviolet resonant Raman spectroscopy of nucleic acid components. Proc. Nat. Acad. Sci. 74,2639. BIazej , D. c., and Peticolas, W. L. (1980). Ultraviolet resonance Raman excitation profiles of pyrimidine nucleotides. J. Chem. Phys. 72, 3134. Brown, E. B., and Peticolas, W. L. (1975). Conformational geometry and vibrational frequencies of nucleic acid chains. Biopolymers 14, 1259. Brown, K. G., Kiser, E. 1., and Peticolas, W. L. (1972). The conformation of polycytidylic acid, polyguanylic acid, polyinosinic acid, and their helical complexes in aqueous solution from laser Raman scattering. Biopolymers 11, 1855. Bushaw, T. H., Lytle, F. E., and Tobias, R. S. (1978). The frequency doubled, synchronously pumped dye laser as a source for resonance Raman spectroscopy of nucleic acid constituents. Appl. Spectrose. 32, 585. Bushaw, T. H., Lytle, F. E., and Tobias, R. S. (1980). The determination of Raman excitation profiles of adenine derivatives in the 285 to 320 nm region. Appl. Spectrosc. 34, 52!. Chen, M. C., and Thomas, Jr., G. J. (1974). Raman spectral studies of nucleic acids. XI. Conformations of years tRNAPhe and E. coli ribosomal RNA in aqueous solution and in the solid state. Biopolymers 13, 615. Chen, M. c., Geige, R., Lord, R. c., and Rich, A (1975). Raman spectra and structure of yeast phenylalanine transfer RNA in the crystalline state and in solution. Biochemistry 14,4385. Chen, M. c., Giege, R., Lord, R. c., and Rich, A (1978). Raman spectra of ten aqueous transfer RNAs and 5S RNA Phenylalanine transfer RNA. Biochemistry 17, 3134. Chinsky, L., and Turpin, P. Y (1978). Ultraviolet resonance Raman study of DNA and of its interaction with actinomycin D. Nucleic Acids Res. 5, 2969. Chinsky, L., and Turpin, P. Y. (1980). Ultraviolet Raman spectroscopy of polyribouridylic acid: Excitation profile of the hypochromism induced by order-disorder transition. Biopolymers 19, 1507. Chinsky, L., Turpin, P. Y., Duquesne, M., and Brahms, J. (1975). Resonance Raman study of actinomycin D interaction with DNA and its models. Biochem. Biophys. Res. Commun. 65, 1440. Chinsky, L., Turpin, P. Y, and Duquesne, M. (1976). Resonance Raman spectra of highly fluorescent molecules: Free and DNA-bonded ethidium-bromide. In Proc. Int. Conf. Raman Spectrosc., 5th, E. D. Schmid, 1. Brandmueller, and W. Kiefer, Eds., Hans Ferdinand Schulz Verlag, Freiburg/Br., Germany, pp. 196-7. Chinsky, L., Turpin, P. Y., Duquesne, M., and Brahms, J. (1977). Structural investigation of poly d(BrU-A) by ultraviolet resonance Raman spectroscopy. Biochem. Biophys. Res. Commun. 75, 766. Chinsky, L., Turpin, P. Y., and Duquesne, M. (1978a). Nucleic acid derivatives studied by preresonance and resonance Raman spectroscopy in the ultraviolet region. Biopolymers 17, 1347. Chinsky, L., Turpin, P. Y., Duquesne, M., and Brahms, J. (1978b). Ultraviolet resonance Raman studies of nucleic acid complexes with proteins. In Proc. Sixth Int. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 92-99. Chou, C. H., and Thomas, Jr., G. J. (1977). Raman spectral studies of nucleic acids. XVI.. Structures of polyribocytidylic acid in aqueous solution. Biopolymers 16, 765. Chou, C. H., Thomas, Jr., G. J., Arnott, S., and Smith, P. 1. C. (1977). Raman spectral studies of ~ nucleic acids. XVII. Conformational structures of polyinosinic acid. Nucleic Acids Res. 4, 2407. .
I
j'
References
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Chrisman, R. W., Mansy, S., Peresie, H. 1., Ranade, A, Berg, T. A, and Tobias, R. S. (1977). Heavy metal-nucleotide interactions. IX. Raman difference spectroscopic studies on the binding of CH 3 Hg(II) to I-methylthymine, thymidine-5'-monophosphate, DNA models and native DNA Bioinorganic Chem. 7,245. Chu, G. Y. H., Mansy, S., Duncan, R. E., and Tobias, R. S. (1978). Heavy metal nucleotide interactions. II. Stereochemical and electronic effects in the electrophilic attack of cis- and trans-diamineplatinum (II) on 5'-guanosine monophosphate and polyguanylate in aqueous solution. J. Am. Chem. Soc. 100, 593. Delabar, 1.-M., and Guschlbauer, W. (1979). Raman spectroscopic study of 2H_ and 15N_ substituted guanosines: Monomers, gels, and polymers. Biopolymers 18, 2073. Erfurth, S. c., and Peticolas, W. L. (1975). Melting and premelting phenomenon in DNA by laser Raman scattering. Biopolymers 14, 247. Erfurth, S. c., Kiser, E. 1., and Peticolas, W. L. (1972). Determination of the backbone structure of nucleic acids and nucleic acid oligomers by laser Raman scattering. Proc. Nat. Acad. Sci. 69, 938. Erfurth, S. C., Bond, P. 1., and Peticolas, W. L. (1975). Characterization of the A .,. B transition of DNA in fibers and gels by laser Raman spectroscopy. Biopolymers 14, 1245. Fabian, H., Boehm, S., Welf1e, H., Reich, P., and Bielka, H. (1981). Laser Raman studies of rat liver ribosomal 5 S RNA FEBS Lett. 123, 19. Fairclough, D. P., Fawcett, V., Long, D. A, Taylor, L. H., and Turner, R. L. (1978). Raman spectroscopic studies of the interaction between DNA and the oncolytic agent bleomycin. In Proc. Sixth 1m. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 90-91. Forrest, G. (1976). Deuterium exchange in pyridine dinucleotide coenzymes. Raman spectroscopic evidence for a modified amide charge distribution in p-dihydronicotinamide adenine dinucleotide. J. Phys. Chem. SO, 1127. Forrest, G., and Lord, R. C. (1977). Laser Raman spectroscopy of biomolecules. X. Frequency and intensity of the phosphodiester stretching vibrations of cyclic nucleotides. J. Raman Spectrosc. 6,32. Goodwin, D. C., and Brahms, J. (1978). Form of DNA and the nature of interactions with proteins in chromatin. Nucleic Acids Res. 5,835. Gramlich, V. and Schmid, E. D. (1978). Raman studies on ribodinucleoside monophosphates in aqueous solution. In Proc. Sixth 1m. Conf. Raman Spectrosc., E. D. Schmid. R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 82-83. Gramlich, V., Klump, H., Herbeck, R., and Schmid, E. D. (1976a). A Raman investigation into the self-association of 5'-GMP in neutral aqueous solution. FEBS Lett. 69, 15. Gramlich, V., Klump, H., Herbeck, R., and Schmid, E. D. (1976b). A Raman investigation into the self-association of 5'-GMP in neutral aqueous solution. In Proc. Int. Conf. Ramall Spectrosc., 5th, E. D. Schmid. 1. BrandmueIIer, and W. Kiefer, Eds., Hans Ferdinand Schulz Verlag, Freiburg/Br., Germany, pp. 194-195. Hadjiliadis, N., and Theophanides, T. (1976). Platinum purine nucleosides. II. Interaction of K 2PtX 4 (X = CI, Dr) with inosine and guanosine. Inorg. Chim. Acta 16,77. Herbeck, R., Schlenker, P., Gramlich, V., and Schmid, E. D. (1976). Raman intensity measurements on nucleotides. In Proc. Int. Conf. Raman Spectrosc., 5th. E. D. Schmid, J. BrandmueIIer, and W. Kiefer, Eds., Hans Ferdinand Schulz Verlag, Freiburg/Br., Germany, pp. 192-193. Hernandez, L. A, Vieta, R. S., and Jao, T.-e. (1980). Laser Raman spectroscopic studies of poly(r-cytidylic acid) and poly(r-adenylic acid) complexes with poly(L-lysine) and poly(Larginine). Biopolymers 19, 1715. Herskovits, T. T., and Brahms, J. (1976). Structural investigations on DNA-protamine complexes. Biopolymers 15,687.
170
Nucleic Acids
King, T. c., Schlessinger, D., and Milanovich, F. (1980). Laser Raman studies of RNA backbone ordering in E. coli ribosomes. Biophys. J. 32 (abstract). King, T. c., Rucinsky, T., Schlessinger, D., and Milanovich, F. (1981). Escherichia coli ribosome unfolding in low Mg 2+ solutions observed by laser Raman spectroscopy and electron microscopy. Nucleic Acids. Res. 9, 647. Kiser, E. 1. (1976). Application of Raman spectroscopy to determination of nucleic acid conformations. Diss. Abstr. Int. B 36, 3189. Klainer, S. M., and Frazer, J. W. (1975). Discussion paper: Raman spectroscopy of molecular species during exposure to lOO-MHz radiofrequency fields. Ann. N. Y. Acad. Sci. 247, 323. Koglin, E., Sequaris, J. M., and Valenta, P. (1980). Surface enhanced Raman spectra of adenine monocucleotides adsorbed at a silver electrode. In Proc. Vllrh Int. Conf. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, p. 628. Lafleur, 1., Rice, J., and Thomas, Jr., G. J. (1972). Raman studies of nucleic acids. VII. Poly A . Poly U and Poly G . Poly C. Biopolymers 11,2423. Lane, M. 1., and Thomas, Jr., G. J. (1979). Kinetics of hydrogen-deuterium exchange in guanosine 5'-monophosphate and guanosine 3':5'-monophosphate determined by laser-Raman spectroscopy. Biochemistry 18, 3839. Lanir, A., and Yu, N.-T. (1979). A Raman spectroscopic study of the interaction of divalent metal ions with adenine moiety of adenosine 5'-triphosphate. J. BioI. Chern. 254, 5882. Lewis, A., Nelson, N., and Racker, E. (1975). Laser Raman spectroscopy as a mechanistic probe of the phosphate transfer from adenosine triphosphate in a model system. Biochemistry 14, 1532. Lippert, B. (1979). Uracil and thymine monoanions in solution: Differentiation of tautomers by laser Raman spectroscopy. J. Raman Spectrosc. 8, 274. Livramento, J., and Thomas, Jr., G. 1. (1974). Detection of hydrogen-deuterium exchange in purines by laser-Raman spectroscopy. Adenosine 5'-monophosphate and polyriboadenylic acid. J. Am. Chern. Soc. 96, 6529. Lord, R. c., and Thomas, Jr., G. 1. (1967a). Raman studies of nucleic acids. II. Aqueous purine and pyrimidine mixtures. Biochim. Biophys. Acta 142, 1. Lord, R. C., and Thomas, Jr., G. J. (\967b). Raman spectral studies of nucleic acids and related molecules-I. Ribonucleic acid derivatives. Spectrochimica Acta 23A, 2551. Lord, R. c., and Thomas, Jr., G. J. (\968). Spectroscopic studies of molecular interaction in DNA constituents. Develop. Appl. Spectrose. 6, 179. Lu, K. c., Prohofsky, E. W., and Van Zandt, 1. 1. (\977). Vibrational modes of A-DNA, B-DNA, and A-RNA backbones: An application of a green-function refinement procedure. Biopolymers 16, 2491. Luoma, G. A., and Marshall, A. G. (\978a). Laser Raman evidence for a new cloverleaf secondary structure for eucaryotic 5 S RNA. J. Mol. BioI. 125, 95. Luoma, G. A, and Marshall, A. G. (\ 978b). Laser Raman evidence for new cloverleaf secondary structures for eukaryotic 5.8 S RNA and prokaryotic 5 S RNA. Proe. Nat. Aead. Sci. 75, 4901. Majoube, M. (1978). Raman spectra of 15N(I,3) and deuterated hypoxanthine. In Proc. Sixth Int. Conf. Raman Spectrosc. In E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 8-9. Makrigiannis, G., Papagiannakopoulos, P., and Theophanides, T. (1980). Raman study of metalguanosine-5'-monophosphate aqueous solutions. Inorganica Chimiea Acta 46, 263. Manfait, M., Alix, A. J. P., Jeannesson, P., Jardillier, J. c., and Theophanides, T. (1980). Resonance Raman spectroscopic studies of the interaction between adriamycin and DNA. In Proc. Vllth Int. Conf. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 636-637. Manfait, M., Alix, A. J. P., Butour, J., Labarre, J., and Sournies, F. (1981). Raman studies on anticancer inorganic ring-DNA interactions. J. Mol. Struct. 71, 39.
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Mansy, S., and Peticolas, W. L. (1976). Detection of the sites of alkylation in DNA and polynucleotides by laser Raman spectroscopy. Biochemistry 15, 2650. Mansy, S., Chu, G. Y. H., Duncan, R. E., and Tobias, R. S. (1978). Heavy metal nucleotide interactions. 12. Competitive reactions in systems of four nucleotides with cis- or trallsdiamineplatinum(II). Raman difference spectrophotometric determination of the relative nucleophilicity of guanosine, cytidine, adenosine, and uridine monophosphates as well as the analogous bases in DNA J. Am. Chern. Soc. 100,607. Marshall, A G., and Smith, J. L. (1980). Raman and 19F1H nuclear Overhauser evidence for a rigid solution conformation of Escherichia coli 5-fluorouracil 5S ribonucleic acid. Biochimstry 19, 5955. Martin,1. C. (1979). A study of conformational features of distamycin-DNA and netropsin-DNA complexes by Raman spectroscopy. Diss. Abstr. Int. B 39,3388. Martin, 1. c., Wartell, R. M., and O'Shea, D. C. (1978). Conformational features of distamycinDNA and netropsin-DNA complexes by Raman spectroscopy. Proc. Nat. Acad. Sci. 75,5483. Marzilli, L. G., de Castro, B., Caradonna, J. P., Stewart, R. c., and Van Vuuren, C. P. (1980). Nucleoside complexing. A Raman and 13C NMR spectroscopic study of the binding of hard and soft metal species. J. Am. Chem. Soc. 102, 916. Medeiros, G. c., and Thomas, Jr., G. 1. (l97Ia). On the tautomeric structure of inosine. Biochim. Biophys. Acta 238, I. Medeiros, G. c., and Thomas, Jr., G. 1. (197Ib). Raman studies of nucleic acids. IV. Vibrational spectra and associative interactions of aqueous inosine derivatives. Biochim. Biophys. Acta 247,449. Millard, M. M., Macquet, J. P., and Theophanides, T. (1975). X-ray photoelectron spectroscopy of 06(Gua) . N 7 (Gua) chelation of DNA with cis-dichlorodiamine platinum(II). Biochim. Biophys. Acta 402, 166. Moller, M. R., Bruck, M. A, O'Connor, T., Armatis, Jr., F. J., Knolinski, E. A, Kottmair, N., and Tobias, R. S. (1980). Heavy metal-nucleotide interactions. 14. Raman difference spectrophotometric studies of competitive reactions in mixtures of four nucleotides with the electrophiles methylmercury(II) perchlorate, cis-dimethylgold(III) perchlorate, dichloroethylene-diaminepalladium(I1), tralls-dichlorodiaminepalladium(I1), cis-dichlorodiamineplatinum(II), transdichlorodiamineplatinum(II), tetra-waceto-dirhodium(I1), and aquopentaaminecobalt(III) perchlorate. Factors governing selectivity in the binding reactions. J. Am. Chem. Soc. 102, 4589. Morikawa, K., Tsuboi, M., Takahashi, S., Kyogoku, Y., Mitsui, Y., Iitaka, Y., and Thomas, G. J. (1973). The vibrational spectra and structure of poly(rA-rU) . poly(rA-rU). Biopolyrners 12, 799.
.....
Nishimura, Y., Hirakawa, A Y., Tsuboi, M., and Nishimura, S. (1976). Raman spectra of transfer RNAs with ultraviolet lasers. Nature 260, 173. Nishimura, Y., Hirakawa, A Y., and Tsuboi, M. (1978). Resonance Raman Spectroscopy, Vol. 5, R. H. H. Clark and R. E. Hester, Eds., Heyden, London, Philadelphia, Rheine, pp. 217-275. O'Connor, T., and Scovell, W. M. (1981). pH-Dependent Raman spectra and thermal melting profiles for polycytidylic acid. Biopolymers 20, 2351. O'Connor, T., Johnson, c., and Scovell, W. M. (1976a). Raman pH profiles for nucleic acid constituents. I. Cytidine and uridine ribonucleosides. Biochim. Biophys. Acta 447, 484. O'Connor, T., Johnson, c., and Scovell, W. M. (I 976b). Raman pH profiles for nucleic acid constituents. II. 5'-AMP and 5'-GMP ribonucleotides. Biochirn. Biophys. Acta 447, 495. Painter, P. c., Mosher, L., and Rhoads, C. (1981). Low-frequency modes in the Raman spectrum of DNA Biopolymers 20, 243. Patrick, II, D. M., Wilson, J. E., and Leroi, G. E. (1974). A Raman and infrared spectroscopic study of 3-carbonyl group of pyridine nucleotide coenzyme and related model compounds. Biochemistry 13, 2813.
172
Nucleic Acids
Peticolas, W. L. (1971). Raman spectroscopy of polynucleotides and nucleic acids. Proc. Nucleic Acid Res. 2, 94. Peticolas, W. L. (1975). Applications of Raman spectroscopy to biological macromolecules. Biochimie 57,417. Peticolas, W. L., and Lippert, J. L. (1973). Spettroscopia Raman di molecole di tipo biologico. La Nuova Chimica 8, 28. Peticolas, W. L., Small, E. W., and Fanconi, B. (1971). Characterization of Biological Polymers by Laser Raman Scattering, Polymer Characterization: Interdisciplinary Approaches, C. D. Craver, Ed., Plenum, pp. 47-77. Pezolet, M., Yu, T.-J., and Peticolas, W. L. (1975). Resonance and preresonance Raman spectra of nucleotides using ultraviolet lasers. J. Raman Spectrosc. 3, 55. Pham, V. H. and Giege, R. (1981). Conformation and Raman spectra of transfer ribonucleic acid, tRNAAsP. Biochimie 63,921. Prescott, B., Gamache, R., Livramento, J., and Thomas, Jr., G. J. (1974). Raman studies of nucleic acids. XII. Conformations of oligonucleotides and deuterated polynucleotides. Biopolymers 13, 1821. Prescott, B., Chou, C. H., and Thomas, Jr., G. J. (1976). A Raman spectroscopic study of complexes of polylysine with deoxyribonucleic acid and polyriboadenylic acid. J. Phys. Chern. SO, 1164. Rice, Jr., L. L., and Thomas, Jr., G. J. (1972). Raman studies of nucleic acids. VII. Poly A . Poly U and Poly G . Poly C. Biopolymers 11, 2423. Rice, J., Lafleur, L., Medeiros, G. c., and Thomas, Jr., G. J. (1973). Raman studies of nucleic acids. IX. A salt-induced structural transition in poly(rG). J. Raman Spectrosc. 1, 207. Rimai, L., Maher, V. M., Gill, D., Salmeen, 1., and McCormick, J. J. (1974). The temperature dependence of Raman intensities of DNA evidence for premelting changes and correlations with ultraviolet spectra. Biochim. Biophys. Acta 361, 155. Ringland, R. L., Milanovich, F. P., Weiss, M. A., and Balhom, R. (1979). Chromatin structure: kinetics of deuterium exchange at the C-8 hydrogen of purines. Fed. Proc. 38, 1391. Rodgers, E. G. and Peticolas, W. L. (1980). Selective ultraviolet resonance Raman excitation of individual chromophores in nicotinamide adenine dinucleotide. J. Raman Spectrosc. 9, 372. Savoie, R., Klump, H., and Peticolas, W. L. (1978). A Raman and calorimetric investigation of 3', 5'-GpG. Biopolymers 17, 1335. Schmid, E. D., and Gramlich, V. (1979). Ramanspektroskopische Untersuchungen an Nukleinsauren. Acta Physica Austriaca XX (Suppl.), 75. Scovell, W. M., Amamath, V., and Broom, A. D. (1979). Raman spectral studies of poly(l-methylinosinic acid). Nucleic A cids Res. 6, 1049. Siiman, 0., and Callaghan, R. (1981). Resonance Raman studies of ferric NADH transients. J. Am. Chern. Soc. 103, 5526. Small, E. W., and Peticolas, W. L. (l97Ia). Conformational dependence of the Raman scattering intensities from polynucleotides. Biopolymers 10, 69. Small, E. W., and Peticolas, W. L. (l97Ib). Conformational dependence of the Raman scattering intensities from polynucleotides. III. Order-disorder changes in helical structures. Biopolymers 10, 1377. Small, E. W., Brown, K. G., and Peticolas, W. L. (1972). Structural changes in t-RNA from changes in the Raman scattering intensities. Biopolymers 11, 1209. Smulevich, G., Angeloni, L., and Marzocchi, M. P. (1980). Raman excitation profiles of actinomycin D. Biochim. Biophys. Acta 610, 384. Strobel, J. L., and Scovell, W. M. (1980). Laser Raman spectroscopy of a complementary base pair in the Hoogsteen configuration. Biochim. Biophys. Acta 608,201.
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Sturm, J., Savoie, R., Edelson, M., and Peticolas, W. (1978). Fast Raman spectroscopic detection of rapidly disordering macromolecules. Indian J. Pure Appl. Phys. 16, 327. Theophanides, T. (1979). Vibrational spectroscopy of metal nucleic acid systems. In Infrared and Raman Spectroscopy of Biological Molecules, T. Theophanides, Ed., Reidel, Dordrecht, HoIland; Boston; London; pp. 205-223. Theophanides, T. (1980). Cancer and platinum coordination compounds. Chern. in Canada 32,30. Thomas, Jr., G. J. (1975). Structural studies of nucleic acids and polynucleotides by laser-Raman spectroscopy. In Structure and Conformation of Nucleic Acids and Protein-Nucleic Acid Interactions, M. Sundaralingam and S. T. Rao, Eds., University Park Press, Baltimore, pp. 253-281. Thomas, Jr., G. 1., and Hartman, K. A. (1973). Raman studies of nucleic acids. VIII. Estimation of RNA secondary structure from Raman scattering by phosphate-group vibrations. Biochim. Biophys. Acta 312, 311. Thomas, Jr., G. 1., and Lane, M. J. (1980). A kinetic isotope effect in purinic 8-CH exchange of guanine nucleotides: Measurement by Raman spectroscopy. J. Raman Spectrosc. 9, 134. Thomas, Jr., G. 1., and Livramento, J. (1975). Kinetics of hydrogen-deuterium exchange in adenosine 5'-monophosphate, adenosine 3':5'-monophosphate, and poly(riboadenylic acid) determined by laser-Raman spectroscopy. Biochemistry 14, 5210. Thomas, Jr., G. J., Medeiros, G. c., and Hartman, K. A. (1971). The dependence of Raman scattering on the conformation of ribosomal RNA. Biochem. Biophys. Res. Commun. 44, 587. Thomas, Jr., G. J., Medeiros, G. c., and Hartman, K. A. (1972). Raman studies of nucleic acids. VI. Conformational structures of tRNAfMET , tRNA Val and tRNA 2 Phe. Biochim. Biophys. Acta 277, 71. Thomas, Jr., G. 1., Chen, M. c., and Hartman, K. A. (1973a). Raman studies of nucleic acids. X. Conformational structures of Escherichia coli transfer RNAs in aqueous solution. Biochim. Biophys. Acta 324, 37. Thomas, Jr., G. J., Chen, M. c., Lord, R. C., Kotsiopoulos, P. S., Tritton, T. R., and Mohr, S. C. (l973b). Transfer RNA: Change of conformation upon aminoacylation determined by Raman spectroscopy. Biochem. Biophys. Res. Commun. 54, 570. Thomas, Jr., G. J, Prescott, B., and Hamilton, M. G. (1980). Raman spectra and conformational properties of ribosomes during various stages of disassembly. Biochemistry 19, 3604. Tobias, R. S., Chrisman, R. W., Chu, G. Y. H., English, J. c., and Moller, M. R. (1976). Interactions of nucleotides and polynucleotides studied by automated Raman difference spectrophotometry. In Proc. Int. Con/. Raman Spectrosc., 5th. E. D. Schmid, 1. Brandmueller, and W. Kiefer, Eds., Hans Ferdinand Schulz Verlag, Freiburg/Br., Germany, pp. 188-189. Tobias, R. S., Bushaw, T. H., and English, 1. C. (1978). Raman difference spectroscopy in the study of reactions of biological molecules. Indian J. Pure Appl. Phys. 16,401. Tomlinson, B. L., and Peticolas, W. L. (1970). Conformational dependence of Raman scattering intensities in polyadenylic acid. J. Chern. Phys. 52, 2154. Tu, A. T, and Heller, M. J. (1974). Structure and stability of metal-nucleoside phosphate complexes. In Metal Ions in Biological Systems, H. Sigel, Ed., Dekker, New York, pp. 1-49. Tu, A. T., and Reinosa, 1. A. (1966). The interaction of silver ion with guanosine, guanosine monophosphate, and related compounds, determination of possible sites of complexing. Biochemistry 5, 3375. Turpin, P. Y., and Chinsky, L. (1978), DNA and DNA-actinomycin D interaction studied by visible and ultraviolet resonance Raman spectroscopy, In Proc. Sixth Int. Con/. Raman Spectrosc. E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 86-89.
CHAPTER
NucleoproteinsVirus and Chromosome
Many nucleic acids are present in nature as nucleoproteins. Nucleic acids are usually highly elongated chains linked by phosphodiester bonds. The phosphodiester bond is highly susceptible to nonspecific phosphodiesterases and to specific phosphodiesterases such as DNase and RNase. Cleavage of one phosphodiester bond in a long chain of a nucleic acid immediately reduces the molecular size of the nucleic acid, sometimes rendering it biologically inactive. Therefore, nucleic acids preserve their intact structure and biological function by combining with protein to form stable nucleoproteins.
1.
VIRUSES
1
From the chemical viewpoint, viruses are nucleoproteins that contain proteins and either RNA or DNA. .Infectivity resides in the nucleic acid portion and not in the protein moiety, but the protein coat usually protects nucleic acids from being hydrolyzed by surrounding cellular nucleases. Depending on the
j
1
.•.•.•
Viruses
175
type of host, the virus can be classified as an animal, plant, or bacterial virus (bacteriophage). The application of Raman spectroscopy to viruses is relatively new. This unique technique can examine the structure of RNA and viral protein within an intact virus particle. Analysis of proteins and nucleic acids in a virus follows a strictly empirical approach. The backbone conformations of both proteins and nucleic acids have been well studied by Raman spectroscopy by using a variety of proteins and nucleic acids. The microenvironment of some amino acid residues and the state of base pairing and stacking have also been well studied by Raman spectroscopy. However, obtaining spectra of intact virus presents some problems. For example, the nucleic acid content of the virus must be considered. If the content is low, the Raman spectrum of a virus will be similar to that of a protein. However, when the nucleic acid content is high, Raman bands and their intensities result from the contribution of both components. Thus one can analyze the states of proteins as well as nucleic acid components in an intact virus particle. One should pay attention to the following special bands for the nucleic acid structure in a virus: 1.
Ordered structure is detected from the 815-cm - I band and will shift to a lower frequency upon disordering. 2. Some bands show Raman hypochromism and others show Raman hyperchromism with changing temperature or pH. Analysis of these bands indicates the state of the bases in viral nucleic acids, for example, base-base hydrogen bonding, base-base stacking, and base-protein interactions. 3. The intensity of C=O stretching vibration band of uracil, guanine, cytosine, and thymine bases is usually strong. Shifts in this band resulting from a change in pH or temperature may involve the keto group of these bases. One also should compare this band in free nucleic acids and in nucleic acids within a virus particle. 1.1.
PLANT VIRUSES
Plant viruses are extensively used in virus research because they are not infectious to humans, and they can be used as a good model for the transfer of genetic information. Among the many plant viruses, tobacco mosaic virusTMV-is one of the most extensively studied. In this section we will mainly use TMV as an example to illustrate how Raman spectroscopy can be used to analyze intact virus particles and their components. TMV is a very stable virus containing only 5% RNA. It is a very large elongated particle with typical dimensions of 300 X 15 nm (Figure 6.1). TMV is a hollow cylinder; the diameter of the empty hole is 4 nm. The RNA is
176
Nucleoproteins-Virus and Chromosome
FIGURE 6.1. Electron micrograph of tobacco mosaic virus (TMV) shadow-cast with uranium. The spherical object is a polystyrene ball with a diameter of 80 nm, used as an internal standard to compare with the size of the elongated TMV particles. The electron micrograph was made by the author.
embedded inside the virus particle and has a diameter of 8 nm. The threedimensional model is shown in Figure 6.2. The coat protein has a molecular weight of 12,000, and the subunit protein readily aggregates into a rod-shaped particle at proper pH and salt concentration. The aggregated protein· rod is indistinguishable from a native virus when it is viewed under the electron microscope, but it is noninfectious because it lacks RNA. The infectious RNA can be isolated from TMV by the phenol or the SDS method. The Raman spectrum of intact TMV particles appears to be dominated by the contribution from the protein component rather than from the RNA (Fox et al., 1979). This is understandable, as TMV consists of 95% protein and only 5% RNA. The ionic-phosphate-group symmetrical-stretching vibrational band at 1100 cm- I is masked and appears only as a shoulder at 1093 cm- I . Also the phosphodiester vibrational band at 815 cm - I becomes unclear (Figure 6.3). The amide I band at 1656 cm -I of the intact virus is much more intense than the amide I band of the coat protein (Figures 6.3 and 6.4). This is probably due to the contribution from the C=O stretching vibration of viral RNA in TMV. Nevertheless, it suggests the presence of a-helical structure in the protein. The spectrum of the isolated coat protein also suggests the presence of a-helix. X-ray crystallographic studies of the TMV protein by Stubbs et al. (1977) show five regions of a-helical structure with a total content of 41 % a-helix. Structural prediction analysis by Chou and Fasman's method indicates the presence of 32% a-helix, 31 % ,B-sheet, and 22% ,B-tum. TMV protein contains 1 mol of free sulfhydryl groups per mole of protein, and does not contain any disulfide bonds. The lack of disulfide bonds is demonstrated in the Raman spectrum by the absence of disulfide stretching vibrations in the region of 500-600 cm -I. However, an SH stretching vibration
...
Viruses
177
G (\.00.··. \)0
O\J
+
o
Prolein Subunits
Intermediote Step in TMV Formation
FIGURE 6.2. Diagram indicating how TMV coat-protein subunits and the viral RNA strand combine to form a virus particle.
band is seen at 2567 cm-] (Figure 6.5), which confirms the presence of a free sulfhydryl group in the protein. The sulfhydryl group in TMV is known to be masked, and many chemical reagents are unable to react with this group, yet Raman spectroscopy can detect it. A direct comparison of the Raman spectra of solid and aqueous coat protein is interesting. The isolated protein shows the amide I band at 1650 and 1657 cm -] in aqueous solution and at 1649 and 1659 cm - I in the solid state. From this evidence it can be concluded that the peptide-backbone configuration of the coat protein is the same in both states. However, differences in the tyrosine bands at 850 and 830 cm -] suggest that the tyrosine side chains may have different conformations in the two phases. Judging from the 1815 /11100 ratio, the isolated infectious viral RNA has 51 % ordered structure compared with 85% for transfer RNA and ribosomal RNA (Figure 6.6). Turnip yellow mosaic virus (TYMV) is a plant virus with a high molecular weight (2 million), about one-third of which is RNA. It consists of 180 molecules of a coat protein that is relatively small, with a molecular weight of 20,100. Unlike TMV, TYMV has a much higher RNA content, and the Raman spectrum of TYMV shows vibrational frequencies characteristic of both RNA
>-
1686
RNA
[WAT E RI
"; c
G)
1050
c
"-C
888
tV
E tV
a:
D
j..o-
497
1126 I
\j 1600
1400
1200
'H20
noo
800
600
Wave number
(eM-')
FIGURE 6.3. Raman spectrum of TMV in aqueous solution. The figure was obtained from Fox et al. (1979) by permission of the copyright owner, Academic Press.
-,>"0 C
G)
C
C
~
E
!
CH
1124
13~58303 I
1273
I
~
499 H,D
FIGURE 6.4. Raman spectrum of viral coat protein (A-protein) from TMV. The figure was obtained from Fox et al. (1979) by permission of the copyright owner, Academic Press.
10.
Viruses
179
SH
>-
2567
CIJ C
Q)
c c
~
E ~ a: 2720 2660 2560 2500 leM-11
Wave number
FIGURE 6.5. Raman spectrum of TMV coat protein in the region of 2500-2660 cm -) , indicating the presence of a sulfhydryl group. The figure was reproduced from Fox et aI. (1979) by permission of the copyright owner, Academic Press.
and protein. Like TMV, TYMV has no disulfide bond, but it contains four sulfhydryl groups. It shows a typical SH stretching vibration at 2569 cm -1 that shifts to 1868 cm-( on deuteration (Turano et aI., 1976). The RNA within a TYMV particle has approximately 77% ordered structure compared with 51 % for RNA within a TMV particle (Hartman et aI., 1978). The coat protein contains 9% a-helix, 43% ,8-sheet, and 48% irregular structure in the isolated form or in the encapsulated form. Raman spectroscopy can be used to determine not only protein secondary structure, but also the microenvironment
>-
CIJ
TMV [WATER]
C
Q)
c
c:
~
E ~
a:
FIGURE 6.6. Raman spectrum of viral RNA isolated from TMV. The figure was reproduced from Fox et aI. (1979) by permission of the copyright owner, Academic Press.
180
Nucleoprotelns-Virus and Chromosome
of selected side chains. Raman data indicate that the tryptophan and cysteine residues of the coat protein appear to be in contact with the solvent, whereas only one of three tyrosines per coat protein is available for hydrogen bonding of its p-hydroxyl group with water molecules. For the viral RNA, both cytosine and adenine bases are protonated near pH 4.5. This means that the interaction between the protonated bases of the RNA and the amino acid side chains of the coat protein is partly responsible for maintaining the intact virus particle. This conclusion is reasonable, as it is known that the stability of TYMV is increased as the pH of the solution is lowered from 7 to below 6 (Hartman et aI., 1978).
1.2.
BACTERIOPHAGES
Phage Rl7 is small, spherical RNA-containing virus. A protein coat (capsid) is found on the surface of an isosahedron, and the RNA residues are inside the sphere. Raman spectra of isolated RNA and of the viral RNA within the phage are similar, indicating that the RNA structure is similar in either the virus particle or the free state. However, judging from Raman lines, the configuration of guanine residues within the phage and in the isolated RNA is different. This suggests that guanine may be important in the interaction of RNA and protein to maintain a virus particle (Hartman et aI., 1973). MS2 is a spherical bacteriophage that contains RNA. The molecular weight of one viral RNA strand is about one million, and that of each protein subunit, 38,000. The intact virus consists of one RNA strand and 180 identical coat-protein subunits. For a virus, MS2 is relatively small, with a diameter of approximately 26 nm. MS2 phage is stable up to 55°C. The protein coat has two sulfhydryl groups but no disulfide bridge. Thus no S-S vibrational frequency can be found in the 500-600-cm- I region in the Raman spectra of either intact phage or the isolated protein. Unlike the intact virus, isolated RNA is relatively heat labile, as evidenced from the melting point of the RNA detected from many Raman lines (786, 665, 672, 1531 cm- I) (Thomas et aI., 1976). There is substantial disordering of the RNA secondary structure with breaking up of base pairing with increased temperature to 50°C; however, the unstacking of purine and pyrimidine bases takes place mainly above 50°C. The study suggests that the viral capsid stabilizes the secondary structure of MS2 RNA. The coat protein has a high ,B-sheet content and random-chain structure. In this aspect MS2 is quite different from the filamentous DNA phages fd and Pfl, in which the coat proteins are mainly a-helical (Thomas et aI., 1976). P22 is a DNA-containing bacteriophage. Raman spectra show that P22 DNA exists in the B form both inside the phage head and in the free state. The major coat protein (gp5) is composed of 18% a-helix, 20% ,B-sheet, and 62% irregular conformations. The gp8 scaffolding protein in the phage prohead contains more a-helix than gp5 protein (Fish et aI., 1980). The intensity of Raman scattering by nucleic acids can be enhanced by using uv laser excita-
~
\;,.. Viruses
181
tion. The Raman scattering of the P22 DNA is more prominent than that of phage protein when 363.8-nm light is used (Li et al., 1981). The phage fd also has a long filamentous structure of about 900 nm length with 6 nm diameter and contains about 2700 coat-protein molecules. Bacteriophage Pfl is an elongated virus that infects gram-negative bacteria and is structurally similar to fd phage. The viral DNA is coated with protein subunits in the a-helix conformation (Figure 6.7). Pfl and fd phages contain only 12% DNA; therefore, their Raman spectra are dominated by the contributions from the protein components. Pfl and fd have different amino acid compositions, and the Raman spectra of the two viruses reflect this difference. Depending on conditions, the phage fd coat protein can assume three distinct conformations (Thomas and Murphy, 1975; Fodor et al., 1981). Biological assay shows no loss of viral activity due to laser illumination; thus the Raman data reflect the properties of biologically active bacteriophages. The DNA bases have considerable stacking, as shown by the low intensities of the pyrimidine line at 785 cm - I and the purine line near 1580 cm- I. The ratio I s5o/Is3o for fd in H 20 is 10/2.7, indicating that the tyrosine residues are buried. This agrees well with the pH titration and uv spectroscopic studies, which show that the tyrosine residues do not become deprotonated unless the phage becomes disrupted (Dunker et al., 1979). Both Pfl and fd coat proteins have the same a-helical secondary structures, and viral-DNA backbones do not exhibit the A-type geometry, according to
(a)
(b)
(e)
FIGURE 6.7. Structure of bacteriophage PO. The a-helical segments in PO viral protein subunits are represented as cylinders around the central core, occupied by DNA in the virus particles. (A) Two axial repeats of the inner layer of rod segments. (B) 2.5 repeats of both layers. (C) Two repeats of the outer layer. The figure was obtained from Crowther (1980) by permission of the copyright owner, Macmillan Journals Ltd.
182
Nucleoprotelns- Virus and Chromosome
evidence obtained from Raman spectra (Thomas and Murphy, 1975). PB is also a filamentous phage, and its coat protein is predominantly a-helical; this is similar to the case with the subunits of phages Pfl and fd. However, the subunits of PB are converted to ,8-sheet structures when the temperature is raised. When the subunits are cooled, the ,8-structure reverts to an a-structure identical to the native phage protein (Thomas and Day, 1981; Thomas et al., 1981). 2.
CHROMOSOME
The cell nucleus contains a substance that appears dense when observed under a microscope. This is a collection of nucleoprotein fibrils generally called chromosomes. Genetic information is stored in the base sequence of DNA within the chromosome. The chromosome is composed of many subunits, including some called nucleosomes. Our knowledge of nucleosome structure is still incomplete. Basically, these complex nucleoproteins are maintained by the ionic interaction between acidic DNA and basic proteins called histones. Nucleosomes consist of regions of double-helical DNA about 200 base pairs in length wrapped around a histone protein core. It is believed that nucleosomes are coiled into a super-helical structure. Histones are a complex mixture of basic proteins. They are usually subdivided into groups called lysine rich (HI), moderately lysine rich (H2A and H2B), and arginine rich (H3 and ·H4). Chromatin also contains nonhistone proteins, which may playa role similar to that played by the histones. 2.1.
HISTONES
Chromatin is composed of subunits called nucleosomes (nu) bodies that are complexes of DNA with five types of histones to form an ordered structure. Histone fractions are called HI, H2A, H2B, H3, and H4. DNA consists of about 140 nucleotide base pairs enveloping a protein core (inner histones) that is composed of 2 mol each of H2A, H2B, H3, and H4. Adjacent nucleosomes bodies ar~ connected by a stretch of DNA (30 to 70 base pairs), to which the lysine-rich histone H I are bound. Histones can be separated from DNA in nucleosomes in 2 M NaCl. Judging from the amide I band at 1656 em- 1 and the amide III bands at 1274, 1252, and 1234 em-I, the purified inner histone has a high a-helix content. The amide I and III bands of nucleosomes are similar to those of isolated histones, suggesting that the secondary structure of the inner histones remains the same, whether they are associated with DNA or not (Thomas et al., 1977). However, pezolet et al. (1980) analyzed the arginine-rich histones H3 and H4 and concluded that they are rich in ,8-sheet content (about 40%) with less a-helix content (about 20%). On the other hand, lysine-rich histones HI, H2A, and H2B are found to be predominantly disordered, without any substantial ,8-sheet conformation (Guillot et al., 1977).
....
Chromosome
2.2.
183
DNA-HISTONE INTERACTIONS IN NUCLEOSOMES
The important question is how DNA and histone components interact with each other to form the biologically integral unit of the nucleosome. DNA is acidic and histone is basic; therefore, acidic and basic interactions naturally play a role. However, the mechanism of the interaction is more complicated than a simple acid-base combination. The DNA double helix has two types of grooves, one minor and one major (Figure 6.8). A minor groove lies between the C-I' of deoxyribose on two strands, whereas a major groove is found on the opposite side of the helix. It is believed that both the major and minor grooves are sites of protein interaction with DNA. In the study of nucleosome structure, two Raman bands at 1490 and 1580 cm- I are used. The l490-cm- 1 band of guanine is sensitive to substitution. Methylation or protonation at the N-7 position of guanine affects DNA interactions at this position in B-type DNA, in which this position is exposed to the large groove. The evidence suggests that a hydrogen bond involving guanine occurs at the N-7 position (Mansy et al., 1976). One approach used to study this is to look at the Raman spectra of nucleosomes that differ in their nonhistone-protein contents. The 1490-cm - I band is sensitive to substitution, methylation, or protonation at the N-7 position of guanine and therefore reflects DNA interaction at this position, which in B-type DNA is exposed at the large groove. The
NUCLEOSOME CORE
FIGURE 6.8. Attachment of proteins to DNA in a nuc1eosome. Nonhistone proteins (NHP) probably attach to the major groove, and histones attach to the minor groove.
184
Nucleoproteins-Virus and Chromosome
1580-cm - I band is due to the adenine-base vibration near the N-3 position, which lies in the minor groove. Therefore, by observing the intensity of these two bands in spectra of nucleosomes of different nonhistone-protein content, it is concluded that histone attaches in the minor groove, whereas nonhistone protein attaches in the major groove (Figure 6.8). The evidence for interaction is that the intensity of the 1490-cm- 1 band is low compared with that of DNA when spectra are obtained from nucleosomes and chromatin extracted from tissues such as rat liver that are relatively rich in nonhistone proteins. In contrast, in nucleosomes that are very low in nonhistone protein such as in chicken erythrocytes and calf thymus, the intensity of this band is essentially similar to that of DNA. HI is not responsible, since its presence or addition has no influence on the intensity of this band. Further evidence is the low intensity of the 1580-cm- 1 band that is observed with calf-thymus nucleosomes that have no detectable nonhistone protein, and show no significant reduction in the intensity of the 1490-cm- 1 band (Goodwin et aI., 1979). Another approach is to use histone fragments. The N-terminal fragment of H4 (peptide 1-53) and the globular-portion fragment (peptide 54-102) were isolated and their interactions with DNA were investigated by Raman spectroscopy. It was concluded that the globular part of histones is the important site for structural interactions with DNA. The N-terminal fragment probably assists to find the right location on the DNA so that the histone can attach in a precise manner. The Raman spectra of reconstituted and native nucleosomes show a decrease in the intensity of the 1578-cm - I band, which is the vibrational mode involving the N-3 of adenine. There is no intensity change in the 1490-cm- 1 band, which originates from the vibration involving the N-7 of guanine. This means that the site of interaction for the N-terminal fragment involves the N-3 of adenine, which is exposed in the minor groove, whereas the large groove remains free (Couppez et aI., 1980). DNA-base recognition is not random. The lysine residue tends to interact with the G-C bases and the arginine residue with the A-T bases (Laigle et eI., 1981). Nucleosome conformation is believed to be flexible and therefore can be altered in active transcription. In order to understand the flexible nature of nucleosomes, the effect of pH and organic solvents on the conformation was investigate.d by Raman spectroscopy. The nucleosome conformation was found to be stable over a pH range of 5-8. The conformation can also be influenced by organic solvents, and the changes are irreversible. The solvents affect DNA conformation much more than the a-helical structure of histone (Zama et aI., 1978). Since Raman spectroscopy can detect different conformations of DNA, one can detect the conformation of DNA in nucleosomes directly. The characteristic lines for specific DNA conformation are summarized here: Type A TypeB TypeC
807 cm- I 835 cm-t, 787 cm- I 865-870 cm- I j
.
References
185
The Raman spectrum of mononucleosomes from calf thymus shows the band at 835-840 em-I, indicating the B form DNA (Goodwin and Brahms, 1978). From IR study, it is established that the DNA in native rat-liver chromatin and nucleosomes remains in the B conformation at low relative humidity (Liquier et aI., 1979). The deoxyribose in chromatin DNA has a C-3'-endo puckering conformation (Brahms et aI., 1981).
REFERENCES Brahms, S., Brahmachari, S. K., Angelier, N., and Brahms, J. G. (1981). Conformation of DNA in chromatin reconstituted from poly[d(A-T)] and the core histones. Nucleic Acid Res. 9, 4879. Couppez, M., Sautiere, P., Brahmachari, S. K., Brahms, 1., Liquier, 1., and Taillandier, E. (1980). Site and role of the N-terminal fragment of the nuclesomal core histones in their binding to deoxyribonucleic acid as determined by vibrational spectroscopy. Biochemistry 19, 3358. Crowther, R. A (1980). Structure of bacteriophage Pfl. Nature 286, 440. Dunker, A K., Williams, R. W., and Peticolas, W. L. (1979). Ultraviolet and laser Raman investigation of the buried tyrosines in fd phage. J. BioI. Chem. 254,6444. Fish, S. R., Hartman, K. A, Fuller, M. T., King, 1., and Thomas, G. J. (1980). Investigation of secondary structures and macromolecular interactions in bacteriophage P22 by laser Raman spectroscopy. Biophys. Discuss. 32, 234. Fodor, S. P. A, Dunker, A K., Ng, Y C, Carsten, D., and Williams, R. W. (1981). Lipid-tail group dependent structure of the fd gene 8 protein. Prog. Clin. BioI. Res. 64, 441. Fox, 1. W., Lee, 1., Amorese, D., and Tu, A T. (1979). Application of Raman spectroscopy to a complex biochemical system: Tobacco mosaic virus and its components. J. Appl. Biochem. 1, 336. Goodwin, D. C, and Brahms, 1. (1978). Form of DNA and the nature of interactions with protein in chromatin. Nucleic Acids Res. 5, 835. Goodwin, D. C, Vergne, 1., Brahms, J., Defer, N., and Kruh, 1. (1979). Nucleosome structure: Sites of interaction of proteins in the DNA grooves as determined by Raman scattering. Biochemistry 18, 2057. Guillot, J. G., Pezolet, M., and Pallotta, D. (1977). Laser Raman spectra of calf thymus histones HI, H2A, and H2B. Biochim. Biophys. Acta 491,423. Hartman, K. A, Clayton, N., and Thomas, Jr., G. J. (1973). Studies of virus structure by Raman spectroscopy. 1. R17 virus and RI7 RNA Biochem. Biophys. Res. Commun. 50, 942. Hartman, K. A, McDonald-Ordzie, P. E., Kaper, 1. M., Prescott, B., and Thomas, Jr., G. J. (1978). Studies of virus structure by laser-Raman spectroscopy. Turnip yellow mosaic virus and capsids. Biochemistry 17, 2118. Laigle, A, Chinsky, L., Turpin, P. Y, Liquier, J., and Taillandier, E. (1981). Specificity of DNA-peptides and DNA-histones interactions by ultraviolet resonance Raman spectroscopy. Biochimie 63, 831. Li, Y, Thomas, Jr., G. 1., Fuller, M., and King, J. (1981). Investigations of bacteriophage P22 by laser Raman spectroscopy. Prog. Clin. BioI. Res. 64, 271. Liquier, J., Gadenne, M. C Taillandier, E., Defer, N., Favatier, F., and Kruh, J. (1979). Conformation of DNA in chromatin protein-DNA complexes studied by infrared spectroscopy. Nucleic Acids Res. 6, 1479. Mansy, S., Engstrom, S. K., and Peticolas, W. L. (1976). Laser Raman identification of an interaction site on DNA for arginine containing histones in chromatin. Biochem. Biophys. Res. Commun.68, 1242.
186
Nucleoproteins-Virus and Chromosome
Pezolet, M., Savoie, R., Guillot, 1.-G., Pigeon-Gosselin, M., and Pallotta, D. (1980). Conformations of calf thymus and rye histones H3 and H4 in aqueous solution by laser Raman spectroscopy. Can. J. Biochem. 58, 633. Stubbs, G., Warren, S., and Holmes, K. (1977). Structure of RNA and RNA binding site in tobacco mosaic virus from 4-}\ map calculated X-ray fiber diagrams. Nature 267, 216. Thomas, Jr., G. J., and Day, L. A. (1981). Conformational transitions in Pf3 and their implications for the structure and assembly of filamentous bacterial virus. Proc. Nat. Acad. Sci. USA, 78, 2962. Thomas, Jr., G. J., and Murphy, P. (1975). Structure of coat proteins in PO fd virions by laser Raman spectroscopy. Science 188, 1205. Thomas, Jr., G. J., Prescott, B., McDonald-Ordzie, P. E., and Hartman, K. A. (1976). Studies of virus structure by laser-Raman spectroscopy. II. MS2 phage, MS2 capsids and MS2 RNA in aqueous solutions. 1. Mol. BioI. 102, 103. Thomas, Jr., G. 1., Prescott, B., and Olins, D. E. (1977). Secondary structure of histones and DNA in chromatin. Science 197, 385. Thomas, Jr., G. 1., Prescott, B., Boyle, P. D., and Day. L. A. (1981). Structural transitions in bacteriophage Pf3 and XF. Prog. Clin. BioI. Res. 64, 429. Turano, T. A., Hartman, K. A., and Thomas, Jr., G. 1. (1976). Studies of Yirus structure by laser-Raman spectroscopy. 3. Turnip yellow mosaic virus. J. Phys. Chern. SO, 1157. Zama, M., Olins, D. E., Prescott, B., and Thomas, Jr., G. J. (1978). Nucleosome conformations: pH and organic solvent effects. Nucleic Acids Res. 5, 3881.
CHAPTER
Lipids and Biological Membranes The use of Raman spectroscopy as a probe of membrane structure and function is a rapidly growing field. Membrane fluidity is largely due to rotational isomerism of fatty acid hydrocarbon chains, and this can be detected by Raman spectroscopy. This chapter first discusses some basic aspects of membranes, then a detailed description of the vibrational aspects of fatty acids and phospholipids is given. The third part of the chapter deals with interactions between lipids and different compounds. Finally, the various membrane structures that have been studied by Raman spectroscopy are summarized. There are brief review articles on Raman spectroscopy and membranes in French (Pezolet and Duchesneau, 1977) and Japanese (Koyama and Kyogoku, 1977; Harada, 1980). Comprehensive review articles on membranes have been written by Wallach et al. (1979) and Lord and Mendelsohn (1981). 1.
BRIEF REVIEW OF LIPIDS AND MEMBRANES
Fatty acids, lipids, and phospholipids contain both hydrophobic and hydrophilic moieties within a single molecule (Figure 7.1). Such compounds are
188
Lipids and Biological Membranes
called amphipathic molecules. In polar solvents, such as water, these amphipathic molecules form a variety of structures. It is important to understand the various structures formed in aqueous solution, so they are briefly described before discussion of the Raman spectroscopic properties of lipids and membranes. Lipids in water can aggregate to form monolayer, bilayer, multilayer, micelle, and emulsion forms, which are shown in Figure 7.2. In biological membranes, the structure of a bilayer is much more complicated because it contains not only phospholipids but also proteins, glycoproteins, carotenoids, and other molecules. Basically biological membranes are a modification of the bilayer structure (Figure 7.3). A biological membrane separates a cell from the outside, or it separates cell components from the surrounding medium, effectively compartmentalizing them. Yet biological membranes neither are static structures nor are they functionally inert. Membranes have a high degree of lateral fluidity and actively participate in various biological functions. There are a variety of biological membranes. The membrane of a sectioned capillary endothelial cell is shown in Figure 7.4. Liposomes are frequently used as a model for biological membranes. Liposomes are artificially produced roughly spherical particles enclosed by a lipid bilayer (Figure 7.2B). There are several ways to make liposomes by suspending phospholipids in water. Liposomes are also called lipid vesicles; they have an inner compartment that contains water. Many ions and hydrophilic compounds such as protein can be trapped inside the liposome compartment.
Structure
Symbol
Fatty Acids .¥
Hydrophl1ic moiety
r····-···,)"r -.- -... -----..-....-.------.... :
;
II :
/C~
~~~ii~id~·_·-·-····~·;:;~:;h::~,-:;:t~·-·· ,..._-_._--- --, -- -----_..--!':'.----------_._........ :
O~
:
IIY-...AAAA/:
iH2COC'l
,! I
0
1
~
V'
IlkV
:HCO~
!
I
A
V
V
V
V
At..
V'
0-
i :
V
A
V
A
V
l\
Hydrophi 1fe J.(yd~Phob1C
/!
;
II
mfe~
~e~
0=
··0-·····-···---·-·················1
:
+:
II
i H2C-0-P-0-CH2CH2NICH313 ! c •• -•••.
1...
Q=..... ..._.....__...._.....1 ~
Hydropl1111c
motety
FIGURE 7.1. Hydrophobic and hydrophilic properties of fatty acids and phospholipids. The open circle represents the head group, a relatively hydrophilic portion of a molecule, while the tail represents a hydrophobic portion of the hydrocarbon backbone.
10..
@~~~1T~
~a~
air
-.:.:.:.!k~J1:~~g~g7Y~.:.:-.
~ 0::::
?)
~
:::::0
:::::0
'@
:3:b
~J~g~%~
-·:·:-:·::·~-:·:-:·:·:·:-w~:(r:··'-:·:·:-,:-:-:":·:··
(a) Monolayer
(b) Bilayer (Liposome)
•
( d) Miceli.
(c) Multilayer
~~
I
~~~
~~~ (.) Emulsion
FIGURE 7.2. Different structures of phospholipids in water. (A) monolayer; (B) bilayer, or called liposome or vesicle; (C) multilayer; (D) micelle; (E) emulsion.
Transmembrane
Peripheral FIGURE 7.3. Schematic representation of biological membranes: Channel protein, a protein inserted across a membrane in such a way as to allow some ion to pass through; integral protein, a protein partially or wholly imbedded into the lipid bilayer; peripheral protein, a protein loosely attached to the inner or outer surface of the lipid bilayer membrane; and transmembrane protein, a protein that is inserted across the lipid bilayer membrane, but does not form a channel.
190
Lipids and Biological Membranes
FIGURE 7.4. Electron micrograph of an endothelial cell of a capillary tube from mouse. The outside membrane is the plasma membrane, consisting of a lipid bilayer. The electron micrograph was obtained by Dr. C. Ownby of the author's laboratory.
2.
VIBRATIONS OF FATTY ACIDS AND PHOSPHOLIPIDS
Our objective is to use Raman spectroscopy to study biological membranes at the molecular level without perturbing the samples. For this purpose we must know the assignment of Raman lines, especially those that are structurally sensitive or provide molecular information. Raman spectroscopy, the only probe to be discussed in this chapter, is actually only one of many physical techniques used for the study of membrane structure (Andersen, 1978). Numerous vibrational bands can be observed in IR and Raman spectra for fatty acids, lipids, and phospholipids. The frequency of these vibrations may reach as low as 100 cm - I and as high as 3000 cm - t. In earlier days, before Raman spectroscopy became more common, IR was mainly used (see Parker, 1971). The IR and Raman bands of phospholipids have very similar frequencies, as is expected (Wallach, 1972; Akutsu and Kyogoku, 1975). The most prominent use of Raman spectroscopy in the study of lipids is its application to membrane systems. However, its use is not restricted to the membrane. For instance, Raman spectroscopy can be used to differentiate positional isomers of furan-containing fatty acid esters (Jie et al., 1981). The higher-frequency vibrational modes are more complicated and are more directly correlated to structural and phase changes of lipids than low-frequency modes.
..
Vibrations of Fatty Acids and Phospholipids
2.1.
191
LOW-FREQUENCY VIBRATIONS
The low-frequency « 400 cm -I) vibrations represent acoustic modes. The motion involved in this type of mode is the symmetrical stretching of the entire molecule (Figure 7.5). Such acoustic accordionlike motion can be observed in free fatty acids or in fatty acids in phospholipids (Lippert and Peticolas, 1972; Brown et aI., 1973; Verma and Wallach, 1976; Vogel and Jahnig, 1981) (Figure 7.6). It is not easy to measure low-frequency Raman scattering because of being very close to the area of Rayleigh scattering. In aqueous solutions, there is strong Raman scattering below 200 cm- 1 with a broad background of water near 150 cm- I, and this can mask the true low-frequency bands of a compound under examination. Assignments of low-frequency Raman bands are frequently obtained from model compounds. Triclinic paraffin (n-C 24 H 50 ) shows eight bands from 15 to 102 cm - I. The observed polarization of the bands corresponds to the transverse acoustic modes of the triclinic polyethylene lattice. This suggests an appreciable mixing between the in-plane and out-of-plane vibrations (Kobayashi et aI., 1979). In crystals, a lattice vibration can also appear in the low-frequency region. Crystalline lauric acid at -160°C shows bands at 119 and 171 cm- 1 that may be due to lattice modes, because as the acid melts, these bands disappear (Mendelsohn et aI., 1975). 2.2.
STRUCTURALLY SENSITIVE RAMAN BANDS
As temperature is increased, phospholipids undergo an abrupt change in organization. The temperature that causes this change is called the transition temperature, melting temperature, thermotropic transition, or midtransition temperature and is frequently expressed as Tm (Jain, 1972). Below this temperature the phospholipids are in the gel phase. Above Tm the phospholipids are
c
+-
c
c
c
c
/\/\/\/\---+ c c c c
c
---+/ " c/ c
c
c
" c/
c
" c / , .C- .
FIGURE 7.5. Longitudinal acoustic vibrations of extended chains such as those of fatty acids. The diagram represents the symmetrical stretch of the entire molecule.
192
lipids and Biological Membranes
LAURIC ACID. ·12·C
:
E~~\.
./
STEARIC ACID. 21·C
....
.
15
•
.... .. ;
LAURIC ACID. 51· C
STEARIC ACID,
n·c
•N
FIGURE 7.6. Low-frequency vibration of hydrocarbon and fatty acids. The spectra are reproduced from Lippert and Peticolas (1971, 1972).
transformed to the liquid crystalline phase. Liquid crystalline originally referred to liquids possessing a high degree of order, which can be shown using X-ray diffraction patterns. For phospholipids the liquid crystal phase refers to the phase containing more gauche conformation. Thus by measuring the Tm one can deduce lipid fluidity and phospholipid organization within the biological membrane. The possible molecular arrangement of the phospholipid liquid crystalline phase is shown in Figure 7.7. The gel-to-liquid-crystal phase change can be detected by Raman spectroscopy. This is because the transition involves changes in the fatty acid hydrocarbon backbone (Figure 7.8). The acyl backbone consists of many C-C bonds. In the gel phase they tend to have more of the trans configuration. As the temperature increases, trans configuration
.
m:~: RIJUJ! !!I!H!I~ n nllllll
nl~Hnu
.
Gel Phase
\7n
Increase in Temperature
Liquid Crystal
FIGURE 7.7. Molecular arrangement of phospholipid gel phase and liquid crystalline phase.
Phase
Side View
Cross Sectional View
~
C I
I Trans
C) I
~
~5
~
I Gauche
FIGURE 7.8. Acyl backbone chain is flexible and can take trans or gauche conformation in
C-C-C-c.
194
Lipids and Biological Membranes
transforms to gauche configuration and changes bilayer structure and fluidity. The C-C and C-H stretching vibrations of trans and gauche conformations differ, and they appear at different frequencies in Raman spectra. 2.2.1.
C-C Stretching Vibrations
The acyl C-C backbone in phospholipids has greater mobility at temperatures above the chain-melting phase transition. The C-C and C- H stretching vibrations reflect this property.
A
A,
~
'-----.---T
J
I
,
~
on
it
iii
~ i
,
,
a
b
c
d
I
,
,
,
~
I
I
I
!§
I
I
I
I
§!
I
I
I
§
I
.-, em-I
FIGURE 7.9. Raman spectra of 1l00-cm - I region of 20% DL-dipalmitoylphosphatidylcholine sonicates in water at (A) 20°C, (B) 30°C, (C) 40°C, (D) 50°C. The figure was reproduced from the paper of Lippert and Peticolas (1971).
Vibrations of Fatty Acids and Phospholipids
195
Structurally sensitive bands can be found in the region of 1000-1140 cm - I. As the temperature is increased, the intensities of the 1066- and 1l30-cm- 1 bands decrease, whereas the intensity of the 1090-cm-] band increases (Figure 7.9). This decrease in the 1066- and 1l30-cm-] bands is attributed to a decrease in the amount of all-trans crystal structure, whereas the intensity increase in the 1090-cm- 1 band is due to the appearance of structures containing several gauche rotamers in the melted paraffin. Therefore, the intensity ratio of these bands can be used to monitor the phase transition of a mainly trans gel to a mainly gauche fluid configuration (Lippert and Peticolas, 1971; Bulkin and Krishnamachari, 1972; Spiker and Levin, 1975; Yellin and Levin, 1977a, b; Susi et al., 1980). The Raman band changes associated with structural changes that occur as temperature is increased are illustrated in Figure 7.10. However, one should be aware that the 1130 cm-] band intensity is not entirely linearly related to the concentration of gauche bonds (Snyder et al., 1982). The II OO-cm-] PO; symmetrical stretching bands are shifted to a lower frequency upon phase transition. This frequency shift is explained in terms of a
All Trans
Mixture of Trans
a
!
Gauche
~
L
Irons
+
gauche ~
J
Irons
Mostiy Gauche
FIGURE 7.10. Transition of all-trans to mainly gauche configurations. The I ]()()-cm -] band shifts to 1124 cm -I, and the intensity decreases. The intensity of the 1062-cm-] band decreases. The intensity of the 1090-cm- 1 band increases.
196
Lipids and Biological Membranes
change in the exposure of the PO; group to the solvent upon melting (Brown et al., 1973). Thus it can be concluded that the 1090-cm - I band is a superposition of the C-C stretching vibration of gauche rotamers and the O-P-O- symmetrical stretch of ionic phosphate. A theory of chain conformation as a function of temperature and rules for the assignment of Raman scattering intensity at the 1130-cm - I line were developed by Pink et al. (1980). Readers who are interested in the theoretical aspects are advised to read this article. 2.2.2.
CH Stretching Vibrations
The CH stretching frequency bands are extensively used for phase-transition studies of membranes and phospholipid bilayers. The lateral motion (the motion along the axis of fatty acid chains) is strongly restricted in the gel state, as the layer is tightly packed. As the temperature increases, the phospholipid bilayers or membranes undergo phase transition, and the lateral packing becomes loosened; this change can be reflected in the intensities of the CH stretching bands. The CH stretching vibration region (2800-3000 cm- I ) has many bands, and their exact assignment is difficult. C- H stretching is complicated by the interaction, enhanced by Fermi resonance, between C- H stretching fundamentals and overtones of HCH deformation modes (Snyder et al., 1978). Origin. The HCH deformation mode usually appears in the region of 14001500 cm-I, or about half of the C-H stretching vibration region. The C-H deformation vibrations vary considerably with chain configuration. Therefore, the C- H stretching vibration region gives very complicated vibrational bands that are composed of fundamental vibrations, overtones, and resonance interactions. The condition for such interaction is indeed very favorable. The methylene CH 2 deformation bands are at 1418 and 1436 cm- I • The symmetrical and antisymmetrical C-H stretching vibrations of solid stearic acid are at 2843 and 2873 cm- I. These values are very close to the values of 1418 X 2 and 1436 X 2. The intensities of these bands are due to Fermi resonance between the in-phase CH 2 stretching fundamental and the overtone of the CH 2 scissoring mode. Thus a small change in the frequency of the CH 2 scissoring fundamental occurs upon the formation of gauche isomers. This would remove the resonance condition, resulting in an intensity decrease in this band. The C- H stretching vibrations at 2843 and 2873 cm- I are slightly lower than those of the fundamentals, and their intensities are greater, because of Fermi resonance interaction with HCH deformation overtones. Structurally Sensitive C- H Bands. The 2890-cm- I Raman band originates from C-H stretching modes, and the intensity of this band decreases at higher temperature as intramolecular chain disorder (trans-gauche isomeriza-
Vibrations of Fatty Acids and Phospholipids
197
tion) leads to a loss of chain symmetry (Bunow and Levin, 1977a). On the other hand, the 2850-cm -1 Raman band, which is the C- H symmetrical stretching mode, is relatively constant in intensity under different temperature conditions. The intensity ratio of 2890- and 2850-cm - 1 bands is frequently used to monitor the transition temperature (melting temperature) of lipids and phospholipids (Brown et al., 1973; Larsson, 1973; Mendelsohn, 1973; Faiman and Long, 1975)(Figure 7.11). Actually the C- H stretching vibration region is more sensitive to structural changes than the C- C stretching vibration region (1066 and 1130 cm- ]). Both methods give the same information and reveal whether phospholipids are in the gel or liquid crystalline form. The ratio 12890/12850 is also a measure of lateral packing (Gaber and Peticolas, 1977; Snyder et al., 1980). The 2890-cm- 1 band is due to Fermi resonance between C- H units on adjacent hydrocarbon chains. Thus the 2890-cm-] intensity is reduced when lateral packing between chains is disrupted. Although the 12890/12850 method is commonly used to determine the phase of phospholipids, it has been proposed that use of CD2 stretching vibration line widths of the deuterated molecule at 2103 cm-] is more sensitive and accurate. The 2890- and 2850-cm-] bands are C- H stretching vibration modes, and it is argued that bands at these wavelengths include significant contributions from nonlipid components (Mendelsohn et al., 1976a). A similar opinion was expressed by Forrest (1978), who felt that protein C-H stretching may obscure the lipid C- H bands; thus relying on C- D vibration measurement is better. Three Raman bands due to C-H stretching vibrations of the terminal CH 3 group are found around 2965, 2934, and 2873 cm-] regardless of the chain
1.5
1289°1.0 1 2850
0.5
o
50
°c
100
FIGURE 7.11. Ratio of 2890-cm- I band to-2850 cm - I band in phosphatidylethanolamine as a function of temperature. The figure was reproduced from Brown et aI. (1973).
198
Lipids and Biological Membranes
length (Sunder et al., 1976a, b; Okabayashi and Kitagawa, 1978). The 2964em - I band is an asymmetrical vibration that occurs in the skeletal plane. The 2952-cm- I band was identified to be the out-of-plane asymmetrical C- H stretching vibration (Wallach et al., 1979). Detailed Assignment From Deuterated Study. The deuterated hydrocarbon chain is able to provide more-precise information about the nature of the CH 2 vibration. There are many C- H stretching vibration bands clustered in the 2800-3000-cm -I region; thus it is difficult to assign one band to one particular C- H stretching vibration. Exchange of hydrogen with deuterium at a known position within the molecule provides accurate assignments, since this C-H line shifts to a lower frequency. Moreover, C-D stretching modes occur in the region of 2100 em -1 instead of 2900-3000 em-I. There is little overlap in this region by Raman bands from other membrane components (Bunowand Levin, 1977c; Ljusberg-Wahren and Larsson, 1981; Sunder et al., 1981). From the study of deuterated fatty acids, it has been found that not all -CH 2 - residues are equivalent; especially the CH 2 residue adjacent to the CH 3 terminus, the one adjacent to the double bond -CH=CH-, and the CH 2 adjacent to the carboxyl residue (Verma and Wallach, 1977a, b). The vibration of the CH 2 group adjacent to the carboxyl residue is known to appear at 2920 em-I. Similarly, Bansil et al. (1980) concluded that· the frequency of the CD 2 stretching modes depends on the position of the CD2 group and is strongly influenced by the charge distribution of the polar carboxyl group. Instead of the 12890/12850 ratio, the variation in the line width of the C-D stretching vibration near 2100 em -1 can be used to monitor the phase transition of phospholipids (Mendelsohn et al., 1976a, b; Mendelsohn and Taraschi, 1978; Sunder and Bernstein, 1978; Mendelsohn and Koch, 1980; Bansil et al., 1980). Mendelsohn and Maisano (1977) studied the Raman spectra of binary mixtures of deuterated dimyristoylphosphatidylcholine (deuterated in the fatty acid portion) and nondeuterated distearoylphosphatidylcholine. Such an experiment 'directly monitors the conformation and phase behavior of each component (Gaber et al., 1978c). The results show two distinct transitions: a lower transition occurring at 22°C due to the melting of the shorter-chain fatty acids, and a higher transition at 47°C resulting from the melting of the longer chains. Weidekamm et al. (1978) used w-deuterated dipalmitoylphosphatidylcholine liposomes to study the detailed mechanism of valinomycin-induced phase transition. According to this study, trans-to-gauche hydrocarbon-backbone conformation changes mainly originated from CH 2 rather than terminal -CH 3 groups. .J
Vibrations of Fatty Acids and Phospholipids
199
TABLE 7.1. Assignments of CH Vibrations of Fatty Acids, Lipids, and Phospholipids
Frequency (em-I)
1400-1500 2850 2890 2920 2928 2945 2952 2964
Assignment Deformation Symmetrical CH stretching vibration of methylene (-CH 2 - ) Asymmetrical CH stretching vibration of methylene (-CH 2 - ) Stretching vibration of CH 2 residue adjacent to the carboxyl residue Symmetrical CH stretching vibration of CH 3 Symmetrical CH stretching vibration of CH 3 Out-of-plane asymmetrical CH stretching vibration of CH 3 In-plane asymmetrical CH stretching vibration of CH 3
Summary of C-H Vibrations
1. 2.
3.
2.3.
The deformation vibration appears in the region of 1400-1500 em-I. The stretching vibration has complicated band patterns and is in the region of 2800-3000 em-I. Many fine small bands arise from Fermi resonance between the CH 2 fundamental stretching vibrations and overtones of the CH 2 deformation vibration. The ratio 12890/12850 is useful as an indicator of lipid conformation and lateral packing. The different C- H vibrational modes are summarized in Table 7.1. UNSATURATED FATTY ACIDS
Unsaturated fatty acids contain C=C in their hydrocarbon chains. The frequencies of C=C stretching vibrations are different for the cis and trans isomers (Figure 7.12). The bands from cis isomers have lower frequencies than the bands from trans isomers (Table 7.2). Therefore, judging from the position
C
'C=C
/
/
'c
C, /
/C
,
C=C cis
trans c-c stretching vibration v· 1670-1680
FIGURE 7.12.
c=c
stretching vibration V='
1650-1665
C=C stretching vibrations of cis and trans isomers.
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Lipids and Biological Membranes
TABLE 7.2.
Raman Frequencies of Some Unsaturated Fatty Acids
C=C Stretch Compound Methyllaurate Methyloleate Triolein Methylelaidate Trielaidin Methyllinoleate Trilinolein Methyllinoelaidate Trilinoelaidin Methyllinolenate Trilinolenin
Class
(cm;;;~)
saturated cis cis trans trans cis, cis cis, cis trans, trans trans, trans cis, cis, cis cis, cis, cis
1655.1 1653.8 1669 1668.5 1656.1 1656.4 1671.5 1670.5 1657 1656
of the C=C stretching vibration, one can readily distinguish the cis and trans isomers. By measuring the intensities of C=C stretching fundamentals near 1656 and 1670 cm - I, Bailey and Horvat (1972) analyzed the amount of cis and trans isomers of edible vegetable oil. The 1650-1670 cm - 1 band is very sharp and unequivocal in the case of fatty acids and phospholipids, but it is often obscured by the protein amide I band in this region in membranes (Milanovich et al., 1976a). The degree of unsaturation can be measured from the ratio 11303/11267' The 1303-cm- I band is assigned to an in-phase CH 2 twisting motion, whereas the 1267-cm -1 is attributed to a =CH in-plane deformation. As the degree of unsaturation in
2
11~0~ 11267 1
0'
o
I
I
5
10
Ratio (CH2/=C-HI
FIGURE 7.13. Relation between the Raman intensity ratio [1303//1267 cm - I and the ratio of methylene to vinyl groups for a series of fatty acids and fatty acid methyl esters of 18, 20, and 22 carbons with one to four cis double bonds.
Vibrations of Fatty Acids and Phospholipids
H
H
H, / C.....
\~c
....C \
/
C....
C=C
/ H
\
....H
H
H,.....C/ C.....
\
/
H
201
H _
\..,.-H C.... /
....C
C-C
\H
H
A
B
FIGURE 7.14. Stable conformations around a pair of sp2, C-C axes; (A) skew, skew, and (B) skew, skew'. The figure was redrawn based on the figure of Koyama and Ikeda (1980).
the chain increases, the Raman intensity at 1267 cm - I increases, whereas that at 1303 cm-] decreases. As can be seen in Figure 7.13, the ratio of the peak intensities at 1303 cm-] to the peak intensities at 1267 cm - I varies linearly with the ratio of the number of CH 2 to =CH groups. Thus the comparison of the peak intensities at 1303 and 1267 cm - I may be useful as a means of estimating the degree of unsaturation of fatty acids in lipids and membranes (Butler et aI., 1979). There are detailed assignments of Raman bands of cis unsaturated fatty acids by Koyama and Ikeda (1980). They also concluded that C=C stretching at 1650 cm-] and C-H deformation at 1270 cm-] provide a good probe for the conformation around the Sp2, C-C axis. The frequencies are high for the skew-skew' conformation and low for the skew-skew conformation (Figure 7.14).
2.4. 2.4.1.
PHASE TRANSITION (MELTING BEHAVIOR) Hydrocarbon Backbone
Raman spectroscopy is a good tool to monitor phase transition of lipids and phospholipids, as it can follow the change from trans to gauche conformations of hydrocarbon backbones by following the acyl group, either the band of C-C stretching or the band of C-H stretching vibrations. Since both the 1130- and 2890-cm-] bands are sensitive to structural changes, it is possible to follow the endothermic melting behavior of phospholipids going from the gel to the liquid crystal phase. Dispersions of dipalmitoylphosphatidylcholine (DPPC) examined in these regions are found to have two melting phenomena. The pretransition temperature is 34.2°C, and the main transition occurs at 37°C. These values agree very well with the data obtained from calorimetric and fluorescence methods (Table 7.3). Prior to the transition from the gel to the liquid crystal phase of DPPC, there is a noncooperative formation of four to five gauche rotamers per chain (Mendelsohn and Taraschi, 1978).
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Lipids and Biological Membranes
TABLE 7.3. Transition Temperature for Dispersions and Vesicles of Dlpalmltoylphosphatidylcholine
Dispersion Pretransition Main transition Vesicles Source:
Raman CC)
Calorimetry (0C)
Fluorescence (0C)
34.2 41.5 37
35.4 41.2 37
25.2-33.9 41.1 37
The table was reproduced from the paper of Gaber and Peticolas (1977).
The premelting is characterized by a pronounced change in the lateral interaction between the hydrocarbon chains, whereas the melting is marked by a substantial increase in the number of gauche rotations about the C-C bonds (Gaber and Peticolas, 1977; Gaber at al., 1978a). Deuterated fatty acyl chains are very useful for the study of phospholipid or membrane structure. Deuteration of the acyl chains of the lipid molecule does not change the phase behavior of the phospholipid dispersion other than by lowering the temperatures of the premelting and melting transitions by 5°C (Sunder et al., 1978). In multilayers of dimyristoylphosphatidylcholine and distearoylphosphatidylcholine, there are two transitions. Using the deuterated myristroyl moiety, it is found that the first transition is mainly due to the melting of the shorter chain component (Mendelsohn and Maisano, 1977). Fatty acyl chains attached to positions 1 and 2 on the glycerol backbone are not structurally equivalent. Gaber et al. (1978c) synthesized two types of phospholipids; one contained deuterated fatty acid at position I and the other at position 2. Raman difference spectra taken between samples of the two compounds under identical conditions showed significant features. Below the pretransition temperature, the conformation of chain 2 is slightly less all-trans than is the chain at position 1. There is also some evidence that the conformations of the terminal methyl groups of the two chains are significantly different at low temperature. The results obtained from Raman spectroscopy confirm the results from NMR. Seelig and Seelig (1975) proposed that fatty acyl chain 1 is relatively perpendicular to the plane of the bilayer, but that chain 2 exists in either of two conformations of equal stability in which the first two carbons near the glycerol backbone are nearly parallel to the plane of the bilayer. By using the ratio 11130//1089' it was concluded that the hydrocarbon chains in dipalmitoylethanolamine are in a predominantly trans conformation (crystalline), whereas those in egg phosphatidylcholine are in a liquid state (Mendelsohn, 1972). Phospholipid hydrocarbon backbones can also be perturbed by organic solvents. By monitoring the 11130//1089 ratio, Szalontai (1976) found that benzene promotes a gel-to-liquid-crystalline transition. The
Vibrations of Fatty Acids and Phospholipids
203
transition temperatures detected by this method for dipalmitoylphosphatidylcholine and dimyristoylphosphatidylcholine multilayers are 39 ± 1°C and 23 ± 1°C respectively (Spiker and Levin, 1976a, b). Using a similar method, Mendelsohn et al. (1975) found that below Tm the hydrocarbon chains of 1,2-dilauroylphosphatidylethanolamine are ordered in the solid phase. The spectra of spingomyelin show significant formation of gauche isomers below Tm , indicating a less-rigid environment for this molecule in the solid state. When lipids are in the liquid state, the symmetrical-stretching vibrations of the fatty acid CH 2 groups at about 2850 cm -I dominate. When lipids are in a solid state, with close packing of planar zigzag chains, the intensity of CH 3 stretching vibrations at 2885 cm - I dominates. Therefore, if the lipid is melted or dissolved in an organic solvent, the peak near 2850 cm - 1 increases in intensity. At a given temperature, whether gauche or trans conformations predominate depends on the particular lipid. Sphingomyelin has a considerable fraction of gauche configuration below the melting point, whereas dilauroylphosphatidylethanolamine has little or no gauche conformation in the solid phase (Menddsohn et aI., 1975; Sunder et aI., 1976a, b). Sonication of phospholipid dispersions is widely used to transform large coarse aggregates to smaller particles. In order to see whether sonication has any effect on hydrocarbon chains of egg phosphatidylcholine and dipalmitoylphosphatidylcholine, Raman spectra were examined before and after sonication. Analysis of Raman spectra in the C-C (1000-1300 cm- I ) and C-H (2800-3000 cm- I ) stretching vibration regions reveals that there is no change in the relative population of trans and gauche isomers (Mendelsohn et aI., 1976b; Gaber and Peticolas, 1977). Using the ratios 12890/12850 and 11130//1100' one can follow the phase change of phospholipids. The transition temperature Tm for dimyristoylphosphatidylcholine and dipalmitoylphosphatidylcholine are 23.5 and 42°C, respectively. For the mixture of these two compounds, the transition curve is broadened, and Tm is at 33.5°C, which is about the intermediate value of the individual components (Knoll, 1978). Similarly, the phase-change transition temperature for dipalmitoylphosphatidic acid was found by this method to be 57°C, which is in good agreement with the value derived from polarized fluorescence measurements (Hartmann and Galla, 1978). For sphingomyelin, Faiman (1979) showed that the ratios of peak intensities at 1063/1128, 1087/1128, and 1105/1128 cm- J all are sensitive to the phase change. For Raman spectra analysis, band assignment is very important; the assignments for various lipids and phospholipids are summarized in Table 7.4. 2.4.2.
.
Glycerol Moiety
Deuteration of the alkyl chains reveals the C- H vibrations of the head group (glycerol moiety). An observation of change in these bands at the melting temperature indicates that there is an alteration of the glycerol-backbone conformation upon melting (Gaber et aI., 1978b).
TABLE 7.4.
Frequency (em-I)
-218 } 224 720 725 860 - 890 870 872 875 960 967 1065 1070} 1075 1080 1085 1092 1097 1101 1125 1130 1230 1200-1350 1298 1299 1300 1340 1370 1421 1440 1440 1445 1446 1457 1460 1470 1471 1658 1731 1732
Assignment of Raman Frequency for Lipids and Phospholipids
Compound a
Assignment h
Reference
DLPE
Longitudinal acoustical Mendelsohn et al. (1975) mode Head group def. Mendelsohn (1972) PC C-N symmetric stretching Akutsu (1981) PE O-P-O sym. str. Akutsu and Kyogoku (1975) PE C-C str. involving acyl carbon Bicknell-Brown et al. (1981) C-N symmetric stretching Akutsu (1981) PC Mendelsohn (1972) CH 2 def. PC C-C-N+ sym. str. PE Akutsu and Kyogoku (1975) Sphingomyelin C-C stretch Verma et al. (1975) PC =CH def. Mendelsohn (1972) (out of plane) C-C SIr. DLPE Mendelsohn et al. (1975) Sphingomyelin C-C and C-N str. Verma et al. (1975) PE
C-O and C-O-C sym. str.
Akutsu and Kyogoku (1975)
DLPE PC Sphingomyelin PE DLPE PE DLPE PE Tripalmitin PC DLPE Sphingomyelin Endoplasmic reticulum PC DLPE Sphingomyelin PE PC DLPE DLPE Sphingomyelin PE DLPE PC PC DLPE
C-C str. C-C str. C-C and C-N str. PO; sym. str. O-P-O sym. str. C-C str. C-C str. PO; asym. str. CH 2 wag. CH 2 twist CH 2 twist CH 2 rocking CH def.
Mendelsohn et al. (1975) Mendelsohn (1972) Verma et al. (1975) Akutsu and Kyogoku (1975) Mendelsohn et al. (1975) Akutsu and Kyogoku (f975) Mendelsohn et al. (1975) Akutsu and Kyogoku (1975) Larsson (1973) Mendelsohn (1972) Mendelsohn et al. (1975) Verma et al. (1975) Verma et aI. (1975)
CH 2 wag. CH 2 bend. CH 2 and CH 3 def. CH 2 sci. CH 2 def. CH 2 bend. CH 2 bend. CH 2 and CH 3 def. CH 2 sci. CH 2 bend. C-C SIr. Ester C-O str. C=O str.
Mendelsohn (1972) Mendelsohn et al. (1975) Verma et al. (1975) Akutsu and Kyogoku (1975) Mendelsohn (1972) Mendelsohn et al. (1975) Mendelsohn et aI. (1975) Verma et aI. (1975) Akutsu and Kyogoku (1975) Mendelsohn et al. (1975) Mendelsohn (1972) Mendelsohn (1972) Mendelsohn et aI. (1975)
'., '\
...
Vibrations of Fatty Acids and Phospholipids
TABLE 7.4.
Continued
Frequency (em-I) Compound U 2847 2848 2850
2855 2880 2882 2883 2885 2890 2890 2960 2930 2937 2940
2960 2965 2967
205
DPPC Stearic acid DLPE Tripalmitin PC Sphingomyelin PE Tripalmitin DPPC Stearic acid DLPE PC Sphingomyelin PE PC Stearic acid Endoplasmic reticulum, sphingomyelin PE PC DLPE Stearic acid
Assignment b
Reference
Sym. CH 2 str. Sym. CH 2 str. Sym. CH 2 str. Sym. CH 2 str. Sym. CH 2 str. Sym. CH 2 str. Sym. CH 2 str. Sym. CH 3 Asy;n. CH 2 str. Asym. CH 2 str. Asym. CH 2 str. Asym. CH 2 str. Sym. (CH 2 and CH 3 ) str. Sym. NHj deg. str., Sym. CH 3 str. Sym. CH 3 str. Sym. CH 3 str. Asym. CH 2 str.
Spiker et aI. (1976) Sunder et al. ( 1976a. b) Mendelsohn et al. (1975) Larsson (1973) Spiker and Levin (1976a) Verma et al. (1975) Akutsu and Kyogoku (1975) Larsson (1973) Spiker et al. (1976) Sunder et al. (1976a, b) Mendelsohn et al. (1975) Spiker and Levin (1976a) Verma et al. (1975) Akutsu et al. (1975)
Sym. CH 3 str. Asym. CH 3 str. Asym. CH 3 str. Asym. CH 3 str.
Akutsu and Kyogoku (1975) Spiker and Levin (I 976a) Mendelsohn et aI. (1975) Sunder et aI. ( I 976a, b)
Spiker and Levin (1976a) Sunder et aI. (I 976a, b) Verma et aI. (1975)
UDLPE, dilauroylphosphatidylethanolamine: pc, phosphalidylcholine: PE, phosphatidylethanolamine: DPPC, dipalmitoylphosphalidylcholine. hAbbreviations: bend, bending: deL, deformation: deg.. degenerate: sci., scissoring: str., stretch: sym.. symmetrical: wag., wagging: asym.. asymmetrical.
2.4.3.
Cerebrosides
Cerebrosides are glycosphingolipids having the general structure shown in Figure 7.15. Kerasin contains lignoceric acid (C 24 , saturated), whereas phrenosine contains cerebronic acid (2-hydroxy derivative of lignoceric acid). When the degree of lipid disorder increases (or the temperature increases), the intensity of the scattering at 1090 cm- I increases, whereas at 1060 and 1130 cm- 1 it decreases. Thus the peak ratios 11090//1060 or 11090/11130 express the gauche/trans conformers (conformation isomers) ratios. When these ratios were measured, kerasin was found to have lower values than phrenosine, indicating that kerasin has less gauche content than phrenosine. When the enthalpy of the gel-to-liquid-crystalline phase transition was measured by
206
Lipids and Biological Membranes
fatty aci d CH20H
O
HO
OH
NHCOR O-CH2-t-CH-CH=CH-ICH21 - CH3 I I 12 H OH
,
I
OH
I
sphingosine
Galactose FIGURE 7.15.
Structure of cerebrosides.
calorimetric methods, kerasin was found to have ~H value of 15.8 kcal/mol compared with 7.0 kcal/mol for phrenosine. Thus both Raman spectroscopic and calorimetric methods show that kerasin has a higher degree of order than phrenosine in the hydrocarbon chains. The high transition enthalpy for kerasin is thus ascribed to a lesser accommodation of gauche conformers just below the transition temperature (Bunow, 1979). Occasionally two carbonyl stretching vibration bands appear near 1700 cm - I for anhydrous crystalline phospholipids. This is due to different configurations at the two acyl linkages. Monitoring of the Pc=o may serve as a valuable means to analyze the conformation about the acyl linkages in phospholipids (Bicknell-Brown et aI., 1980).
2.5.
QUANTITATIVE ESTIMATION OF CONFORMATION
Using the intensity of the 1BO-cm - I band, one can measure the amount of the trans configuration. Such an equation has been developed by Gaber and Peticolas (1977) by assuming that the intensity contribution per trans bond in a segment is constant.
s trans
=
(1 1133 / I ref )observed
(1 1133 / I ref )solid DPPC
DPPC refers to dipalmitoylphosphatidylcholine. S is the order parameter of the trans configuration; S = 1 indicates the highest possible order and S = 0, no order. However, as Cornell et al. (1978) pointed out, there is no simple relationship between Strans and the number of trans bonds in a lipid chain. The effect of various anions and cations on the structure of the DPPC multibilayer system was investigated (KarvalY and Loshchilova, 1977; Loshchilova and Karvaly, 1978). It was found that the different trans order parameters are sensitive to ion-polar-head-group interaction, and thus they cannot give unequivocal information on the trans-gauche isomerization of hydrocarbon chains of phospholipids. As mentioned previously, the symmetrical methylene C-H stretching modes, /2890//2850' is sensitive to the packing order of the acyl
Interaction
0' lipids
207
chains in the liquid crystalline state, decreasing in the following order: lamellar liquid> hexagonal or cubic liquid crystal> micellar solution> solution in organic solvent (Larsson and Rand, 1973). Based on this principle, a semiquantitative estimate of the lateral crystal-like order between the chains was eventually developed by Gaber and Peticolas (1977). The order parameter for the lateral interaction Slat is expressed as
_ 1CH,(sample) S lat
-
-
0.7
1.5
where 1 CH , = 12890/12850' So Slat""= 1 when a phospholipid is in the crystal state (ICH, = 2.2), and Slat = 0 in the liquid state (ICH, = 0.7). As pointed out by Gaber and Peticolas, this parameter is only approximate and must be considered semiquantitative. In contrast to the restricted applicability of Strans, the lateral parameter was proved to be suitable for quantitative studies even in the case of ion-head-group interaction (Karvaly and Loshchilova, 1977). Using this equation, the degree of order in lipid multilayer assemblies was examined (Lis et aI., 1977). Fatty acid multilayers are more ordered than phospholipid samples but less ordered than crystalline samples. This suggests that there is a high degree of acyl-chain order in the multilayers of fatty acids compared with crystalline hexadecane. Electron spin resonance is an extensively used technique in membranestructure studies. The spin-labeling method can give much information. Order parameters obtained by Raman spectroscopy and by the spin-label method were correlated by Horvath et aI. (1980).
3.
INTERACTION OF LIPIDS
Phospholipids are major components of biological membranes; however, many other compounds are also constituents of these membranes. How these components interact with each other within a membrane is a matter of great interest. Raman spectroscopy is one way to look at this problem by observing C-C and C-H stretch regions. An interaction of one component with phospholipids in the lipid matrix can be reflected in certain Raman lines. 3.1. 3.1.3.
LIPID-LIPID INTERACTION Intramolecular Interaction
Gel formation from liquid crystalline lipid is an example of lipid-lipid interaction. As the lipid matrix solidifies from its fluid, specific lipid conformational-change and packing rearrangement take place. As temperature is
208
Lipids and Biological Membranes
increased, there is an increase in lateral-chain-packing disorder. The lateralpacking (intermolecular) and trans-gauche (intramolecular) disordering processes can be monitored by observing the C-C (1000-1190 cm- I ) and C- H (2800-3000 cm -I) stretching regions. Tg denotes the transition temperature for trans-gauche isomerization in the gel. When the Tg was examined for C 14 ' C 16 ' and C I8 fatty acids in phosphatidylcholine lipid bilayers, it was found to be -40, -40, and 5°C, respectively. The higher Tg value for distearoylphosphatidylcholine is associated with increasing interactions between the terminal-chain areas of the individual monolayers forming the bilayer unit (Yellin and Levin, 1977a). 3.1.2.
Effect of Cholesterol
The liquid-crystalline-to-gel phase transItIOn of phospholjpids in aqueous medium can be detected by different physical techniques, including Raman spectroscopy. The transition is influenced by many factors such as the length and degree of unsaturation of the hydrocarbon chains of phospholipids. Addition of cholesterol to the membrane has many effects, including a reduction of membrane permeability, a disturbance of hydrocarbon-chain packing, a change in lateral diffusion rates for both lipids and proteins, and an alteration in the gel-to-liquid-crystal phase transition (Spiker and Levin, 1976b). Phospholipids in micelles can also be considered as another model for lipid-lipid interactions. In micelles there is substantial disorder in the hydrocarbon chain conformations (Burns et aI., 1982). Dipalmitoylphosphatidylcholine-cholesterol mixtures show more fluid properties than the pure lipid at a temperature below the transition temperature. However, above this temperature, the mixture shows more rigidity (Lippert and Peticolas, 1971). In other words, cholesterol increases the phospholipid fluidity in the gel phase and decreases it in the liquid crystalline phase. The reason for the marked increase in the chain rigidity of fatty acid is that the addition of cholesterol inhibits the formation of certain gauche isomers (Mendelsohn, 1972). However, in the case of egg phosphatidylcholine, cholesterol restricts the mobility of the fatty acid chain in the bilayer both above and below the chain transition temperature (Verma et aI., 1975). The effect of cholesterol addition to phospholipids is not as simple as had been thought. It depends on the chain length of the fatty acids as well as the degree of saturation. Using 12890/12850 as a measure of conformational change, researchers have found that the incorporation of cholesterol into the lipid bilayer exerts a greater effect on hydrocarbon-chain mobility in the case of the saturated chains than unsaturated (Faiman et al., 1976; Pink et al., 1981). Cholesterol affects the acyl region more than the head group. Evidence for this is obtained form the C=O stretching band, which shifts to 1720 em-I from 1737 em-I, whereas the C-N+ (CH 3 )3 stretching vibration at 718 cm- I remains the same after the addition of cholesterol to a phosphatidylcholine bilayer interface (Brown and Bicknell-Brown, 1980).
Interaction of Lipids
3.2.
209
LIPID-PROTEIN INTERACTIONS
Biological membranes are composed mainly of lipids and proteins. The ratio of lipids to proteins in membranes varies, depending on their source. Less metabolically active membranes usually contain less proteins. For instance, myelin has a relatively low protein content (18%). Metabolically active membrane such as the internal membrane of mitochondria may contain as much as 75% protein. Neither lipids nor proteins are inert within biological membranes but exert a remarkable influence on each other. It is also known that the activity of membrane-bound enzymes depends on the state of phospholipid structures. In order to visualize the active interaction between lipids and proteins, one should study it under well-defined conditions. Raman spectroscopy can be used in such a study because it can monitor trans-gauche transitions as well as lateral interactions of acyl chains of phospholipids. Any effect protein may exert on phospholipid structures can readily be followed by careful observation of Raman spectra. Lipid-protein interactions have been extensively studied by other techniques. For instance, spin-labeled phospholipids have frequently been used to study protein-lipid interactions. Unlike the ESR technique, Raman spectroscopy does not require chemical modification of proteins or lipids. When a protein interacts with sodium monodecylphosphate, a model compound for fatty acid, the intensity of the 2850-cm -I line increases. This can be explained by the hydrocarbon-chain -CH 2 - interaction with protein hydrophobic side chains; thus the hydrocarbon backbone has more gauche than trans conformation. By broadening the analogy, one can say that when interacting, with a protein, lipid loses more crystalline character and gains more characteristics of the liquid state (Larsson and Rand, 1973). Interaction of phospholipids with proteins is quite common. Even albumin and fibrinogen can influence the lipid hydrocarbon-chain conformation, which can be detected in Raman spectra near 2900 cm- I, a region of C- H stretching vibrations (Lis et al., I 976a). Both C-C and C- H stretching vibration regions are very useful to probe the state of lipids, as these bands reflect the hydrocarbon-chain randomness or rigidity. With the addition of arginine, histidine, and lysine, the intensity of the 2930-cm - I band increases. This can be explained by the fact that the lipid hydrocarbon-chain environment becomes more polar. The intensity ratio 11064/11089 also decreases upon the addition of amino acids, indicating an increase in the gauche hydrocarbon-chain character of phosphatidylcholine (Lis et al., 1976b). 3.2.1.
Heme Proteins
The interaction of cytochrome c and cytochrome c oxidase with lipids is well recognized. From ESR and electron-diffraction studies, it is suggested that cytochrome c interacts in an extrinsic fashion with phosr-hotidylcholine, whereas
210
Lipids and Biological Membranes
cytochrome c oxidase is an intrinsic transmembrane protein (Griffith et aI., 1973; Vanderkooi et aI., 1972). Cytochrome c decreases the ratio 12850 /1 2930 both above and below the liquid crystalline transition temperature. Cytochrome c oxidase decreases the Raman ratio 12890/12850 above Tm . This ratio remains constant below the liquid crystalline transition of the lipid. Therefore, both cytochrome c and cytochrome c oxidase affect the phospholipid hydrocarbon backbone, but their manner of interaction is different (Lis et aI., 1976a). 3.2.2.
Glucagon
The glucagon-dimyristoylphosphatidylcholine (DMPC) interaction was studied by Raman spectroscopy using the C-C and C-H vibrational regions (Taraschi and Mendelsohn, 1979). The study indicates that there is a strong interaction between the phospholipid hydrocarbon chain and the hydrophobic regions of glucagon. Thus the phospholipid forms two or three additional gauche rotamers, but these gauche isomers are immobilized by the protein. Moreover, the addition of the protein causes the loss of interchain lateral interactions. By observing the conformationally sensitive band of the C-H stretching vibration (12890/12850)' Mendelsohn et al. (1979) showed that the lateral interactions between phospholipid hydrocarbon chains are greatly disrupted in the DPPC-glucagon complex. 3.2.3.
Insulin
A DPPC-cardiolipin complex undergoes a broad phase separation and shows a lowered transition temperature relative to DPPC. One of the reasons is the destabilization of hydrocarbon chains by the presence of excess negative charge on the cardiolipin head group, which facilitates the formation of gauche rotamers. Insulin has a net positive charge of + 5, which neutralizes the negative charges on the cardiolipin. Thus insulin restores more order for the DPPC-cardiolipin system (Mendelsohn et aI., 1979). 3.2.4.
Melittin
Melittin is the primary constituent of honeybee venom. It is a basic polypeptide (26 amino acid residues) and has many biological effects. One effect is to cause the lysis of erythrocytes. It appears, therefore, that melittin has some kind of effect on biological membranes. This was carefully studied by Raman spectroscopy in a model membrane system of phosphatidylcholine liposomes (Verma and Wallach, 1976b). Particular attention was focused on the C-H stretching vibrations at 2885 and 2930 cm- I. The 2885-cm - 1 band is assigned to the asymmetrical methylene-stretching vibration, whereas the 2930-cm - I band is assigned to the symmetrical methyl C-H stretching vibration (see
Interaction of Lipids
211
Section 2.2.2 in this chapter). In the presence of melittin, the transItion temperature, as determined from 12885/12850' is shifted to much higher temperatures than in the liposomes without melittin. The shift in transition temperature as measured by 12930/12850 is smaller. From these results it can be concluded that the hydrophobic portion of melittin forms a complex with phospholipids, but the effect of the peptide moiety is greater on the methylene segments than on the methyl termini of acyl hydrocarbon chains of liposome phospholipids. As mellitin is inserted into liposome membranes, several phospholipid molecules around the mellitin actually bind to it. This can explain the two order-disorder transitions in the temperature profile for the liposome-melittin complex. The first is the main gel-to-liquid-crystalline phase transition and the second is the melting behavior of lipids bound to melittin (Lavialle et al., 1980). The effect of melittin on lipid membranes is phase dependent. Below the phase-transition temperature (gel phase) melittin decreases the conformational order, whereas above phase-transition temperatures (fluid phase) the conformational order is increased (Vogel, 1981). 3.2.5.
POly(L-Lys)
In order to follow phospholipid liposome-polypeptide interaction more quantitatively, sometimes a relative-intensity parameter 1R can be used. 1R is defined as 1 R
= (I1062/
1715 )
(11062/1715 )
Iiq N
2
At liquid-nitrogen temperature, lipids are assumed to have 100% trans conformation (Susi et al., 1979). As the temperature increases, the intensity at 1062 cm- I decreases because of an increasing amount of gauche rotamer. Poly(L-Lys) interacts with the polar groups of phospholipid liposomes without penetrating extensively into the hydrocarbon domain. The addition of poly(L-Lys) increases the transition midpoint by 4°C, suggesting that it stabilizes the phospholipid bilayer. 3.2.6.
Phage Coat Protein
The interaction of phage coat protein with DPPC was investigated by Dunker et al. (1979). B-protein is the major coat protein obtained from the filamentous phage fd. It is extremely hydrophobic and forms tight associations with lipids. The protein has a very dramatic effect on the organization of the lipid bilayer. The phase-transition temperature for pure DPPC is 41°C. However, for the equimolar mixture of B-protein and DPPC, the melting transition is lowered to 21°C.
212
Lipids and Biological Membranes
3.2.7.
Lysozyme
Interaction of lysozyme with mixed liposomes was also studied by Raman spectroscopy. Liposomes were prepared by mixing 1,2-dipalmitoyl-L-phosphatidic acid and 1,2-dimyristoyl-L-phosphatidylcholine. At temperatures below 27°C, the lipid is in the all-trans configuration, and the helical content of lysozyme increases slightly at the expense of random configuration. At temperatures above 30°C, considerable ,B-sheet is irreversibly formed. The onset of ,B-sheet formation appears to coincide with the formation of disordered lipid side chains in the acidic component of the lipid. Moreover, the 782-cm - 1 band for the O-P-O stretching mode is much more intense at all temperatures for the lipid-protein mixture than for the lipid alone. These results are interpreted to mean that lysozyme at temperatures below the order-disorder transition is essentially excluded from the hydrophobic portion of the lipid. At higher temperatures, the interaction involves the lipid side chains of dipalmitoylphosphatidic acid in the disordered state, and the lysozyme conformation changes (Lippert et al., 1980). Thus Raman spectroscopy provides evidence that the change in protein conformation is related to the change in lipid configuration in the phospholipid-protein complex. 3.2.8.
Glycoprotein, Glycosphingolipids
Glycophorin is an erythrocyte membrane glycoprotein, that contains 60% carbohydrates. The presence of even a small amount of glycophorin perturbs the phase-transition behavior of DPPC. This is interpreted to mean that DPPC in the immediate vicinity of the protein is unable to undergo a cooperative transition because of the perturbation by glycophorin. The Raman data show that these lipid molecules are in the gauche conformation (Taraschi and Mendelsohn, 1980). Glycophorin can be isolated from erythrocyte membranes and reconstituted with phospholipids into unilamellar vesicles. Apparently the mobility of the acyl chains is increased by the addition of the protein, whereas interchain lateral interactions are disrupted (Mendelsohn et al., 198 I). Glycosphingolipids can be found in the white matter of the brain. Judging by RamaJ;} spectral analysis, glycosphingolipids possess a highly ordered structure in the gel phase (Bunow and Levin, 1980). The long-chain character of the fatty acyl groups, in addition to the extensive hydrogen-bonding capacity of the galactose head group and the adjacent polar region of the molecule, may confer stability on the bilayers of these lipids. 3.2.9.
Others
Proteolipid apoprotein is the major protein found in the myelin sheath. The protein stabilizes the acyl chains of phospholipids against melting, as measured by the 12880 /1 2850 ratio (Verma and Wallach, 1978). Apoprotein of human plasma lipoproteins and DMPC form a complex. From Raman spectroscopic analysis, it has been shown that the physical state
Interaction of Lipids
213
of lipid molecules in the complex is different from that in DMPC multilamellar liposomes. It has also been shown by the large change in the I340-cm- I band for apoprotein and the complex that the protein hydrophobic side chains are immobilized by lipid binding (Gilman et al., 1981). 3.3.
LIPID-ION INTERACTIONS
It is known that biological functions of membranes are influenced by the ionic media. The effect is caused by the alteration of lipid lamellar systems by interaction of lipids and ions. Effects of various cations and anions on phospholipid layer structure were investigated by Lis et al. (1975) using the intensity ratio of 1064- to 1089-cm- 1 bands. The intensity of the lO64-cm- 1 band is a measure of all-trans C-C conformations whereas the 1089-cm- 1 band is due to a mixed mode of O-P-O stretch and gauche C-C vibrations (disordered structure of lipids). Divalent cations have more influence in decreasing the proportion of gauche character than monovalent cations, which do not affect the Raman spectra of phosphatidylcholine dispersions. Many anions such as Br- , Cl- , 1- , Cl04- , CNS- , and SOJ- do not affect the Raman intensities. However, a mixture of KI and 12 has a great effect on increasing the intensity of the 1089-cm - I band; this is indicative of random lipid chains. It was known that the conductivity of bimolecular lipid membranes drastically increased in the presence of iodine and iodide. Using the structurally sensitive bands of C-C and C-H vibrations, Loshchi10va and Karvaly (1977) studied this problem. Iodine increases the disorder of phospholipid hydrocarbons, as witnessed by the increase of ratios f 109O/ f lO66 and f 109O / f l128 and also by the increase of the C- H band region detected by the ratio f 2850 / f 2882 • Molecular iodine rather than ionic iodide interacts with phospholipids. Increase of flexible gauche conformation is apparently responsible for the change in conducting of membranes. It is known that calcium ion, but not magnesium ion, induces fusion of acidic phospholipid vesicles and phase separation in mixed vesicles. It is also known that calcium ions interact with acidic phospholipids but not with neutral phospholipids. Raman spectroscopic studies indicate that Ca(II) increases both the number of trans bonds and the lateral order while Mg(II) has much smaller effects (Hark and Ho, 1979; Susi, 1981). All these results suggest that Ca(II) induces the formation of a complex composed of almost anhydrous bilayers, whereas Mg(II) causes a milder perturbation of the bilayer structure. 3.4.
LIPID-ANTIBIOTIC INTERACTIONS
How proteins are arranged in biomembranes has been a subject of extensive investigation. Some proteins are situated on the outside of membranes, whereas others are in the interior of membranes. Some integral proteins even span the lipid bilayer.
214
Lipids and Biological Membranes
Our knowledge of the roles of proteins in biomembranes is greatly advanced by the study of membrane-bound antibiotics of peptide nature such as gramicidin A and valinomycin. Interaction of liposomes and drugs attained significant attention recently as a better system for drug delivery. It is important to administer drugs in a manner so as to localize the drugs at high concentrations for maximum drug effect. A drug encapsulated in liposomes can reach selective target tissues or organs and be diffused out slowly. The retention of the liposomes in the system depends on the charge and size of the liposome. Large liposomes are found to accumulate in organs high in reticuloendothelial cells such as the lung, spleen, and liver. Smaller liposomes have a more general distribution. Release of a drug from the liposome is highly dependent on the polarity and transition temperature of the liposome and each drug has a different effect on the polarity and the transition temperature. Only at or above this temperature, the efflux of encapsulated drugs from liposome can occur. It is thus important to determine the effect of different drugs on the phospholipid transition temperature. Some antibiotics have a great effect on the ion permeability of membranes. Some of these antibiotics are polypeptides (gramicidin S, A, B, C, valinomycin) and some are nonpeptide (amphotericin B). This action is due to the specific interaction of the antibiotic with membrane lipids, probably at the hydrophobic core of the bilayers. Raman spectroscopy has contributed greatly to the understanding of this mechanism. 3.4.1.
Gramicidin A
Gramicidin A is a linear pentadecaptide (15 residues) isolated from Bacillus brevis. Gramicidin A is believed to have a 'IT(L, D) helix, which is made possible by the existence of D and L residues. It is flexible and forms four different conformations (Iqbal and Weidekamm, 1980). This type of helix can form head-to-toe dimers by an intermolecular-hydrogen-bond pattern similar to that of the antiparallel ,B-pleated sheet. The external hydrophobic residues are in contact with the hydrocarbon domain of the phospholipids, and the peptide carbonyl groups line the central hole at the axis of the helix. When it is introduced into natural membranes or artificial lipid bilayers, the membranes become permeable to monovalent cations. Gramicidin A probably forms transmembrane channels through lipid membranes (Figure 7.16). It forms transmembrane channels by the hydrogen-bonded association of two helical molecules. Gramicidin A has an unusually high amide I frequency at 1685 cm- I in dimethylsulfoxide. In chloroform, dioxane, and in powder form, the amide I band is in the normal range of 1665 cm- I (Rothschild and Stanley, 1974). Weidekamm et al. (1977) investigated the effect of gramicidin A on DPCC liposome by measuring the transition temperature of phospholipids from the intensity ratio at 2936/2850 and 2883/2850 cm- I. As mentioned before (see Section 2.2 of this chapter), the band around 2928-2945 is attributed to the symmetrical C-H stretching vibration of CH 3 , whereas the
Interaction of Lipids
-
...".,-
.".
215
~
-----
FIGURE 7.16. Proposed mechanism of the interaction of gramicidin A with a biological membrane. Gramicidin A forms a pore that facilitates the transport of an ion across a lipid bilayer. The figure was reproduced from the paper of Rothschild and Stanley (1975).
2883-cm ~ I band originates from the antisymmetrical C- H stretching vibration of the methylene group. The transition-temperature curve obtained from the 1 2883 /12850 ratio is broadened but not the curve obtained from the 12936/12850 ratio. The 12936/12850 ratio should reflect the thermotropic lipid transitions for the CH 3 termini, in other words, the interior of the lipid bilayer. The 1 2883 /12850 ratio should reflect the transition of the CH 2 groups of the hydrocarbon chain. This suggests that gramicidin A does perturb the phospholipid bilayer membrane laterally, but the degree of perturbation is not uniform at varying depths of the bilayer. Liposomes prepared from a mixture of phosphatidylcholine and cholesterol are unaffected by gramicidin A. It was found that below the phase-transition temperature, the 1R (relative intensity parameter) value is 10 to 20% lower than for the pure phospholipid liposomes; above this temperature, it is about 20 to 25% higher. In other words, gramicidin A increases the fluidity of phospholipid liposomes in the gel phase and decreases their fluidity in the liquid crystalline phase (Susi et a1., 1979). Similarly, the antibiotics nonactin, monactin, and dinactin complex selectively with a wide variety of cations. Again the ester C=O vibrational bands change on complex formation. According to X-ray crystallographic studies, the cation is located at the center of the complex. The cation is coordinated with four ester and four ether oxygen atoms (Figure 7.17). The Raman spectroscopic results agree well with the proposed structure (Asher et a1., 1977). Raman spectra of nonactin in crystalline form and in solution are different, especially at the ester C=O (1700-1740 em-I) and C-H (2800-3000 em-I)
216
Lipids and Biological Membranes
• !0
5
6
8
---~~ ----1----- 0 -*- 0
0
1407910 ,3
*~0-r6--f° 0'. i
(0 )
FIGURE 7.17. (A) Chemical structure of nonactin. (B) X-ray crystallographic structure of the nonactin-K+ complex. The figure was reproduced from Asher et al. (b)
(1977).
stretching vibration regions. It is known that the C- H vibration mode is sensitive to crystal packing. The difference in spectra apparently reflects the effects of crystalline contact forces (Asher et aI., 1974a). 3.4.2.
Gramicidin D
Gramicidin D is a mixture of three gramicidins, A, B, and C, which are all polypeptides. All side chains of gramicidins are hydrophobic types. It is believed that gramicidins interact with hydrophobic regions of the membrane bilayer. Raman difference spectra of the phospholipid-gramicidin complex and phospholipids show two bands centered at 1090 and 1130 cm - I that exactly correspond to the C-C stretching vibrations of gauche and trans conformations. This indicates a change in the phospholipid hydrocarbon chain backbone ~ue to interaction with the antibiotics (Faiman and Long, 1976). 3.4.3.
Gramicidin S
Gramicidin S is a cyc1ic-decapeptide antibiotic also produced from Bacillus brevis. Polar groups are tied up through hydrogen bonding in the skeletal backbone of the molecule. The hydrophobic groupings are outside on the periphery. Addition of gramicidin S to membranes causes a substantial lowering of the transition temperature and a great broadening of the transition region. The relatively small and compact gramicidin S molecules are apparently embedded in the hydrophobic domain in a less orderly way. Thus they decrease the cooperativity and the melting temperature of the hydrocarbon chain.
Interaction of Lipids
3.4.4.
217
Valinomycin
Valinomycin is cyclododecadepsi-peptide antibiotic produced by Streptomyces fulvissimus. It is composed of 3 mol each of L-valine, D-a-hydroxysovaleric acid, D-valine, and L-lactic acid connected by ester and peptide bonds (Figure 7.18). It is an effective antibiotic for tuberculosis. Some cyclic-peptide antibiotics selectively bind certain types of cations. For instance, valinomycin binds to potassium ions and facilitates their transport across mitochondrial membranes. When valinomycin complexes with potassium ion, the band due to the ester C=O stretch becomes a single narrow peak at 1750-1775 em- I (Asher et al., 1974b). This is explained by the fact that all six ester C=O groups coordinate with an enclosed potassium ion (Duax et al., 1972). Similar conclusions were obtained from Raman data (Rothschild and Stanley, 1975). 3.4.5.
Amphotericin B
Amphotericin B is a polyene antibiotic produced by the streptomycete culture M4575, obtained from soil of the Orinoco River region in Venezuela. The compound forms channels through lipid bilayers provided that the bilayers contain a suitable sterol such as cholesterol. In the absence of sterol, the antibiotic does not penetrate the membrane nor increase the characteristic cation permeability. Amphotericin B contains a heptaene backbone with an associated'lT -> '1T* electronic transition (Figure 7.19), which can be resonance excited to give enhanced Raman bands. In a channel, amphotericin B molecules are probably oriented with the ionizable mycosamine and carboxylic functions at the lipid-water interface
H3 C /CH 3 CH 'CH f) / 3 0' CH-CH 3 \\ NH-CH-C-O--c'H CH ~3 /C' l D -c~o I 3 /CH N'Z /CH-CH 3
?
HC
3\
c~o
I
,
CH-- CH
f H3 C
D~
l
O-C
0
D
f
NH
l
I
H C 3 \
I
CH-CH
NH I HC--
D
\0 3
l
J
, NH
/CH
H3 C-CH CH 3
I
CH
3
3
CH
d \
'C 0"""" \ /
CH-CH
\ C~O I
o""'c
f CH
\
l
D
/
0
I ~
CH 3
l /CH '#C.... D /~, 'CH-CH 0' FH_O_c_CH_NH 3 CH II \ CH 3 3 O/C~ CH 3 CH 3
FIGURE 7.18.
"0
,
Chemical structure of valinomycin.
218
Lipids and Biological Membranes H", OH
,'H COOH
H\~OH
NH2
o
OH CH3
FIGURE 7.19.
Structure of amphotericin B.
and are probably aligned with the polyene axis normal to the plane of the bilayer. Interaction of amphotericin B with phosphatidylcholine multilamellar and single-wall vesicles was investigated by Raman spectroscopy by Bunow and Levin (l977b, 1978). The amphotericin B C=C stretching vibration at 1556 cm- I is sensitive to the molecular changes that occur during the phase transition in DMPC-cholesterol multilayers. The C-H stretching region of lipids (2800-2950 em -I) proved sensitive to the inclusion of amphotericin B and provided information about the degree of· penetration of amphotericin B into the bilayer in the presence or absence of sterol. The ratio 12880/12850 temperature was monitored, and it was found that there is a large change in the packing of the lipid chains, which ordinarily occurs during the gel-to-liquid-crystalline phase transition. The transition was spread over a large temperature interval, rather than being suppressed, by the addition of cholesterol. Since amphotericin B can be induced to move from the polar interface to the interior of the lipid bilayer by the addition of a suitable sterol, amphotericin B can mimic both an extrinsic and an intrinsic membrane component. The Raman spectroscopic results are consistent with those of spin-label studies, in which a 15% change in molecular surface area is sensed by the spin label through the phase transition in DPPC-cholesterol multilayers (Marsh, 1974). Nystatin is also a polyene antibiotic, structurally similar to amphotericin B. When nystatin interacts with phospholipid-cholesterol multilayers, the intensities of the C=C and C-C modes of nystatin do not change, indicating that nystatin remains in the planar and trans form (Iqbal and Weidekamm, 1979).
3.5.
INTERACTIONS WITH OTHER COMPOUNDS
It is known that uncouplers of mitochondrial oxidative phosphorylation induce ultrastructural changes in the inner membrane. In order to clarify this phenom-
enon, liposome interaction with the uncouplers has been investigated by
Biological Membranes
219
Raman spectroscopy (Pasquali-Ronchetti et aI., 1980; Fini and PasqualiRonchetti 1981). It was found that the artificial membrane increases in membrane fluidity with the addition of uncouplers. This can be readily detected from the decrease in the ratio of intensity at 2890 to 2850 cm - I and also from the ratio of intensity at 1130 to 1090 cm -1. The decreases in the ratios indicate the increase of the gauche conformer or an increase in the rotational freedom of the fatty acid hydrocarbon backbone and also a decrease in lateral chain interactions. Even water molecules have an effect on lipid bilayer systems. This is studied by monitoring the Raman spectra of an anhydrous phospholipid lattice and the effect of hydration. Apparently the water molecules increase the mobility of the head-group glycerol and carbonyl moieties, as the hydration causes a shift in the CN symmetrical and PO; asymmetrical stretching modes (Bush et aI., 1980a, b). The difference in the Raman spectra of dipalmitoylphosphatidylethanolamine in solid and in aqueous solution is believed to be due to the effect of water. The lack of a Raman line at 1420 cm- 1 is explained in terms of less tightly packed hydrocarbon chains because of hydration (Akutsu et al., 1981). Proton-donor compounds such as monochloroacetic, dichloroacetic, and trichloroacetic acids affect the lateral packing of the acyl chains of phospholipids because of hydrogen-bond formation (Bertoluzza et al., 1981). Calcium ion is known to have a variety of effects on biological membranes, such as fusion of membranes and changes in the phase diagram. Raman spectroscopy shows that some artificial phospholipid membranes do fuse with the addition of calcium ions (Hark and Ho, 1980). 4.
BIOLOGICAL MEMBRANES
The plasma membrane of a cell is a multicomponent system contammg phospholipids, proteins, and carbohydrates as glycoproteins. It forms a boundary between the living and nonliving worlds. Raman spectroscopy has contributed greatly to the study of biological membranes in situ because of its nature as a nondestructive probe. Raman spectroscopy provides insights into the membrane structure where other probes fail. It provides information about the state of phospholipids and proteins within the membranes and their mutual interactions. Following is a review of some of these applications. 4.1.
ERYTHROCYTE MEMBRANES
We have discussed in detail the vibrational modes of fatty acids and phospholipids. C-C and C- H stretching vibrational modes are sensitive to structural changes. The techniques of measuring these changes can be used directly for the study of membranes. Because of the presence of fJ-carotene, which isa major constituent of carotenoids in biological membranes, the conjugated-polyene groups -C=C and =C-C= can be resonance enhanced (see Chapter 9).
220
lipids and Biological Membranes
4.1.1.
Membrane Fluidity
Raman spectra of hemoglobin-free human-erythrocyte membranes were first obtained by Bulkin (1972). Good-quality spectra of erythrocyte-ghost membranes were obtained by Lippert et al. (1975). Using the C-C stretching vibrational bands at 1130, 1090, and 1060 cm - I, they estimated that erythrocyte membranes contain 55 to 65% all-trans rigid configuration. The same conclusion was reached using much better quality Raman spectra from rabbiterythrocyte ghosts (Figure 7.20). The ratio of intensity at 1131 to 1087 cm - I is particularly useful, as these lines reflect the conformation of lipid hydrocarbon side chains and hence can serve as a measure of membrane fluidity. The I 131-cm-) line originates from the trans conformation of the C-C stretch, whereas the I087-cm -1 line is due to the gauche conformation of the C-C stretch plus PO; symmetrical stretching. For normal rabbit erythrocytes, this ratio is 0.71. The ratio is approximately 2.0 for solid DPPC, which is predominantly in the trans conformation. Thus the rabbit-erythrocyte ghost contains significant amounts of gauche conformation and hence has a considerable degree of fluidity (Milanovich et al., 1976a). This agrees with current concepts
1672
Ir
1659
1440
I
1005
I
1300
I
1131 I
1087
I
879 I
1600
1400
1200
1000
em
800
600
400
-1
FIGURE 7.20. Raman spectrum from 300 to 1700 crn - I of Dutch Belt rabbit erythrocyte ghosts in water. The figure was reproduced from Milanovich et al. (1976a) by permission of the copyright owner, North-Holland Publishing Company.
Biological Membranes
221
of model membrane structure. The amide I band is indicated by the band and shoulder at 1634, 1645, and 1659 cm - I and suggests the presence of considerable a-helix. The 1672-cm- I band is probably largely contributed by C=C in the unsaturated fatty acids. Assignments of Raman bands for the membrane phospholipids are shown in Table 7.4. Like -C-C- stretching vibrations, C-H stretching vibrational bands (2800-3100 cm- I ) can also be used to study membrane fluidity. The 2935-cm- 1 line becomes more dominant as polarity increases, the 2890/2850 cm -1 ratio decreases as fluidity increases. Thus a strong 2935-cm - I line (Figure 7.21) indicates that there is significant lipid-protein interaction within the membrane. A low ratio 12890/12850 is also an indication of substantial fluidity, which is in good agreement with the analysis of the C-C stretching region. When the ratios of intensity at 2880/2850 and 2930/2850 cm - I are plotted against temperature, there are several discontinuities. One appears between - 4 ~ - 8°C and the others at 17 and about 35-40°C. The low-temperature discontinuity originates from lipid phase changes in regions containing cholesterol (for cholesterol effect, see Section 3.1.2 of this chapter). The transition near 17°C is due to a restricted, cholesterol-poor lipid domain. The transition above 35°C, which is pH dependent, is due to a phase change involving protein and lipid interactions (Wallach and Verma, 1976 Verma and Wallach, 1976a, c).
3000
2800
em-'
FIGURE 7.21. Raman spectrum from 2800 to 3100 cm - I of Dutch Belt rabbit erythrocyte ghosts in water. The figure was reproduced from Milanovich et aI. (1976a) by permission of the copyright owner, North-Holland Publishing Company.
222
Lipids and Biological Membranes
Oxidative phosphorylation uncouplers alter the interchain interactions, resulting in variation of the lateral packing. This evidence was obtained from the fact that the Raman intensity of the 2890-cm - I band decreases upon the addition of uncouplers (Pasquali-Ronchetti et al., 1978). Uncouplers, however, are not able to induce a liquid crystalline phase transition corresponding to the complete disordering of the hydrocarbon chains. Phloretin is a membranetransport inhibitor and inhibits the transport of sugar, urea, anions, and the Na+ K+ pump. Phloretin binds to human red-cell membranes and the concomitant structural changes can be detected by Raman spectroscopy. A lack of the 1085-cm- 1 band in the phloretin-modified membrane is explained by less fluidity in the PO; and C-C regions of phospholipids (Zimmer et al., 1981). 4.1.2.
The Role of the Membrane Proteins
Membrane proteins have been studied much less than membrane lipids. Upon the addition of ATP and magnesium ion to erythrocyte ghosts, there is a change in IR spectra, indicating some protein-conformation changes from predominantly a-helix to f3-sheet (Wallach, 1972). Erythrocyte membranes are not static structures, but have dynamic properties and are sensitive to pH, cation and anion concentrations, and temperature. For example, an ESR study indicates that a pH-sensitive thermal transition occurs at 37.5-40.5°C. Proton NMR shows that methyl side chains of membrane proteins become more mobile as the temperature is raised (Sheetz and Chan, 1972). Such dynamic properties of the membrane can also be detected by Raman spectroscopy. The change in Raman spectra of the erythrocyte membrane upon varying pH and temperature is attributed to alteration in both lipid and protein components (Verma and Wallach, 1975; Mikkelsen et al., 1978a). When peripheral proteins are specifically removed from erythrocyte membranes, there is little change in Raman spectra in the region of 900-1700 cm- I. This is explained by the fact that removal of the peripheral proteins affects lipid portions only slightly. However, there is considerable change in the C-H stretching vibration region (2800-3000 cm -I), which is understood to mean that the integral-protein environment is affected markedly (Goheen et al., 1977). The presence of hemoglobin in erythrocytes will interfere with Raman spectroscopy because of intense resonance Raman scattering; therefore, hemoglobin must be removed from the erythrocytes. An erythrocyte without hemoglobin is called an erythrocyte ghost. Often erythrocyte membranes will be resealed (sealed erythrocyte ghosts) or sometimes not completely sealed, (unsealed erythrocyte ghosts) depending on the conditions. It has been observed using unsealed erythrocyte ghosts that the imposition of transmembrane cation gradients increases the intensity of Raman scattering in the CH 3 stretching band at 2930 cm- I. It is also known that the imposition of a membrane potential (negative inside) also increases the intensity of the 2930-cm - I band
.J
Biological Membranes
223
when sealed erythrocyte ghosts are used. This suggests that a transmembrane potential and cation gradient can energize membranes by compression of the apolar regions and transfer of protein methyl residues into polar regions (Mikkelsen et al., 1978b). 4.1.3.
Presence of Carotenoids
Human-erythrocyte membranes contain !J-carotene, as indicated by the two sharp, intense bands at 1530 and 1165 cm- I . These bands arise from the -C=C- and =C-C= stretching vibrations of the conjugated-polyene system of !J-carotene. When the membrane structure is perturbed by treatment with trypsin or lysophosphatidylcholine, or lowering of the pH, these two bands increase in intensity. Thus analysis of carotenoid bands can provide a probe for membrane structure (Verma and Wallach, 1975; Wallach and Verma, 1975). In this connection, it is worthwhile to mention the use of carotenoid as a resonance Raman probe. Pure phospholipids lack chromophore groups; therefore, their Raman spectra are not resonance enhanced. By introducing an external chromophore to the membrane phospholipids, one can monitor the membrane order-disorder transition using resonance Raman spectroscopy. Zeaxanthine, a carotenoid, was used by Mendelsohn and Van Holten (1979) for such a purpose. When zeaxanthine is inserted into a phospholipid dispersion and the latter is heated through the gel-liquid-crystal phase transition, large changes are observed in resonance Raman and absorption spectra of the carotenoid molecule. These changes are attributed to the fact that the carotenoid aggregates in the phospholipid gel, whereas it forms a monomer in liquid crystal phases. By following the changes in Raman intensity and absorption spectra, one can monitor the phase change of phospholipids. 4.2.
SARCOPLASMIC RETICULUM MEMBRANES
Raman spectra are readily obtainable from actively transporting sarcoplasmic reticulum membranes. Of particular interest is the appearance of strongly resonance-enhanced bands at 1160 and 1527 cm- I due to membrane-associated carotenoids. The C=C stretching vibration and the amide I band overlap each other. However, it is very likely that the l658-cm - I band is contributed by cis C=C rather than the trans form. This suggests that the sarcoplasmic reticulum membrane retains a considerable degree of membrane fluidity (Milanovich et al., 1976b). ATPase is the major protein component of the sarcoplasmic reticulum. The important question is whether the change in ATP hydrolysis activity at 18-20°C is due to a change in the lipid moieties or in the protein conformation. The protein conformation as studied by Raman spectroscopy indicates that 18°C is the transition temperature for sarcoplasmic ATPase while the lipid conformation is maintained in the range of 1O-37°C.
224
lipids and Biological Membranes
Thus the activity changes induced by both increased temperature and increased Ca(II) and Mg(II) concentrations are associated with changes in protein conformation (Lippert et aI., 1980; 1981). 4.3.
STREPTOCOCCUS FAECALIS MEMBRANE
Measurement of a transmembrane potential is important in understanding the function of cells. The measurement is usually done with microelectrodes. Frequently, cells are too small for the use of microelectrodes and indirect techniques must be used. One way is to use a dye whose uptake depends on a transmembrane potential. One example is the measurement of uptake of quinaldine red by Streptococcus jaecalis, a nonhemolytic species. Energization has an effect on the spectral behavior of quinaldine red in cells. It is suggested that the spectral change of quinaldine red reflects a change in membrane structure upon energization. How the membrane structure changes as a result of energization has not yet been elucidated (Koyama et al., 1979). 4.4.
NERVES
Raman optical responses can be observed during action-potential propagation in the sciatic nerve. Several carotenoids are natural constituents of biological membranes. Such carotenoids have two intense bands, at 1520 ~ 1525 and 1158 cm- I. The l525-cm- I band originates from C=C stretching vibration, whereas the 1I58-cm- 1 band results from a mixture of C-H bending and =C-C= stretching vibrations. During excitation, the intensity of the C=C stretching vibration at 1520 em -1 is lowered. The decreasing Raman intensity is explained in terms of changing bond order. This suggests that the membrane-bound carotenoids may be involved in the conduction of nerve impulses (Szalontai et aI., 1977). Protein-lipid complexes (proteolipid) are commonly found in a variety of nerve tissues, mitochondria, and sarcoplasmic reticulum. The protein component (apoprotein) of a proteolipid is unusually hydrophobic and soluble in organic solvents. In myelin multilayers, it is suggested that the apoprotein interacts with hydrocarbon chains of phospholipids. Freeze-fracture electron microscopy indicates that the protein of human myelin is deeply embedded in the hydrocarbon region of the lipid bilayer. One way to understand the combination of phospholipids and proteins in myelin proteolipids is to study how the phospholipids are perturbed by the protein components, using reconstitution experiments. Raman spectroscopic study indicates that the apoprotein has a considerable effect on the structure of hydrocarbon chains of DMPC. Such perturbation can be detected by observing the C-C stretching vibrational bands in the 10001I50-cm- I region and the C- H stretching region of 2800-3000 em-I (Curatolo et al., 1978). Below the lipid order-disorder transition temperature,
Biological Membranes
225
the number of acyl-chain trans conformers is reduced in the presence of apoprotein. Above the transition temperature, the acyl chains are prevented by the apoprotein from completely attaining extensive gauche conformation. The same result was obtained from a calorimetric study on the same system, so the two independent methods confirm each other (Curatolo et a1., 1977). In reconstitution experiments, myelin apoprotein perturbed saturated phospholipids more than unsaturated phospholipids (Lavialle and Levin, 1980).
4.5. 4.2.1.
OTHER PLASMA MEMBRANES
Thymocytes
Some cell membranes, such as the nuclear membranes from lymphocytes, undergo thermotropic segregation of their lipid and protein moieties. This segregation is observable by freeze-fracture electron microscopy. According to a study using the 12890/12850 ratio as a function of temperature, thymocyte plasma membranes demonstrate a thermotropic lipid transition near 23°C. Judging from the amide I and III bands, Verma et a1. (1975) concluded that thymocyte plasma membranes contain appreciable f3-structure. Concanavalin A is a lectin isolated from jack bean that causes mitogenic stimulation of rabbit thymocytes. It modifies many of the plasma-membrane functions. Raman spectroscopy was used to compare the membranes of resting thymocytes and those of mitogenically stimulated cells (Schmidt-Ullrich et a1., 1976). Major alterations were found in the C-H stretching (/2890/12850) CH in-plane deformation (1400-1500 cm- I ), and C-C stretching (1130, 1190, 1060 cm -I) regions. From this study it was concluded that there is a different lipid architecture of the membranes of activated cells. Moreover, concanavalin A lowers the transition temperature of sealed plasma-membrane vesicles of rabbit thymocytes. The modification of the lipid phase transition is probably due to unfolding of transmembrane proteins induced by concanavalin A (Verma et a1., 1980). 4.2.2.
Lymphocytes
The Raman spectra of plasma membranes from SV40 (simian virus 40)-transformed GD 248 lymphocytes were compared with the spectra of the membranes from normal cells over the range of 100-3010 cm- I. Striking differences between the two membrane categories were observed in the thermal response of the C- H stretching and acoustical regions. Analysis of the C-H stretching shows that the membranes of normal cells exhibit a thermal transition centered at 7°C and about 5°C wide. The membranes of GD 248 cells, in contrast, show a lipid transition centered at -5°C and 12-18°C wide. Analysis of the acoustical region shows the same result (Verma et al., 1977).
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232
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Verma, S. P., and Wallach, D. F. H. (1975). Carotenoids as Raman-active probes of erythrocyte membrane structure. Biochirn. Biophys. Acta 401, 168. Verma, S. P., and Wallach, D. F. H. (1976a). Multiple thermotropic state transitions in erythrocyte membranes. A laser-Raman study of the CH-stretching and acoustical regions. Biochirn. Biophys. Acta 436, 307. Verma, S. P., and Wallach, D. F. H. (1976b). Effect of melittin on thermotropic lipid state transition in phosphatidylcholine liposomes. Biochirn. Biophys. Acta 426,616. Verma, S. P., and Wallach, D. F. H. (1976c). Erythrocyte membranes undergo cooperative, pH-sensitive state transitions in the physiological temperature range: Evidence from Raman spectroscopy. Proc. Nat. Acad. Sci. 73, 3558. Verma, S. P., and Wallach, D. F. H. (1977a). Changes of Raman scattering in the CH-stretching region during thermally induced unfolding of ribonuclease. Biochern. Biophys. Res. Cornrnun. 74,473. Verma, S. P., and Wallach, D. F. H. (I 977b). Raman spectra of some saturated, unsaturated and deuterated C 18 fatty acids in the HCH-deformation and CH-stretching regions. Biochirn. Biophys. Acta 486,217. Verma, S. P., and Wallach, D. F. H. (1978). Interactions of proteolipid apoprotein with various lipids. In Proc. Sixth Jnt. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 110-111. Verma, S. P., Wallach, D. F. H., and Schmidt-Ullrich, R. (1975). The structure and thermotropism of thymocyte plasma membranes as revealed by laser Raman spectroscopy. Biochirn. Biophys. Acta 394, 633. Verma, S. P., Schmidt-Ullrich, R., Thompson, W. S., and Wallach, D. F. H. (1977). Differences between the structural dynamics of plasma membranes of normal hamster lymphocytes and lymphoid cells neoplastically transformed by simian virus 40 as revealed by laser Raman spectroscopy. Cancer Res. 37, 3490. Verma, S. P., Schmidt-Ullrich, R., and Wallach, D. F. H. (1980). State modifications of thymocyte plasma membrane proteins and lipids by mitogenic doses of concanavalin A: A Raman study on isolated membrane vesicles. J. Recept. Res. I, 1. Vogel, H. (1981). Determination of the conformational order of lipid membranes from Raman spectroscopy. Ber. Bunsenges. Phys. Chern. 85, 5 J 8. Vogel, H., and Jahnig, F. (1981). Conformational order of the hydrocarbon chains in lipid bilayers. A Raman spectroscopic study. Chern. Phys. Lipids 29, 83. Wallach, D. F. H. (1972). Infrared and laser Raman spectroscopy in membrane analysis. Chern. Phys. Lipids 8, 347. Wallach, D. F. H., and Verma, S. P. (1975). Raman and resonance-Raman scattering by erythrocyte ghosts. Biochirn. Biophys. Ac/a 382, 542. Wallach, D..F. H., and Verma, S. P. (1976). Structural dynamics of mammalian cell plasma membranes, studies by Raman and resonance Raman spectroscopy. In In/. Conf. Raman Spec/rosc. Proc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rhine, pp. 178-179. Wallach, D. F. H., Verma, S. P., and Fookson, J. (1979). Application of laser Raman and infrared spectroscopy to tbe analysis of membrane structure. Biochirn. Biophys. Acta 559, 153. Weidekamm, E., Bamberg, E., Brdiczka, D., Wildermuth, G., Macco, F., Lehmann, W., and Weber, R. (1977). Raman spectroscopic investigation of the interaction of gramicidin A with dipalmitoyl phosphatidylcholine liposomes. Biochirn. Biophys. Acta 464, 442. Weidekamm, E., Bamberg, E., Janko, K., and Weber, R. (1978). Laser-Raman study of valinomycin-doped and w-CD)-Iabeled phosphatidylcholine liposomes. Arch. Biochern. Biophys. 187, 339.
References
233
Yellin, N., and Levin, I. W. (1977a). Hydrocarbon chain disorder in lipid bilayers. Temperature dependent Raman spectra of 1,2-diacyl phosphatidy1choline-water gels. Biochim. Biophys. Acta 489, 177. Yellin, N., and Levin, I. W. (1977b). Cooperative unit size in the gel-liquid crystalline phase transition of dipalmitoyl phosphatidy1choline-water multilayers: An estimate from Raman spectroscopy. Biochim. Biophys. Acta 468, 490. Zimmer, G., GUnther, H. 0., and Schmidt, H. (1981). Interaction of phloretin with the human red cell membrane and membrane lipids: Evidence from infrared, Raman and ESR spectroscopy. Z. Naturforsch., C: Blosci. 36C, 586.
CHAPTER
Carbohydrates
Like proteins and nucleic acids, macromolecular carbohydrates have complex conformations that help to determine their physical, chemical, and biological activities. Few studies of carbohydrates have been made by Raman spectroscopy, but extensive studies have been made by IR absorption spectroscopy. Raman spectra of carbohydrates are usually sharper than the IR spectra of corresponding compounds and thus provide more-precise information. Some vibrational modes, such as those of C=N, C=S, C-C, and S-H, are manifested strongly in Raman spectra, whereas the corresponding bands are weak in IR spectra. Vibrational modes of carbohydrates are complex, and the interpretation of Raman spectra is often difficult. Many different carbohydrates are actually isomers; sometimes the only difference is the position of a hydroxyl group. Being polyhydroxyl compounds, carbohydrates give extremely complex Raman spectra involving different types of OH, C-H, C-C, C-C-O, C-O, C-O-C, and C-O-H vibrations. In this chapter some related IR data is briefly mentioned. Excellent reviews are available on IR studies of carbohydrates (Tipson, 1968; Parker, 1971). Because of relatively recent application of Raman spectroscopy to carbohydrates, no review article on Raman spectroscopy has appeared.
~
Glycosidic Linkage
1.
235
ASSIGNMENT OF RAMAN BANDS
Exact assignment of Raman bands for carbohydrates is difficult, despite their relatively low molecular weight. Some assignments of Raman bands for glucuronic acid, D-glucose, deuterated D-glucose, glucosamine, deuterated glucosamine, N-acetylglucosamine, and deuterated N-acetylglucosamine are summarized in Table 8.1 (She et aI., 1974). Observed band frequencies of a-D-glucose were compared with the frequencies calculated by normal coordinate analysis and were found to be in fairly good agreement (Vasko et aI., 1972; Cael et aI., 1974). Many carbohydrates, including simple monosaccharides, show a number of low-frequency bands. These are believed to be pyranose-ring-related vibrations, but these Raman bands have not been assigned to specific vibrational modes. However, IR absorption low-frequency bands (70-500 cm~ I) of f3-Dglucose have been assigned by the normal coordinate calculation (Hineno, 1977).
2.
GLYCOSIDIC LINKAGE
Determination of anomeric configuration is an important aspect of structural carbohydrate chemistry and is frequently achieved by proton NMR, measurement of optical rotation, or IR absorption spectroscopy. As IR spectroscopy is closely related to Raman spectroscopy, a brief review of salient features of the IR spectra of anomers is given. 2.1.
IR ABSORPTION SPECTROSCOPY
It is generally accepted that several absorption bands in IR spectra are characteristic for a- and f3-anomers, and they have been grouped as types 1, 2a, 2b, and 3.
a-Anomers f3-Anomers
Type 1 (em-I)
Type 2 (a or b) (em-I)
Type 3 (em-I)
917 ± 13 920 ± 5
844 ± 8 891 ± 7
766 ± 10 744 ± 9
The origin of the type I band is "asymmetrical ring vibration mode," similar to that of 1, 4-dioxane (Barker et aI., 1954a, b; Mateescu, 1971). Type 2 bands originate from the anomeric C-H deformation (bending vibration): equatorial C-H vibration in a-D-anomers in the 4C I(D) conformation. Type 3
.
TABLE 8.1.
Glucuronic Acid
533
Continued
Deuterated GlucosamineHCl
N-Acetyl Glucosamine
Deuterated N-Acetyl Glucosamine
533 525
533
533
533
513
513
460
460
v-Glucose
Deuterated v-Glucose
GlucosamineHCl
533
533 513
468 460 441 425
382 353 340
405 397 382 353
Probably due to skeletal modes
513 482 460
513 482 460
425
430 405
415 397
416
408 382 353
382 353
382 353
381 353 335
381 353 335
328 303 290 260 245 220
303 290 260 245 220
190 165
190 165
130
130
416
Assignments
310 278 260
310 290 260
310 290 260
310 291 265
310 291 265
220
220
220
220
220
190 175 140 130
190 175 140
190 175 140
190 165 140
190 165 140
408
Probably due to skeletal modes
Probably due to torsional modes
TABLE 8.2.
Characteristic, Anomeric, C-H Bands in the Laser-Raman Spectra of a Variety of Carbohydrates
Carbohydrate Glucosyl-S-CHzCOzH ({3) Solid Aqueous solution Galactosyl-S-CH 3 ({3) Solid Aqueous solution Galactosyl-S-CHzCOzH ({3) Solid Aqueous solution Glucosyl-S-(CHzhCONHCHzNHCOCH=CHz ({3) Solid Aqueous solution GlcNAc-S-CH zCN ({3) Solid Aqueous solution a-Methyl-o-glucoside (solid) {3- Methyl-o-glucoside (solid) Deuterated a methyl a-Oglucoside (solid) Deuterated a methyl {3-0glucoside (solid) a-o-glucose (solid)
Type 2a Band
Type 2b Band
Reference
895 898
Tu et al. (1978) Tu et al. (1978)
884 883 (Sh)
Tu et al. (1978) Tu et al. (1978)
895 902
Tu et al. (1978) Tu et al. (1978)
890
Tu et al. (1978)
883 887
Tu et al. Tu et al. Tu et al. Tu et al.
842 890
Tu et al. (1977)
837 850 842 840
(1978) (1978) (1977) (1977)
Tu et al. (1977) She et al. (1974) Vasko et al. (1972)
TABLE 8.2.
Continued
Carbohydrate Deuterated a a-D-glucose (solid) A mixture of 2-amino-2-deoxy-aand -P-D-glucose (solid) A mixture of deuterated a 2-amino2-deoxy-a- and -P-D-glucose (solid) 2-Acetamido-2-deoxy-a-Dglucose (solid) a-Lactose (solid)
Type 2a Band 842
Chondroitin-6-sulfate Chondroitin-4-sulfate MaItose MaItriose Cyclohexaamylose aAll hydroxyl groups were isotopically exchanged to give -O-D
Reference She et al. (1974)
865
896
She et aI. (1974)
845
905
She et al. (1974)
890 898
She et al. (1974) Susi and Ard (1974) Cad et aI. (1974) Susi and Ard (1974) Cael et aI. (1974)
890
Susi and Ard (1974)
896 899 884 891 905
She et aI. (1974) Tu et al. (1977) Bansil et al. (1978) Bansil et aI. (1978) Bansil et al. (1978) Tu et aI. (1979)
865 850 840
p-Lactose (solid) A mixture of a- and p-Iactose (aqueous solution) Deuterated a 2-acetamido-2-deoxya-D-glucose Hyaluronic acid
Type2b Band
855 857
850 854 851-860 865
Tu et aI. (1979) Tu et aI. (1979)
Glycosidic Linkage
~H
~
H20HO
OR
H
I
()( anomer
241
anomer
FIGURE 8.1. C/- and ,B-anomers of hexose. In the a-anomer, the C-H bond is equatorial, whereas the ,B-anomer has an axial C-H bond.
vibrational bands are the result of "symmetrical ring vibration" of the hexopyranose ring. These three types of absorption bands of a- and p-anomers overlap considerably, and only type 2 bands can be used to differentiate the anomeric glycosidic linkages. 2.2.
RAMAN SPECTROSCOPY
Type 2 bands are useful for discerning anomeric linkages by Raman spectroscopy, as they are with IR spectroscopy. As can be seen from Table 8.2, the type 2 bands differentiate a- or p-anomers in monosaccharides and disaccharides or the glycosidic linkages in polysaccharides. The rule is equally applicable to the normal glycosidic linkage (C-O-C) as well as to the I-thioglycosidic linkage (C-S-C). This is reasonable, because type 2a and 2b bands arise from the equatorial C(l)-H and the axial C(I)-H bending vibrational mode respectively and not from the glycosidic bond itself (Figure 8.1). Thus the p-anomer of I-thioglycosides and the p-anomer of normal glycosides show a type 2b band around 890 ± 10 cm - I (Figure 8.2). Hyaluronic acid consists of repeating unit of glucuronic acid and N-acetylglucosamine connected through P-I, 3 and P-I, 4 linkages (Figure 8.3). Hyaluronic acids show only a type 2b band at 896 cm - I, as shown in Table 8.2 (Tu et al., 1977). Again, this is reasonable, as both P-l, 3 and P-I, 4 linkages associate with the identical axial C- H at the C-I position. Similarly, chondroitin sulfate contains two types of p-Iinkages, but it gives only the type 2b band at 884 cm-) for chondroitin 6-sulfate and at 889 cm -2 for chondroitin 4-sulfate (Bansil et al., 1978). Actually, a-lactose is distinguished from p-lactose by Raman spectroscopy using this principle (Susi and Ard, 1974). The proportions of anomers found from Raman intensity H20HO
~
SR
~~~
~OR H
H
f3 FIGURE 8.2. bond.
anomer
f
anomer
Both ,B-thiohexopyranoside and normal ,B-hexopyranoside have the axial C-H
242
Carbohydrates
H
----
~~ O·
o
HO
~ •••••
0
0
HO
NH C
0 COOH
o
C~
G FIGURE 8.3. acetylglucose.
N
G
Repeating units of hyaluronic acid where G is glucuronic acid and N is N-
ratios for glucose and sucrose are similar to those found by other techniques (Mathlouthi and Luu, 1980a).
3. 3.1.
FUNCTIONAL GROUPS OH VIBRATIONS
The OH stretching vibration appears in the 3100-3500 em-I region. In o-glucose, there is a sharp 3400-cm - I band and a broad band in the region of 3265 em-I (She et al., 1974). When the OH group is isotopically substituted with deuterium to make -0-0, the hydroxyl-group vibrational band shifts to 2450-2555 cm- I (Figure 8.4). The assignment of each Raman band to specific hydroxyl groups is not an easy task, even for glucose. Glucose has five hydroxyl groups, with two bands at 3400 and 3265 em -1. The conversion of one OH group in glucose to -O-CH) leaves four other OH groups identical to those in the original glucose (Figure 8.5). Yet a-methyl-o-glucoside has three new bands at 3250, 3350, and 3455 em - I and f3-methyl-o-glucoside has one sharp band at 3550 em - I and a broad band from 3100 to 3350 em -I (Figure 8.6). The C-OH deformation modes usually appear in the region of 970-1250 cm- 1 (Galat, 1980). Some C-OH deformation bands appear in IR but not in Raman scattering bands. The complexity of OH vibrational bands in carbohydrates is due to the complex pattern of hydrogen bonding. A slight change in OH modification or conformation induces a change in the hydrogen-bonding pattern. Further complications are that the hydrogen bonding is not only intramolecular but also involves intermolecular effects. Hydrogen bonding in various compounds has been a subject of intense study, especially by IR spectroscopy (Badger and Bauer, 1937; Mitchell, 1968; Hadzi and Bratos, 1976; Novak, 1979). Hydrogen bonding usually shifts the OH stretching bands to lower frequency. Another reason for the complexity of the vibrational spectra of carbohydrates is that the fine structure in the 0- H stretching region arises from coupled vibrations rather than from separate vibrations of individual 0 - H groups. Carbohydrates
",0 'it '" N 01 N
o
A
III <Xl
;e, '2960
1?~8CH
-00 ~
.,. '"
~
~ ...
~jl\ IIMI'2995 '" 0 I
'"
010 CD 'it "'01
'"
B
~---l
.. _...",
~1\11-CH II~ ,,,,
2910
-OHo
o
'" CD
o
--~ 2500
2000 2200
'it
'"
2800 (II
3400 3600
3100
= em-I)
FIGURE 8.4. Raman spectra of (A) deuterated D-glucose; (B) D-glucose. See the frequencies of OH before and after deuteration. The figure was reproduced from She et al. (1974) by permission of the copyright owner, Elsevier Scientific Publishing Company.
,F-O-J
CH20H
0
O
.OH HO
H~"""CH3 HO
O_CH 3
1 H
HO
a-Methyl-D-Glucoside
8-Methyl-D- Glucoside
HO
H
.
H(E)
oCH
HU
roCH3 (I)
H
(A)
3
(A)
FIGURE 8.5.
Structures of
Cl-
and {l-methyl-D-g1ucoside.
243
244
Carbohydrates ,~
o
o
11;g~
~I I/I~
It)
(b)
(\J
It)
,;g m
m
o
CD
~
(\J
(a)
2200
2500
2800 /I
3100
3400
3600
=cm-1
FIGURE 8.6. Raman spectra of (A) a-methyl-D-glucoside; (B) ,B-methyl-D-glucoside. The figure was reproduced from Tu et a!. (1977).
are polyhydroxyl compounds and ideal ones to study hydrogen bonding using vibrational spectroscopy. Unfortunately, very few such studies have been made using Raman spectroscopy.
3.2.
C-H VIBRATIONS
The C-H stretching vibration of carbohydrates gives a complex pattern in the region of 2800-3050 em-I. The complexity arises from the presence of different types of CH-containing groups such as -CH 3 , -CH 2 , and C-H. a-Methyl-D-glucoside and its anomer have identical primary structures except the position of -OCH 3 , but their C-H vibration patterns are not the same (Figure 8.6). Whether the CH vibration has any sensitivity to structural change in carbohydrates as in lipids is not yet known. The Raman bands at 1457, 1337, 1219, and 1011 cm- I for a-D-glucose are assigned to CH 2-re1ated modes,
Functional Groups
245
whereas those at 1404, 1360, 1250, 1076, 1047,911, and 836 cm- I are assigned to C-H-related modes (Vasko et al., 1971). 3.3.
N-H VIBRATIONS
Many carbohydrates, such as glucosamine, galactosamine, and N-acetylglucosamine contain primary and secondary amino groups. The N-H stretching vibration mode usually appears in 3000-3200 cm - I, slightly lower than that of the OH group (She et al., 1974). On some occasions, when the Raman bands appear in the borderline area of OH- and NH-vibration frequencies, the differentiation of NH from OH may become a problem. Deuteration in D2 0 does not solve the problem, because both NH and OH are readily exchangeable with deuterium. Hyaluronic acid, which contains N-acetylglucosamine, shows a broad band from 3100 to 3600 cm - ), without any particular discrete bands for OH and NH (Tu et al., 1977). In chitin, there is a broad IR absorption band, but there are two identifiable bands, from which it ean be concluded that the 3100-3300 cm- I region is due to NH stretching and the 3450-3480 cm - I region is due to 0- H stretching vibrations (Hackman and Goldberg, 1974). 3.4.
CARBOXYL AND ACETYL GROUPS
The -COOH and CH 3CO- groups are common in naturally occuring carbohydrates. The strongest band is the C=O stretching vibration of the carboxyl or the acetyl group. The carboxylic C=O usually appears at higher frequency than that of the acetyl group (Table 8.3). In the free carboxyl group, the band from the symmetrical -COO- vibrational mode (Figure 8.7) appears at 1410 cm -) (Bansil et al., 1978). The band from the carboxylate-ion asymmetrical vibration appears at 1605 :±: 5 cm - I in IR spectra (Casu et aI., 1978).
3.5.
SULFATES
The sulfate group is common in carbohydrates. For instance, chondroitin sulfate and heparin contain sulfate. The sulfate can be attached to an OH group as -OSO; or to an amino group as -NHSO; . The sulfate group in carbohydrates has a variety of stretching vibrational modes, as shown in Figure 8.8. The symmetrical-vibration mode of the -SO; group shows the most prominent peak near 1060 em-I. Using this band, it is quite easy to detect sulfate-containing carbohydrates. Several Raman bands associated with the sulfate group are summarized in Table 8.4. Symmetrical -SO; vibration with N-sulfate appears at somewhat lower frequencies (~ 1040 em -) than those of O-sulfates (~ 1060 em -) (Cabassi et al., 1978).
TABLE 8.3.
Comparison of C=O Stretching Vibration Frequencies for Carboxylic Group and N-Acetyl Group
Compound
f3-D-G1ucosy1-SCH 2 COOH D-G1ucuronic acid Hyaluronic acid Deuterated glucosamine 2-Acetamido-2-doxy-D-mannopyranose N-Acety1-D-g1ucosamine Hyaluronic acid N-Acety1-D-g1ucosy1 SCH 2 CN 2-Acetamido-2-deoxy-D-g1ucopyranose Chitin (Antarctic krill, and crayfish) Chitin (lobster, crayfish, honey bee)
Frequency Carboxyl Group 1675 1707 1735 N-Acetyl Group 1620 1620-1630 1632 1650 1659 1650-1660 1660 1660
Reference
Tu et al. (1978) She et al. (1974) Tu et al. (1977)
She et al. (1974) Galat and Popowicz (I 978a) She et al. (1974) Tu et al. (1977) Tu et al. (1978) Galat and Popowicz (1978a) Galat and Popowicz (1978a) Galat and Popowicz (1978b)
o
Ott 1=$ -C" ~~asy
II
-c,oSl
~O
/0
/0 S
"
1240 em- 1
C-O-S
S
C-O-S
\0
0
aSYl1111etrie
sym
FIGURE 8.7. Symmetrical and asymmetrical vibrations of free carboxylate ion. Sym, symmetricalstretching vibration; asy, asymmetrical-stretching vibration.
sYl1111etri e
aSYl1111etri e
1060 em- 1
980 em
-1
sYl1111etrie 820 m- 1
FIGURE 8.8. Vibrations associated with the C-O-S0 3 group.
TABLE 8.4.
Raman Frequencies Associated With Sulfate-Group Vibrations
Compound -Chondroitin 6-sulfate Chondroitin 4-sulfate a-Methyl-D-glucopyranoside 2-sulfate D-Glucose 6-sulfate 2-Deoxy-2-sulfoamino-Dglucose
Asymmetrical Stretch 1237 1240 1232 1240 1226 1216 1182
Dextran sulfate Dermatan sulfate Heparin
1240 1225 1222
Heparin sulfate N-Desulfated heparin COOH-reduced heparin Heparin-OD Keratan sulfate
1220 1225 1220
Symmetrical Stretch
~
Reference
1062 1065 1079 1067
Bansil et al. (1978) Cabassi et al. (1978) Bansil et al. (1978) Cabassi et al. (1978)
1050
Cabassi et al. (1978) Cabassi et al. (1978)
1040 (H 2 O) 1046 (solid) 1069 ~ 1065 1062 (H 2 O) 1065 (solid) - 1055
1060 1066
Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al. Cabassi et al.
(1978) (1978) (1978) (1978) (1978) (1978) (1978) (1978) (1978) (1978) (1978)
248 4.
Carbohydrates
COMPARISON OF SOLID AND AQUEOUS PHASES
Because of the relatively weak background due to water, the Raman spectra of many compounds can be measured in aqueous solution; this is usually considered to be the greatest advantage of Raman over IR spectroscopy. However, in this regard, caution must be exercised in the interpretation of Raman spectra of carbohydrates. Although the water background may be weak, its OH vibration is still evident in Raman bands (Figure 8.9). The OH stretching vibration of a water molecule has a broad band from 2800 to 3800 cm- I . Detailed assignment of water vibrations has been made by Walrafen (1964, 1970, 1971). For carbohydrates, the CH and OH stretching vibration bands are also located in this region; thus the water background completely obscures the carbohydrate bands, and this renders the Raman spectra of carbohydrates in this region almost useless. On the other hand, Raman spectra of carbohydrates can be useful in the region of 1800-3000 cm- I. However, even here, extreme caution is necessary, as the spectra are considerably influenced by the water background. For instance, the C=O stretch vibration of N-acetylglucoseSCH 2CN appears at 1659 cm- I for a solid sample but it appears at 1648 cm- I in aqueous solution. The Raman spectrum of pure water shows one intense band at 1643 cm- 1 (Figure 8.9C). Therefore, the apparent C=O band of N-acetylglucose-SCH 2 CN at 1648 cm - I is misleading (Tu et al., 1978). In addition to problems created by the overlapping of water bands, spectra of
jI
A
~
13861
I
357
<Xl C\l
"
c c o
E o
0:
B
C
1643
1700
1500
1300
1100
900
700
500
300
Wavenumber (cm- 1 )
FIGURE 8.9. Raman spectra of cyanomethyl-2-acetamide-2-deoxy-l-thio-fJ-D-glucopyranoside GluNAc-S-CH 2CN(fJ) in solid form (A), aqueous solution (B), and water (C). The figure was reproduced from Tu et aI. (1978) by permission of the copyright owner, Elsevier Scientific Publishing Company.
Conformation
249
solid carbohydrates and aqueous solutions show differences due to hydrogenbonding-pattern differences.
5.
CONFORMATION
Like proteins, macromolecular carbohydrates have complex conformations that help in determining their physical, chemical, and biological activities (Rees, 1977). Unfortunately, the conformations of carbohydrates have been studied less than those of other biological macromolecules. Vibrational modes of carbohydrates are complex, and the interpretation of spectra is often difficult. In view of this complexity, it is advisable to examine the structural studies of the simpler carbohydrates first. Cyclohexaamylose (Schardinger a-dextrin), a cyclic compound composed of six a-D-glucopyranosyl residues, was examined by Raman spectroscopy (Tu et al., 1979). As the compound is cyclic, there is no free, hemiacetyl hydroxyl group, and all of the D-glucosidic linkages involved are a-D. There is the possibility of a number of rotamers along the C(1)-O-C(4) axis for maltose, but not for cyclohexaamylose, as all of the D-glucopyranosyl residues are locked into a ring structure. If there are free rotations along the glycosidic-linkage axis, a great number of rotamers are possible for maltotriose (see Figure 8.10), and it would be expected to give a Raman spectrum more complicated than that of cyclohexaamylose. For cyclohexaamylose, there is probably only one conformation because of the restriction of rotation due to its cyclic nature. However, the Raman spectrum of maltotriose is no more complex than that of cyclohexaamylose. Careful examination of the three carbohydrate spectra showed that the spectrum of maltotriose is, overall, more similar to that of cyclohexaamylose than that of maltose, suggesting that the trisaccharide probably exists in a conformation very similar to that of cyclohexaamylose. Instead of a D-glucose unit rotating freely around the C(1)-O-C(4) axis, maltotriose probably has a somewhat-fixed conformation, like that of half cyclohexaamylose (Figure 8.10). Cellulose is the primary polysaccharide component of plant cell walls and consists of ~-l, 4-linked glucose polymers. In cellulose, the methine CH bonds are axial and the pyranose rings are equatorial with respect to the individual anhydroglucose residues. Raman intensities of cellulose depend on the fiber-axis orientation in respect to the incident exciting radiation. In particular, the 1098-cm - I skeletal band is very intense in the parallel mode and much reduced in the perpendicular mode. For the methine stretching band at 2920 cm -I, the intensity is greater for the perpendicular orientation than the parallel mode. This reflects the fact that the orientation of methine CH bonds is, essentially, perpendicular to the direction of the cellulose chain. All these results suggest a preferential orientation of the glucose units in tangential planes (Atalla, 1980; Atalla et a1., 1980). The ratio /874//826 was used by Mathlouthi and Luu
250
Carbohydrates CH20H
r!r
HOH C 2 HOHO
OH
rr
°7~'\
A
HO~
OH
H:H2~~O~CH20H
o
HOH 2C
OH / H X
'oH
HO
0
B
HO
OH
OH
HO
p~~ -----~-------------------O-----
c
o FIGURE 8.10. Structure of (A) maltose, (8) maltotriose, (C) cyclohexaamylose. Note that a maltotriose molecule is exactly half the size of cyclohexaamylose, and it is very likely that it has a conformation similar to that of half of cyclohexaamylose. The figure was reproduced from Tu et a1., (1979) by permission of copyright owner, Elsevier Scientific Publishing Company.
(l980b) as a measure of fructose furanoid ring (five-membered ring) and pyranoid ring (six-membered ring). Amylose is a homopolysaccharide composed of only glucose units with A, B, C, and V forms, which have different X-ray diffraction patterns. Two types of amylose were extensively studied by Cael et al. (1973). The B-form amylose occurs in tuber starches. Amylose can be isolated from starch by precipitation using organic solvents. The precipitated V-form amylose is complexed with solvent and has a different X-ray diffraction pattern from that of B-form amylose. The V form converts to B-form amylose in water. Raman lines at
Lipid-Carbohydrate Interactions
251
1263 and 946 cm- 1 of V-form amylose shift to 1254 and 936 cm- 1 on conversion into the B form. There is also an intensity change for the lines at 2940 and 1334 cm -1. These Raman changes are associated with an extension of the helix and changes in the intramolecular hydrogen bonding. Raman spectra of other polysaccharides have been obtained; these include hyaluronic acid (Tu et al., 1977), chondroitin sulfate (Bansil et al., 1978), chitin (Galat and Popowicz, 1978a, b), and heparin (Cabassi et al., 1978). However, these studies have not been extended to the conformational properties. One unique way to study the conformation of carbohydrates is using the fact that some cations interact more strongly with one conformer than the other. This difference can be detected in the Raman spectra of the complexes (Williams and Atalla, 1981). 6.
LIPID-CARBOHYDRATE INTERACTIONS
Monoglyceride forms a complex with the V form of amylose. The C-H stretching vibration bands of lipids are sensitive to structural change, yet there
FIGURE 8.11. Diagram showing a complex of monostearin and amylose. The figure was reproduced from Carlson et al. (1979).
252
Carbohydrates
is a considerable overlapping of different bands. The interaction was studied with deuterated monoglyceride so that the C-D stretching vibration bands could be examined; in this case there is no ambiguity because of no interference from other vibrational modes. Monoglyceride forms a complex with the V form of amylose. The monoglyceride is believed to be inside the loop of the amylose helix as shown in Figure 8.11. When the complex is formed, the affinity of the amylose to iodine is reduced to zero, indicating that all of the I 2-reactive sites of the amylose molecule are complexed with monoglyceride molecules. The hydrocarbon chain conformation inside the amylose helix seems to be ordered, as in the crystalline state, and the geometry of the amylose-V-form of this complex corresponds to nearly three turns of the helix per hydrocarbon chain (Carlson et aI., 1979).
7.
OTHER STUDIES
Sucrose forms two types of complexes with CaCl 2 in NaOH. The formulas of these complexes are C I2 H 220\l . 3CaO and C12H160\lCa3. It is possible to distinguish two types of complexes by comparing their Raman spectra (Francotte et aI., 1979). Inclusion type complexes are possible for cyclic oligoamylose. A hypoglycemic drug, tolbutamide [p-CHd'S02NHCONH(CH2kCH3] can be included in the cavity of ,B-cyc1odextrin (cycloheptaamylose). With the inclusion a change occurs in the C-H out-ofplane vibrational bands of the drug's phenyl-moiety (Veda and Nagai, 1981). An antifreeze glycoprotein obtained from the sera of an Antarctic fish causes an anomalous decrease in the freezing point of water. The hydroxyl group may play an important role, as there is a difference in Raman spectra of the hydroxyl group for active and inactive glycoprotein components. It may be that hydrogen bonding of the carbohydrate hydroxyls of the active glycoprotein at the ice-solution interface physically prevents the growth of the ice lattice; thus water does not freeze (Tomimatsu et aI., 1976). It is well known that the iodine-starch complex exhibits a blue color, whereas the isolated I) anion absorbs mainly in the uv region. It is believed that the iodine-starch complex contains elongated polyiodide chromophores absorbing in the visible region (Tasumi, 1972; Heyde et aI., 1972). Raman spectra of the iodine-starch complex always contain fundamental vibrations at 162 and 112 cm- I (Mulazzi et aI., 1980). The iodide ion may also exist as I~- , I~- , and Ifo- , and these ions can combine with the amylose lattice. The lattice-vibration band for the iodidestarch complex is found at 55 and 27 cm - I (Handa et aI., 1980). When carbohydrates are dissolved in water, there is an interaction between solute and water molecules. As the concentration of solute increases, the CH 2 bending frequency for sucrose and glucose changes. Also, depending on the concentra-
References
253
tion of solute, the water-molecule bending vibration at 1640 cm - I varies in frequency and intensity (Mathlouthi et a1., 1980; Walrafen and Abede, 1978). REFERENCES Atalla, R. H. (1980). Raman spectral evidence of molecular orientation in native cellulosic fibers. In Proc. VlIth Int. Con/. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 618-619. Atalla, R. H., Whitmore, R. E., and Heimbach, C J. (1980). Raman spectral evidence for molecular orientation in native cellulosic fibers. Macromolecules 13, 1717. Badger, R. M., and Bauer, S. H. (1937). Spectroscopic studies of the hydrogen bond. II. The shift of the O-H vibrational frequency in the formation of the hydrogen bond. 1. Chern. Phys. S, 839. Bansil, R., Yannas, I. V., and Stanley, H. E. (1978). Raman spectroscopy: A structural probe of glycosaminoglycans. Biochim. Biophys. Acta 541, 535. Barker, S. A, Bourne, E. J., Stacey, M., and Wiffen, D. H. (I 954a). Infrared spectra of carbohydrates. Part I. Some derivatives of o-glucopyranose. J. Chem. Soc. 171, 171. Barker, S. A, Bourne, E. J., Stephens, R., and Whiffen, D. H. (I 954b). Infrared spectra of carbohydrates. Part II. Anomeric configuration of some hexo- and pento-pyranoses. J. Chem. Soc. 171, 3468. Cabassi, F., Casu, B., and Perlin, A S. (1978). Infrared absorption and Raman scattering of sulfate groups of heparin and related glycosaminoglycans in aqueous solution. Carbohydr. Res. 63, I. Cael, J. J., Koenig, J. L., and Blackwell, J. (1973). Infrared and Raman spectroscopy of carbohydrates. Part Ill. Raman spectra of the polymorphic forms of amylose. Carbohydr. Res. 29, 123. Cael, J. J., Koenig, J. L., and Blackwell, J. (1974). Infrared and Raman spectroscopy of carbohydrates. Part IV. Identification of configuration and conformation-sensitive modes for D-glucose by normal coordinate analysis. Carbohydr. Res. 32, 79. Carlson, T. L.-G., Larsson, K., Dinh-Nguyen, N., and Krog, N. (1979). A study of the amylosemonoglyceride complex by Raman spectroscopy. Starch/Starke 31,222. Casu, B., Scovenna, G., Cifonelli, A J., and Perlin, A S. (1978). Infrared spectra of glycosaminoglycans in deuterium oxide and deuterium chloride solution: Quantitative evaluation of uronic acid and acetamidodeoxyhexose moieties. Carbohydr. Res. 63, 13. Francotte, C, Vandegans, J., Jacqmain, D., and Michel, G. (1979). Etude spectroscopique de complexes ca\ciques du saccharose. La Sucrerie Beige 98, 137. Galat, A (1980). Study of the Raman scattering and infrared absorption spectra of branched polysaccharides. Acta Biochim. Polonica 27, 135. Galat, A, and Popowicz, J. (1978a). Study of the infrared spectra of chitins. Bull. de L'Academie Polonaise des Sciences, serie des sciences biologiques, Cl, II, XXVI, No.5, 295. Galat, A, and Popowicz, J. (1978b). Study of the Raman scattering spectra of chitins. Bull. de L'Academie Polonaise des Sciences, serie des sciences biologiques, Cl, II, XXVI, No.8, 519. Hackman, R. H., and Goldberg, M. (1974). Light-scattering and infrared-spectrophotometric studies of chitin and chitin derivatives. Carbohydr. Res. 38, 35. Hadzi, D., and Bratos, S. (1976). Vibrational spectroscopy of the hydrogen bond. In The Hydrogen Bond Il, P. Schuler, G. Zundel and C Sandorfy, Eds., North-Holland, New York. Handa, T., Yajima, H., and Kajiura, T. (1980). On the blue color of triiodide ions in starch and starch fractions. III. Resonance Raman spectra of bluing species in amylose. Biopolymers 19, 1723.
254
Carbohydrates
Heyde, M. E., Rimai, L., Kilponen, R. G., and Gill, D. (1972). Resonance-enhanced Raman spectra of iodine complexes with amylose and polyvinyl alcohol, and some iodine-containing trihalides. J. Am. Chern. Soc., 94, 5222. Hineno, M. (1977). Infrared spectra and normal vibrations of ,B-o-glucopyranose. Carbohydr. Res. 56,219. Mateescu, G. H. (1971). Infrared Spectroscopy: Applications in Organic Chemistry, Wiley, New York. Mathlouthi, M., and Luu, D. V. (l980a). Laser-Raman spectra of o-glucose and sucrose in aqueous solution. Carbohydr. Res. 81, 203. Mathlouthi, M., and Luu, D. V. (1980b). Laser-Raman spectra of o-fructose in aqueous solution. Carbohydr. Res. 78, 225. Mathlouthi, M., Luu, c., Meffroy-Biget, A. M., and Luu, D. V. (1980). Laser-Raman study of solute-solvent interactions in aqueous solutions of o-fructose, o-glucose and sucrose. Carbohydro Res. 81, 213. Mitchell, A. J. (1968). Spectra in the O-H stretching region of mono- and oligo-saccharides at low temperatures. Aust. J. Chern. 21, 1257. Mulazzi, E., Piseri, L., Pollini, I., and Tubino, R. (1980). Raman scattering in resonance with the electronic transitions of the solid amylose/iodine complex. In Proc. VIlth Int. ConI Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 634-635. Novak, A. (1979). Vibrational spectroscopy of hydrogen bonded systems. In T. M. Theophanides, Ed., Infrared and Raman Spectroscopy of Biological Molecules, Reidel, Dordrecht, Holland, pp. 279-303. Parker, F. S. (1971). Applications of Infrared Spectroscopy in Biochemistry, Biology and Medicine, Plenum, New York. Rees, D. A. (1977). Polysaccharide Shapes, Wiley, New York. She, C. Y, Dinh, N. D., and Tu, A. T. (1974). Laser Raman scattering of glucosamine, N-acetyl-glucosanllne, and glucuronic acid. Biochim. Biophys. Acta 372, 345. Susi, H., and Ard, J. S. (1974). Laser-Raman spectra of lactose. Carbohydr. Res. 37, 351. Tasumi, M. (1972). Resonance Raman spectrum of the starch-iodine complex. Chern. Lett. 1, 75. Tipson, R. S. (1968). Infrared Spectroscopy of Carbohydrates, a Review of Literature, National Bureau of Standards Monograph //0, U.S. Gov!. Printing Office, Washington, D. C. Tomimatsu, Y, Scherer, J. R., Yeh, Y, and Feeney, R. E. (1976). Raman spectra of a solid antifreeze glycoprotein and its liquid and frozen aqueous solutions. J. Bioi. Chern. 251, 2290. Tu, A. T., Dinh, N. D., She, C. Y., and Maxwell, J. (1977). Laser Raman scattering of a- and ,B-methyl-o-glucosides and hyaluronic acid. Stud. Biophys. 63, 115. Tu, A. T., Lee, J., and Lee, Y. C. (1978). Laser-Raman spectroscopic study of carbohydrates: 1-thio-,B-o-hexopyranosides. Carbohydr. Res. 67, 295. Tu, A. T., Lee, J., and Milanovich, F. P. (1979). Laser-Raman spectroscopic study of cyclohexaamylose and related compounds: Spectral analysis and structural implications. Carbohydr. Res. 76, 239. Ueda. H., and Nagai, T. (1981). Solid-state nuclear magnetic resonance spectroscopy and Raman spectroscopy of the inclusion compound of Tolbutamide with ,B-cyclodextrin. Chern. Pharm. Bull. 29, 2710. Vasko, P. D., Blackwell, J., and Koenig, J. L. (1971). Infrared and Raman spectroscopy of carbohydrates. Part I: Identification of O-H and C-H-related vibrational modes for o-glucose, maltose, cellobiose, and dextran by deuterium-substitution methods. Carbohydr. Res. 19, 297. Vasko. P. D., Blackwell, J., and Koenig, J. L. (1972). Infrared and Raman spectroscopy of carbohydrates. Part II: Normal coordinate analysis of a-o-glucose. Carbohydr. Res. 23, 407. Walrafen, G. E. (1964). Raman spectral studies of water structure. J. Chern. Phys. 40, 3249.
References
255
Walrafen, G. E. (1970). Raman spectral studies of the effects of perchlorate ion on water structure. 1. Chem. Pilys. 52, 4176. Walrafen, G. E. (1971). Raman spectral studies of the effects of solutes and pressure on water structure. J. Cilem. Pilys. 55, 768. Walrafen, G. E., and Abede, M. (1978). Raman studies of the bending and vibrational bands from water and ice VI to - 12 kbar at 32°e. J. Cilem. Pilys. 68,4694. Williams, R. M., and Atalla, R. H. (1981). Interactions of group II cations and borate anions with nonionic saccharides. Am. Cilem. Soc. Symp. Ser. 15C,317-330.
CHAPTER
Carotenoids and Flavins
Carotenoids and flavins are structurally unrelated compounds but both are common, naturally occurring pigments. Carotenoids are responsible for a reddish orange color, whereas flavins give a yellow color. Both compounds give rise to resonance Raman spectra.
1.
CAROTENOIDS
Carotenoids are orange-to-reddish-colored pigments widely distributed in both the plant and animal kingdoms. The best examples of carotenoids are the pigments of the tomato (lycopene) and the carrot ((X- and fJ-carotene). Vitamin A is closely related chemically to fJ-carotene; they are both polyenes and derivatives of isoprene. Resonance Raman spectroscopy is a useful analytical tool in the study of carotenoids, since these compounds give resonance-enhanced Raman spectra in very dilute solution. Since the absorption maxima of carotenoids are
Carotenoids
257
different from those of other biological pigments, we can selectively obtain the resonance Raman spectra without interference from other compounds present in living tissues. Thus carotenoid Raman spectra have been obtained from live carrot, tomato fruit, bottled tomato sauce, and canned carrot juice! The reason one can selectively obtain carotenoid spectra even in a mixture is the polyenic structure (high degree of unsaturation) of these compounds. Resonance Raman spectra of carotenoids are characterized by intense bands due to C=C (1500-1600 em-I) and C-C (1100-1200 em-I) stretching modes. Some of the C-C stretching modes are coupled with C-C- H bands. Extensive theoretical as well as experimental studies have been made on C=C stretching modes of carotenoids using resonance Raman spectroscopy (Salares et al., 1977b; Inagaki et al., 1974; Warshel and Dauber, 1977; Huong, 1978; Rimai et al., 1971, 1973; Dallinger et al., 1979; Jensen, 1980). An interesting correlation has been found between C=C stretching vibration frequencies and the absorption maxima of carotenoids. When Pc=c is plotted against I/A max , a nearly linear relation is obtained. This is because as the polyene chain length is increased, it increases delocalization of 7T-electrons and thus decreases the force constants of the C=C bonds, and a bathochromic shift takes place (Heyde et al., 1971; Rimai et al., 1973). This correlation does not apply to cyclic-polyene compounds. The excited triplet state of all-transfJ-carotene shows six transient Raman bands as determined by time-resolved resonance Raman spectroscopy. The spectra suggest that the C=C bond order is decreased, and the molecule may be substantially twisted in the triplet state (Jensen et al., 1980). Raman spectra of carotenoids can also be obtained by a special technique called coherent anti-stokes Raman scattering (CARS) (Carreira et al., 1977). CARS is discussed in Section 3.2.1 of Chapter 2. Trans and cis types of fJ-carotenes can be differentiated by resonance Raman spectroscopy (Saito et al., 1980). 1.1. CAROTENOIDS FROM PLANTS, MICROORGANISMS, AND BLOOD
Carotenoids are formed in higher plants, blue-green algae, and photosynthetic bacteria, and are distributed in both the animal and plant kingdoms. Since animals cannot synthesize carotenoids, these compounds are taken in the diet; carotenoids can be found in the liver, blood, and visual pigments of the retina. Carotenoid pigment can be detected in intact plant tissue by the resonance Raman technique. A classical example is the resonance Raman spectra of lycopene and fJ-carotene, obtained by examining live carrot root and tomato fruit by Gill et al. (1970). Extremely high quality spectra were obtained without much interference from other compounds present in the tissue (Figure 9.1). Similarly, fJ-carotene, Iycopene, and xanthophyll pigments were detected in blood plasma. The resonance Raman spectra are shown in Figure 9.2; very little interference from plasma is noticed (Rein et al., 1976).
LIVE
1527
CARROT
~ 457
ROOT
11,59
965 1005 1 .
I
CANNED CARROT JUICE
~
A
1160 115B
1179 1199
~21,7'
1
B
1527
A
P-CAROTENE
n-HEXANE 1158
~~ANE
---
1,600
1,500
1,400
c
, "
_-,--
1,200
1,300
1,100
1.000
900
em-I
LIVE
15225 I
TOIIIATO FRUIT
1158
~
119~' t001 I
I
-~
9~3
~
o
BOTTLED TOMATO SAUCE 1156 I
1522 I
~
~
E
LYCOPENE IN n-HEXANE
1516 I
1158
A HEXANEI~04 ~E ~~~ 1,500
1,459
1,200
1,100
F
1,000
em-'
FIGURE 9.1. Resonance Raman spectra of (A) live carrot root, (B) canned carrot juice, (C) fJ-carotene in n-hexane, (D) live tomato fruit, (E) bottled tomato sauce, (F) lycopene in n-hexane. These are good examples that carotenoids ill situ can be examined by resonance Raman spectroscopy. The figure was reproduced from Gill et al. (\979) by permission of the copyright owner, Macmillan Journals Ltd.
j 'lI&::a
Carotenoids
259
1523 DRIED PETROLEUM ETHER EXTRACT of DENATURED BLOOD PLASMA io CHLOROFORM
1157
t P-CAROTENE io CHLOROFORM (eluted from alumIna with he ...one-e'hanol)
1006
1500
1400
1500
1200
900
em-I
FIGURE 9.2. Resonance Raman spectra of blood plasma and ,a-carotene. The figure was reproduced from Rein et aI. (1976) by permission of the copyright owner.
In oceanographic studies, it is important to estimate the phytoplankton concentration and the total organic carbon in seawater. The total carotenoid concentration in phytoplankton is believed to be a good measure of both of these quantities. This analysis can be accomplished by the use of resonance Raman spectroscopy on the carotenoids in acetone extracts of marine plankton (Hoskins and Alexander, 1977). The carotenoid content of tobacco leaves can
260
Carotenoids and Flavins
be measured by the resonance Raman technique. When carotenoid scattering is selectively enhanced with 488- or 5l4.5-nm laser excitation, the presence of chlorophyll does not interfere with the analysis. The main carotenoids in tobacco leaves are fJ-carotene and lutein. The band frequencies for fJ-carotene and lutein are identical because of their similar polyene structure. As a result, the carotenoids must be separated from each other for quantitative analysis. Using this analytical method it was found that carotenoid content decreases during the curing process (Forrest and Vilcins, 1979). Spheroidene is a carotenoid structurally closely related to fJ-carotene (Figure 9.3). Some carotenoids, including spheroidenes, attach to photosynthetic reaction centers of different species of photosynthetic bacteria. Most of the spheroidenes have all-trans conformation. Spheroidene from Rhodopseudomonas spheroides is an unusual carotenoid because it has cis conformation. Evidence of this was obtained from a comparison of resonance Raman spectra of 15, lY-cis-fJ-carotene (Lutz et al., 1978). Resonance Raman spectra of the carotenoid~reaction-center complex indicate that the carotenoid is bound as a cis isomer (Agalidis et al., 1980). In chromatophores (pigmentary cells) from R. spheroides, an artificially generated diffusion potential causes small shifts in the position of the carotenoid absorption. With 473-nm excitation, the intensities of the two most prominent resonance Raman lines at 1526 and 1157 cm ~ 1 respond very differently to small shifts in the absorption maxima. Thus the intensity ratio of these two lines is a sensitive probe for absorption shifts or for membrane potential (Koyama et al., 1979). Simple lipids and phospholipids do not give resonance Raman spectra. However, the presence of even a small amount of carotenoids in lipid structures can give strong resonance Raman spectra. It has been found that human blood platelets contain carotenoids. This finding was made using the resonance Raman spectra of platelets illuminated by a 488-nm laser beam. The characteristic vibrational modes of =C-C= and -C=C- bonds were detected at 1160 and 1530 cm ~ 1. The usefulness of resonance Raman in detecting a low concentration of carotenoids in membrane was well demonstrated (Aslanian et al., 1979). When isolated platelet membranes are used, the resonance Raman peak of carotenoids is greatly enhanced, suggesting that the carotenoids are strongly bound to the platelet membrane (Aslanian et al., 1980).
YVVVV~ OCH 3
FIGURE 9.3.
Structure of spheroidene.
261
Carotenoids
Phycocyanin is the light-harvesting pigment present in blue-green bacteria. It is a conjugated protein containing phycocyanobilin. There is evidence that carotenoids interact with phycocyanin in biomembranes. Raman intensity decay constants at 1525 cm- I (C=C vibration) and at 1160 cm- I (=C-C= vibration mode) are different for the cyanobacterium Anacystis nidulans containing phycocyanin and without phycocyanin. Since the Raman spectrum is obtained by resonance enhancement of membrane-bound carotenoids, the difference in the decay constants suggests that there is an interaction between carotenoids and phycocyanin in the biomembrane (Szalontai and Csatorday, 1980). 1.2.
LOBSTER-SHELL CAROTENOPROTEINS
The shell of the lobster (Homarus americanus) contains two types of proteins that are responsible for the color of the shell. Blue crustacyanins absorb in the 600-nm region, and a yellow protein absorbs at 410 nm; both contain astaxanthin as the chromophore (Figure 9.4). The color of lobster shells appears blue, yellow, green, or mottled depending on the relative amount of yellow and blue proteins. The absorption maximum of free astaxanthin is at 480 nm, the blue crustacyanins absorb at 600 nm, and a yellow protein at 410 nm. Two types of pigment proteins cause large shifts in the absorption of the free chromophore. The shift in the absorption spectrum for the yellow protein is attributed to chromophore-chromophore rather than to protein-chromophore interactions. When the astaxanthin aggregates or binds to the yellow protein, the absorption maximum shifts. The resonance Raman spectra indicate that the main change occurs in the C=C stretching vibration bands, suggesting that a large perturbation of the electronic excited state takes place, whereas only a minimal perturbation of the ground state occurs (Salares et al., 1977a). The presence of an astaxanthin-bearing pigment in situ can be detected from resonance Raman spectrum (Nelson and Carey, 1981). Another protein, ovoverdin, occurs in lobster eggs and has two absorption maxima, one at 480 nm, essentially the same as that of free astaxanthin, and
o
"
OH
'Y'V'Vo/~ HO
II
o FIGURE 9.4.
Structure of astaxanthin. It is very similar to the structure of ,B-carotene.
262
Carotenolds and Flavlns
the other at 640 nm, resembling the absorption found in the crustacyanins. The Raman spectrum of intact eggs is identical to that of isolated ovoverdin, suggesting that the carotenoid sites are unperturbed by protein isolation (Salares et al., 1979). 1.3.
EXCITATION PROFILES
Excitation profiles refer to the vanatlOn of Raman intensity with excitmg wavelength. It has been reported that excitation profiles can be used as a probe of vibrational mode in absorption spectra (Gaber et al., 1974; Inagaki et al., 1974; Rimai et al., 1970). In order to examine these reports, Salares et al. (1976) selected astaxanthin. It was found that the spacing between 0-0, 0-1, 0-2. .. transitions of astaxanthin cannot be resolved from the excitation profiles at room temperature (Salares et al., 1976). Interpretation of the excitation profile of the carotenoprotein ovorubin led to estimates of the elongation of the C=C and shrinkage of the C-C bonds in the resonant excited state (Clark et al., 1980). Raman intensities are closely related to the absorption profile of a molecule, and shifts in absorption bands are reflected in increases or decreases in Raman intensities. For example, changes in membrane potential change the absorption spectrum of carotenoids. Thus using this property, one can use Raman spe,troscopy to see the change in the membrane potential. The resonance Raman spectrum of intact Halobacterium halobium cells is dominated by the 1507-cm-] (C=C stretching) and 1150-cm-] (C-C stretching) bands, which originate from carotenoids. Raman intensity changes of these bands are correlated with the red shift of the carotenoid absorption, which is caused by an increase in membrane potential (Szalontai, 1981).
2.
FLAVINS
Flavins are yellow pigments and vitamins; they are constituents of coenzymes or prosthetic groups of many flavoenzymes. Flavins also give resonanceenhanced Raman spectra when the sample is illuminated with an excitation line within the absorption band. 2.1.
FLAVINS AND FLAVOENZYMES
Both flavinadeninedinucleotide (FAD) and flavinmononucleotide (FMN) give similar resonance Raman spectra, indicating that resonance enhancement is limited to the isoalloxazine ring. The only significant changes in the resonance Raman spectrum of FAD on binding to fatty acyl-CoA occurs in the 1250-cm -1 region, which is associated with the bending motion of N-3 (Benecky et al., 1979). Resonance Raman spectra of riboflavins are similar to those of glucose oxidase (Nishina et al.. 1978).
J
Flavins
263
Kitagawa et aI. (l979a) also concluded that the 1252-cm- 1 band involves C-2 and N-3 ring vibration strongly coupled with the N(3)-H bending mode. Moreover, they concluded that this band can be used as an indicator for interaction involving a hydrogen bond between N(3)- H and a protein. It is known that oxidized flavins form colored complexes with electron donors such as phenols, purines, and indols, giving a characteristic chargetransfer band. Raman spectra of a riboflavin-phenol charge-transfer complex were examined (Nishina et aI., 1980a). The 1585- and 1550-cm - I bands were found to be particularly enhanced. These bands involve the vibrational displacement of the N-5 and C-4a atoms of isoalloxazine (Figure 9.5). The ring I mode is little enhanced, whereas ring III modes are moderately enhanced. From this it is concluded that the 7T-electrons of the benzene ring of phenol interact as a whole with the isoalloxazine ring III; two rings are arranged in parallel so as to produce maximum 7T-7T interaction. The site of nucleophilic attack is probably at N-5 and C-4a. The band is also assigned to a ring mode localized in the N-3 region including N-H in-plane bending (Dutta et aI., 1977). However, more-recent work indicates that many Raman bands are associated with highly delocalized aromatic-framework vibrations rather than each being associated with well-defined localized vibration. When positions 7 and 8 (ring I) of the isoalloxazine are modified, significant shifts occur in the bands previously assigned to rings II and III (Schopfer and Morris, 1980). This probably is best explained from the fact that the isoalloxazine contains a substantial contribution from a quinonoid form, as shown in Figure 9.5 (Schopfer et aI., 1981). Because of the similarity in the spectra of FAD and enzyme-bound FAD, it can be concluded that no covalent bonding is involved between the enzyme and flavin (Benecky et aI., 1979). The line at 1620 cm - I is reported to be useful in determining the redox state of flavin (Nishina et aI., 1980b). R 9
110
1
H3C~~ 9 N~N~O 7IIn
H3C ~
6a
6
N~. 5
ml3
NH 4
o
Isoalloxazine
R
R
I
X~NyN'f0 .
~~NH N
FIGURE 9.5.
I
• X~NJCN~'~r"'O
o"
~ I N
NH
0"
Benzenold form Quinonoid form Structure of isoalloxazine ring of FAD and FMN (top). Benzenoid and quinonoid
forms of isoalloxazine (Jower). X renresent.s - N{("H .. '-
_("H..f
l\.Tl-J
nr
_<::H
264
Carotenoids and Flavins
Under natural conditions, flavin binds to protein to form a variety of f1avoproteins. It is important to know how flavin interacts with protein in these f1avoproteins. Ring I is probably exposed to the solvent, as little vibrationalfrequency change due to ring I can be observed in flavodoxin. Instead N-5 in ring II, C(4)=O, N(3)-H, and C(2)=O in ring III (Figure 9.5) vibrational modes shift their frequencies in glucose oxidase protein, suggesting the importance of ring III and possibly ring II in binding to proteins. For flavodoxin, binding of flavin to the protein is very likely at C(2)=O and N-5. In lumazine
po,; I
CH 2 I
9
(CH H)3 CH2
A.
CH
3(rr~~:J~O
CH3
0..
I
4.... 3N
5""'-'::;
N
I
Ag_
CH2 [CHOH] 3CH20H
B.
H3CXX~!fYOI
H C::::---... 3
N
I""'-:;::N
L
H20 Ag-O
O-Ag
I
!
C.
N~'C-N~
I
II
........C~ ....... C-N RI N
5
CH
I
R2
FIGURE 9.6. (A) Structure of Ag(I)-flavin phosphate based on Raman spectroscopy proposed by Benecky el aI. (1980). (8) Structure of Ag(I)-flavin proposed by Bamberg and Hemmerich (196\). (C) Structure of Ag(l)-guanosine and related compounds proposed by Tu and Reinosa (1966). Note the common penta coordination structure involving Ag-O-C-C-N.
! j
Flavins
265
protein, the frequencies of the C(4)=0 and C(2)=0 bonds are observed, indicating that the chromophore sits at the surface of the protein; the binding is probably mainly by its ribityl side chain (Irwin et aI., 1980). It seems that each flavoprotein has a specific way of binding flavin to proteins. Raman spectroscopic analysis indicates that the isoalloxazine ring of riboflavin interacts with the histidine NH of the apoprotein (Zak et aI., 1977). It is known that flavin compounds interact with Ag(I) and that upon complexing, the lines responsible for the C(4)=N(5) stretching vibration and the N(3)-H bending vibration change. It is, therefore, proposed that the metal complexation occurs at N-5 and 0-4 (Figure 9.6). It is of interest that the metal complex tends to form a five-membered ring as shown in Figure 9.6 (Benecky et aI., 1980). When a ,flavoenyzme combines with a substrate, the intermediate shows a broad absorption band at longer wavelength than that of the enzyme. It has been suggested that the intermediate is a charge-transfer complex of the oxidized or the reduced enzyme with a substrate or product. In order to confirm this, "old yellow enzyme," which contains 1 mol of FMN, was interacted with phenol. (In earlier days, FMN-containing flavoprotein was referred to as the "old yellow enzyme," whereas the FAD-containing flavoprotein was referred to as the "new yellow enzyme.") The enzyme-phenol intermediate complex was excited at the long-wavelength band at 568.2 nm. If the complex is the charge-transfer type, the Raman spectrum should contain charge-transfer bands. The enzyme-phenol complex indeed showed many Raman bands not observed for old yellow enzyme and phenol. The 1588-cm - 1 band was also selectively enhanced. The same conclusion was obtained for the 1586 cm -1 band-which is coupled strongly to the charge transfer electronic transition-in the acetoacetyl CoA-fatty acid CoA dehydrogenase complex. This band is due to C(4)-N(5) vibration of isoalloxazine. This suggests that C(4)-N(5) is the most likely site for the charge-transfer interaction (Kitagawa et aI., 1979b; Nishina et aI., 1980a, b; Schmidt et aI., 1981). Recently a new technique, CARS, has been successfully applied to biological compounds including flavin and flavoprotein (see Section 3.2.1, Chapter 2). By this technique, good Raman spectra are obtained without the fluorescence background (Dutta et aI., 1977; Dutta et aI., 1978; Dutta and Spiro, 1980). 2.2.
FLAVOCYTOCHROME cm
Flavocytochrome C m is an example of a multiple-redox enzyme; the enzyme contains two mesohemes that attach to one subunit, and one FAD that attaches to one subunit and one FAD that attaches to another subunit. The enzyme is isolated from purple photosynthetic bacteria, Chromatium vinosum, and has a molecular weight of 72,000. The important question is whether there is a direct heme-flavin interaction during the electron-transport reaction. Resonance Raman spectra of both oxidized and reduced enzymes were obtained by laser excitation at 514.5 nm, which is within the absorption spectrum
266
Carotenolds and Flavins
H
y~; FIGURE 9.7.
Structure of proflavin.
of the enzyme because of one of heme electronic transitions. Raman spectra of both oxidized and reduced enzymes are consistent with typical heme vibrational modes, and there is no indication of a flavin vibrational-mode contribution. From this evidence, it is concluded that there is no direct heme-flavin interaction. If there is a heme-flavin interaction, it must be intermediated through protein (Ondrias et al., 1980). However, different conclusions were reached by Kitagawa et al. (1980), who reported that both isoalloxazine and heme Raman bands are observable, and these bands can be used to monitor the oxidation states of the flavin and heme separately. Moreover, they postulated that in flavocytochrome c-552, an electron transfer occurs between flavin and heme on reduction. 2.3.
PROFLAVIN AND OTHER FLAVIN COMPOUNDS
Proflavin has a structure somewhat similar to that of flavin (Figure 9.7). Proflavin exhibits absorbance deviations from Beer's law, suggesting stacking of the molecules. The Raman laser temperature-jump technique was used to study the solvent effect on proflavin stacking. The results suggest that the solvent effect on dye stacking is determined by specific dye-solvent interactions (Dewey et al., 1979). Lumiflavin has a methyl group at position 10 of the isoalloxazine ring; its Raman spectrum is considerably different from that of riboflavin in the region of 1200-1300 cm -I (Nishina et al., 1980c). Detailed assignments of the Raman lines for 10-methylisoalloxazine (Iumiflavin) have been made using the normal mode calculation (Bowman and Spiro, 1981). Flavocytochrome c, containing both flavin and heme c, can be found in photosynthetic sulfur bacteria. From the same resonance Raman spectrum, it is possible to identify the oxidation states of the flavin and heme separately (Kitagawa et al., 1980). REFERENCES Agalidis, I., Lutz, M., and Reiss-Husson, F. (1980). Binding of carotenoids on reaction centers from Rhodopseudomonas sphaeroides R 26. Biochim. Biophys. Acta 589, 264. Aslanian, D., Vainer, H., Bolard, 1., Guesdon, I.-P., and Balkanski, M (1979). Resonance Raman spectrometric study of human blood platelets. FEBS Lett. 101, 39. Aslanian, D., Vainer, H., Lautie, A., and Guesdon, I.-P. (1980). Further Raman spectrometric studies on human platelet and isolated platelet membranes. IRCS Med. Sci. 8, 421. Bamberg, P., and Hemmerich, P. (1961). Farbe und Konstitution der Isoalloxazin-SilberKomplexe. He/v. Chim. Acta 44, 1001.
References
267
Benecky, M., Li, T. Y, Schmidt, 1., Frerman, F., Watters, K. L., and McFarland, 1. (1979). Resonance Raman study of f1avins and the flavoprotein fatty acyl coenzyme A dehydrogenase. Biochemistry 18, 3471. Benecky, M., Yu, T.-J., Watters, K. L., and McFarland, J. T. (1980). Metal-flavin complexation, a resonance Raman investigation. Biochim. Biophys. Acta 626, 197. Bowman, D. W., and Spiro, T. G. (1981). Normal mode analysis of lumiflavin and interpretation of resonance Raman spectra of f1avoproteins. Biochemistry 20, 3313. Carreira. L. A., Maguire, T. c., and Malloy, Jr., T. B. (1977). Excitation profiles of the coherent anti-Stokes resonance Raman spectrum of ,8-carotene. J. Chern. Phys. 66, 2621. Clark, R. 1. H., D'Urso, N. R., and Zagalsky, P. F. (1980). Excitation profiles, absorption and resonance Raman spectra of the carotenoprotein ovorubin, and a resonance Raman study of some other astaxanthin proteins. J. Am. Chern. Soc. 102,6693. Dallinger, R. F, Guanci, Jr., 1. 1., Woodruff, W. H., and Rodgers, M. A 1. (1979). Vibrational spectroscopy of the electronically excited state: pulse radiolysis/time-resolved resonance Raman study of triplet ,8-carotene. J. Am. Chern. Soc. 101, 1355. Dewey, T. G., Raymond, D. A, and Turner, D. H. (1979). Laser temperature jump study of solvent effects on proflavin stacking. J. Am. Chern. Soc. 101,5822. Dutta, P. K., and Spiro, T. G. (1980). Resonance coherent anti-Stokes Raman scattering spectra of oxidized and semiquinone forms of Clostridum MP f1avodoxin. Biochemistry 19, 1590. Dutta, P. K., Nestor, J. R., and Spiro, T. G. (1977). Resonance coherent anti-Stokes Raman scattering spectra of fluorescent biological chromophores: vibrational evidence for hydrogen bonding of flavin to glucose oxidase and for rapid solvent exchange. Proc. Nat. Acad. Sci. 74, 4146. Dutta, P. K., Nestor, J., and Spiro, T. G. (1978). Resonance coherent anti-Stokes Raman scattering (CARS) spectra of flavin adenine dinucleotide, riboflavin binding protein and glucose oxidase. Biochem. Biophys. Res. Commun. 83, 209. Forrest, G., and Vilcins, G. (1979). Determination of tobacco carotenoids by resonance Raman spectroscopy. J. Agric. Chern. 27, 609. Gaber, B. P., Miskowski, V., and Spiro, T. G. (1974). Resonance Raman scattering from iron(II1)and copper (II)-transferrin and an iron(III) model compound. A spectroscopic interpretation of the transferrin binding site. J. Am. Chern. Soc. 96, 6868. Gill, D., Kilponen, R. G., and Rimai, L. (1970). Resonance Raman scattering of laser radiation by vibrational modes of carotenoid pigment molecules in intact plant tissues. Nature 227, 743. Heyde, M. E., Gill, D., Kilponen, R. G., and Rimai, L. (1971). Raman spectra of Schiff bases of retinal (models of visual photoreceptors). J. Am. Chern. Soc. 93, 6776. Hoskins, L. c., and Alexander, V. (1977). Determination of carotenoid concentrations in marine phytoplankton by resonance Raman spectroscopy. Anal. Chern. 49, 695. Huong, P. V. (1978). Spectres Raman de resonance des carotenoides: torulene et )I-Carotene. C. R. A cad. Sc. Paris 286, 25. Inagaki, F., Tasurni, M., and Miyazawa, T. (1974). Excitation profile of the resonance Raman effect of f3-carotene. J. Mol. Spectrosc. SO, 286. Irwin, R. M., Visser, A J. W. G., Lee, J., and Carreira, L. A (1980). Protein-ligand interactions in lumazine protein and in desulfovibrioflavodoxins from resonance coherent anti-Stokes Raman spectra. Biochemistry 19. 4639. Jensen, N. H. (1980). Photosynthetic pigments and model compounds studied by pulse radiolysis. Risoe Rep.. 415, 93 pp. Risoe National Laboratory, Roskilde, Denmark. Jensen, N.-H., Wilbrandt, R., Pagsberg, P. B., Sillesen, A H., and Hansen, K. B. (1980). Time-resolved resonance Raman spectroscopy: the excited triplet state of all-trans-f3-carotene. J. Am. Chern. Soc. 102,7441.
268
Carotenoids and Flavins
J(jtagawa, T, Nishina, Y., Kyogoku, Y., Yamano, T, Ohishi, N., Takai-Suzuki, A, and Yagi, K (I979a). Resonance Raman spectra of carbon-13-and nitrogen-15-labeled riboflavin bound to egg-white f1avoproteins. Biochemistry 18, 1804. J(jtagawa, T, Nishina, Y., Shiga, K., Watari, H., Matsumura, Y., and Yamano, T (I 979b). Resonance Raman evidence for charge-transfer interactions of phenols with the flavin mononucleotide of old yellow enzyme. J. Am. Chern. Soc. 101,3376. J(jtagawa, T., Fukumori, Y., and Yamanaka. T (1980). Resonance Raman evidence for intramolecular electron transport from flavin to heme in f1avocytochrome c-552 and nature of chromophoric interactions. Biochemistry 19, 5721. Koyama, Y., Long, R. A, Martin, W. G., and Carey, P. R. (1979). The resonance Raman spectrum of carotenoids as an intrinsic probe for membrane potential. Oscillatory changes in the spectrum of neurosporene in the chromatophores of Rhodopseudomonas spheroides. Biochim. Biophys. Acta 548, 153. Lutz, M., Agalidis, 1., Hervo, G., Cogdell, R. J., and Reiss-Husson, F. (1978). On the state of carotenoids bound to reaction centers of photosynthetic bacteria: a resonance Raman study. Biochim. Biophys. Acta 503, 287. Nelson, W. H., and Carey, P. R. (1981). Infrared excited resonance Raman spectra of lobster shell pigments in situ. J. Raman Spectrosc. 11, 326. Nishina, Y., J(jtagawa, T, Shiga, K., Horiike, K., Matsumura, Y., Watari, H., and Yamano, T (1978). Resonance Raman spectra of riboflavin and its derivatives in the bound state with egg riboflavin binding proteins. J. Biochem. (Tokyo) 84, 925. Nishina, Y., J(jtagawa, T, Shiga, K., Watari, H., and Tanabim, T (1980a). Resonance Raman study of f1avoenzyme-inhibitor charge-transfer interactions. Old yellow enzyme-phenol complexes. J. Biochem. 87, 831. Nishina, Y, Shiga, K, Horike, K, Tojo, H., Kasai, S., Matsui, K, Watari, H., and Yamano, T (1980b). Resonance Raman spectra of semiquinone forms of f1avins bound to riboflavin binding protein. J. Biochem. 88, 411. Nishina, Y., Shiga, K., Horike, K., Tojo, H., Kasai, S., Yanase, K., Matsui, K., Watari, H., and Tamano, T (1980c). Vibrational modes of flavin bound to riboflavin binding protein from egg white. J. Biochem. 88, 403. Ondrias, M. R., Findsen, E. W., Leroi, G. E., and Babcock, G. T (1980). Chromophore interactions in f1avocytochrome c552: a resonance Raman investigation. Biochemistry 19, 1723. Rein, A J., Saperstein, D.O., Pines, S. H., and Radlick, P. C. (1976). Blood plasma investigations by resonance Raman spectroscopy: detection of carotenoid pigments. Experientia 32, 1352. Rimai, L., J(jlponen, R. G., and Gill, D. (1970). Excitation profiles of laser Raman spectra in the resonance region of two carotenoid pigments in solution. J. Am. Chem. Soc. 92, 3824. Rimai, L., Gill, D., and Parsons, J. L. (1971). Raman spectra of dilute solutions of some stereoisomers of vitamin A type molecules. J. Am. Chern. Soc. 93, 1353. Rimai, L., Heyde, M. E., and Gill, D. (1973). Vibrational spectra of some carotenoids and related linear polyenes. A Raman spectroscopic study. J. Am. Chern. Soc. 95, 4493. Saito, S., Harada, 1., Tasumi, M., and Eugster, C. H. (1980). Resonance Raman spectra of all-trans and 15, IY-cis-p, p-carotenes. Chern. Lett. 1045. Salares, V. R., Mendelsohn, R., Carey, P. R., and Bernstein, H. J. (1976). Correlation between the absorption spectra and resonance Raman excitation profiles of astaxanthin'a . J. Phys. Chern. SO, 1137. Salares, V. R., Young, N. M., Bernstein, H. 1., and Carey, P. R. (1 977a). Resonance Raman spectra of lobster shell carotenoproteins and a model astaxanthin aggregate. A possible photobiological function (or the yellow protein. Biochemistry 16, 4751. Salares, V. R., Young, N. M., Carey, P. R., and Bernstein, H. 1. (1977b). Excited state (exciton) interactions in polyene aggregates. J. Raman Spectrosc. 6, 282.
References
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Salares, V. R., Young, N. M., Bernstein, H. J., and Carey, P. R. (1979). Mechanisms of spectral shifts in lobster carotenoproteins. The resonance Raman spectra of ovoverdin and the crustacyanins. Biochim. Biophys. Acta 576, 176. Schmidt, 1., Reinsch, 1., and McFarland, 1. T. (1981). Mechanistic studies on fatty acyl-CoA dehydrogenase. 1. BioI. Chem. 256, 11667. Schopfer, L. M., and Morris, M. D. (1980). Resonance Raman spectra of Oavin derivatives containing chemical modifications in positions 7 and 8 of the isoalloxazine ring. Biochemisoy 19,4932. Schopfer, L. M., Haushalter, 1. P., Smith, M., Milad, M., and Morris, M. D. (1981). Resonance Raman spectra for Oavin derivatives modified in the 8 position. Biochernistty 20, 6734. Szalontai, B. (1981). Light induced membrane potential changes in Halobacterium halobium observed with high time resolution by resonance Raman spectroscopy. Biochem. Biophys. Res. Commun. 100, 1126. Szalontai, B., and Csatorday, K. (1980). A resonance Raman study of carotenoid susceptibility to photooxidation in the cyanobacterium Anacystis nidulans. 1. Mol. Struct. 60, 269. Tu, A. T., and Reinosa, J. A. (1966). The interaction of silver ion with guanosine, guanosine monophosphate, and related compounds. Determination of possible sites of complexing. Biochemistry 5, 3375. Warshel, A., and Dauber, P. (1977). Calculations of resonance Raman spectra of conjugated molecules. 1. Chern. Phys. 66, 5477. Zak, Z., Malkiewicz, I., and Pytasz, G. (1977). RiboOavin-apoprotein interactions studied by infrared and Raman spectroscopy. In Flavins Flavoproteins: Physicochemisoy Properties Function, Proc. Int. Meet., 1976, W. Ostrowski, Ed., pp. 39-50.
CHAPTER
o ~
Visual Pigments and Bacterial Rhodopsin
Both rhodopsin (visual pigment) and bacterial rhodopsin absorb light, undergo a series of complicated photo-induced reactions, and eventually regenerate the original compounds. They have totally different biological functions. Rhodopsin is present in the retina where it converts light energy into nervous impulses. Bacterial rhodopsin is found in certain bacterial membranes. It converts light energy into chemical energy as ATP. Yet they both have very similar prosthetic groups and photoactivated reactions.
1.
VISUAL PIGMENTS
The retina is the place in the eye where light is transformed into nerve impulses, which are eventually presented to the brain as a rainbow of images instead of mere shadows (Figure 10.1). The visual pigment responsible for this action is rhodopsin. Rhodopsin is located in vertebrate retinal rod cells; it is a
Rods B Cones
..
...• co ..
l!iil'
Location of Rhodopsin
in Brain
FIGURE 10.1.
Diagram of eye and retina.
272
Visual Pigments and Bacterial Rhodopsin
conjugated protein. The chromophore group is ll-cis-retinal, which is connected to the glycoprotein opsin by a Schiff-base linkage. The absorption maximum of retinal occurs near 370 nm in solution and shifts to longer wavelengths (bathochromic shift) of 500-575 nm when retinal combines with opsin. In the human eye, three different cone cells absorb visible light at maxima of 440, 535, and 575 nm. Thus these cells discriminate among the three primary colors, and the resulting pattern of nerve transmissions is interpreted in the brain as full color vision. A fascinating fact is that all three different cells depend ultimately on the same chromophore: ll-cis-retinal. The ll-cis-retinal is eventually converted into all-trans-retinal as shown in Figure 10.2. Actually the cis form is highly strained, and with light it becomes the unstrained trans form. ll-cis-retinal is regenerated from all-trans-retinal by the enzymatic actions of retinol dyhydrogenase and isomerase. The conversion of ll-cis-retinal to all-trans-retinal involves a series of short-lived intermediates as shown here: 6 X 1O- IZ s
Rhodopsin (500 nm)
. prelumirhodopsin (bathorhodopsin) (545 nm)
5 X 10- 5 S 5 X 10- 8 S - - - - lumirhodopsin . metarhodopsin I (497 nm) (480 nm) - - - - . metarhodopsin II (380 nm)
. all-trans-retinal (387+nm) opsin (apoprotein)
Vitamin A has close structural similarities to all-trans-retinal. At C-15, the former has a CHzOH group, whereas the latter has a CHO group. Also closely related to ll-cis-retinal is 9-cis-retinal, or isorhodopsin. The difference between these two compounds lies in their binding sites to the opsin molecule. 1.1.
VIBRATIONAL ASSIGNMENTS
Rhodopsin and related compounds have been extensively studied with Raman spectroscopy by many investigators. Hence the assignment of various vibrational bands is fairly complete (Table 10.1). The most intense line for ll-cis-retinal is at 1577 cm- I and is identified as the ethylenic mode (Gill et aI., 1971). The most prominent peak in rhodopsin is a band at 1545 cm - I that has an extremely high intensity. This band originates from a C=C stretching vibration, and the frequency varies, depending on the compounds (Table 10.1). The frequency of the polyene stretching vibration has an inverse linear relationship to the wavelength of the absorption maximum.
Visual Pigments CH3
11
9~
2
273
"=::: 12
10
ll-ais-retinal in Rhodopsin
13
3' 14
4
-:?
CH
3
15
O~C""H
J ~15"<:O 11 13 C~ 9~
10
~
12
~
H 14
Al1-t,..a1l8-
retinal
3
4
FIGURE 10.2.
Conversion of II-cis-retinal to all-trans-retinal.
Thus the plot of ;\max against v(C=C) in retinal-containing proteins gives a straight line. This is because a bathochromic shift is caused by the increased delocalization of the electrons of the polyene chain, and this decreases the force constants of the C=C bonds (Callender et aI., 1976). The role of rhodopsin in the visual process is essentially associated with the conversion of one retinal isomer to another; understanding the Raman spectra of the various retinal isomers is important. There is a doublet band in the 1000-1031-cm- 1 region that originates from the vibrational modes of C(5)H 3 , C(9)-H 3 , and C(l3)-H 3 with the conjugated double bond that is characteristic of each individual isomer of all-trans-, 9-cis-, or l3-cis-retinal (Cookingham et aI., 1976). In the spectrum of 11-cis-retina1 there are two lines, at 998 and 1018 em-I, in solution compared with a single line at 1018 cm~l in the crystalline state. These lines have been assigned to C-CH 3 stretching motions. Callender et aI. (1976) assigned the 998-cm- 1 line to l3-CH 3 in 12-S-trans and 12-S-cis forms and the 1018-cm- 1 line to C(9)-H 3 in l2-S-trans and 12-S-cis forms. Thus the existence of two lines is interpreted to mean the existence of a mixture of 12-S-trans and 12-S-cis forms in a solution of 11-cis-retinal. Some major bands of all-trans-retinal have been identified (Rimai et aI., 1973; Warshel, 1977). Their results are briefly summarized here: 1182 em-I 1167,1201 cm- 1 970, 1009 cm- 1
C-C stretching A mixture of C-C stretching and C-H bending C-CH 3 stretching
The frequency of C=O and C=N stretching vibrations is markedly affected by the hydration of retinal. This is probably due to hydrogen bonding of water and the retinal carbonyl group (Allan and Cooper, 1980).
TABLE 10.1.
Wave Number (em-I)
Vibrational Assignment for Rhodopsin-Related Compounds
Assignment
C(14)-H out-of-plane wagging C(IO)-H out-of-plane 875 wagging C(II)-H and C(12)-H 922 out-or-plane wagging C-H bend 954 C-H bend 960 C-C-H bend, out-of963 plane C- H bend C-H bend 965 C-C-H bend, out-of969 plane C- H bend C-C-H bend, out-of970 plane C-H bend C-Hbend 971 C-H bend and some C-CH) stretch 977 C-H bend 996-997 C(9)-CH) stretch and C(13)-CH) stretch 1000 C-CH) stretch 1009 C(9)-CH) stretch and C(13)-CH) stretch 1010 CH) polyene-C stretch 854
1012 1013 1017 1018 1117 1118 1128 1143 1147 1153 1163
Compound
Reference
Bathorhodopsiri
Eyring et al. (1980b, 1982)
Bathorhodopsin
Eyring et al. (1980b, 1982)
Bathorhodopsin
Eyring et al. (1980b, 1982)
Isorhodopsin Isorhodopsin 9-cis- Retinal
Sulkes et al. (1978) Mathies et al. (1976a) Cookingham et al. (1978)
Acid Metarhodopsin Sulkes et al. (1978) 13-cis-Retinal Cookingham et al. (1978) II-cis, trans-Retinal
Cookingham et al. (1978)
Rhodopsin Rhodopsin
Sulkes et al. (1978) Mathies et al. (1976a)
Bathorhodopsin II-cis-Retinal
Sulkes et al. (1978) Cookingham et al. (1978), Cookingham and Lewis (1978) Mathies et al. (I976a) Cookingham et al. (1978), Cookingham and Lewis (1978) Oseroff and Callender (1974)
Rhodopsin 9-cis, trans- Retinal
Rhodopsin, isorhodopsin, bathorhodopsin l3-cis-Retinal Isorhodopsin Rhodopsin Rhodopsin 9-cis- Retinal 13-cis-Retinal II-cis-Retinal ll-cis-Retinal
C(13)-CH) stretch C-CH) stretch C(9)-CH) stretch C-CH) stretch C(14)-C(15) stretch C(14)-C(15) stretch C(14)-C(15) stretch C-C(C-19 to C-l3) stretch C-C(C-9 to C-l3) stretch 9-cis-Retinal C-C stretch and Isorhodopsin C-C-Hbend C-C(C-19 to C-B) l3-cis, trans-Retinal stretch
Cookingham et al. (1978) Mathies et al. (1976a) Cookingham and Lewis (1978) Mathies et al. (1976a) Cookingham et al. (1978) Cookingham et al. (1978) Cookingham et al. (1978) Cookingham et al. (1978) Cookingham et al. (1978) Mathies et al. (1976a) Cookingham et al. (1978)
TABLE 10.1.
Wave Number (em-I) 1193 1198 1201 1206 1207 1216 1216 1219 1219
1240 1241 1270 1271
1272 1274 1282
1295 1316 1318 1329
Continued
Assignment
Compound
C-C stretch, C-9 and C-13 Me rock C-C stretch, C(9) C-C stretch, C-9 and C-13 Me rock
13-cis-Retinal
Cookingham et al. (1978)
trans- Retinal 9-cis- Retinal
Cookingham et al. (1978) Cookingham et al. (1978)
ll-cis-Retinal Isorhodopsin
Cookingham et al. (1978) Mathies et al. (1976a)
Rhodopsin
Mathies et al. (1976)
9-cis- Retinal
Cookingham et al. (1978)
l1-cis- Retinal
Cookingham et al. (1978)
13-cis-Retinal
Cookingham et al. (1978)
Rhodopsin
Mathies et al. (1976a)
Isorhodopsin
Mathies et al. (l976a)
Rhodopsin
Mathies et al. (1976a)
11-cis- Retinal
Cookingham et al. (1978)
trans- Retinal
Cookingham et al. (1978)
13-cis-Retinal
Cookingham et al. (1978)
trans- Retinal
Cookingham et al. (1978)
13-cis-Retinal
Cookingham et al. (1978)
9-cis- Retinal 13-cis-Retinal
Cookingham et al. (1978) Cookingham et al. (1978)
Isorhodopsin
Mathies et al. (l976a)
9-cis- Retinal
Cookingham et al. (1978)
C-C stretch and C-C-Hbend C-C stretch, C-9 and C-13 Me rock C-C stretch, C-9 and C-13 Me rock C-C stretch, C-9 and C-13 Me rock C-C-Hbend + C=C stretch or C-C stretch C-C stretch and C-C-Hbend C-C stretch and C-C-Hbend C-C stretch and C-C-Hbend C-C-Hbend + C=C stretch or C-C stretch C-C-Hbend + C=C stretch or C-C bend C-C-H bend + C=C stretch or C-C bend C-C-H bend + C=C stretch or C-C bend C-C-Hbend + C=C stretch or C-C bend C-C-Hbend + C=C stretch or C-C bend C-13 sym. Me deformation C-C stretch and C-C-Hbend C-13 sym. Me deformation
Reference
........
TABLE 10.1.
Continued
-Wave Number (em-I)
Assignment
C-13 sym. Me deformation C-13 sym. Me deformation C-13 sym. Me deformation 1345 C-13 sym. Me deformation 1352 1358(IR) C-1 gem-Me 2 1360(lR) C-1 gem-Me 2 1362(IR) C-1 gem-Me 2 C-1 gem-Me 2 13-74(CC1 4 ) C-9 sym. Me deformation C-1 gem-Me 2 1375(1R) C-1 gem-Me 2 1379(IR) C-1 gem-Me 2 1380(IR) 1402(CC1 4 ) C-9 and C-13 asym. Me deformation 1431(CC1 4 ) C-9 and C-13 asym. Me deformation CH bending of -CH 3 1440 1337
1446(CC1 4 )
1530 1537 1539 1545 1545 1550 1551 1563
C-9 and C-13 asym. Me deformation C-9 and C-13 asym. Me deformation C-9 and C-13 asym. Me deformation C-9 and C-13 asym. Me deformation C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch
1568 1573 1576 1577 1584 1586
C=C C=C C=C C=C C=C C=C
1448(CC1 4 ) 1448(CC1 4 ) 1448(CC1 4 )
stretch stretch stretch stretch stretch stretch
Compound.
Reference
trans- Retinal 9-cis- Retinal 11-cis- Retinal 13-cis- Retinal 11-cis- Retinal 9-cis- Retinal trans- Retinal 13-cis- Retinal 9-cis- Retinal
Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al.
(1978) (1978) (1978) (1978) (1978) (1978) (1978) (1978) (1978)
11-cis- Retinal 9-cis- Retinal trans- Retinal 9-cis- Retinal
Cookingham et Cookingham et Cookingham et Cookingham et
(1978) (1978) (1978) (1978)
11-cis- Retinal
Cookingham et al. (1978)
Rhodopsin, isorhodopsin, bathorhodopsin 9-cis- Retinal
Oseroff and Callender (1974)
11-cis- Retinal
Cookingham et al. (1978)
13-cis- Retinal
Cookingham et al. (1978)
trans- Retinal
Cookingham et a1. (1978)
Rhodopsin Bathorhodopsin Bathorhodopsin Acid metarhodopsin 11-cis- Retinal Isorhodopsin Isorhodopsin trans- Retinylidene ethanol amine trans- Retinal 13-cis- Retinal 11-cis-Retinal trans- Retinal 13-cis- Retinal 9-cis- Retinal
Su1kes et al. (1978) Sulkes et al. (1978) Oseroff and Callender (1974) Sulkes et al. (1978) Mathies et al. (1976a) Mathies et al. (1976a) Oseroff and Callender (1974) Oseroff and Callender (1974)
al. al. al. al.
Cookingham et al. (1978)
Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al. Cookingham et al.
(1978) (1978) (1978) (1978) (1978) (1978)
~
Visual Pigments
TABLE 10.1.
277
Continued
Wave Number (em-I) 1625 1625 1630
1653 1654 1655 1656 1657 1658 1659 1660 1670
1.2.
Assignment
Reference
Compound
Deuterated Schiff base (C=ND+) Unprotonated Schiffbase vibration Deuterated Schif~ base vibration C=N stretching Protonated Schiffbase vibration Protonated Schiffbase C=N stretch Protonated Schiffbase vibration C=O stretch C=O stretch C=O stretch C=O stretch Protonated Schiffbase C= N stretch C=O stretch
Bathorhodopsin
Eyring and Mathies (1979)
Rhodopsin
Oseroff and Callender (1974)
Rhodopsin
Oseroff and Callender (1974)
Retinal Compounds Rhodopsin
Doukas et al. (l978b) Sulkes et al. (1978)
Isorhodopsin
Mathies et al. (1976a)
Rhodopsin
Oseroff and Callender (1974)
9-cis-Retinal Bathorhodopsin I l-cis- Retinal 13-cis-Retinal I I -cis-Retinal
Cookingham et al. (1978) Eyring and Mathies (1979) Cookingham et al. (1978) Cookingham et al. (1978) Mathies et al. (1976a)
Retinal compounds
Doukas et al. (1978b)
LINKAGE BETWEEN PROTEIN AND PROSTHETIC GROUP
One of the significant contributions of Raman study to our understanding of the retinal mechanism is the proof that opsin and retinal are joined by a protonated-Schiff-base linkage (Rimai et al., 1970; Heyde et al., 1971).
+
(retinal)-CH= N - (CH 2)4- (opsin) H The proton at the protonated Schiff base is labile and should be readily exchanged with deuterium. If so, one should observe an isotopic shift of the protonated-Schiff-base stretching vibration.
0 20
CH=NH+ -CH=NO+ Rhodopsin shows the Schiff-base stretching vibration at 1655 cm- I . Upon deuteration, the frequency is shifted to 1630 cm - I (Oseroff and Callender,
278
Visual Pigments and Bacterial Rhodopsin
1974). Likewise, bathorhodopsin (the first bleaching intermediate) is also a protonated Schiff base (Lewis, 1976: Aton et al., 1980). Bathorhodopsin has
+
the protonated-Schiff-base C= N vibration at 1657 cm -I, which shifts to H 1625 cm - I upon deuteration. Therefore, the frequencies of the protonated Schiff base of rhodopsin and bathorhodopsin are essentially identical, indicating that absorption of light by rhodopsin does not change the state of protonation (Eyring and Mathies, 1979; Aton et al., 1980; Lewis, 1980). Metarhodopsin I is linked to opsin by a protonated Schiff base, whereas metarhodopsin II is linked by an unprotonated Schiff base (Doukas et. al.,
C
n
A. ALL TRANS PROTONATED
n
METARHODOPSIN 1
;'"
N
"'~
. ...
t
~
....
III
:D
'"
l;
~IJ\_ J in z w >-
CD
'"'"-on ....!£on~
~ ~"~~ '"
on-
II
.
i,
$on
o
~
..
:'ri~3=~'"
'" 0
~
:i I B.
'"
....
..
'§
on
ALL TRANS UNPROTOf:jATED
o
::Ii
METARHODOPSIN
n
a:
800
1000
1200
1400
16oocm-' 1
800
1000
1200
1400
FIGURE 10.3. (A) The 1657-cm- band is due to the protonated Schiff base. The model compound used was alI-trans-retinal-n-butylamine hydrochloride. (B) Unprotonated compound lacks the band at 1657 cm -I. The model compound used was all-trans-retinal-n-butylamine. (C) The 1657-cm- 1 band is the indication of the protonated Schiff base for metarhodopsin I. (D) Metarhodopsin II shows a band at 1632 cm -I, indicating that the Schiff base is nonprotonated. The figures were reproduced from Doukas et al. (1978b).
Visual Pigments
279
1978b). Evidence for this is seen in Figure 10.3. The protonated and unprotonated species of the model compound all-trans-retinal-n-butylamine show bands at 1657 and 1632 cm- I , respectively (Figure 1O.3A and B). Metarhodopsin I shows the Raman band at 1657 cm- 1, whereas metarhodopsin II shows it at 1632 cm -1. Rhodopsin from the class Cephalopoda, which contains the nautilus, cuttlefish, octopus, and squid, differs from that of the vertebrates, as it does not dissociate into opsin and retinal. Judging from Raman frequencies of acid and alkaline metarhodopsin, the Schiff base of both compounds is protonated (Kitagawa and Tsuda, 1980). 1.3.
EXPERIMENTAL TECHNIQUES
The example shown here illustrating the use of Raman spectroscopy to examine visual pigments involves considerable ingenuity. The conversion of the cis to the all-trans conformation is very fast and also involves many photosensitive intermediate compounds. The relative proportions of different intermediates are dependent on the irradiation wavelength, because each intermediate has a different absorption maximum. Thus a dual-beam method is used by a number of workers (Oseroff and Callender, 1974; Sulkes et aI., 1978; Eyring and Mathies, 1979). The dual-beam method involves pumping two beams, one to alter the relative concentrations of the intermediates and the other to enhance the Raman spectrum of one particular intermediate. An example of spectra obtained by the dual-beam method is shown in Figures lO.4A and B. One BOVINE
s;:
95K
~!?
... q
AJ PROBE BEAM: 4B2.5 nm
...v.>z
...'"z
..,.., .....
1D.n...._
~
O-N.
""
~ (rl
0" N
N
r ~"I ';~; Y: J ir i' : ' I
q.~ I B·
RI, II
Alli
'
,
.,..,
.
i ~' '
••
., ......
l• 't q q
B B I I
I I I 'B
1&00
'700
FIGURE 10.4. Resonance Raman spectra of rhodopsin obtained by the dual-beam method. The figure was reproduced from Marcus and Lewis (1979).
280
Visual Pigments and Bacterial Rhodopsin
immediately notices the differences between the two spectra. These differences reflect the relative contributions of rhodopsin and bathorhodopsin (Marcus and Lewis, 1979; Aton et a1., 1980). Significant differences between the Raman spectrum of bathorhodopsin and that of its parent compound suggest that a geometric change in the chromophore is an important part of the photobiochemical process involved in vision. Some investigators have used continuous laser beams and obtained resonance Raman spectra of flowing samples of rhodopsin. This is called the jet-stream technique, and it avoids photoisomerization (Callender et a1., 1976; Mathies et a1., 1976a, b, 1977). By this technique the photoisomerized molecules are constantly removed by the highvelocity flow. Raman spectra can also be obtained by using continuous lasers with stationary samples of rhodopsin at a very low temperature to overcome rhodopsin's photolability (Lewis et a1., 1974; Oseroff and Callender, 1974). Another approach is to use visual-pigment analog (Auerbach et a1., 1979). The advantage of using model compounds is that it allows clear-cut assignments of Raman bands to be made, and it simplifies the interpretation of photochemical events, since they do not contain many intermediates. However, the use of model compounds can be helpful only as an initial guide, as these experiments do not provide unequivocal conclusions. Model compounds with trans-retinal covalently linked to aliphatic amines can be readily prepared, but these model compounds do not reproduce the bathochromic shift of the absorption maxima observed with retinal isomers. This is just one example of the difference between model compounds and naturally occurring retinals. 1 .4.
CONVERSION OF CIS TO TRANS
The conclusion of the rhodopsin photochemical event is the conversion of the cis to the trans configuration by a mechanism of photoisomerization at the 11,12 double bond. The first step is the formation of the intermediate compound bathorhodopsin from rhodopsin. In this step, photon energy is converted to chemical energy, which is used for the subsequent reactions. The time required for this conversion is less than 6 ps. The conversion to bathorhodopsin occurs even at very low temperatures, even at 4°K (-269°C) (Sulkes et al., 1978). The same conclusion was obtained by Hayward et al. (1980; 1981), who observed the appearance of all-trans-retinal Raman lines within 30 ps after photolysis of rhodopsin and isorhodopsin. This implies that isomerization is complete within picoseconds of the absorption of a photon. Bathorhodopsin has ·intense Raman lines at 853-856, 875-877, and 920 cm - I that are not found in all-trans protonated-Schiff-base derivatives of retina1. Bathorhodopsin has a protonated-Schiff-base vibration at 1657 cm-" which shifts upon deuteration to 1625 cm - I. Metarhodopsin was also found to have the all-trans protonated-Schiff-base form (Sulkes et a1., 1976). In this case the determination was done by comparing the Raman spectrum of metarhodopsin with those of model compounds of the retiny1idene chromophore in the all-trans conformation. The same conclusion-that the chromophore of
.J
Visual Pigments
281
metarhodopsin has the all-trans conformation-was arrived at by Doukas et al. (l978a, b). The important question is when isomerization first occurs. It is agreed that the first step of the photochemical reaction is proton translocation. The time involved for the conversion of rhodopsin to bathorhodopsin is generally believed to be too short for complete isomerization to the trans form. This assumption is challenged by some investigators (Callender, 1980). Eyring et al. (1980a, b) concluded that the structure of the bathorhosopsin chromophore is twisted all-trans. They considered the 853-, 875-, and 920-cm - I lines to be due to out-of-plane vinyl-hydrogen motions on the chain rather than exomethylene or ring modes. Moreover, they concluded that the 1100-1400cm -I Raman fingerprints are an indication of trans configuration of the double bonds. It was proposed by Callender et al. (1976) that rhodopsin contains the distorted conformation of the l2-S-trans form. Moreover, the 1271-cm- 1 band is much more intense when the C-B position is occupied by a methyl group and the retinal can have a 12-S-trans conformation (Figure 10.5). Therefore, the existence of the 1271-cm- I band is used to indicate that the chromophore in bovine and squid rhodopsin is ll-cis-12-S-trans (Cookingham and Lewis, 1978). Raman spectra can be obtained in a live eye that is stimulated by light. Changes in the vibrational spectrum of the retinylidene chromophore are observed when a neural impulse is generated by the photoreceptor (Lewis, 1975). The protonated Schiff base is the most accepted and reasonable interpretation of the 1655-cm- I line and the shift to 1630 cm - I upon deuteration. A
9
~10
2
11
~12
ll-ais,12-S-trans rhodopsin
lJ
J'
~14
4
15~O
ll-ais,12-S-ais rhodopsi n
lJ
4
14~ ~'15
O~
FIGURE 10.5.
Structures of 12-S-trans- and 12-S-cis-rhodopsin.
282
Visual Pigments and Bacterial Rhodopsin
different explanation was offered by Favrot et al. (1979a, b), who suggested that the nitrogen atom is involved in a hydrogen bond with a proton donor, so that the proton remains closer to the donor. 1.5.
OPSIN
The apoprotein in rhodopsin is opsin. Opsin makes up 80% of the total protein complement in the disk membrane. When rhodopsin undergoes photochemical reaction, not only the retinal portion (chromophore) changes its structure, but the structure of opsin is also altered (Figure 10.6). It is thus important to understand the structure of the opsin moiety too. Highly purified opsin was isolated from retinal by Rothschild et al. (1976). Remarkably fluorescent-free, good-quality Raman spectra were obtained by them. The distinct amide I band at 1660 cm - I and amide III band at 1269 cm - I lead them to conclude that opsin contains a-helical structure, but little ,B-structure. Judging from the ratio
Rhodopsin is II-CIS retinal+a membrane glycoprotein matrix called opsin 498nm psec
thv
•.. ,
Batho Rhodopsin 543nm
nsec ~
}J-
sec ~
••
Lumi Rhodopsin 497nm Meta Rhodopsin I 460nm
msec T
minutes
t
Meta Rhodopsin 380nm
+
n
o
Opsin
FIGURE ]0.6. Overall change of rhodopsin induced by light. Both chromophore and opsin portions change their conformations. The figure was reproduced from Lewis, The Spex Speaker, 2], 3, (1976) by permission of Spex Industries, Inc.
,
J
Bacteriorhodopsin
283
of the tyrosine band doublet 1850/1830' it was concluded that the tyrosine residues are predominantly hydrogen bonded. The optical as well as photochemical events change considerably when retinal binds to opsin (Rosenfeld et al., 1977). Such change is believed to be due to the electrostatic interactions between the charged opsin and the 11-cisretinal chromophore (Honig et al., 1979a, b). 1.6.
SUMMARY
There are two main processes in the photoactivation of rhodopsin. One is the isomerization process, and the other is proton transfer. The important question is which one of these processes comes first. Some investigators consider that isomerization comes first, and then protonation occurs (Favrot et al., 1979a, b; Aton et al., 1980), but other investigators think that protonation is a precondition for isomerization (Sandorfy, 1980). This important question has not yet been resolved conclusively.
2.
BACTERIORHODOPSIN
Bacteriorhodopsin (purple membrane protein) was discovered by Stoeckenius and his co-workers (Oesterhelt and Stoeckenius, 1971; Blaurock and Stoeckenius, 1971). Actually there are two types of bacteriorhodopsin. One is light-adapted (A max = 568 nm) and the other is dark-adapted (A max = 558 nm). Only the light-adapted form has the ability to pump protons. Bacteriorhodopsin is a membrane-bound protein found in Halobacterium halobium and H. cutirubium. Like rhodopsin, light-adapted bacteriorhodopsin contains retinal but for the totally different biological purpose of pumping ions for energy. Unlike rhodopsin, which contains the II-cis isomer, bacteriorhodopsin bR 570 contains the all-trans isomer, and the dark-adapted form of bacteriorhodopsin bRr6~ is a mixture of the 13-cis and the all-trans form, with a ratio of approximately 1 : 1 (Pettei et al., 1977). Narva et al. (1981), nevertheless, suggest that such conclusions from the present Raman results should be viewed as tentative. In this chapter, notations of bacteriorhodopsin intermediates are expressed as bR, bK, bL, bM, and bO; however, some investigators simply use R, K, L, M, and O. Subscripts indicate the wavelength of absorption maxima, since each intermediate has its own characteristic absorption spectrum. Thus bR 570 is a bacteriorhodopsin with its absorption maximum at 570 nm. The dark-adapted form has an absorption maximum at 560-570 nm. Upon illumination with visible light, bacteriorhodopsin undergoes a series of lightinduced conformational changes, and at the same time protons are pumped across the cell membrane, resulting in a gradient. The gradient is believed to drive the phosphorylation of ADP to form ATP. The chlorophylls and
;~ 284
Visual Pigments and Bacterial Rhodopsin
bacteriorhodopsin are the only biological systems capable of converting solar energy to chemical energy. Under anaerobic conditions, bacteria cannot produce energy by the normal oxygen-requiring mechanisms, and therefore life is maintained through other energy-producing mechanisms. In the absence of oxygen, bacterial rhodopsin is produced, serving as an energy converter. H. halobium synthesizes ATP under anaerobic conditions in the presence of light, but not in the dark. It is thus clear that light is an essential factor for ATP production in these bacteria. The bacteria do not ferment carbohydrates nor contain chlorophyll; they are not photosynthetic bacteria. The cycle operates about 200 times per second, pumping protons from inside of the bacterial cell to the outside when light is absorbed. From absorption spectroscopy it is found that bR 570 ~ bK590 is photoreversible (Goldschmidt et aI., 1976). Therefore, bacteriorhodopsin is a biological pump to convert light energy into chemical energy in the form of ATP when the bacteria are deprived of oxygen. On binding to protein, the absorption maximum of retinyledene shifts from 370 to 570 nm, indicating considerable structural changes in the chromophore. The photochemical reaction is very rapid, and in continuous light it undergoes 100-200 cycles/s (Figure 10.7). Like rhodopsin, the time required for photocyclic reactions of bacteriorhodopsin is extremely short. The rapid photochemical reaction of the purple membrane protein was measured by the fluorescence technique (Alfano et aI., 1976). The fluorescence rise time is less than 8 ps, and its lifetime is only 40 ± 5 ps at 90 o K, and is less than 3 ps at room temperature. It is remarkable that such complicated reactions can all take place in such a short time. Using a flow technique, picosecond and nanosecond resonance Raman spectra of bacteriorhodopsin can be obtained. It requires less than 40 ps for the transition from all-trans-retinal to a 13-cis-type configuration. Thus it is clear that more than one species exists in the 40 ps to lOOns time scale (Hsieh et aI., 1981).
e
~r . / bR
570
~10
_cJ:~psec
/
+Jr'
H
bN
10
T,~
bL 550
520
~
bM412
~sec
~N"
+
H
( FIGURE 10.7. Photochemical cycle of bacteriorhodopsin. .J
Bacteriorhodopsin
285
There are good review articles by Stoeckenius and Lozier (1974) and Stoeckenius et aI. (1979) that cover all aspects of bacteriorhodopsin and the purple membrane protein of halo bacteria. 2.1.
VIBRATIONAL ASSIGNMENTS
The biggest difficulty in the study of bacteriorhodopsin and rhodopsin by Raman spectroscopy is .that the spectrum obtained usually arises from a complex mixture of many intermediates. Thus one must use several techniques to attain the largest steady-state concentration of a particular compound before proper assignments can be made. Under continuous laser illumination, bR s70 and bM412 have the largest steady-state concentrations (Mendelsohn et aI., 1974; Lewis et aI., 1974). So practically all the band assignments have been made for these two species (Table 10.2). TABLE 10.2.
Wave Number (em-I) 957 960 972 1005 1007 1009 1010 1051 1115 1121 1134 1136 1168 1170 1173 1184
Vibrational Assignment for Bacteriorhodopsin and Related Compounds
Assignment
Compound
Reference
C-H out-of-p1ane bending (C-C-H) out-of-p1ane bend (C-C-H) out-of-p1ane bend (C-C) stretch + (C-H) bend C-CH 3 stretch C-CH 3 stretch C(9)-CH 3 stretch, C(13)CH 3 stretch C(9)-CH 3 stretch, C(13)CH 3 stretch C-C stretch, C-H bend C(14)-C(15) stretch C(14)-C(15) stretch C(14)-C(15) stretch C-C stretch, C-H bend C(14)-C(15) stretch C-C stretch, C-H bend C-C (C-9 to C-13 stretch) C-CH 3 rock and C-C stretch C-C stretch, CH 3 rock
bR s70
Stockburger et al. (1979)
bR s70
Marcus and Lewis (1978)
bM412
Marcus and Lewis (1978)
bR
Mendelsohn et al. (1974)
bR s70 bR 412 bM412
Stockburger et al. (1979) Stockburger et al. (1979) Marcus and Lewis (1978)
bR s70
Marcus and Lewis (1978)
bR bM412 bM4I2 bR s70 bR bR s70 bR bR s70 bR s70
Mendelsohn et al. (1974) Marcus and Lewis (1978) Marcus and Lewis (1978) Marcus and Lewis (1978) Mendelsohn et al. (1974) Marcus and Lewis (1978) Mendelsohn et al. (1974) Marcus and Lewis (1978) Stockburger et al. (1979)
bM412
Marcus and Lewis (1978)
TABLE 10.2.
Continued
--
Wave Number (cm- I ) 1185 1189 1196 1197 1198 1202 1203 1215 1225
1251 1255
1272 1274 1275 1276 1303 1304 1305 1322 1329 1330 1353 1374 1377 1379 1450
1455
Assignment C-C stretch, CH 3 rock C-C stretch, C-H bend C-C stretch, CH 3 rock C-C stretch, CH 3 rock C-C stretch, C-H bend C-C stretch, CH 3 rock C-C stretch C-C stretch, CH 2 rock Isoprenoid chain motions involving the Schiff-base linkage C-C stretch, C-H bend Isoprenoid chain motions involving the Schiff-base linkage C-C-H bend + C=C or C-C stretch C-C stretch, C-H bend C-C-H bend + C=C or C-C stretch C-C-H bend and C-C-(C=C) stretch C-C-H bend + C=C or C-C stretch C-C stretch, C-H bend C-13 Methyl symmetrical deformation region C-13 Methyl symmetrical deformation region C-C stretch, C-H bend C-13 Methyl symmetrical deformation region C-13 Methyl symmetrical deformation region C-9 Methyl symmetrical deformation C-9 Methyl symmetrical deformation C-C stretch, C-H bend Asymmetrical methyl deformation C-9 and C-13 Asymmetrical methyl deformation
Compound
Reference
bR s70 bR bR s70 bM412 bR bR s70 bR s70 bR s70 bM412
Marcus and Lewis (1978) Mendelsohn et al. (1974) Marcus and Lewis (1978) Marcus and Lewis (1978) Mendelsohn et al. (1974) Marcus and Lewis (1978) Stockburger et al. (1979) Marcus and Lewis (1978) Marcus and Lewis (1978)
bR bR s70
Mendelsohn et al. (1974) Marcus and Lewis (1978)
bM412
Marcus and Lewis (1978)
bR bR s70
Mendelsohn et al. (1974) Marcus and Lewis (1978)
bR s70
Stockburger et al. (1979)
bM412
Marcus and Lewis (1978)
bR bR s70
Mendelsohn et al. (1974) Marcus and Lewis (1978)
bR s70
Marcus and Lewis (1978)
bR bR s70
Mendelsohn et al. (1974) Marcus and Lewis (1978)
bR s70
Marcus and Lewis (1978)
bM412
Marcus and Lewis (1978)
bR s70 bR
Mendelsohn et al. (1974) Mendelsohn et al. (1974)
bM412
Marcus and Lewis (1978)
bR s70
Marcus and Lewis (1978)
.
Bacteriorhodopsin
287
TABLE 10.2. Continued Wave Number (cm- I )
Assignment
Compound
1465 1512 1529
C-9 and C-13 CH 2 bend C=C stretch C=C stretch C=C Stretch
1530
C=C Stretch
bR S70
1537 1551 1554 1556 1567 1568 1568
C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch C=C stretch
bL ssO bL sso bL sso bM412 bM412 bM412
1570 1604 1619
1621 1622
C=C stretch C=N stretch C=N stretch in D 20 C=N Unprotonated stretch Isolated, unprotonated C=N stretch C=N Unprotonated stretch Unprotonated C=N stretch
1626 1640 1642 1652
Schiff-base C=NH stretch C=NH+ stretch C=NH+ stretch C=NH+ stretch
1276
C-C-H bend and C-C-(C=C) stretch
1620
bR bR s70
bR s70
Mendelsohn et al. (1974) Ehrenberg et al. (1980b) Ehrenberg et aI. (1980b) Mendelsohn et aI. (1974), Campion et aI. (I 977a, b) Marcus and Lewis (1978) Campion et al. (1977a, b) Stockburger et aI. (1979) Teener et aI. (1977) Campion et aI. (1977a, b) Marcus and Lewis (1978) Stockburger et aI. (1979) Teener et aI. (1977) Mendelsohn et aI. (1974), Mendelsohn (1976) Campion et aI. (1977a, b) Teener et aI. (1977) Mendelsohn et aI. (1974) Marcus and Lewis (1978) Stockburger et aI. (1979)
bR bR
Mendelsohn et aI. (1974) Lewis et aI. (1974)
bKs90
Teener et al. (1979a) Marcus and Lewis (1978) Marcus and Lewis (1978) Mendelsohn et aI. (1974)
bM412
bR
bR
+
Reference
bM412
bR s70 bR bM412
bR bR s70 N-Protonated retinylidendysine
bR s70
Stockburger et aI. (1979)
The photoinduced reaction is very rapid, and the lifetime of each intermediate is very short. This means that the concentration of an intermediate compound is extremely low at a given time. Therefore, several special techniques are used to obtain Raman spectra of intermediates. One technique is called time-resolved resonance Raman spectroscopy. Using this technique, one can obtain spectra of transient species in milliseconds. A continuous-wave laser is focused onto a rotating chopper with two slits. The slits are adjusted so that
288
Visual Pigments and Bacterial Rhodopsin
one is wide enough to cause considerable photolysis of the sample on each pulse, whereas the other causes minimal photolysis but is sufficient to accumulate a Raman spectrum. The spectrum of bacteriorhodopsin obtained by this method can be assigned to three intermediates in the photochemical cycle: bR 570 , bL 550 , and bM412 (Teener et al., 1977; Teener and El-Sayed, 1978; El-Sayed, 1981). The 1643- and 1620-cm-I bands are assigned with the continuous-wave method to protonated and unprotonated Schiff bases. These assignments are confirmed with the time-resolved technique. Using deuterosubstituted retinals, Mathies et al. (1980) concluded that the intense bands of bathorhodopsin at 835, 875, and 920 cm -I are C- H out-of-plane bending modes of vinyl chains. As Ehrenberg et al. (l980a) pointed out, the 1642- and 1620-cm - I bands are due to vibrational modes of the protonated and unprotonated Schiff bases, yet neither is due to simple, localized C=NH or C=N stretching vibrations. The rate of proton-deuterium exchange for the Schiff base of bacteriorhodopsin is extremely rapid and was found to be 4.7 ms with the rapid-mixing technique (Ehrenberg et al., 1980c) and less than 3 ms with the continuous-flow method (Doukas et al., 1981). 2.2.
LINKAGE BETWEEN PROTEIN AND PROSTHETIC GROUP
With the resonance Raman technique, it was found that the Schiff base in bR 570 is protonated, but the bM412 intermediate is not. The protonated Schiff base can be deuterated, with a subsequent shift of the frequency. The protonated Schiff base of bacteriorhodopsin appears at 1643 cm -I, whereas the unprotonated intermediates show the Schiff-base band at 1620 cm- I (Mendelsohn, 1976; Lewis et al., 1974; Campion et al., 1977a, b; Marcus and Lewis, 1977; Aton et al., 1977; Pande et al., 1981). Although the primary linkage of the Schiff base is well recognized, some investigators report a secondary protein interaction with the protonated Schiff base in bacteriorhodopsin bR 570 (Lewis et al., 1978; Ehrenberg and Lewis, 1980; Marcus and Lewis, 1978; Fischer and Oesterhelt, 1979; Stockburger and Massig, 1980). In order to clarify this secondary linkage, Raman Spectra of I5 N-e-lysine-1abeled and non1abe1ed bacteriorhodopsins bR 570 were compared. It was shown that the 15N isotopic effect on the Schiff-base vibration can be accounted for by 15N labeling only at the Schiff-base nitrogen. Thus there is a linkage of the retinal chromophores with the e-amino nitrogen of lysine (Argade et al., 1981).. More-detailed descriptions of the two compounds are discussed in the following section. 2.2.1.
bR 570
As is the case with rhodopsin, the protonated Schiff base is responsible for conjugating retinal and the protein moiety. This conclusion was not im-
J
Bacterlorhodopsln
289
mediately obvious. Mendelsohn (1973) observed the 1623- and 1646-cm - I bands in water, but the 1646-cm - I band disappeared in D20. Initially the 1646-cm- 1 band was considered to be water-molecule OH bending vibration, and the 1623-cm - I band was assigned to the unprotonated-Schiff-base vibrational mode. Lewis et a1. (1974) made an extensive study of the 1646- and 1623-cm- 1 bands. They found that as the excitation wavelength approaches 550 nm, the intensity of the 1646-cm - I band is resonance enhanced but not the intensity of the 1623 cm- 1 band. They explain that the 1646-cm- I band results from the protonated Schiff base, whereas the 1623-cm- 1 band is of the nonprotonated-Schiff-base (C=N) mode. The absence of a 1646-cm- 1 band in D20 can be explained by the fact that the proton is replaced by deuterium, and the band shifted to 1623 cm - I.
+ D 20 + -CH= N - -CH= N -
I
H 1646 cm- I
I
D 1623 cm- 1
The same conclusion was obtained by Aton et a1. (1977), who used the flow technique to avoid photodecomposition of the sample (Figure 10.7). By this method a pure resonance Raman spectrum of bR 570 was obtained. The DzO experiment solves the assignment of the 1646- and 1623-cm- 1 bands very clearly. A second effect is the appearance of a 974-cm- 1 band and the decrease in the 1255-cm- 1 band. It could be that the 1255-cm- 1 band corresponds to the N - H in-plane bending vibration, or it may be an X- H vibration, where X is some unidentified atom. The effect of isotopic exchange with deuterium is expected to be a frequency decrease by a factor of approximately II. The 974-cm- I wave number is within 9% of the calculated value (Stockburger et a1., 1979). The Schiff-base portion is embedded in a hydrophobic region of the lipid-protein complex, and the C=N bond is inaccessible to chemical reagents such as hydroxylamine or sodium borohydride. Thus the addition of either weakens this Schiff-base-protein interaction and produces changes in absorption maxima as well as in the intensity of Raman bands (Mendelsohn et a1., 1974).
Absorption maximum Raman band intensity
Bacterial rhodopsin
Addition of Diethyl Ether
560nm 1529cm- 1 1568 cm- I
Shift to 460-490 om Decrease Increase
Since the proton is removed from the Schiff base as the photochemical reaction of bacteriorhodopsin proceeds, it is believed that the deprotonation process plays an important role in the proton-pumping function. The pK value
290
Visual Pigments and Bacterial Rhodopsin
of the Schiff-base deprotonation can be measured by kinetic resonance Raman spectroscopy as a function of pH. The pK is higher than 12 before light absorption and decreases to 9.9-10.3 within microseconds after photon absorption. When the tyrosine residue is iodinated, the pH of the Schiff base deprotonation drops to 7 ~ 8. This suggests that tyrosine may be involved in the deprotonation mechanism (Ehrenberg and Lewis, 1978; Gogel and Lewis, 1981). 2.2.2.
bK 590
During the first step bR s70 --7 bKs90 , there are large changes in the fingerprint region. A Raman spectrum of the short-lived intermediate bKs90 was obtained by Teener et al. (l979c), using combined microbeam and flow techniques and computer-subtraction methods. The protonated-Schiff-base vibrational frequency shifts from 1646 to 1626 cm - 1 for bR S70' The 1626-cm- 1 band is due to the C=N vibration of the protonated Schiff base. This is confirmed by deuteration, following which it shifts to 1616 cm -I. The relatively low value of the C=N frequency of the protonated Schiff base of bKs90 is explained in terms of strong hydrogen bonding between the Schiff-base proton and the protein. As indicated in Figure 10.7, the process from bR s70 is reversible. Bacteriorhodopsin's primary photoproduct K has a distorted 13-cis-retinal chromophore (Braiman and Mathies, 1982). 2.2.3.
bL 550
The resonance Raman spectrum of the second intermediate bLsso was obtained by a flow technique (Teener et al., I979b). Actually, light irradiation of bL produces a photoproduct bL', which is photoreversible to bL, and which can thermally relax to bR s70 (Hurley et al., 1978; Narva et al., 1981). At low laser power (0.65 mW), the Raman spectrum is essentially due to pure bR s70 , because the incident laser power is too low to cause photolysis. At higher laser power (to 200 mW), photolysis is increased, and the bLsso spectrum appears superimposed on the bR S70 bands. By a computer-subtraction method, the bL sso spectrum was obtained. It was found that the 1647-cm- 1 band is due to bLsso ; it is a protonated-Schiff-base vibration mode (C=NH+), as confirmed by deuteration studies. This is an important finding, as it had been thought that bLsso was unprotonated. Thus the deprotonation step must occur from bL sso to bM412 (Figure 10.7). 2.2.4.
bM 412 (M 412 )
The compound bM412 is the third intermediate of the photochemical reaction of bacteriorhodopsin. At low temperatures, the bM412 intermediate can be
J
II
N
M
A.
:!2
PMS68
>-
~
VI
.,
~I
0
CllI
0
!::!
~
Z Cl
~
cs: I:ll:
800
A.
1000
1200
1600 em'
1400
n
f:::
M412
:!2
....>VI
Z
w
~
Z
z cs: ~ cs: I:ll:
800
1000
1200
1400
1600 em-I
FIGURE 10.8. Resonance Raman spectra of (A) bacteriorhodopsin bR 570 (PM568) and (B) its intermediate bM412 (M412). From these spectra one can readily see that the original bacteriorhodopsin bR 570 (also called PM568) is protonated at the Schiff base, whereas its intermediate bM412 is not. The figure is reproduced from the paper of Aton et al. (1977) with the permission of the copyright owner, American Chemical Society.
292
Visual Pigments and Bacterial Rhodopsin
trapped so that its Raman spectra can be taken. The spectra show that the band at 1622 cm - I is present and resonance enhanced as the excitation wavelength approaches 412 nm, the absorption maximum of the intermediate (Figure 10.8) (Lewis et al., 1974; Aton et al., 1977). This clearly indicates that the Schiff base in bM412 is unprotonated. Thus the proton is removed before bacteriorhodopsin reaches the bM412 intermediate in its photochemical cycle. Relatively small changes in the fingerprint region of the Raman spectra take place when bLsso converts to bM412 , although this step gives the largest change in absorption characteristics. Braiman and Mathies (1980) also demonstrated that bM412 is an unprotonated Schiff base; moreover, they showed it to be 13-cis-retinal rather than all-trans-retinal.
2.2.5.
b0s 40
The last intermediate b0640 is a component of the photochemical cycle of bacteriorhodopsin, and from b0640 the parent component of bacteriorhodopsin bR s70 is regenerated (Figure 10.7). To observe b0640 , the resonance Raman spectrum of photolyzed bacteriorhodopsin is taken under conditions known to increase the concentration of the b0640 intermediate. Then the spectral intensities of bR s70 and other intermediates are subtracted from the original spectrum by a computer, and the Raman spectrum of b0640 is obtained. The band from the protonated Schiff base is at 1630 cm- 1, and is shifted to 1616 cm - I in DzO. This indicates that the reprotonation of the Schiff base does indeed take place during the bM412 -> b0640 step (Terner et al., 1979c).
2.2.6.
bR 560 (the Dark-Adapted bacteriorhodopsin)
The Raman spectrum of dark-adapted bacteriorhodopsin bR s60 or bR~~ is very similar to that of light-adapted bR s70 with the exception of bands at 1238 and 1187 cm- 1 that appear in the spectrum of bR s60 • As indicated by the presence of the 1642-cm- 1 band, bR s60 has a protonated Schiff base. The 1238- and 1187-cm- 1 bands are found to be the 13-cis chromophore mode. Thus bR s60 is a mixture of 13-cis- and all-trans-retinal. The linkage between the chromophore and protein is chemically the same as that of the visual pigment rhodopsin. Retinal is joined to a lysine residue by a Schiff base, and it is protonated (Marcus and Lewis, 1978; Aton et al., 1979; Terner et al., 1979b). The fingerprint regions of the spectra from the two forms of bR~~, the bR s70 , the bLsso , and the bM41Z intermediates are all different. It can be concluded that there are conformational changes of retinal in the dark-tolight-adapted process (E1-Sayed and Terner, 1979; A1shuth and Stockburger, 1981). By the efforts of many investigators, the linkage between the chromophore and apoprotein has been extensively studied and clarified. All of these results are summarized here.
j
References
293
Schiff-Base Vibration Compounds
In Water
In D2 0
Conclusions About Schiff Base
bR 470 bKs90 bL sso bM412 b0640 bR DA s60 bR DA sso
1644 1626 1647 1623 1630 1644 1644
1622 1616 1619 1620 1616 1622 1622
Protonated Protonated Protonated Unprotonated Protonated Protonated Protonated
REFERENCES Alfano, R. R., Yu, W., Govindjee, R., Becher, B., and Ebrey, T. G. (1976). Picosecond kinetics of the fluorescence from the chromophore of the purple membrane protein of Ha/ohacterium halohium. Biophys. J. 16, 541. Allan, E., and Cooper, A. (1980). Hydration of retinal and the nature of metarhodopsin II. FEBS Lett. 119, 238. Alshuth, Th., and Stockburger, M. (1981). Structural changes in the retinal chromophore of bacteriorhodopsin studied by resonance Raman spectroscopy. Ber. Bunsenges. Phys. Chem. 85, 484. Argade, P. V., Rothschild, K. J., Kawamoto, A. H., Herzfeld, J., and Herlihy, W. C. (1981). Resonance Raman spectroscopy of speci fically [ E- 15 N]lysine-Iabeled bacteriorhodopsin. Proc. Nat. A cad. Sci. USA 78, 1643. Aton, B., Doukas, A. G., Callender, R. H., Becher, B., and Ebrey, T. G. (1977). Resonance Raman studies of the purple membrane. Biochemistry 16, 2995. Aton, B., Doukas, A. G., Callender, R. H., Becher, B., and Ebrey, T. G. (1979). Resonance Raman study of the dark-adapted form of the purple membrane protein. Biochim. Biophys. Acta 576, 424. Aton, B., Doukas, A. G., Narva, D., Callender, R. H., Diour, D., and Honig, B. (1980). Resonance Raman studies of the primary photochemical event in visual pigments. Biophys. J. 29, 79. Auerbach, R. A., Granville, M. F., and Kohler, B. E. (1979). Resonance enhanced Raman spectrum of all-trans anhydrovitamin A.. Biophys. J. 25, 443. Blaurock, A. E., and Stoeckenius, W. (1971). Structure of the purple membrane. Nature New BioI. 233, 152. Braiman, M., and Mathies, R. (1980). Resonance Raman evidence for all-trans to 13-cis isomerization in the proton-pumping cycle of bacteriorhodopsin. Biochemistry 19, 5421. Braiman, M., and Mathies, R. (1982). Resonance Raman spectra of bacteriorhodopsin's primary photoproduct: Evidence for a distorted B-cis-retinal chromophore. Proc. Nat. A cad. Sci. USA 79,403. Callender, R. (1980). The primary photochemistry of visual pigments. In Proc. Vllth lnl. Conf Raman Speclrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York. Callender, R. H., Doukas, A., Crouch, R., and Nakanishi, K. (1976). Molecular flow resonance Raman effect from retinal and rhodopsin. Biochemistry 15, 1621.
294
Visual Pigments and Bacterial Rhodopsin
Campion, A., Terner, 1., and EI-Sayed, M. A. (1977a). Time-resolved resonance Raman spectroscopy of bacteriorhodopsin. Nature 265, 659. Campion, A., EI-Sayed, M. A., and Terner, 1. (1977b). Resonance Raman kinetic spectroscopy of bacteriorhodopsin on the microsecond time scale. Biophys. J. 20, 369. Cookingham, R., and Lewis, A. (1978). Resonance Raman spectroscopy of chemically modified retinals: Assigning the carbon-methyl vibrations in the resonance Raman spectrum rhodopsin. J. Mol. Bioi. 119, 569. Cookingham, R. E., Lewis, A., Collins, D. W., and Marcus, M. A. (1976). Preresonance Raman spectra of crystals of retinal isomers. J. Am. Chem. Soc. 98,2759. Cookingham, R. E., Lewis, A., and Lemley, A. T. (1978). A vibrational analysis of rhodopsin and bacteriorhodopsin chromophore analogs: resonance Raman and infrared spectroscopy of chemically modified retinals and Schiff bases. Biochemis/ry 17,4699. Doukas, A. G., Aton, B., Callender, R. H., and Honig, B. (1978a). Resonance Raman excitation profiles of all-trans retinal: theoretical implications. Chem. Phys. Lell. 56, 248. Doukas, A. G., Aton, B., Callender, R. H., and Ebrey, T. G. (1978b). Resonance Raman studies of bovine metarhodopsin I and metarhodopsin II. Biochemistry 17, 2430. Doukas, A. G., Pande, A., Suzuki, T., Callender, R. H., Honig, B., and Ottolenghi, M. (1981). On the mechanism of hydrogen-deuterium exchange in bacteriorhodopsin. Biophys. J. 33, 275. Ehrenberg, B., and Lewis, A. (1978). The pK of Schiff base deprotonation in bacteriorhodopsin. Biochem. Biophys. Res. Commun. 82, 1154. Ehrenberg, B., and Lewis, A. (1980). The structure of the active site of bacteriorhodopsin and a molecular model of proton pumping. In Proc. Vllth Int. Con! Raman Spec/rosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, p. 562. Ehrenberg, B. Lemley, A. T., Lewis, A., Von Zastrow, M., and Crespi, H. L. (I 980a). Resonance Raman spectroscopy of chemically modified and isotopically labeled purple membranes. I. A critical examination of the carbon-nitrogen vibrational modes. Biochim. Biophys. Acta 593, 441. Ehrenberg, B., Lewis, A., and Crespi, H. L. (1980b). Resonance Raman spectroscopy of chemically modified and isotopically labeled purple membranes. II. Kinetic studies. Biochim. Biophys. Acta 593, 454. Ehrenberg, B., Aaron, L., Porta, T. K., Nagle, J. F., and Stoeckenius, W. (1980c). Exchange kinetics of the Schiff base proton in bacteriorhodopsin. Proc. Nat. Acad. Sci. USA 77, 6571. EI-Sayed, M. A. (1981). Time-resolved resonance Raman techniques for intermediates of photolabile systems. Springer Ser. Opt. Sci. 26, 295. EI-Sayed, M. A., and Terner, J. (1979). Power- and time-resolved resonance Raman studies and conformational changes in bacteriorhodopsin. Ph%chem. Photobiol. 30, 125. Eyring, G., and Mathies, R. (1979). Resonance Raman studies of bathorhodopsin: evidence for a protonated Schiff base linkage. Proc. Nat. Acad. Sci. 76, 33. Eyring, G., Curry, B., Mathies, R., Broek, A., and Lugtenburg, J. (1980a). Assignment of the anomalous resonance Raman vibrations of bathorhodopsin. J. Am. Chem. Soc. 102, 5390. Eyring, G., Curry, B., Mathies, R., Fransen, R., Palings, I., and Lugtenburg, J. (1980b). Interpretation of the resonance Raman spectrum of bathorhodopsin based in visual pigment analogues. Biochemistry, 19, 2410. Eyring, G., Curry, B., Broek, A., Lugtenburg, J., and Mathies, R. (1982). Assignment and interpretation of hydrogen out-of-plane vibrations in the resonance Raman spectra of rhodopsin and bathorhodopsin. Biochemistry 21, 384. Favrot, 1., Vocelle, D., and Sandorfy, C. (1979a). Infrared and Raman studies on some imines and their picrates. Relation to the problem of protonation in visual pigments. Photochem. Ph%biol. 30, 417.
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Favrot, 1., Leclercot, J. M., Roberge, R., Sandorfy, C, and Vocelle, D. (1979b). Intermolecular interactions in visual pigments. The hydrogen bond in vision. Photochem. Photohiol 29, 99. Fischer, U., and Oesterhelt, D. (1979). Chromophore equilibria in bacleriorhodopsin. Biophys. 1. 28,211. Gill, D., Heyde, M. E., and Rimai, L. (1971). Raman spectrum of the II-cis isomer of retinaldehyde. J. Am. Chem. Soc. 93,6288. Gogel, G. and Lewis, A (1981). Effect of iodination on the pK of Schiff base deprotonation and M 412 production in purple membrane. Biochem. Biophys. Res. Commun. 103, 175. Goldschmidt, CR., Ottolenghi, M., and Korenstein, R. (1976). On the primary quantum yields in the bacteriorhodopsin photocycle. Biophys. J. 16, 839. Hayward, G., Carlsen, W., Siegman, A, and Stryer, L. (1980). Picosecond resonance Raman spectroscopy: the initial photolytic event in rhodopsin and isorhodopsin. Springer Ser. Chem. Phys. 14, 377. Hayward, G., Carlsen, W., Siegman, A, and Stryer, L. (1981). Retinal chromophore of rhodopsin photoisomerizes with picoseconds. Science 211, 942. Heyde, M. E., Gill, D. Kilponen, R. G., and Rimai, L. (1971). Raman spectra of Schiff bases of retinal (models of visual photoreceptors). J. Am. Chern. Soc. 93, 6776. Honig, B., Dinur, U., Nakanishi, K., Balogh-Nair, Y., Gawinowicz, M. A, Arnaboldi, M., and Motto, M. G. (1979a). An external point-charge model for wavelength regulation in visual pigments. 1. Am. Chern. Soc. 101, 7084. Honig, B., Ebrey, T., Callender, R. H., Dinur, U., and Ottolenghi, M. (I 979b). Photoisomerization, energy storage, and charge separation: A model for light energy transduction in visual pigments and bacteriorhodopsin. Proc. Nat. Acad. Sci. 76, 2503. Hsieh, C L., Nagumo, M .. Nicol, M., and EI-Sayed, M. A (1981). Picosecond and nanosecond resonance Raman studies of bacteriorhodopsin. Do configurational changes of retinal occur in picoseconds? J. Phys. Chem. 85,2714. Hurley, J. B., Becher, B., and Ebrey, T. G. (1978). More evidence that light isomerises the chromophore of purple membrane protein. Nature 272, 87. Kitagawa, T., and Tsuda, M. (1980). Resonance Raman spectra of octapus acid and alkaline metarhodopsin. Biochim. Biophys. Acta 624,211. Lewis, A (1975). Tunable laser resonance Raman spectroscopy as a probe of primary ion movements in the photoreceptor cells. Biophys. J. 15, I74a. Lewis, A (1976). Tunable laser resonance Raman spectroscopic investigations of the transduction process in vertebrate rod cells. Fed. Proc. 35, 51. Lewis, A (1979). Resonance Raman evidence for secondary protein-Schiff base interactions in bacteriorhodopsin: correlation of the primary excitation mechanism with a model for proton pumping and visual transduction. Phil. TrailS. R. Soc. Lond. A293, 315. Lewis, A (1980). Resonance Raman spectroscopy of rhodopsin and bacteriorhodopsin: a synopsis of the Cornell contribution. In Proc. VIJlh flit. Calif Ramall Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, p. 556. Lewis, A, Spoonhower, J., Bogomolni, R. A, Lozier, R. R., and Stoeckenius, W. (1974). Tunable laser resonance Raman spectroscopy of bacteriorhodopsin. Proc. Nal. Acad. Sci. II, 4462. Lewis, A, Marcus, M. A, Ehrenberg, B., and Crespi, H. (1978). Experimental evidence for secondary proteill-chromophore interactions at the Schiff base linkage in bacteriorhodopsin: molecular mechanism for proton pumping. Proc. Nat. Acad. Sci. 75,4642. Marcus, M. A, and Lewis, A (1977). Kinetic resonance Raman spectroscopy: dynamics of deprotonation of the Schiff base of bacteriorhodopsin. Science 195, 1328. Marcus, M. A, and Lewis, A (1978). Resonance Raman spectroscopy of the retinylidene chromophore in bacteriorhodopsin (bR 57o ), bR 560 , M 412 , and other intermediates: structural
296
Visual Pigments and Bacterial Rhodopsin
conclusions based on kinetics, analogues, models, and isotopically labeled membranes. Biochemistry 17, 4722. Marcus, M. A., and Lewis, A. (1979). Assigning the resonance Raman spectral features of rhodopsin, isorhodopsin and bathorhodopsin in bovine photostationary state spectra. Photochem. Photobiol. 29, 699. Mathies, R., Oseroff, A. R., and Stryer, 1. (I 976a). Rapid-flow resonance Raman spectroscopy of photolabile molecules: rhodopsin and isorhodopsin. Proc. Nat. Acad. Sci. 73, I. Mathies, R., Oseroff, A. R., Freedman, T. B., and Stryer, 1. (1976b). Resonance Raman spectroscopy: application of tunable lasers to the study of the molecular mechanisms and dynamics of visual excitation. Springer Ser. Opt. Sci. 3, 294. Mathies, R., Freedman, T. B., and Stryer, 1. (1977). Resonance Raman studies of the conformation of retinal in rhodopsin and isorhodopsin. J. Mol. Bioi. 109, 367. Mathies, R., Eyring, G., Curry, B., Broek, A., Palings, 1., Fransen, R., and Lugtenburg, J. (1980). Interpretation of the resonance Raman spectrum of bathorhodopsin. In Proc. VlIth Int. Con! Raman Spectrosc. W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 546-547. Mendelsohn, R. (1973). Resonance Raman spectroscopy of the photoreceptor-like pigment of Halobacterium halobium. Nature 243, 22. Mendelsohn, R. (1976). Thermal denaturation and photochemistry of bacteriorhodopsin from Halobacterium cutirubrum as monitored by resonance Raman spectroscopy. Biochim. Biophys. Acta 427, 295. Mendelsohn, R., Verma, A. 1., Bernstein, H. J., and Kates, M. (1974). Structural studies of bacteriorhodopsin from Halobacterium cutirubrum by resonance Raman spectroscopy. Can. J. Biochem. 52, 774. Narva, D. 1., Callender, R. H., and Ebrey, T. G. (1981). Low temperature resonance Raman study of the L intermediate of bacteriorhodopsin. Photochem. Photobiol. 33, 567. Oesterhelt, 0., and Stoeckenius, W. (1971). Rhodopsin-like protein from the purple membrane Halobacterium halobium. Nature New Bioi. 233, 149. Oseroff, A. R., and Callender, R. H. (1974). Resonance Raman spectroscopy of rhodopsin in retinal disk membranes. Biochemistry 13, 4243. Pande, J., Callender, R. H., and Ebrey, T. G. (1981). Resonance Raman study of the primary photochemistry of bacteriorhodopsin. Proc. Nat. Acad. Sci. 78, 7379. Pettei, M. J., Yudd, A. P., Nakanishi, K., Henselman, R., and Stoeckenius, W. (1977). Identification of retinal isomers isolated from bacteriorhodopsin. Proc. Nat. A cad. Sci. 16, 1955. Rimai, 1., Kilponen, R. G., and Gill, D. (1970). Resonance-enhanced Raman spectra of visual pigments in intact bovine retinas at low temperatures. Biochem. Biophys. Res. Commun. 41, 492. Rimai, 1., Heyde, M. E., and Gill, D. (1973). Vibrational spectra of some carotenoids and related linear polyenes. A Raman spectroscopic study. J. Am. Chern. Soc. 95, 4493. Rosenfeld, T., Honig, B., Ottolenghi, M., Hurley, J., and Ebrey, T. G. (1977). Cis-trans isomerization in the photochemistry of vision. Pure Appl. Chem. 49, 341. Rothschild, K. J., Andrew, J. R., De Grip, W. J., and Stanley, H. E. (1976). Opsin structure probed by Raman spectroscopy of photoreceptor membranes. Science 191, 1176. Sandorfy, C. (1980). Spectroscopic aspects of the mechanism of vision. In Proc. VlIth Int. Con! Raman Spectrosc. W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 532-535. Stockburger, M., Klusmann, W., Gattermann, H., Massig, G., and Peters, R. (1979). Photochemical cycle of bacteriorhodopsin studied by resonance Raman spectroscopy. Biochemistry 18, 4886. Stockburger, M., and Massig, G. (1980). RR Studies on the photochemical cycle of bacteriorho-
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dopsin. In Proe. VIlth Int. Con! Raman Speetrose. W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 558-559. Stoeckenius, W., and Lozier, R. H. (\ 974). Light energy conversion in Halobacterium halobium. J. Supramol. Struct. 2, 769. Stoeckenius, W., Lozier, R. H., and Bogomolni, R. A. (1979). Bacteriorhodopsin and the purple membrane of Halobacteria. Biochim. Biophys. Acta 505, 215. Sulkes, M., Lewis, A., Lemley, A. T., and Cookingham, R. (1976). Modeling the resonance Raman spectrum of a metarhodopsin: Implications for the color of visual pigments. Proc. Nat. Acad. Sci. 73, 4266. Sulkes, M., Lewis, A., and Marcus, M. A. (1978). Resonance Raman spectroscopy of squid and bovine visual pigments: the primary photochemistry in visual transduction. Biochemistry 17, 4712. Terner, J., and EI-Sayed, M. A. (\ 978). Time-resolved resonance Raman characterization of the intermediates of bacteriorhodopsin. Biophys. J. 24, 262. Terner, J., Campion, A., and EI-Sayed, M. A. (\977). Time-resolved resonance Raman spectroscopy of bacteriorhodopsin on the millisecond timescale. Proc. Nat. A cad. Sci. 74, 5212. Terner, 1., Hsieh, c.-L., Burns, A. R., and EI-Sayed, M. A. (\979a). Time-resolved resonance Raman spectroscopy of intermediates of bacteriorhodopsin: the bK 590 intermediate. Proc. Nat. A cad. Sci. 76, 3046. Terner, J., Hsieh, c.-L., and EI-Sayed, M. A. (\ 979b). Time-resolved resonance Raman characterization of the bL 550 intermediate and the two dark-adapted bR~66 forms of bacteriorhodopsin. Biophys. J. 26, 527. Terner, J., Hsieh, C.-L., Burns, A. R., and EI-Sayed, M. A. (\979c). Time-resolved resonance Raman characterization of the b0640 intermediate of bacteriorhodopsin. Reprotonation of the Schiff base. Biochemistry 18, 3629. Warshel, A. (1977). Interpretation of resonance Raman spectra of biological molecules. Ann. Rev. Biophys. Bioeng. 6, 273.
CHAPTER
Nonheme Iron Compounds
Among iron-containing proteins, the most well known group is the heme proteins, which include biologically important hemoglobin, cytochrome c, peroxidase, and catalase. The presence of iron is not restricted to the heme proteins; it occurs commonly in other compounds in nature. Hemerythrin is a nonheme oxygen-combining protein found in the blood of some invertebrates. Transferrin and uteroferrin are iron-transporting proteins. A variety of oxygenases and other enzymes contain iron as an integral part. Many nonheme-iron proteins have absorption bands associated with the iron atom in the laser-accessible region; they can be examined by the resonance Raman spectroscopic method. No doubt many more iron compounds will be investigated in the future. So far the main effort is focused on the identification of the Fe-ligand-bond vibrational bands, from which the type of amino acid residues involved in the metal binding can be deduced. In the case of a special nonheme-iron protein that transports molecular oxygen, such as hemerythrin, most effort is directed to the identification of 0-0 and Fe-O stretching
Hemerythrin
299
vibrations. Detailed analysis of such vibrations enables us to understand how the oxygen molecule is attached to the iron atom.
1.
HEMERYTHRIN
Hemerythrin is used to transport oxygen in the blood of some marine invertebrates, for example, the branchiopods, sipunculans, polychaetes, and priapulids. In hemerythrin, two iron atoms are required to bind one molecule of oxygen. Iron is coordinated only to the protein, and the identity of the ligands is a focus of extensive research. Although hemoglobin and hemerythrin have similar names, and both bind to molecular oxygen, their structures and oxygen-binding active cores are quite different. The physiologically active forms of hemerythrin are oxyhemerythrin and deoxyhemerythrin. The physiologically inactive form is methemerythrin, which is an oxidized form. There is a good review article on this topic (Klotz et aI., 1976). Practically all Raman spectroscopic investigations so far have focused on the nature of bound oxygen and the iron-oxygen bond. 1.1.
BOUND OXYGEN
There are two resonance Raman bands at 844 and 500 cm- 1 for oxyhemerythrin. When oxyhemerythrin is prepared with 18 2 , these two bands shift to 798 and 478 cm- I , respectively, indicating that these bands are associated with the 02 vibration. Since the two bands are polarized, they must be associated with totally symmetrical modes. Therefore, the 844-cm - I band is assigned to an 0-0 symmetrical stretch. The 500-510 cm- I band is assigned to the Fe-O vibration (Dunn et aI., 1975; Freier et aI., 1980).
°
Fe(III)···· Fe(UI)-O
\
o
or
Fe (I II) ;t'
o I o ,
Fe(lll)
FIGURE 11.1. Possible manner of oxygen binding to hemerythrin. The figure was redrawn from the paper of Kurtz et al. (1976).
300
Nonheme Iron Compounds
When hemerythrin was mixed with unsymmetrical isotopic ligand 16 0 18 0, a doublet at 825 and 818 cm- I was observed (Kurtz et al., 1976; Kurtz, 1977). These two bands were assigned to the dioxygen stretching motion of Fe_ 18 0_ 16 0 and Fe_ 16 0_0 18 , respectively. The reason the two types of Fe-O-O give different frequencies is the difference in the bending-force constants of Fe- 18 0- 16 0 and Fe- 16 0- 18 0. Based on this spectroscopic finding, it is suggested that oxyhemerythrin possesses the structure shown in Figure 11.1. The 0-0 stretching frequencies of other compounds are listed here for comparison: Compound Molecular oxygen, O2 Superoxide,O-O-H Peroxide, H-O-O-H Oxyhemoglobin Oxyhemocyanin
Frequency (em-I)
Reference
1555 1101 878 1107 742
Herzberg (1945) Milligan and Jacox (1963) Bain and Giguere (1955) Barlow et al. (1973) Loehr et al. (1974)
By comparison with different types of 0-0 stretching vibrations, it is reasonable to conclude that oxygen binds to hemerythrin as a form of peroxide
2-
~
Fe(III)- 0-0 -Fe(III). The 844- and 500-cm - I bands are both maximally enhanced by excitation at 525 nm. A transition at 520 nm is indicated in the CD spectrum of oxyhemerythrin; it is thought to arise from a charge transfer between 01- and Fe(III) (Dunn et al., 1977). Biologically inactive methemerythrin is the oxidized form of hemerythrin containing two ferric irons per subunit. But it reversibly binds to a number of anionic ligands. Because these anions bind to the same or near the oxygenbinding site, use of methemerythrin and anionic ligands is useful in elucidating the active-site structure. Sulfide forms a complex with methemerythrin, and there is only one band in the resonance Raman spectrum, at 444 cm- 1 (Freier et al., 1979). The band is assigned to the Fe-S stretching vibrational mode. It is possible that sulfidomethemerythrin contains a IL-sulfido bridge Fe(III)-S2- -Fe(III). 1.2.
OTHER STUDY
There are two irons in hemerythrin, both are attached to the protein through the functional groups of amino acid residues; the irons are 3.0-3.5 A apart. One iron is coordinated to three His, a bridging Glu, and a bridging Asp. The other iron is coordinated to two His, one Tyr, and the same two bridging ligands, Glu and Asp. There is one natural question: whether the active iron sites are the same for hemerythrins of different species. Comparison of Raman ...
301
Hemerythrln
frequencies and enhancement profiles of hemerythrin from four species suggests that the basic active-site structure is the same (Dunn et aI., 1977). Hemerythrin can exist as an octamer or a monomer, but in the blood of most organisms, hemerythrin exists as an octamer consisting of eight identical subunits. However, there are marked differences among these complexes with respect to their exchange behavior in H 2 18 0. This difference can be due to subunit interactions in the oligomer or to local differences in structures at the functional site of each monomer. The ligand-Fe-bond vibrational frequencies of monomer and octamer are found to be the same and appear at 510 cm- 1. The Fe-O frequencies are thus not dependent on the quaternary structure of hemerythrin. It seems that the functional site hemerythrin exists in at least two conformational states (Duff et aI., 1981). The oxidation state of iron is not involved in the maintenance of the octametric ensemble. The interrelationship of deoxyhemerythrin, oxyhemerythrin, and methemerythrin and their polymeric and monomeric forms are illustrated in Figure 11.2.
~~ c;oIor....
-
,0-
II
~
(••SHrF.2»' . Mol •• t . - 107,000
s.o.-7.
Q
..,,~
e- 0
"\ ""f: ~p"
O'"
»,
(--HrF. I 01 Mol .• t. 107,000 SlOw -7,
.
met.
"
~
'
""e~ C)
\,,(F,ICNloI
O III
8 RH9SHrF.I Mol.• t. 13,500
SlOW- 2, ID (--SHrF.I Mol.•t. 107,000 51011-7,
»,
FIGURE 11.2. Macromolecular properties and interrelationships between deoxyhemerythrin, oxyhemerythrin, and methemerythrin, and their monomeric subunit. The figure was reproduced from Klotz et al. (1976) by permission of the copyright owner, American Association for the Advancement of Science.
302 2.
Nonheme Iron Compounds
TRANSFERRIN AND UTEROFERRIN
2.1.
TRANSFERRIN
Transferrin is an iron-transport protein that provides iron for the biosynthesis of hemoglobin and other iron-containing proteins. Only ferric ion can combine with the apoprotein to form transferrin. Transferrin can be found in egg white, milk, and serum (Aisen and Listowsky, (980). The binding constant is extremely high: 10 24 . Such a high binding constant enables transferrin to protect its bound iron from chelation with other proteins, yet transferrin is an iron-transport protein, so it releases the iron when the need arises. Upon encounter with a reticulocyte, the binding constant changes dramatically, and iron is released to a membrane-bound iron-receptor site for eventual incorporation into hemoglobin. Because of its high affinity for ferric iron, lactoferrin (milk transferrin) may serve as a bacteriostatic agent. Despite its high binding constant, the binding site is flexible enough to accommodate other metal ions forming Cu(II)-, Cr(lII)-, Mn(III)-, and Co(III)-transferrin complexes. Ferric transferrin has an absorption maximum at 460 nm, so the resonance Raman spectrum can be obtained by the use of the excitation lines at 488.0 or 514.5 nm. Judging from the resonance Raman bands, either tyrosine or histidine is involved in iron binding in ovotransferrin (Tomimatsu et aI., 1973; Carey and Young, 1974). Bicarbonate is essential for specific iron binding, but it is not involved at the iron-binding site. This is shown by the fact that an 18 0 substitution in bicarbonate does not shift the band frequencies (Gaber et aI., 1974). The binding of iron increases the net negative charge of transferrin due to the following reaction: Fe(III)
+ 3H+ (from transferrin) + HCOj --->
Fe(III)-transferrin-HCO;
+ 3H+
The Raman bands at 300 and 360 cm- I in the spectrum of the iron-transferrin-bicarbonate complex are assigned to HCO; -Fe(III) vibration (Van Kreel et aI., 1972). By comparing the Raman spectra of model compounds it has been shown that tyrosine is a ligand in ovotransferrin and serum transferrin (Tomimatsu et aI., 1976). Lactoferrin occurs in high concentration in milk and has a molecular weight of 80,000, which is similar to that of other transferrins. However, lactoferrin can be differentiated immunologically from other transferrins. Lactoferrin shows resonance Raman spectral bands at 1605, 1505, 1275, and 1175 cm- I ; these bands are characteristic of tyrosine coordination with metal ions. The fact that the peak at 1270 cm - I is particularly affected by a change in the metal ion indicates that the tyrosine coordination of metals is the same in lactoferrin as in ovotransferrin and serum transferrin ~
Oxygenases
303
(Ainscough et aI., 1980). This confirms the results of other studies (EPR and fluorescence) showing that there is close similarity between lactoferrin and serum transferrin. 2.2.
UTEROFERRIN
Uteroferrin is an iron-containing protein isolated from uterine fluid that plays a role in iron transport to the fetus. It contains one high-spin ferric ion per 35,000 daltons of polypeptide. It possesses a purple color, with an absorption maximum at 545 nm. The resonance Raman spectrum of uteroferrin is strikingly similar to that of Fe(III)-transferrin, suggesting that the iron-binding sites of the two proteins must be quite similar. The visible absorption spectrum of uteroferrin results from a phenolate-to-Fe(III) charge transfer (Gaber et aI., 1978, 1979). The optical absorptions of transferrin, ovotransferrin, lactoferrin, and uteroferrin are very similar. Uteroferrin must therefore have an iron core structure similar to those of these compounds.
3.
OXYGENASES
Oxygenases are enzymes that incorporate molecular oxygen into an organic compound. There are two types of oxygenases: dioxygenases and monooxygenases. Dioxygenases incorporate two atoms of oxygen, whereas the others incorporate only one atom. Those most studied by Raman spectroscopy are protocatechuate dioxygenase and pyrocatechase. 3.1.
PROTOCATECHUATE 3, 4-DIOXYGENASE
Protocatechuate 3,4-dioxygenase (Ee 1.13.1.3) contains high-spin Fe(III) and is an aromatic-ring-cleaving enzyme that catalyzes the intradiol oxygenation of 3,4-dihydroxybenzoate (protocatechuate) to fi-carboxy-cis-cis-muconic acid (Figure 11.3). The enzyme is red, with a broad absorption around 450 nm due to chelation of the ferric ion. There are four principal resonance-enhanced peaks at 1602, 1503, 1263 and 1171 cm- 1 with a 5l4.5-nm excitation line (Keyes and Loehr, 1978). These frequencies are associated with ring-mode vibrations of one or COO-
~~ OH
FIGURE 11.3.
,
A+ lL_
2H+
!OOCOO-
Reaction catalyzed by protocatechuate 3,4-dioxygenase.
304
Nonheme Iron Compounds
more tyrosine residues coordinated with the Fe(III). The fact that there are great similarities between the resonance Raman spectra of this oxygenase and those of the transferrins suggests the existence of a class of proteins characterized by Fe(III)-tyrosine coordinations. The similarity of resonance-enhanced bands can be seen in Table 11.1. The iron atom of the oxygenase chelates with the tyrosine residue, as evidenced by the resonance Raman spectrum due to the coordination between the p-hydroxyphenyl group and iron. Moreover, Fe-ligand vibrations can be seen at 465,423, and 371 cm- I . An important question concerns where O2 attaches to the substrate and to the enzyme. When the enzyme is attached to the substrate, there are no changes in the tyrosine-ring vibrational modes at 1263 and 1174 cm -I. However, in the ternary complex of the 02-substrate-enzyme system, the tyrosinate vibrational modes shift to 1252 and 1165 cm- I . Judging from the Raman spectra, O2 attaches only to the substrate. However, there is no spectral evidence for the presence of dioxygen in the metal-ligand chromophore. If bound dioxygen is a peroxide ion, it should give 740- and 850-cm -1 vibrational bands, a superoxide-ion band at 1107 cm -!, and a Fe-O vibration band between 500 and 570 cm -I. Lack of such bands suggests that the iron is not coordinated to peroxide or superoxide in the enzyme-substrate-oxygen complex. This may suggest that the substrate has already reacted with dioxygen to form some form of intermediate (Keyes et al., 1979).
TABLE 11.1. Resonance Raman Frequencies of Phenolate Ring Vibrations in Iron-Tyrosine Protein and Model Compounds
Compounds Ovotransferrin Serum transferrin Lactoferrin Uteroferrin Protocatechuate 3,4-dioxygenase From Ps. acruginosa From Ps. putida Pyrocatechase p-Creso1-Fe3+ ,pH 7.0 p-Creso1, pH 14 Fe(EDDHA)Fe(salhishC104, pH 7.0 Assignments
Source:
Frequencies 1605 1613 1604 1607
1504 1508 1500 1504
1270 1288 1272 1293
1170 1174 1170 1173
1605 1605 1605 1618 1607 1600 1625 1605 Ring Stretch
1505 1504 1505 1488 1490 1482 1476 1452 Ring Stretch
1265 1270 1289 1222 1276 1286 1337 1310 CO Stretch
1176 1175 1175 1180 1176 1168 1159 1132 CH Bend
The table was reproduced from Que et al. (1980). ..ill
Oxygenases
305
Similarities in the Raman spectra of protocatechuate 3,4-dioxygenase and transferrin are well recognized; therefore, it is generally recognized that the iron in the two proteins binds to tyrosine phenolic oxygen. Yet there is some evidence that they may have different chromophore structures. The absence of strong low-frequency bands suggests the involvement of an additional ligand in protocatechuate 3,4-dioxygenase chromophore that is not present in transferrin (Bull et al., 1979). It is fairly well recognized that iron binds to the tyrosine residue. However, the anionic phenolate ion, rather than undissociated phenolic OH, of the tyrosine residue is the real ligand. When the iron is removed, the spectrum of the apoenzyme does not show the bands (1177,1265, 1505, and 1605 cm- I ) shown in the spectrum of the native enzyme. The original bands in the native enzyme do not shift in a D 2 0 solution, indicating that the vibrational bands originated from groups that do not contain replaceable hydrogen. p-Cresol at pH 7.0 and 14 shows four lines. Thus it can be concluded that the phenolate ion coordinates with the iron atom in protocatechuate 3, 4-dioxygenase (Tatsuno et al., 1978). There is no change in tyrosine ligation after binding to substrate or inhibitor, since there are no detectable changes in the Raman spectra. This indicates that the tyrosine ligands present in the resting enzyme are not displaced (Tatsuno et al., 1978; Felton et al., 1978; Keyes et al., 1979). Enzyme-substrate and enzyme-inhibitor complexes have different Raman spectra, yet retain the tyrosine peaks observed in the native enzyme, suggesting that the tyrosine ligation is not altered upon substrate or inhibitor binding (Felton et al., 1978; Que and Epstein, 1981). 3.2.
PYROCATECHASE
Pyrocatechase (EC 1.13.1.1) is also a dioxygenase, fixing two atoms of oxygen into the product from 2 mol of molecular oxygen (Figure 11.4). Like protocatechuate 3,4-dioxygenase, pyrocatechase catalyzes the cleavage of intradiols such as catechol to cis, cis-muconic acid. A visible absorption band (450 nm) arises from a tyrosinate-to-iron charge transfer, and the enzyme can be resonance enhanced due to tyrosinate vibrations. The Raman spectrum of pyrocatechase has peaks at 1173, 1293, 1505, and 1605 cm - I that are assigned to tyrosine. Therefore, like protocatechuate 3, 4-dioxygenase, pyrocatechase is an enzyme with iron coordinated to tyrosine (Que and Heistand, 1979). Felton et al. (1978) found addition of catechol (substrate) and 4-nitrocatechol (inhibitor) markedly changes the Raman spectra. Que and Heistand (1979) suggest that catechol does bind to the active-site Iron.
I
OC ?'
~
OH
+ OH
FIGURE 11.4.
18
O2
.
pyrocatechase
(
18
?'
eOOH
~
dbOH
Reaction catalyzed by pyrocatechase.
306
Nonheme Iron Compounds
Many iron-tyrosinate proteins and model compounds have similar resonance Raman vibrational bands due to the phenolate ring. The band frequencies of these compounds were summarized by Que et aI. (1980) and are shown in Table 11.1. In pyrocatechase, two tyrosines coordinate with the active-site iron. The two tyrosines exhibit different C-O stretching vibrations at 1260 and 1290 em-I. The 1260-cm -1 line has an excitation profile different from that of the l290-cm -1 line. One tyrosine is more susceptible to changes in the active-site environment. This suggests that there are two distinctly different charge-transfer interactions (Que et aI., 1980). Pyrocatechase is the only example thus far among iron-tyrosine proteins in which the tyrosines coordinating with the iron are distinguishable in Raman spectra.
4.
FERREDOXIN AND RELATED COMPOUNDS
One of the well-known examples of a nonheme-iron compound is ferredoxin. Ferredoxin and other related compounds are iron-sulfur proteins that form sulfur-atom bridging ligands. Although there are many iron-sulfur proteins, the manner of arrangement of the Fe and S atoms is different. So far three well-defined types of Fe-S centers in such proteins have been found. One is a binuclear cluster, as shown in Figure 11.5; this type represents the Fe-S clusters of plant-type ferredoxins, hydroxylase ferredoxins, and chloroplast ferredoxins. The second type is found in rebredoxins and is a tetrahedron shape, as shown in Figure 11.6. The third type of Fe-S cluster can be found in
s
s (Cys)
s (Cys)
(Cys)
FIGURE n.s. Binuclear-cluster-type Fe-S linkages in plant ferredoxins, hydroxylase ferredoxins, and chloroplast ferrodoxins.
...
Ferredoxin and Related Compounds
307
o
(Cys.g)
Approximately tetrahedral
FIGURE 11.6. Tetrahedron-type Fe-S linkages in rebredoxins.
bacterial ferredoxin; it has a cubane-type shape in which the iron and sulfur atoms occupy alternating corners (Figure 11.7). Some iron-sulfur proteins do not belong to any of these three groups; they have complex iron-sulfur arrangements that have not yet been well studied. When ferredoxin is acidified, H 2 S is released, indicating that the sulfur atoms are labile. Adrenoredoxin, putidaredoxin, and chloroplast ferredoxin
tS\ ~}
FIGURE 11.7. Cubane-type Fe-S linkages in bacterial ferredoxins.
308
Nonheme Iron Compounds
contain two sulfur and two iron atoms, whereas some bacterial ferredoxin contains at least four iron and four sulfide atoms, and possibly more. The ferrodoxins are involved in the electron-transport chains. One of the most interesting physical properties of such Fe-S compounds is the antiferromagnetic interactions between two iron atoms. Antiferromagnetism refers to the antiparallel spin of a neighboring iron atom; therefore, these compounds have no magnetism. Raman spectroscopy has been applied to iron-sulfur proteins only recently, and the majority of studies in this field have been carried out with other physical techniques. In this section only the Raman spectroscopic investigation is summarized. Readers who wish to see a more general view and detailed information are directed to other articles (Aisen and Listowsky, 1980; Averill and Orme-Johnson, 1978; Sweeney and Rabinowitz, 1980) or the book on iron-sulfur proteins by Lovenberg (1973). 4.1.
FERREDOXIN
The resonance Raman spectrum of Clostridium pasteurianum ferredoxin shows two peaks at 345 and 360 cm -1. The lower frequency is assigned to bridging Fe-S and the higher frequency to the terrninal-S vibrational modes (Tang et a1., 1975). For oxidized spinach ferredoxin, 284-, 330-, and 395-cm - I bands are assigned to Fe-S stretching frequencies. Reduced ferredoxin shows only the 330-cm -1 band in this region (Blum et a1., 1977). Other Raman bands are seen at 365, 770, 1080, 1475, and 1930 cm- 1 for oxidized ferredoxin and at 590, 770, 1070, and 1480 cm- 1 for the reduced form at 300°K. These observed bands are close to the values predicted from the Gibson model. The 2840- and 2950-cm- 1 bands are assigned to a cysteinyl C- H stretching vibration. This assignment is reasonable, as the cysteine residues are involved in terminal Fe-S bridging. Some ferredoxins contain unusual types of iron-sulfur clusters. For instance, a ferredoxin from Desulfovibrio gigas contains three iron atoms per monomer, and a ferredoxin from Azotobacter vinelandii contains seven iron atoms. In the latter case, two types of Fe-S clusters are found; one is the normal 4Fe-4S type, whereas the other is a noble type of 3Fe-3S. The Raman spectrum of the 3Fe-3S type of clusters is markedly different from those of rubredoxins or the normal 4Fe-4S type of clusters (Johnson et aI., 1981). 4.2.
RUBREDOXIN
Originally rubredoxin was isolated from C. pasteurianum during the isolation of ferredoxin, but it is also present in many aerobic and anaerobic bacteria. Ferredoxin is dark brown, whereas rubredoxin is red. Rubredoxin contains only one iron atom, which is coordinated with four cysteinyl sulfur atoms
Ferredoxin and Related Compounds
309
(Figure 11.6). The Fe-S bonds of many metalloproteins are actually involved in electronic transitions, so the resonance Raman spectra should reflect the Fe-S vibrations of cysteine and inorganic-sulfur ligands. The Raman bands associated with the iron-containing moiety of rubredoxin are, indeed, intense. Understanding the Fe-S bond of many iron-sulfur-containing compounds is very important, as this bond is a primary factor in unique metabolic and electron-transfer reactions. The resonance Raman spectrum of both amorphous and crystalline samples of rubredoxin shows only two bands at 311 and 365 em ~ I. These bands are assigned to the symmetrical and antisymmetrical stretching modes of the Fe-S4 tetrahedron (Long and Loehr, 1970). All four vibrations of the tetrahedron type bonds were assigned by Long et al. (1971). They are as follows: Vibration Stretching vibration Bending vibration Stretching vibration Bending vibration
Wave Number (em-I) 314 126 368 150
The Raman spectrum of rubredoxin in solution is very similar to that of the solid. This indicates that the tetrahedral coordination about the iron atom by four cysteinyl sulfurs known to exist in the crystalline form is maintained in aqueous solution. Oxidized rubredoxin has a high-spin (8 = 5/2) iron, and on reduction it is converted to a ferrous high-spin (8 = 4/2) state with little or no change in coordination geometry. The iron(III) can be substituted with Co(II) with retention of some enzymatic activity. Substitution of iron by cobalt is useful in identifying metal-binding groups and the nature of coordination at the active site. Cobalt rubredoxin is less active than native rubredoxin but is much more stable toward denaturation and metal dissociation than the native enzyme. The cobalt-rubredoxin has two resonance Raman bands at 343 and 419 cm- I compared with iron-rubredoxin, which has bands at 313 and 365 cm-I. The 343- and 419-cm- 1 bands are Co(II)-S(Cys) stretching vibrational bands. The substituted rubredoxin contains a Co(II)S04 core in a distorted tetrahedral shape (May and Kuo, 1978).
4.3.
ADRENODOXIN
Adrenodoxin activates oxygen to hydroxylate steroids in mitochondria of the adrenal cortex, and it transfers an electron from reduced flavoprotein and to oxidized cytochrome P-450. It is believed that each iron is tetrahedrally coordinated with two labile sulfur atoms and two cysteine sulfur atoms.
310
Nonheme Iron Compounds
The intensity of the resonance Raman band at 995 cm -) depends on the excitation wavelength. This suggests that the optical absorption (at 400500 nm) bands are indeed labile sulfur-iron charge-transfer bands. Adrenodoxin shows three bands at 397, 350, and 279 cm -) that are absent in apoenzymes. The 397- and 279-cm- 1 bands are associated with vibrations of the labile sulfur atoms; the 350-cm- 1 band is for Fe-S(Cys) bonds. Selenium substitution also shows three bands at 355, 350, and 263 cm - I. The Fe-Se charge-transfer bands are observed at 438 and 480 nm fOf the oxidized form and at 580 nm for the reduced form (Tang et al., 1973). The 2944-cm -1 band is assigned to the C-H stretch of cysteinyl residues that are involved in the Fe-S bridge. The 995-cm -) band is assigned to the transition from the antiferromagnetically coupled ground state to the S = 1 first excited state. The Gibson model predicts the next transition to the second excited state to be at 3 X 995 = 2985 cm -I. The band at 2975 cm - 1 agrees almost exactly with this frequency_ The dependence of the 995-cm -) band on excitation wavelength suggests that the optical absorption bands are labile sulfur-iron charge-transfer bands (Adar et aI., 1977). The Fe-S bond vibration is very important in elucidating the active site of iron-sulfur proteins. Observed S-S and Fe-S frequencies in iron-sulfur proteins and synthetic analogs are shown in Table 11.2.
TABLE 11.2. Observed S-S and Fe-S Frequencies in Iron-Sulfur Proteins and Synthetic Analogs O Molecule S,Fe,(CO).
Structure
S-s
.,-./
'- F.
"(em-I)
Assignment
Reference
554 329
S-S Fe-S
Scovell & Spiro (1974)
350
Fe-S
Scovell & Spiro (1974)
/Fe, (SCH,),Fe,(CO).
CH,S
'-F./
cus"
5CH,'
/SC)'1
F. /'-SCr'
oxidized rubrtdoxin (c. pasreurianum) eylS
365 311
oxidized adrenodoxin (beef adrenal glands) oxidized ferredoxin (spinach)
C)'IS,
/sc,.
/'"
h
F,
/ ' -s/ ' -SCp
CysS
oxidized ferredoxin (C. pasteur;tlnum)
397 297 350 395 284 330 360 345
1./) IJ I
of Fe-5.. tetrahedron of Fe-S. tetrahedron
Fe-Slabile Fe-Slabile
Long el al. (1971)
Tang el a!. (1973)
Fe-S eYi Ft-Slabile
Blum el a!. (1977)
Fe-Slabll. Fe-SCy. Fe-S efs Fe-SCYS
TanJ el a!. (1975)
Fe-SCy.
Tang e' a!. (1975)
c)"\ reduced hi(th-potential iron protein (tNomatum)
Fe.S.(SCH, Ph). ,-
F.-S
I's-{'F.
/SC,.
s,J-F,I-sc" F.-S
365 338 250 332 275
Fe-Smercaptidt> fe-SlabUe
Tang.1 a!. (1975)
451 509
S-S S-S
Janz el a!. (1976) Nicki... (1968)
/
FC'-ScYt
Fe-SlAbU.
F.S Na.S. H,S,
"This table was reproduced from Freier et aI. (1979).
TABLE 11.3. Typical Metal- Ligand Vibrational Frequencies for Divalent and Trivalent Metal Ions and Various Ligands
Type of Binding
Ligand
Carboxylate
°II
Range of p(M - L), cm- 1 Fe(II)
M-O-C-
350-530
M-O~C_ M-O.;7
400-560
Phenolate
M - O - @ -545-560
Aqua
M-OH 2
Hydroxo (methemoglobin)
M-OH
310-405
Fe(III)
References
McConnell and Nuttall (1977)
Percy and Stenton (1976) 490-540
Adams (1968)
497
Asher et al. (1977)
500-580
Asher et al. (1977)
363 (sym.) 538 (sym.)
Burke et al. (1978) Hewkin & Griffith (1966)
503
Dunn et al. (1973)
H Hydroxobridged p.-Oxobridged
M
/0"
480-560 M
M-O--M M
/,0"
M
Peroxo (oxyhemerythrin)
M-O-O
Superoxo (oxyhemoglobin)
M-O-O
Amine
M-NH 2
Histidine (oxyhemocyanin)
A
M-ioi
Histidine (azurin) Cysteinato (azurin) Cysteinato (Fe-S proteins) -
M-S
300-440
570
Brunner (1974)
510-590
Adams (1968) Nakamoto (1978)
265-285
Freedman et al. (1976)
400-425
Thamann (1980)
369
Thamann (1980)
300-400
Spiro and Loehr (1975)
Source: The table was reproduced from Sjoberg et aI. (1980).
311
312
Nonheme Iron Compounds
s. OTHERS Ribonucleotide reductase converts ribonucleotides to their corresponding deoxyribonucleotides. It has been isolated from E. coli and consists of two nonidentical subunits, proteins Bland B2; the subunit itself has no enzymatic activity. Protein B2 contains two iron atoms per mole of enzyme and at least one tyrosyl free radical that is essential for the enzymatic activity. The iron center in the native B2 protein consists of an antiferromagnetically coupled pair of high-spin Fe(III) atoms. In this respect it is similar to the iron center of hemerythrin. With a chelating agent, the iron atoms can be removed from protein B2, resulting in the disappearance of the radical. Thus B2 becomes an iron-, and radical-free apoprotein B2. Both the native (no tyrosyl radical) and the radical-free enzyme exhibit a resonance-enhanced Raman band at 496 em -I, which is assigned to the Fe-O vibrational mode. This is evidenced by the isotopic shift from 496 em-I to 481 em-I when the enzyme is suspended in H 2 18 0 converting FeO to Fe 180. Since there are no tyrosinate-ring modes, the possibility of tyrosinate oxygen as a ligand is ruled out. By comparison with Fe model compounds, it was concluded that a carboxylate from either an aspartic or glutamic acid residue or an oxygen-containing group from the solvent, such as H 20, OH- , or a p.-oxo bridge (similar to the iron center in hemerythrin), is a ligand (Table 11.3) (Sjoberg et al., 1980, 1982). Nitrogenase from Azotobacter vinelandii contains not only iron but also molybdenum, sulfur, and coenzyme A. It is important to know its structure and function in order to understand the mechanism of biological nitrogen fixation. As a first step toward this goal Fe, Mo-cofactors, which may be involved in the nitrogenase active site, were investigated by Raman spectroscopy. The Fe-S stretching vibrational band can be observed at 368 em-I. There are also intensive bands at 1600 and 1400 em-I, which are asymmetrical and symmetrical stretching vibrations of the carboxylate ion (-COO-). The result is interpreted as the presence of coenzyme A as a component of the Fe, Mo-cofactors of nitrogenase (Levchenko et al., 1980).
REFERENCES Adams, D. M. (1968). Metal-Ligand and Related Vibrations, St. Martin's Press, New York. Adar, F., Blum, R., Leigh, Jr., J. S., Ohnishi, T., and Salerno, J. (1977). Anti-ferromagnetic exchange in beef adrenodoxin as measured by resonance Raman spectroscopy. FEBS Lett. 84, 214. Ainscough, E. W., Brodie, A. M., Plowman, 1. E., Bloor, S. 1., Loehr, J. S., and Loehr, T. M. (1980). Studies on human lactoferrin by electron paramagnetic resonance, fluorescence, and resonance Raman spectroscopy. Biochemistry 19, 4072. Aisen, P., and Listowsky, 1. (1980). Iron transport and storage proteins. Ann. Rev. Biochem. 49,
357.
.
j
References
313
Asher, S. A, Vickery, L. E., Schuster, T. M., and Sauer, K. (1977). Resonance Raman spectra of methemoglobin derivatives. Selective enhancement of axial ligand vibration and lack of an effect of inositol hexaphosphate. Biochemistry 16, 5849. Averill, B. A, and Orme-Johnson, W. H. (1978). Iron-sulfur proteins and their analogs. In Metal Ions in Biological Systems, H. Sigel, Ed., Dekker, New York, pp. 127-183. Bain, 0., and Giguere, P. A (1955). Hydrogen peroxide and its analogues. VI. Infrared spectra of HzO z , DzO z , and HD01. Can. 1. Chem. 33,527. Barlow, C. H., Maxwell, J. c., Wallace, W. J., and Caughey, W. S. (1973). Elucidation of the mode of binding of oxygen to iron in oxyhemoglobin by infrared spectroscopy. Biochem. Biophys. Res. Commun. 55, 91. Blum, H., Adar, F., Salerno, J. C., and Leigh, Jr., J. S. (1977). Exchange coupling in spinach ferredoxin determined by resonance Raman spectroscopy Biochem. Biophys. Res. Commun. 77,650. Brunner, H. (1974). Identification of the iron-ligand vibration of oxyhemoglobin. NatUlwissenscha/ten 61, 129. Bull,
c., Ballou, D. P., Salmeen, I. (1979). Raman spectrum of protocatechuate dioxygenase from Pseudomonas putida; new low frequency bands. Biochem. Biophys. Res. Commun. 87, 836.
Burke, J. M., Kincaid, J. R., Peters, S. Gagne, R. R., Collman, J. P., and Spiro, T. G. (1978). Structure-sensitive resonance Raman bands of tetraphenyl and "picket fence" porphyrin-iron complexes, including an oxyhemoglobin analogue. 1. Am. Chem. Soc. 100,6083. Carey, P. R., and Young, N. M. (1974). The resonance Raman spectrum of the metalloprotein ovotransferrin. Can. J. Biochem. 52, 273. Duff, L. L., Klippenstein, G. L., Shriver, D. F., and Klotz, I. M. (1981). Multiple conformations at functional site of hemerythrin: Evidence from resonance Raman spectra. Proc. Nat. A cad. Sci. USA 78,4138. Dunn, J. B. R., Shriver, D. F., and Klotz, I. M. (1973). Resonance Raman studies of the electronic state of oxygen in hemerythrin. Proc. Nat. A cad. Sci. 79, 2582. Dunn, J. B. R., Shriver, D. F., and Klotz, I. M. (1975). Resonance Raman studies of hemerythrinligand complexes. Biochemistry 14, 2689. Dunn, J. B. R., Addison, A W., Bruce, R. E., Loehr, J. S., and Loehr, T. M. (1977). Comparison of hemerythrins from four species of sipunculids by optical absorption, circular dichroism, nuorescence emission, and resonance Raman spectroscopy. Biochemistry 16, 1743. Felton, R. H., Cheung, L. D., Phillips, R. S., and May, S. W. (1978). A resonance Raman study of substrate and inhibitor binding to protocatechuate-3,4-dioxygenase. Biochem. Biophys. Res. Commull. 85, 844. Freedman, T. B., Loehr, J. S., and Loehr, T. M. (1976). A resonance Raman study of the copper protein, hemocyanin. New evidence for the structure of the oxygen-binding site. J. Am. Chern. Soc. 98, 2809. Freier, S. M., Duff, L. L., Van Duyne, R. P., and Klotz, I. M. (1979). Resonance Raman studies and structure of a sulfide complex of methemerythrin. Biochemistry 24, 5372. Freier, S. M., Duff, L. L., Shriver, D. F., and Klotz, 1. M. (1980). Resonance Raman spectroscopy of iron-oxygen vibrations in hemerythrin. Arch. Biochem. Biophys. 205,449. Gaber, B. P., Miskowski, V., and Spiro, T. G. (1974). Resonance Raman scattering from iron(III)and copper(II)-transferrin and an iron(III) model compound. A spectroscopic interpretation of the transferrin binding site. J. Am. Chern. Soc. 96, 6868. Gaber, B. P., Sheridan, J. P., and Roberts, R. M. (1978). Raman scattering from progesteroneinduced glycoprotein. In Proc. Sixth Jill. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 112-113.
314
Nonheme Iron Compounds
Gaber, B. P., Sheridan, J P., Bazer, F. W., and Roberts, R. M. (1979). Resonance Raman scattering from uteroferrin, the purple glycoprotein of the porcine uterus. J. BioI. Chem. 254, 8340. Hawkin, D. J, and Griffith, W. P. (1966). Infrared spectra of binuclear complexes. J. Chem. Soc. (A), 472. Herzberg, G. (1945). Infrared and Raman Spectra of Polyatomic Molecules. Van Nostrand, New York. Janz, G. J, Roduner, E., Coutts, J. W., and Downey, Jr., J R. (1976). Raman studies of sulfur-containing anions in inorganic polysulfides. Barium trisulfide.lnorg. Chem. IS, 1751. Johnson, M. K., Hare, J. W., Spiro, T. S., Moura, J J G., Xavier, A V., and LeGall, J (1981). Resonance Raman spectra of three-iron centers in ferredoxins from Desulfovibrio gigas. J. Bioi. Chem. 256, 9806. Keyes, W. E., and Loehr, T. M. (1978). Raman spectral evidence for tyrosine coordination of iron in protocatechuate 3,4-dioxygenase. Biochem. Biophys. Res. Comm. 83, 941. Keyes, W. E., Loehr, T. M., Taylor, M. L., and Loehr, J. S. (1979). Protocatechuate 3,4dioxygenase. Resonance Raman studies of the oxygenated intermediatc. Biochem. Biophys. Res. Comm. 89,420. Klotz, 1. M., Klippenstein, G. L., and Hendrickson, W. A (1976). Hemerythrin: Alternative oxygen carrier. Science 192, 335. Kurtz, Jr., D. M. (1977). Structural investigations of the active sites of hemcrythrin and hemocyanin by isotopic vibrational analyses of resonance Raman spectra. Diss. Abstr. Till. B, 38(9), 4205. Kurtz, D. M., Shriver, D. F., and Klotz, I. M. (1976). Resonance Raman spectroscopy with unsymmetrically isotopic ligands. Differentiation of possible structure of hemerythrin complexes. J. Am. Chem. Soc. 98, 5033. Levchenko, L. A, Roschupkina, O. S., Sadkov, A P., Marakushev, S. A, Mikhailov, G. M., and Borod'ko, Yu. G. (1980). Spectroscopic investigation of FeMo-Cofactor. Coenzyme A as one of the probable components of an active site of nitrogenase. Biochem. Biophys. Res. Commun. 96, 1384. Loehr, J S., Freedman, T. B., and Loehr, T. M. (1974). Oxygen binding to hemocyanin: A resonance Raman spectroscopic study. Biochem. Biophys. Res. Commun. 56, 510. Long, II, T. V., and Loehr, T. M. (1970). The possible determination of iron coordination in nonheme iron proteins using laser-Raman spectroscopy. Rubredoxin. J. Am. Chem. Soc. 92, 6384. Long, II, T. V., Loehr, T. M., Allkins, JR., and Lovenberg, W. (1971). Determination of iron coordination in nonheme iron proteins using laser-Raman spectroscopy. II. Clostridium pasteurianum rubredoxin in aqueous solution. J. Am. Chern. Soc. 93, 1809. Lovenberg, W. (1973). Iron-Sulfur Proteins, Vol. II, Molecular Biology Series, Academic, New York. May, S. W., and Kuo, J-Y. (1978). Preparation and properties of cobalt (II) rubredoxin. Biochemistry 17, 3333. McConnell, A A, and Nuttall, R. H. (1977). The vibrational spectra of EDTA complexes of divalent tin and lead. Spectrochim. Acta 33A, 459. Milligan, D. E., and Jacox, M. E. (1963). Infrared spectroscopic evidence for the species H0 2 . J. Chem. Phys. 38, 2627. Nakamoto, K. (1978). Infrared and Raman Spectra of Inorganic and Coordination Compounds, 3rd ed., Wiley, New York. Nickless, G. (1968). Inorganic Sulfur Chemistry, Elsevier, Amsterdam. Percy, G. c., and Stenton, H. S. (1976). Infrared and electronic spectra of N-salicylideneglycinate complexes of cobalt and nickel. Spectrochim. Acta 32A, 1615. .J
References
315
Que, Jr., L., and Epstein, R. M. (1981). Resonance Raman studies on protocatechuate 3,4dioxygenase-inhibitor complexes. Biochemistry 20, 2545. Que, Jr., L., and Heistand, II, R. H. (1979). Resonance Raman studies on pyrocatechase. J. Am. Chern. Soc. 101, 2219. Que, Jr., L., Heistand, II, R. H., Mayer, R., and Roe, A. L. (1980). Resonance Raman studies of pyrocatechase-inhibitor complexes. Biochemistry 19, 2588. Scovell, W. M., and Spiro, T. G. (1974). Vibrational and Raman intensity analysis of a ferredoxin model: S2Fe2(COk Inorg. Chern. 13,304. Sjoberg, E.-M., Graslund, A., Loehr, 1. S., and Loehr, T. M. (1980). Ribonucleotide reductase: A structural study of the dimeric iron site. Biochem. Biophys. Res. Commun. 94, 793. Sjoberg, B.-M., Loehr, T. M., and Sanders-Loehr, 1. (1982). Raman spectral evidence for a p.-oxo bridge in the binuclear iron center of ribonucleotide reductase. Biochemistry 21, 96. Spiro, T. G., and Loehr, T. M. (1975). Resonance Raman spectra of heme proteins and other biological systems. In Advances in Infrared and Raman Spectroscopy, Vol. I, R. 1. H. Clark and R. E. Hester, Eds., Heyden, London, New York, Rheine, pp. 98-142. Sweeney, W. V., and Rabinowitz, 1. C. (1980). Proteins containing 4Fe-4S clusters: An overview. Ann. Rev. Biochem. 49, 139. Tang, S.-P. W., Spiro. T. G., Mukai, K., and Kimura, T. (1973). Resonance Raman scattering and optical absorption or adrenodoxin and selenaadrenodoxin. Biochem. Biophys. Res. Commun. 53,869. Tang, S.-P. W., Spiro, T. G., Antanaitis, c., Moss, T. H., Holm, R. H., Herskovitz, T., and Mortensen, L. E. (1975). Resonance Raman spectroscopic evidence for structural variation among bacterial ferredoxin, Hi PIP, and Fe4S4(SCH2Ph)~- . Biochem. Biophys. Res. Commun. 62, I. Tatsuno, Y., Saeki, Y., IWaki, M., Yagi, T., Nozaki, M. (1978). Resonance Raman spectra of protocatechuate 3, 4-dioxygenase. Evidence for coordination of tyrosine residue to ferric iron. J. Am. Chern. Soc. 100,4614. Thamann, T. 1. (1979). Structural investigations of the active sites of azurin, hemerythrin, and hemocyanin, and vibrational analyses of the copper(II) and copper(III) complexes of biuret and oxamide. Diss. Abstr. Int. 41, No. 8109791. Tomimatsu, Y., Kint, S., and Scherer, J. R. (1973). Resonance Raman spectroscopy of iron(III)ovotransferrin. Biochem. Biophys. Res. Commun. 54, 1067. Tomimatsu, Y., Kint, S., and Scherer, 1. R. (1976). Resonance Raman spectra of iron(III)-, copper(II)-, cobalt(lII)-, and manganese(III)-transferrins and of bis(2, 4, 6trichlorophenoJato)diimidazolecopper(II) monohydrate, a possible model for copper(II) binding to transferrins. Biochemistry 15, 4918. Van KreeJ, B. K., van Eijk, H. G., Leijnse, B., and van der Maas, 1. H. (1972). Laser-Raman spectroscopy of the iron-transferrin-bicarbonate complex. Z. Klin. Chern. Klill. Biochem. 10, 566.
CHAPTER
2
Hemes and Porphyrins
The heme of heme proteins can be studied by resonance Raman spectroscopy without interference from scattering by the protein because the intense heme optical absorptions strongly enhance the scattering by heme vibrations. By careful analyses of empirical results, it is sometimes possible to predict the oxidation and spin states of heme iron and the type of axial ligands. It may also be possible to study certain peripheral substituents, as for example, the carbonyl at position 8 in heme a of cytochrome oxidase. This is possible because effects on the chromophore can be reflected in the optical absorption spectra as well as in the resonance Raman spectra. The resonance Raman spectra are due primarily to porphyrin vibrations. However, both oxidation and spin states, as well as quarternary structure and ligand, have indirect effects on the porphyrin. Oxidation and spin states, for example, are properties of the iron atom and not of the porphyrin. There are many good review articles on resonance Raman spectroscopy of heme compounds; readers are advised to see Spiro and Stein (1977), Yu (1977), Kitagawa et al. (1978), and Spiro (1978) for further information.
Vibrational Modes
1.
317
VIBRATIONAL MODES
Heme compounds have two distinct electronic transitions, giving rise to bands in the absorption spectrum near 400 nm (Soret band) and around 500-550 nm (a- and ,B-bands) (Figure 12.1). Resonance-enhanced Raman scattering is observed from the vibrational modes that couple to these electronic transitions. The Raman lines vary in relative intensities depending on the wavelength of laser excitation. The frequency of a vibrational mode cannot change with the excitation. But sometimes two modes with nearly the same frequency may be enhanced differently with different excitations, causing an apparent shift in frequency of an observed band. Raman spectra of heme compounds differ depending on whether they are excited at the Soret or a-band wavelengths (Salmeen et aI., 1973; Spiro and Strekas, 1974). Since the Raman spectra of heme compounds are due to the vibrations of heme rather than those of the protein, heme compounds with identical hemes usually give similar spectra (Strekas and Spiro, 1972a, b; Brunner, 1973; Loehr and Loehr, 1973). Hemeundecapeptide can be obtained from cytochrome c by proteolytic digestion. The Raman spectrum of the hemeundecapeptide is very similar to that of cytochrome c. Raman spectra of many b-type cytochromes
Sorel
,-. , ,, I I I I I I
,,
,, ,, ,
,
I
,,
~ ,.\
I
I~ " '~'''''' '
'
ft
\
400
\._------450
I
I
\
\
"' ...., - , '_ _\
500
550
'
~
_
600
Wavelength (nm)
FIGURE 12.1. Typical absorption spectrum of ----ferrous (reduced) and forms of heme proteins.
ferric (oxidized)
318
Hemes and Porphyrins
are very similar to each other (Adar and Erecinska, 1974; Bullock and Myer, 1978). Using the characteristic Raman lines, b-type can be differentiated from c-type cytochrome (Kitagawa et al., 1975a). For a mixed heme complex such as succinate-cytochrome c reductase, which contains both b- and c-type cytochromes, both hemes contribute to the resonance Raman spectrum. Because of differences in the absorption spectra of the two types of cytochromes, Raman spectra of the reductase depend on the excitation wavelength. At 514.5-nm excitation, the spectrum of reduced succinate-cytochrome c reductase is due to the spectrum of both b- and c-types of cytochromes. The spectrum obtained using light at 568.2 nm is due mostly to b-type cytochrome because of the proximity of the excitation wavelength to its a-absorption band (Adar and Erecinska, 1974). A similar example is found with cytochrome oxidase, which contains both cytochromes a and a 3 • With proper control of the excitation wavelength, two independent hemes can be distinguished by resonance Raman spectroscopy (Woodruff et aJ., 1981). For oxidized cytochrome oxidase, excitation at 441.6 nm enhances the vibrations of only cytochrome a, whereas excitation at 413.1 and 406.7 nm enhances both low-frequency vibrations of cytochrome a 3 and high-frequency vibrations of a and a 3 (Babcock and Salmeen, 1979; Ondrias and Babcock, 1980). Each heme compound has its own characteristic Raman lines. In mixed hemes sometimes these characteristic lines deviate from the line positions of an individual heme. Such deviation can sometimes be interpreted as heme-heme i3teraction (Adar et al., 1981).
1.1.
EXCITATION PROFILES
One way to study heme compounds by Raman spectroscopy is to measure the intensity of Raman scattering as a function of the excitation frequency. When the wavelength of the laser used to excite the Raman spectrum is near an electronic transition of the molecule, an enormous enhancement of the scattered light is observed (see Chapter 1, see Section 5). This enhancement is due to cancellation of terms in the denominator of the first term in equation 5.1 in Chapter 1 when the laser is tuned through the energy of the electronic transition. In general, a number of peaks in the Raman spectrum will result, since the electronic absorption band is made up of many overlapping vibrational-electronic (vibronic) transitions. That is, the vibrational structure of the electronic transition complicates the absorption band. Resonance with these overlapping vibronic levels gives rise to a complicated dependence of the intensity of a Raman line on a wavelength of the laser light used to excite the spectrum. A plot of the intensity of a Raman line versus the exciting light frequency is called an excitation profile. Improved excitation profiles can now be obtained as continuous-range wavelengths of lasers are becoming more available. Excitation profiles are also called Raman dispersion. Each of the Raman lines has an excitation profile.
,.
~:,:
Vibrational Modes
319
A great deal of detailed information about the excited states of heme can be learned from the excitation profiles of its absorption bands. For example, the vibrational frequencies in the ground and excited electronic states may differ. The Raman spectrum gives the vibrational frequencies of the ground electronic state. In contrast, the vibrational frequencies in an excited electronic state may be obtained from the peak positions in the excitations profiles of that electronic state. Thus the vibrational frequencies of heme after absorbing light into the a-band can be determined from the excitation profiles of the a, ,B-band region by measuring the separation of the two peaks in the excitation profile of each Raman line. This procedure has shown the vibrational frequencies to differ only slightly from the ground-state frequencies. Information about the interaction between the electronic and vibrational motion (vibronic coupling) can also be learned from the excitation profiles. For example, quantitative calculations from Raman excitation profiles show that the strength of the vibronic coupling can be quite different for different porphyrins (Cheung et aI., 1978; Shelnutt and O'Shea, 1978, 1980; Shelnutt et al., 1977). Such calculations can also separate the contributions to the Raman intensity from several possible vibronic-coupling mechanisms. For example, intensity in a Raman line can arise from a single electronic state or from vibrational coupling of electronic states of different energies. In metalloporphyrins and heme proteins, all of these mechanisms can contribute to the excitation profiles, so they are quite complicated. Nevertheless, excitation profiles are powerful tools for studies of excited electronic states of heme and should be considered complementary to absorption spectroscopy for that purpose. By using this technique, one can readily see the difference between Fe(II)cytochrome c and Fe(II)-cytochrome bs. The difference is probably due to different mechanisms of energy transition for the two compounds (Friedman et al., 1977). Using Raman-excitation difference spectra, one can determine if there is a difference in the environments of the chromophores in the two heme proteins (Shelnutt, 1980a). 1.2.
ORIGINS OF VIBRATIONS
The vibrational modes of heme and porphyrins are extremely complicated and have been the subject of many investigations (Ogoshi et al., 1972; Stein et al., 1975; Susi and Ard, 1977; Rimai and Salmeen, 1978). Detailed vibrational modes of four types of symmetry and their frequencies can be seen in Figure 12.2 for Ni-octaethylporphyrin. The frequencies of these vibrational modes can be influenced by the iron spin state, the coordination geometry, and the peripheral substituents; therefore, the actual frequencies of different heme compounds vary slightly. Resonance-Raman-active vibrations of heme compounds can be grouped into A 19' A 2g , BIg' and B 2g , square-planar D 4h heme-molecule symmetry. Strictly speaking, hemes are not D 4h symmetrical, but most analyses start with D4h for convenience. As can be seen from
320
Hemes and Porphyrins
A,g
"'6
806
'i6
'is
A2g
l
674
7S1
l-J8 -
J.j7
~ ~ ~
~3
8 29
"32
785
~3
~
~4
-
"35
-
FIGURE 12.2. Vibrational modes of Ni-octaethylporphyrin. The figure was reproduced from Abe et aI. (1978).
Figure 12.2, the A 1g and Big vibrations are symmetrical about the C2 axis of a pyrrole ring. In the A 1g type, the pyrrole rings vibrate in phase, but they are out of phase in the BIg type. A 2g and B 2g refer to the antisymmetrical vibrations in relation to the C2 axis of each pyrrole ring. In the A 2g type, the adjacent pyrrole rings vibrate in phase, they vibrate out of phase in the B 2g type (Abe et aI., 1978). There are many bands from 600 to 1700 em- I that arise from vibration modes of the heme chromophore. Most of these bands are due to vibrational modes of the conjugated C=C, C-C, and C=N bonds present in the porphyrin ring. There are some low-frequency bands (below 600 em-I), which usually involve the metal-ligand bonds. Low-frequency modes are well enhanced with Soret excitation (Rimai and Salmeen, 1978).
....
321
Vibrational Modes
A 1g
$+** '2
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1602
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~
1519
1383
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FIGURE 12.2. Continued.
1.3.
EFFECT OF METALS
Raman frequencies of a metalloporphyrin can vary, depending on the particular central metal atom. When Zn(II)-, Cu(II)-, Co(lI)-, and Ni(II)-octaethylporphyrins were examined, Raman frequencies of bands in the region of 1450-1700 cm -] were proportional to the frequencies of a-absorption bands of the different metal compounds (Kitagawa et al., 1975b). Gouterman (1959) demonstrated that the frequency of the a-absorption band in the spectrum of various metal tetraphenylporphyrins is proportional to the electronegativity of the metal ion. Raman spectra of Fe(III)- and Mn(III)-azide myoglobin are quite different from each other (Yu and Tsubaki, 1980). The intensity of Raman bands for various platinum-hematoporphyrin complexes like those for heme compounds, depends on the wavelength of excitation (Berjot and Theophanides, 1980). Cobalt myoglobin can bind to oxygen, and the 0-0 stretching frequencies can be observed at 1103, 1137, and 1153 cm- I . The Co - 0 vibration is also identified at 539 cm -I, which is considerably lower than the Fe-O stretching frequency of oxymyoglobin (Tsubaki and Yu, 1981). Resonance Raman spectra of metalloporphyrins originate from porphyrinring vibrations. and the Raman scattering is enhanced bv resonance between
322
Hemes and Porphyrins
the incident laser frequency and the porphyrin 7T-tO-7T* transItIOn. Since the metal and porphyrin orbitals interact, each metal exerts different degrees of perturbation on the porphyrin 7T-orbitals. Therefore, porphyrins show different optical- and vibrational-band frequencies depending on the type of metal present in the center. When there is a variation in the 7T-electron density of the porphyrin ring, some Raman bands shift. For instance, Cu(II)-uroporphyrin I forms complexes with various aromatic compounds, and this complex formation induces frequency shifts of Raman lines in the 1300~ 1700 cm - I region (Shelnutt, 1981).
2.
SIDE CHAINS
Different peripheral substituents of heme may affect the resonance Raman spectra. Sometimes the presence of a particular side chain shifts the absorption bands; then the excitation profiles will also be shifted. Another reason that side chains affect resonance Raman spectra is that the side-chain electrons interact with the porphyrin-ring 7T-electron system (Alben et aI., 1973). For instance, tetraphenylporphyrin shows strong phenyl-group Raman bands because the phenyl group and the porphyrin are conjugated (Fuchsman et aI., 1978). The phenyl rings also have a big effect on the eXyitation profiles (Shelnutt and O'Shea, 1978). We also must consider the case when peripheral substituents are the same; thus heme proteins contain the same heme but the manner of attachment to the protein moiety is different. In some heme compounds, the conjugation of heme to apoprotein is made through a thioether bridge. Cytochrome c, which has a thioether bridge, shows C-S stretch lines at 690 and 700 cm- I . No such lines can be seen for cytochrome bS62 ' which does not contain such a bridge (Srivastava et aI., 1981). Another example can be cited. Horseradish peroxidase contains heme IX, but it is attached to the apoprotein through a noncovalent type of interaction. The type of heme present in intestine peroxidase has not been identified, but heme IX is suspected. However, the Raman spectrum of intestine peroxidase is quite different from that of horseradish peroxidase, especially in the lowfrequency region, which is particularly sensitive to the attached peripheral groups. The most logical explanation is that heme IX probably attaches covalently to the apoprotein at position 4 of the heme periphery (Kimura et aI., 1981).
2.1.
VINYL GROUPS
Due to the presence of 7T-orbital electrons, vinyl groups exert considerable influence on the resonance Raman spectra. The effects of some side-chain
Side Chains
323
vinyl-group modifications are shown here: Cytochrome c -CH-CH 3 Mesoheme -CH 2 -CH 3 (em-I)
Deuteroheme -H (em-I)
Protoheme -CH=CH 2 (em-I)
1313
1324
1548
1545
1306 1342 1538 1564
I
S (em-I) 1313-1315 1548
In protoheme the side chain is -CH=CH 2 ; in mesoheme it is -CH 2 -CH 3 (Figure 12.3). The splitting of Raman bands in protoheme relative to deuteroheme occurs because the vinyl 'IT-electrons are coupled to the porphyrin 'IT-orbitals (Adar, 1975). In order to have such interactions, the vinyl side chain must be coplanar with the porphyrin ring. X-ray diffraction study indicates that the vinyl carbons do indeed lie in the porphyrin plane in Fe(III)-protoporphyrin myoglobin, whereas the ethyl side chains of Co(II)-mesoporphyrin myoglobin are out of plane (Padlan and Eaton, 1975). The replacement of a hydrogen atom with an alkyl chain decreases the frequency of the 1324-cm- 1 band of cytochrome c to 1313-1315 em-I. On the other hand, when the vinyl group is twisted out of the porphyrin plane, as in the case of Co- and Cu-protoporphyrin, then the vinyl group decreases its conjugation with the macro ring (Verma, 1976). Cytochromes C 558 and cm are believed to have one vinyl side chain and one c-type thioether linkage (Pettigrew et aI., 1975). Raman spectra of these cytochromes excited at 514.5 or 568.2 nm show the characteristic bands for the vinyl group as well as for the thioether-type side chain. C 557
C558
(em-I)
(em-I)
Remarks on Side Chains
1312 1339 1543
1319 1339 1543
1560
1558
Cytochrome c type Protoheme type Frequency between protoheme type and cytochrome c Frequency between protoheme type and cytochrome c
From these data, Adar (1977) concluded that cytochromes C557 and C 558 both contain one vinyl group and one thioether linkage (to the protein) on the heme. This example illustrates the usefulness of Raman spectroscopy for structural determination.
324
Hemes and Porphyrins
R2
H 3C
H 3C
CH2
CH2
I
I CH2 I COOH
CH2
I
COOH
Compound
Side Chain R] -H
Mesoheme
-CH -CH Z 3
-H
Protoheme IX
-CH=CH
Isospi rographi s
-CH=CH
Z Z
- CH -CH Z 3 -CH=CH
-CHO
-CH=CH
Z,4-0i fontlYl heme
-CHO
-CHO
R]:
Z
Cr-CHZ-CHz-CH=y-CHZ-CHz- CH=f-CH z -CH z -CH=f-CH OH
R : Z
Z
-CHO
Spirographis
Heme ~
CH 3
CH 3
CH=CH ; Pos i ti on 8: -CHO Z
FIGURE 12.3.
2.2.
RZ
Oeuteroheme
3
CH
3
Structures of some hemes.
FORMYL GROUP
The formyl group has a powerful electron-withdrawing property. This may regulate the heme a iron reactivity in cytochrome oxidase, which contains two nonequivalent hemes. When the Raman spectrum of cytochrome a 3 is examined, the carbonyl stretching vibration is observed at 1670-1675 cm -I. This suggests that there is strong coupling between the side-chain C=O 7T-electrons and the porphyrin 7T-system. This means that the formyl group in cytochrome a 3 is nearly coplanar to the prophyrin plane. The Raman spectra of reduced and oxidized cytochrome a do not show well-defined bands at 1670-1675 cm ~ I. Thus in this case the formyl group may be much less coplanar to the
Cu- Etioporphyrins Solid ,n KBr
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I c-c
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'H M
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/ C-, c-!., /c-c
'\
H,
"'c_ I C
I
E/C-C
" j! c._ I I 'C-C'E HI
,
H-
l
-c/
J_c.. . . N'c_J
,,/
~_/
,I
III
'\
".H
'"
'H
/
c-c I \ /H c-c c-c
H,
,
I
'H/
I
'c-\ I le- cI'" I .M . . c_ I I 'c_ el H - C.-H
C
!_C",N,C_C 11/
I c-c I \, M \
IV
'H
,
,
M
FIGURE 12.4. Identification of Cu-etioporphyrins by resonance Raman spectroscopy. M, methyl; E, ethyl, and H, hydrogen atom. The figure was reproduced from Sunder et aI. (1975).
326
Hemes and Porphyrins
porphyrin plane (Babcock and Salmeen, 1979; Babcock et al., 1979a, b; Ondrias and Babcock, 1980; Babcock et al., 1981). The vibrational frequencies of the formyl group can be influenced by factors such as the spin state of the heme iron and the type of solvents used (Van Steelandt-Frentrup et al., 1981). The formyl C=O band appears at 1660 cm- I for reconstituted myoglobin whose heme is replaced by spirographis, isospirographis, or 2,4-diformylheme (Tsubaki et al., 1980). This frequency is slightly lower than that of cytochrome a 3 • Normal myoglobin, which contains protoheme IX without the formyl group, does not give the 1660-cm- I keto band. Cyanide ion often serves as an axial ligand, and it may also react with side-chain formyl groups. Likewise, NaHS0 3 may react with the formyl group; this too can be detected by Raman spectroscopy (Kitagawa et al., 1977b). However, this should be considered as a tentative interpretation of the experiments. There is other evidence showing that CN does not react with side-chain formyl (personal communication with Salmeen, 1981). 2.3.
IDENTIFICATION OF ISOMERS
Different side chains and their substitution at different positions may produce unique fingerprint regions in Raman spectra. Using these characteristic fingerprint regions, Raman spectroscopy can be used as an an:dytical tool. Many porphyrin isomers differ little and are not easily distinguished by conventional physicochemical methods. For instance, etioporphyrin has methyl and ethyl side chains. Isomers of etioporphyrins I, II, III, and IV differ only in the relative positions of these substituents. The resonance Raman spectra of the Cu complexes of these four isomers differ markedly in the 650-850 cm- 1 region (Figure 12.4) (Verma and Bernstein, 1974a; Sunder et al., 1975). Isomers of many other porphyrins have also been distinguished from each other by Raman spectroscopy (Verma and Bernstein, I974a).
3.
SPIN STATE
The Raman spectra of some heme proteins show characteristic spin-state marker bands, from which it is sometimes possible to determine the spin state of the iron. However, the effect of the spin state on resonance Raman spectra is an indirect one, which occurs via the perturbation of the porphyrin-ring 'IT-electron system. Thus sometimes Raman spin marker bands may not apply to all heme compounds. In this chapter the effects of spin state, ligands, and oxidation state will be discussed in separate sections. However, one should keep in mind that they are all interrelated. For instance, the spin state of iron can be altered by changes in ligand species.
,,.
Spin State
327
Before discussing the relationship between the spin state of heme iron and resonance Raman spectra, some basic information about spin state is briefly reviewed. 3.1.
SPIN
The electron has an inherent angular momentum called spin. Depending on the direction of spin of the electron, the Z-spatial component of the spin quantum number is assigned either + 1 or - 1. There are four quantum numbers to describe the state of an electron in an atom. Three designate a particular orbital, and the fourth designates one of two values of a spatial component of the spin angular momentum. Each electron must have a unique set of quantum numbers; therefore, only two electrons are allowed to occupy each set of orbitals, one with + 1 and one with spin - !. When all orbitals are occupied by paired electrons, the total spin state is zero. This is because the spin angular momenta of the two electrons with opposing spin quantum numbers of + ! and - ! cancel each other. If there is only one electron in an orbital, then the spin number is 1, and the atom shows paramagnetism. 3.2.
ELECTRONS IN THE COORDINATED IRON
Iron has an atomic number of 26 and contains 26 electrons, which are distributed in different electronic orbitals. They are the Is 2, 2s 2, 2 p6, 3s 2, 3p6, 3d 6 , and 4s 2 orbitals. When the iron atom loses two or three electrons it becomes Fe(II) or Fe(III) ion, respectively. 3d
Fe(II) Fe(III)
@CDCDCDCD CD CD CD CD CD
4s
o
o
4p
000 000
When iron forms a coordination complex, the electrons in the metal are relocated. The s orbital is spherical, and the p and d orbitals are shown in Figure 12.5. When a complex is formed, metal and ligand orbitals interact to form moleculer orbitals. For simplicity, only the interaction of the ligand (J orbitals with metal orbitals is discussed (Figure 12.6). Examples of such complexes include FeFl- and Co(NH3)~+ . The iron atom provides five 3d, one 4s, and three 4p orbitals. The f 2g (d XY ' d yz ' d zx ) orbitals will not interact with ligand a orbitals, so they are occupied by the electrons from the metal. The six ligands provide six orbitals that contain two electrons in each orbital. The interaction leads to the formation of bonding (e g , flU' a lg ) and antibonding (flU.' a]u.' ego) molecular orbitals. The electrons from ligands will not occupy antibonding molecular orbitals, but they will fill bonding orbitals. The
328
Hemes and Porphyrins z
5 -
orbital y
z
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~
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*
y
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x
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y
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Py
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z
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d-orbitals y dxz
y
y
x dx 2_ y2
FIGURE 12.5.
dz 2
Shapes of s, p, and d electron orbitals.
egO and l2g molecular orbitals have more metal character than other orbitals, and the l2g orbitals are occupied by electrons belonging to the iron atom. The six bonding molecular orbitals (e g , a'g' llu) actually retain considerable ligand character. The electrons (12 altogether) from ligands occupy the newly formed molecular orbitals e g , flu' and a'g' The electrons (6 for ferrous ion, 5 for ferric ion) from the metal fill the l2g and egO orbitals. If the energy difference between these two orbitals is large (the strong-field case), then all the electrons will fill the f 2g orbital first; only then will the egO orbital be filled. This will produce a low-spin complex, that is, 8 = t for Fe(III) or 8 = 0 for Fe(II). When the energy difference is small (the weak-field case), then the electrons enter the l2g and egO orbitals with their spins parallel (same sign). Thus a high-spin complex is formed (8 = 1 or 8 = 2 for Fe(III) and Fe(II), respectively).
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330
Hemes and Porphyrins
The interaction of metal with (J- and 7T-bonding ligands is much more complicated. Examples of such complexes include Fe(CN)~- and Cr(CO)6' The newly formed molecular orbitals are shown in Figure 12.6B. 3.3.
SPIN STATES OF HEME COMPOUNDS
The spin state of heme originates from the electron distribution of the iron electrons in the molecular orbitals, and it has important biological significance. The low-spin Fe(II) of hemoglobin accounts for the diamagnetism of oxyhemoglobin. The redox potential of the cytochromes depends on the spin state of the iron atoms involved. An Fe(III) compound may have one, three, or five unpaired electrons, depending on its spin state. A high-spin state produces a large magnetic moment. The diamagnetic state occurs when all the electrons in the d orbitals are paired. Highly simplified diagrams of low-spin and high-spin heme iron are shown in Figure 12.7. Empirically, Raman spectra have been found to show spin-characteristic bands for a given heme compound. Therefore, the spin states of the iron in heme compounds are determined by the axial ligands. Any ligand that affects the electron system of porphyrin and iron, therefore, may cause the frequency of some vibrations to change. This can be illustrated by the example of Fe(III)-octaethylporphyrin. In coordination with strong ligands, it becomes a low-spin complex, but with weaker ligands such as MezSO or acetone, it forms a hexacoordinated high-spin complex (Ogoshi et al., 1980). Similarly, the spin state of myoglobin heme iron can be influenced by the sixth ligand. MbOH, MbCN, MbN3 , and MbIm give low-spin ferric ion, whereas MbF, MbOCN, and MbHzO give high-spin ferric ion. Likewise Fe(III)-cytochrome P-450 has high-spin iron in the presence of diphenylmethane, but it has low-spin iron in the presence of phenanthrene (Anzenbacher et al., 1980).
Ferric
00
eg
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S= 1/2 d z2
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S= 5/2
d xt. y2
00
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d yz dxz
Low Spi n S=O
High Spin S=2
FIGURE 12.7. Distribution of electrons in d orbitals and spin state of oxidized and reduced forms of heme iron.
....
Spin State
331
Cytochrome c peroxidase has a molecular weight of 43,200 and has two heme c. It was proposed from Raman analysis that there is one low-spin and one high-spin iron for the native oxidized enzyme (Ronnberg et aI., 1980).
3.4.
SPIN STATE AND RAMAN BANDS
There are several spin-sensitive Raman bands that are characteristic of particular classes of heme compounds; these are considered the spin marker bands. The empirical correlation is determined from the spin state of the heme protein iron as determined by magnetic susceptibility or EPR spectroscopy and band frequency. The correlation is then used to infer the spin state of heme proteins not studied by EPR or magnetic susceptibility, or for heme proteins for which EPR and susceptibility data are controversial. Using the spin marker band, Champion et aI. (1978) concluded that cytochrome P-450CAM has low-spin ferric heme in the free state and a mixture of high- and low-spin ferric hemes in the substrate complex. The 1584-cm- 1 band for hemoglobins and myoglobin, the 1586-cm -I band for Fe(III)-cytochrome c, and the I590-cm -I band for Fe(II)-cytochrome c are low-spin marker bands. This low-spin marker band region of 1584-1590 cm- I shifts to 1566 cm - I as the iron changes to the high-spin state. The 1584-1590- and 1566-cm - I bands are also highly sensitive to the wavelength of laser excitation (Kitagawa et aI., 1975a). It is known that cytochrome c heme undergoes a structural transition at pH 2.5 from the native low-spin heme to a high-spin form with concomitant displacement of the axial ligands by two water molecules (Lanir and Aviram, 1975). Raman spectra of cytochrome c change markedly as the pH is lowered; this reflects structural and environmental changes around the heme. As the spin state of reduced horseradish peroxidase (aquo complex) iron changes from the low- to the high-spin state, there are band shifts from 1622 to 1610 cm- I , from 1494 to 1476 cm- I , and from 1367 to 1359 cm- I (Loehr and Loehr, 1973). These variations may also be due to apparent shifts or different vibrational modes, both of which are Raman active in the two spin states. The high intensity of the 1637-cm- 1 band of ferric ion in the low-spin state in heme proteins is associated with the methine-bridge stretching vibration (Kitagawa et al., 1975a, c; Stein et aI., 1975; Ozaki et aI., 1976a). Of course, this conclusion depends on the validity of the normal-coordinate analysis used to interpret the spectra. Heme Q, which is often used as a model compound in the study of cytochrome oxidase, has vibrational bands in the 1540-1660-cm -I region that are sensitive to iron spin state. From this and from excitation-profile data, Babcock and Salmeen (1979) concluded that the Raman spectrum of oxidized cytochrome oxidase obtained with 441.6-nm excitation is due primarily to vibrations of low-spin Fe(III)-cytochrome Q. The same conclusion was obtained by exciting at 590 nm (Babcock et aI., 1982). Reduced cytochrome
332
Hemes and Porphyrins
P-450 excited at 488 nm exhibits spectra of the ferrous high-spin type, whereas the oxidized compound shows the ferric low-spin type of spectra (Ozaki et al., 1976b; 1978). On some occasions the spin states of hemes may be temperature dependent. Chloroperoxidase is a heme glycoprotein, isolated from Caldarimyces fumago, that contains 1 mol of Fe(III)-protoporphyrin IX. It catalyzes the peroxidative formation of a carbon-halogen bond in the presence of suitable halogen donors, Cl- , Br- , or 1- , at low pH. At room temperature, the enzyme heme is mainly in the high-spin ferric form, but the enzyme contains a small fraction of low-spin heme that becomes the dominant species at low temperature (Remba et al., 1979). Although there is extensive empirical correlation between the frequencies of some Raman bands and the spin state of the iron atom, this is not always the case. So-called spin marker bands do not fit very well for high-spin ferric chloroperoxidase (Champion et al., 1976b). They pointed out that the spinmarker-band scheme works quite well for high- and low-spin ferrous as well as for low-spin ferric heme proteins, but it does not work well for high-spin ferric heme proteins. There are also cases in which the Raman spectra of ferrous low-spin derivatives (for instance, oxyhemoglobin) are more like those of the ferric type (Yamamoto et al., 1973; Kitagawa et al., 1976b). One also should be cautious, as some Raman bands are sensitive not only to spin state but also to oxidation state and whether heme is pentacoordinated or h~xacoordinated (Sievers et al., 1979). An example is Fe(III)-cytochrome c'; at low pH it is expected to have high-spin iron, yet the Raman spectrum is more like that of the low-spin type (Strekas and Spiro, 1974). Quaternary structure may affect spin-state equilibrium. The addition of inositol hexaphosphate (IHP) to methemoglobin lowers the intensity of the low-spin marker bands still further relative to the high-spin bands (Ferrone and Topp, 1975). A similar conclusion was reached by Scholler and Hoffman (1979a, b), who showed that the high-spinjlow-spin equilibrium of azidomet-Hb is changed when the quaternary structure is changed by pH or IHP. Many of the spin states seem to be responding to the core expansion that accompanies spin-state changes. In Section 9 of this chapter, the core-expansion theory originally proposed by Spaulding et al., (1975) is discussed in more detail.
4.
EFFECT OF LIGANDS
Sometimes axial ligands influence frequencies and intensities of some of the Raman lines, and several "ligand-sensitive Raman lines" have been identified. The extent of the ligand's effect is determined by how much the ligand perturbs the iron-porphyrin electronic interaction.
.
.
j~-
Effect
4.1.
0' Ligands
333
LIGAND-SENSITIVE RAMAN BANDS
Cytochrome c is one of the most extensively studied heme compounds. It is, therefore, appropriate to discuss the results from cytochrome c first. Under natural conditions, the fifth and sixth coordination positions are occupied by histidine N and methionine S atoms, respectively. A change of the sixth ligand in cytochrome c has an effect on the Raman intensity of some lines, as shown below: Compound Fe(III)-cytochrome c Fe(II)-cytochrome c DicarboxymethylmethionylFe(II)-cytochrome c Fe(II)-cytochrome c3
Line (cm- I )
Reference
1565 1540 1545 (pH 3.9) 1533 (pH 9.7) 1541 (pH 9) 1536 (higher pH)
Kitagawa et al., 1977d Kitagawa et aI., 1975a, b Ikeda-Saito et al., 1975 Ikeda-Saito et al., 1975 Kitagawa et al., 1977d Kitagawa et al., 1977d
Different ligands give slightly different ligand-sensitive Raman lines. When the axial ligand is lysine or a lysine-like nitrogenous base, it gives a lowerfrequency 1540-cm- 1 band than histidine or its analog (Kihara et al., 1978a). Researchers using the ligand-sensitive Raman band concluded that lysine is the sixth ligand at pH 9.3, a conclusion that agrees well with the results of photometric titration (Kihara et al., 1978b; Smith and Millett, 1980). By a similar technique, lysine was also identified as the sixth ligand in cytochrome c' (Kitagawa et al., 1977c). Cytochrome c' is found in purple photosynthetic and denitrifying bacteria and is similar to cytochrome c. Cytochrome cm is found in thermophilic bacteria and chromatium-a purple sulfur bacterium. It has a molecular weight of 15,000, with a high isoelectric point: 10.8. By analyzing the ligand-sensitive Raman lines of this compound at different pH, it was concluded that ferric cytochrome cm at neutral pH has the axial ligands histidine and methionine. The fifth ligand of the corresponding ferrous compound is histidine, and the sixth ligand is either histidine or lysine (Kihara et al., 1978a; Hon-Nami et al., 1980). Cytochrome b562 from Escherichia coli consists of 110 amino acids and contains a single noncovalently bonded protoheme. Judging from Raman spectra of the three pH forms, the axial ligands of the acidic and neutral forms are methionine and histidine. At higher pH, the axial ligands are either lysine-histidine or histidine-histidine (Myer and Bullock, 1978). For reduced cytochrome t, obtained from the blue-green alga Spirulina platensis, the ligand-sensitive band at 1545 cm- I is very similar to that of cytochrome c. From this fact, it was concluded that the fifth and sixth ligands are histidine and methionine, just as in cytochrome c (Kitagawa et al., 1975a).
334
Hemes and Porphyrins
As mentioned in the previous section, many axial ligands affect the spin state of iron, which in turn affects the Raman spectra. The best examples are hemoglobin and myoglobin. The Raman spectrum of chloroperoxidase is also influenced by its complexing with cyanide, fluoride, chloride, and hydroxide, which in turn influences the spin state of the iron atom. The cyanide ligand achieves a complete conversion to the low-spin state, as evidenced by the strong peaks at 1503 and 1633 cm- I (Remba et al., 1979). 4.2.
CARBON MONOXIDE LIGAND
Many hemoproteins bind strongly to carbon monoxide. The best example is the so-called poisoning of hemoglobin by carbon monoxide. The oxygen molecule attaches to heme as a bent ligand instead of in a linear fashion. In the microenvironment of the heme, the polypeptide chain folds in such a way that the sixth coordination position is surrounded by nonpolar side chains, which do not permit a linear ligand to sit directly on top of the central iron atom. This may be an evolutionary mechanism to discriminate against the attachment of the small amount of carbon monoxide linear ligand, but it allows the oxygen molecule to attach to the iron as a bent ligand. Discrimination against carbon monoxide that attaches to the iron in linear fashion is essential in order to prevent poisoning from endogenous CO, which is produced by the breakdown of bile pigments. Moreover, such a protein microenviromaent around the sixth coordination bond is beneficial to the Fe(II) atom, which is protected from oxidation. Carbon monoxide is frequently used as a special ligand to study the mechanism of oxygen binding or electron transport of various heme compounds. In this section we review the effect of carbon monoxide on the Raman spectra of heme compounds. From IR study it is firmly established that the C-O stretching vibration in carboxyhemoglobin is in the region of 1950-2000 cm- I . No such band was observed in Raman spectra (Strekas and Spiro, 1972b); this is because of the dissociation of CO from carboxyhemoglobin caused by the laser beam. Using nanosecond transient Raman spectroscopy, researchers showed that carboxyhemoglobin is indeed photolysed through different steps (Woodruff and Farquharson, 1978; Lyons et al., 1978; Terner et al., 1980; Terner et aI., 1981). Before HbCO undergoes a change in its quaternary structure, there must be a change in the Fe-imidazole bond since the Fe-N band frequency changes within 0.3 p.s of the photolysis (Stein et al., 1982). The photolysis occurs within picoseconds, but the recombination of CO and deoxyhemoglobin takes place in 65 ns. Mb-CO also undergoes photolysis, but fast recombination does not take place. It is likely that variation in protein structure rather than differences in the electronic state determine the photolytic mechanism, because protein rearrangement is believed to require much longer times (Friedman, 1980; Friedman and Lyons, 1980). Within 6 ns of HbCO dissociation, the iron atom moves out of the heme plane. The 1584- and 1631-cm -1 bands are characteristic for in-plane iron. Because no such bands were observed in the CARS
Effect of Ligands
335
spectra (see Chapter 2, Section 3.2), it was concluded that the iron atom is not in the heme plane (Dallinger et al., 1978). The Raman spectrum of the photolysis product of Hb-CO is similar to that of deoxyhemoglobin (Terner et aI., 1980), but small frequency shifts are observed (Friedman and Lyons, 1980). The shifts are very similar to the static Rand T (see Section 7) frequency differences observed (Shelnutt et al., 1979b). However, with the excitation wavelength at 406.7 nm, carbon monoxide vibrations (Fe-C-O) were finally observed at 507, 578 and 1951 cm- 1 for HbCO and 512, 577 and 1944 cm- I for MbCO. The first two bands for HbCO and MbCO are sensitive to CO isotope substitution (Tsubaki et al., 1982). 4.3.
OXYGENATION
Oxygenation can be considered as a reaction between the ligand O2 and the heme iron atom in ferrous state. This ligand attachment is an important phenomenon, especially in the oxygen-transport molecules hemoglobin and myoglobin. Oxygenation of hemoglobin induces many changes; a few examples are shown here: Hb
Hb02 --
Sixth ligand Spin state Distance from Fe(II) to the center of porphyrin ring Distance from Fe(II) to pyrrol N
None High
O2
0.60 A 2.09 A
0.05 2.01
Low
A A
When the oxygen molecule is removed from oxyhemoglobin, the intense band at 1376 cm- 1 is shifted to 1355 cm- I • These bands can be used to detect the oxygenation state of hemoglobins (Brunner et al., 1972, 1974; Yamamoto et al., 1973: Albrecht and Breitinger, 1976). Considering the geometry of Oz-Fe in oxyhemoglobin, two different linkages are possible (Figure 12.8). When hemoglobin is oxygenated with asymmetrical isotopic oxygen 16 0_ 18 0, one would expect to see Fe_ 16 0 and Fe- 18 0 vibrations for the Pauling and Coryell model (Figure 12.8). On the other hand, one would expect a single frequency corresponding to a symmetrical Fe-O stretch somewhere between those for Fe- 16 0 and Fe- 18 0 for the Griffith model (Figure 12.8). Experiments indicated that there were two bands at 567 and 540 cm- I (Figure 12.9). Thus oxygen attaches to hemoglobin in the end-on geometry of the Pauling and Coryell model (Duff et al., 1979). The experiment by Duff et al. agrees well with IR studies, which show that the 0-0 stretching frequency corresponds to a bending type, as shown in Figure l2.8A (Barlow et al., 1973;
336
Hemes and Porphyrins
/0
o
0-0
-Fe-
-Fe-
I
A
\ I
FIGURE 12.8. Two possible attachments of molecular oxygen to hemoglobin iron. (A) Pauling and Coryell model; (B) Griffith model.
B
Maxwell et al., 1974). Although the evidence may be indirect, Griffith structures have not been found in nonprotein iron-containing model compounds of dioxygen carriers (Jameson et a1., 1978). Soybean leghemoglobin also combines with oxygen; however, the spectra indicate no significant difference between the strength of the Fe-O in oxy-Ieghemoglobin and oxy-myoglobin (Irwin et a1., 1981). ul eT>
VI..,
~l m
-
I
i
(D
a tSl za
v
w
VI ocCl)
weT> CLrVI t-
Z
5l'i Vm VI
ul (D N
FIGURE 12.9. Resonance Raman spectrum in the Fe-O stretching vibration region of oxyhemoglobin. The geometry of Oz-Fe in hemoglobin was proved using the mixed isotope 16 0 18 0. Vertical lines a and b show the VFe-O peak positions for 160Z- and 180z-oxyhemoglobin. Line c shows the calculated value (555 em-I) for structure B in Figure 12.8. This figure was reproduced from Duff et al. (1979) by permission of the copyright owner, Academic Press.
~.
Effect of Ligands
337
°
Some model compounds form a complex with 02' For instance, mesotetra(0:, 0:, 0:, o:-O-pivaloylamidophenyl) porphyrin combines with 2, The Fe-O stretching band appears at 568 cm - 1, which is very close to the value of the oxyhemoglobin vibration (Burke et al., ! 978). Similarly, the Fe-O stretching frequency appears at 572 cm - 1 for (dioxygen)(porphyrinato)(hindered imidazole) iron(II) complexes (Walters et al., 1980).
The time required for oxygenation or deoxygenation of hemoglobin is very short, but using a picosecond time-resolved resonance Raman method, one can obtain the transient spectra. From this study it was found that the time required for the reorganization of the porphyrin structure is less than 30 ps. As the oxygen ligand departs from the porphyrin core, there is a change in the electronic distribution, which is manifested in the transient picosecond resonance Raman spectrum of Hb02 (Coppey et al., 1980). Photolysis of oxyhemoglobin indicates that iron is converted to a high-spin state within a few picoseconds and remains closer to the heme plane on the picosecond and nanosecond time scale (Nagumo et al., 1981).
4.4.
Fe-AXIAL-liGAND VIBRATIONS
Introduction or replacement of the axial ligands has considerable effect on the biological and chemical properties of a compound. For instance, deoxyhemoglobin has an empty sixth coordination position and is high spin; when an oxygen molecule occupies the sixth position, Hb becomes Hb02, which is low spin. The redox potential of the heme iron in cytochrome c and its related compounds is also influenced by the axial ligands. From these examples it is obvious that the effect of axial ligands is important. It is worthwhile to review axial-ligand vibrations in Raman spectroscopy. The Fe-axial-ligand vibration can be observed at relatively low-frequency regions (200-600 em-I). Assignments of such bands are usually made with the use of ligands containing different isotopes. The axial-ligand stretching vibrations are summarized in Table 12.1. There are many low-frequency porphyrinring vibrations that are not Fe-axial-ligand vibrations; these are summarized in Table 12.2. Differentiation of Fe-N(pyrrole) and Fe-N(axial ligand) vibrations is not an easy matter. The assignments made by the original investigators are summarized in this table; however, readers should not consider these data final assignments. Desbois et al. (1981) made an interesting observation. A 14N -> 15N substitution of the four pyrrole nitrogens in Fe(II)etioporphyrin was carried out, and it was found that the bands at 204, 257, 334, 343, 390, and 408 cm- I are sensitive to N(pyrrole) isotopic substitution. This experiment suggests that these bands originate from the stretching vibrations of the Fe-N(pyrrole) bonds. However, one still must be cautious, as Fe -N(axial) and Fe-N(pyrrole) vibrations may have a coupling effect.
TABLE 12.1.
Frequency (em-I) 205
220
220 244 274 279 290 363 364 373
374
408 409 409 411 411 411 412 412 413 413 413 413 413 413 414 416 443
':l':lA
Axial Ligand Vibrations Involving the Central Iron Atom
Assignment Fe-N
Compound
Reference
2-Methylimidazole Stein et al. (1980) adduct of octaethylporphine Fe-N 2- Methylimidazole Stein et aI. (1980) adduct of Fe(III) protoporphyrin IX Kitagawa et al. (1979) Fe-N(His F8) Deoxy-Mb Fe-N(Proximal His) Ferro-horseradish Teraoka and Kitagawa Peroxidase (1981 ) Fe-N (Proximal His) Ferric-horseradish Teraoka and Kitagawa Peroxidase (1981) Fe-Br Fe(III)-OctaethylKitagawa et al (I 976a) porphyrin Fe-N(imidazole) Fe(III)-OctaethylKitagawa et al. (I 976a) porphyrin Fe-O-Fe Fe(III)-Octaethy1Kincaid and Nakamoto porphyrin (1976) Fe-CI Fe(III)-OctaethylKitagawa et al. (1976a) porphyrin Heme in-plane Mb0 2 Desbois et aI. (1979) Deformation Fe-N,(Histidine) Heme in-plane MbNO Desbois et al. (1979) Deformation Fe-N.(histidine) Fe-N.(histidine) Mb Desbois et aI. (1979) Fe-N.(histidine) Mb0 2 Desbois et al. (1979) Fe- N.(histidine) MbNO Desbois et aI. (1979) Mb+N 3Fe-N(histidine) Desbois et al. (1978) Mb+H 2 O Fe-N(histidine) Besbois et aI (1978) Fe-N FeN3(metmyoglobin Tsubaki et aI. (1981) azide) Mb+HCOOFe-N.(histidine) Desbois et al. (1979) Fe-N Deoxy-Hb Brunner and Sussner (1973) Mb+OCNFe-N.(histidine) Desbois et aI. (1979) Mb+lmFe-N(histidine)] Desbois et aI. (1979) Mb+CNFe-N.(histidine) Desbois et al. (1979) Mb+OHFe-N(histidine) Desbois et aI. (1979) Mb+CNFe-N(histidine) Desbois et aI. (1979) Fe-N(azide) Met-Hb Derivative Asher et aI. (1977) Mb+SCNFe-N,(histidine) Desbois et aI. (1979) Mb+FFe-N.(histidine) Desbois et aI. (1979) Fe-F Met-Hb(fluoride) Asher et aI. (1977)
.;;
Effect of Ligands
TABLE 12.1
Frequency (cm- I )
339
Continued
Assignment
461
Fe-F
466
Fe-F
468 471 471
Fe-F Fe-F Fe-F
490 497 549 551 567 572 581
Fe-O Fe-O Fe-N(nitroso) Fe-N (nitric oxide) Fe-O Fe-O Fe-F
606
Fe-F
Compound Met-Mb Mb(fluoride) a-Subunit of metMb Hb(fluoride) Met-Hb(fluoride) fi-Subunit of metMb Met-Mb Met-Hb Derivative NO-Hb Hb-NO Hb-0 2 Hb-0 2 Fe(III)-Octaethyl porphyrin Fe(III)-Octaethyl porphyrin
Reference Asher and Schuster (1981) Asher et al. (1981) Asher and Schuster (1981) Asher et al. (1981) Asher et al. (1977) Asher and Schuster (1981) Asher and Schuster (1979) Asher et al. (1977) Chottard and Mansuy (1977) Tsubaki and Yu (1982) Brunner (1974) Brunner and Sussner (1973) Kincaid and Nakamoto (1976) Kitagawa et al. (I 976a)
Source: The table was compiled from data published by different investigators. Assignments of the vibration are listed as they were presented in the original papers. For the basis of these assignments, readers should look at the original references.
The 0-0 stretching vibration of oxyhemoglobin can be observed by IR spectroscopy but not by Raman spectroscopy. However, the Fe-O stretching vibration is detected by Raman spectroscopy. Deoxyhemoglobin does not show any line between 500 and 650 cm- I (Figure 12.10, top). Oxyhemoglobin shows a distinct band at 567 cm - I (Figure 12.10, middle). When hemoglobin is saturated with the 18 0 2 , the 567-cm- 1 line is shifted to the lower frequency 540 cm- I (Figure 12.10, bottom). This is a clear indication that the 570-cm- J band originally observed in oxyhemoglobin is due to the vibration of the oxygen molecule and the heme iron, that is P Fe - O (Brunner, 1974). On some occasions, isotopic iron 54Pe is used rather than 18 0 in order to determine the Fe-ligand vibration. For instance, Desbois et a1. (1978) used 54Fe to assign a number of Pe-N bond vibrations (Table 12.3). On some occasions, the effect of ionization of amino acid residues can be transmitted to the proximal ligand through the Fe-NHis bond. In reduced horseradish peroxidase, only the 244-cm - I line is pH dependent (Teraoka and Kitagawa, 1980a).
TABLE 12.2. Low-Frequency Vibrations of Heme Compounds Other Than Axial Ligand- Metal Vibrations.
Frequency (em-I)
Assignment
Compound
Reference
214 214 214 214 216 217 217 218 218 218 218 222 249 249 249 251 252 252 254 254 254 255 255 255 339
Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe - N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe- N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe- N(pyrro1e) Fe- N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N(pyrro1e) Fe-N
Mb+H 2 O Mb+ HCOOMb+ SCNMb+OCNMb+lmMb+N 3Mb+CNMb0 2 MbNO Mb0 2 MbNO Mb Mb+ SCNMb+H 2O Mb+FMb+ H 2 O Mb+FMb+HCOOMb+N; MB+CNMb+N; Mb+OCNMb+OHMb+lmDeoxy-Hb
351
Fe-N
Hb-02
351
Fe-N
cyt.
352
Fe-N
364
Fe-N
Fe(III)-cyt. c Fe(II)-cyt. c Deoxy-Hb
380
Fe-N
Hb-0 2
392 404 424
Fe-N Fe-N Fe-N
Fe(II)-cyt. c Fe(III)-cyt. c Hb-0 2
441
Fe-N
Fe(I1)-cyt. c
Desbois et al. (1979) Desbois et al. (1979) Desbois et aL (1979) Desbois et al. (1979) Desbois et aL (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et aL (1979) Desbois et aL (1979) Desbois et al. (1979) Desbois et aL (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et aL (1979) Desbo;s et al. (1979) Desbois et aL (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et al. (1979) Desbois et aL (1979) Desbois et al. (1979) Brunner and Sussner (1973) Brunner and Sussner (1973) Champion and Gunsalus (1977) Brunner (1973) Brunner (1973) Brunner and Sussner (1973) Brunner and Sussner (1973) Brunner (1973) Brunner (1973) Brunner and Sussner (1973) Brunner (1973)
~4n
P450 cam
Effect of Ligands
341
>
l-
e;; Z W
I-
~ Z 04:
~
04:
a:
FIGURE 12.10. Proof of Fe-O stretching band using 18 0 2 isotope. The figure was drawn from Brunner (1974).
800
The hemoglobin and myoglobin Fe-axial-ligand vibrations may be selectively enhanced by exciting at 600-640 nm. This may be due to charge-transfer transitions from a ligand to the ferric ion (Asher et a1., 1977; Asher and Schuster, 1979). The vibrations of other metal-ligand bonds of metalloporphyrins also appear at low frequency. For instance, the Mg-N bond vibration of Mgetioporphyrin I appears at 254 cm - I (Thoai et a1., 1979). TABLE 12.3. Low-Frequency Porphyrin-Ring Vibrations (In-Plane Deformation) Other than Fe-Ligand Bond
Frequency
Source:
Frequency
(em-I)
Compound
(em-I)
Compound
341 341 342 344 344 345 346 347 347 347 347 347
Mb+ SCNMb+OCNMb+N1 Mb0 2 MbNO Mb Mb+H 2 0 Mb+HCOOMb+FMb+OHMb+CNMb+Im-
349 (sh) 354 (sh) 373
Mb+SCNMb+OCNMb Mb+ SCNMb+CNMb+FMb+OCNMb+H 2 0 Mb+HCOOMb+N1 Mb+OHMb+Im-
377
378 378 378 378 378 379 379 384
This table was reconstructed from the table of Desbois et aI. (1979). Sh., Shonlder.
342 4.5.
Hemes and Porphyrins
PENTACOORDINATION AND HEXACOORDINATION
Porphyrin is essentially a square-planar compound, but with the iron atom it becomes an octahedral or square pyramid. The central iron atom always chelates some kind of ligand to form a square pyramidal or octahedral complex (Figure 12.11). Whether a heme compound takes on a five- or six-coordinate complex depends on the oxidation state, the spin state, and the type of ligands. It is possible to postulate the type of coordination in heme compounds empirically from Raman spectroscopy (Scholler et a1., 1979). This is possible because many Raman bands sensitive to the oxidation state, the spin state, and the type of ligands are empirically correlated to the coordination number. Fe(III)-meso-tetra-(a, a, a, a-pivaloylamidophenyl) porphyrin shows 390-, 1360-, and 1560-cm- I Raman bands that are associated with characteristic oxidation and spin states (see the sections on oxidation and spin, section 5 and section 3 of this chapter). For instance, the 1560-cm- 1 band is characteristic of the high-spin Fe(II) compound. Low-spin Fe(II) in cytochrome c and hemoglobin appears at the higher frequency of 1584 cm- I (see section 3). Similarly, the 1360- and 390-cm - I bands are associated with high-spin ferrous compounds. From these band characteristics, it was concluded that the compound is a high-spin five-coordinate complex (Burke et a1., 1978). On many occasions the Raman lines of less-well-known compounds are compared with those of well-studied compounds or model compounds for judgment 0f the coordination number. For instance, ferric cytochrome P-450 was judged to have pentacoordination in a high-spin ferric state by comparison with the Raman spectrum of a model compound (Anzenbacher et al., 1981). The ferric form of high-spin cytochrome a 3 is believed to be a six-coordinated complex. In general, a change from a five-coordinate, high-spin to a six-coordinate, low-spin complex results in a: frequency change of approximately 10 cm- I in the band around 1580 cm- I . A change from a six-coordinate, high-spin to a six-coordinate low-spin complex results in a change in this
CI N-N
N~NI
N
/-,\~N ~\bN _
F~'N N_
N--;'N N
B
C FIGURE 12.11. Porphyrin with or without iron. The 'shape of the prophyrin ring and two types of heme compounds are shown. (A) Square-planar (porphyrin); (B) square-pyramid (heme chloride); and (C) octahedron (pyridine hemechrome). A
Oxidation States
343
band of approximately 20 cm- I. Thus the l8-cm -I shift from 1572 to 1590 cm - I that occurs upon conversion of high-spin Fe(III)-cytochrome a j to low-spin Fe(III)-cytochrome aj-CN suggests that in the resting enzyme, highspin Fe(II)-cytochrome a j is six-coordinate (Babcock et al., 1980). Upon conversion of nitrosylhemoglobin, NOHb, from the R to the T structure, the l636-cm- 1 line splits into 1645- and l637-cm- 1 lines. This indicates that hexacoordinated NOHb becomes a mixture of pentacoordinate and hexacoordinate forms. This implies that the Fe-N(His) bond is disrupted in the T structure of NOHb (Nagai et al., 1980b). The Rand T structures of hemoglobin are discussed in detail in Section 7 of this chapter. Methemoglobin and metmyoglobin have histidine at the fifth coordination position and a water molecule at the sixth position (Stryer et al., 1964; Deatherage et al., 1976). In the cases of cytochrome e peroxidase, horseradish peroxidase, horseradish peroxidase isozyme C 2 , and cytochrome e', the axial ligands are not definitely known, but it is believed that histidine is the fifth ligand. The close similarity in the Raman spectra of these peroxidases to those of the pentacoordinated-heme model compounds indicates that they are pentacoordinated without the sixth extraplanar ligand (Rakshit and Spiro, 1974; Strekas and Spiro, 1974; Sievers et al., 1979; Osterlund et al., 1979). Cytochrome e' shows an unusual resonance Raman spectrum. This is interpreted as due to the carbonyl or carboxyl oxygen's occupying the sixth position, with histidine at the fifth coordination position (Teraoka and Kitagawa, 1980b).
5.
OXIDATION STATES
Different oxidation state of heme compounds usually show lines at different frequencies because the electronic configurations of ferric and ferrous irons result in different bonding to the porphyrin ring. Therefore, the relation between Raman frequency and oxidation state is by no means direct; nevertheless, some Raman lines can be used to probe the oxidation state of the heme iron empirically (Table 12.4). One should keep in mind that Raman frequencies can be influenced by many factors such as the type of axial ligands. For example, the 1370-cm -1 line for leghemoglobin a is not only an oxidation-sensitive line, but it is also ligand sensitive (Osterlund and Sievers, 1981). 5.1.
CYTOCHROME
c
Raman spectra of cytochrome e and hemeundecapeptide are sensitive to the oxidation states of the iron atoms (Strekas and Spiro, 1972a; Loehr and Loehr, 1973; Adar, 1978). The characteristic frequency for the reduced cytochrome e lies between 1355 and 1362 cm- 1 (Yamamoto et al., 1973). For the oxidized compound, the characteristic frequency is between 1369 and 1375 cm - I (Table
344
Hemes and Porphyrins
TABLE 12.4.
Iron-Oxidation-State Indicator Raman Bands for Hemoprotein
Compound Cytochrome c Complex III (cytochromes b + c 1) Cytochrome bs Horseradish peroxidase Native Native (high spin) +CN (low spin) Compound II (low spin) Cytochrome P-450 Cytochrome c peroxidase Native Compound I Cytochrome oxidase
Cytochrome c3 cytochrome c' Hemoglobin Myoglobin Heme a (Imh Heme IX (CNh Heme IX (Imh Cytochrome P-450
Fe(II) (em-I)
Fe(I1I) (em-I)
Fe(IV) (em-I
1361
1372
Yamamoto et aI. (1973)
1359 1359
1370 1373
Yamamoto et aI. (1973) Yamamoto et aI. (1973)
1359
1378 1375 1375
Yamamoto et aI. (1973) Rakshit et aI. (1976) Rakshit et aI. (1976) Rakshi t et aI. (1976) Felton et aI. (1976) Shimizu et al. (1981) Ozaki et aI. (l976b, 1978)
1382 1342 1346
1362
1359 1355 1356 1355 1360 1360 1359
1371-1373
Reference
1373
Sievers et aI. (1979)
1372
Babcock and SaImeen (1979) Kitagawa and Orii (1979) Carter et aI. (1981) Kitagawa et aI. (1976b) Kitagawa et aI. (1976b) Yamamoto et aI. (1973) Yamamoto et aI. (1973) Kitagawa and Orii (1979) Kitagawa and Orii (1979) Kitagawa and Orii (1979) Anzenbacher et aI. (1980)
1374 1373 1372 1371 1371 1375 1373 1374 1368 and 1372
.
12.4). Raman spectra of reduced and oxidized cytochrome c are shown in Figure 12.12. When cytochrome c is reduced, there is a transient reduced state before it becomes a stable reduced form. The lifetime of the transient reduced state is in the range of 20 ms to a few seconds. As cytochrome c is reduced, the 1567 cm- 1 band momentarily shifts to 1533 cm- 1 and then to the stable reduced state band of 1547 cm -1 (Cartling and Wilbrandt, 1981). The Raman oxidation-state marker band originates from the ring vibration of pyrrole (Ca - N symmetrical stretching vibration) mixed with in-phase displacement of the four pyrrole nitrogen atoms toward the central iron atom. This conclusion is based on evidence from the spectroscopic study of IsN_ enriched derivatives (Kitagawa et al., 1977a) and also from the normal coordinate calculation (Abe et aI., 1976).
345
Oxidation States 1584
.. I~
C''':i~ ,IllrJ\"l) ~~'
698
f
c.~ ~vwd\W\
o
75t
E
'h
'589
361
o
0::
Cytochrome C (II.)
690
NL ~ 1228 it32it74
,,,e~
~....J'J
~
~
~'489t547
I AEX =406.7nm
300 400 500 600 700 SOO 9001000 ilOO 120013001400 1500 1600
Wove Number (eM-I)
FlGURE n.n. Resonance Raman spectra of Fe(III)- and Fe(II)-cytochrome c excited at 406.7 nm (Soret band). The figure was reproduced from Adar (1978) with copyright permission of American Chemical Society.
5.2.
CYTOCHROME OXIDASE (CYTOCHROME
c OXIDASE)
The resonance Raman spectrum of cytochrome oxidase originates from both heme a and heme a 3 and does not involve copper (Salmeen et a1., 1973, 1978a, b; Adar and Erecinska, 1979). Oxidized cytochrome oxidase is photoreduced under laser illumination (Adar and Yonetani, 1978; Salmeen et a1., 1978a). Fully oxidized cytochrome oxidase has an oxidation marker band at 1372-1374 cm- I , whereas the marker band of reduced form is observed at 1362 cm- 1 (Kitagawa and Orii, 1979; Carter et a1., 1981). These bands are reversible, depending on the oxidation state of cytochrome oxidase iron. This suggests that fully oxidized and reduced cytochrome oxidase have different heme configurations. This is generally believed to be due to the presence of two nonequivalent hemes in the enzyme, as suggested by Salmeen et a1. (1978a, b; Babcock and Salmeen, 1979; and Kitagawa and Orii, 1978). The low-frequency bands observed in the resonance Raman spectrum of oxidized cytochrome oxidase (excitation wavelength of 600 nm) do not disappear upon reduction of the protein. From this it is concluded that resonance Raman bands excited at 600 nm are entirely due to heme (Chan et a1., 1979; Bocian et a1., 1979). Photoreduction can be avoided by flowing the sample
346
Hemes and Porphyrins
(Babcock and Salmeen, 1979). When cytochrome c oxidase is excited at 441 nm, the laser induces photoreduction, which can be slowed down by cooling the sample to - 100e. The slowdown of the reduction process makes it possible to study the various states of reduction by analyzing the Raman spectra. The relative intensities of the 1609- and 1623-cm- I doublet are affected by reduction. From the reductive titration, it was concluded that the two hemes are nonequivalent but interacting (Adar and Yonetani, 1978; Adar and Erecinska, 1979). 5.3.
OTHERS
In spectra of yeast cytochrome c peroxidase, the bands at 1500 and 1480 cm- I are sensitive to oxidation state (Sievers et al., 1979). Other compounds, such as cytochrome bs and horseradish peroxidase, also give valence-state-dependent bands (Yamamoto et al., 1973). These are summarized in Table 12.4. A number of heme proteins show some characteristic features with excitation in the Soret region. For instance, Fe(II)-heme proteins always show their strongest band between 1356 and 1361 cm -1, whereas in ferric proteins it occurs between 1370 and 1378 cm - I. The strongest band of oxyhemoglobin occurs at 1375 cm -I, which is the typical band for Fe(III)-heme proteins. From this evidence, Yamamoto et al. (1973) proposed that the iron in oxyhemoglobin is in the formal low-spin ferric state, Fe(III)-O; , instead of the normally accepted Fe(II) . O2 , Resonance Raman spectra of iron tetraphenylporphyrin indicate that there are lines sensitive to spin and oxidation state just as in natural heme proteins (Burke et al., 1978; Chottard and Mansuy, 1980).
6.
POLARIZATION
Laser light is plane polarized, so the depolarization ratio can take any value from zero to the maximum of 0.75. When the depolarize.tion is 0.75, the light is depolarized. This occurs when the vibrating atoms are asymmetrical. If the vibration is totally symmetrical the depolarization ratio becomes zero, and the light is referred to as polarized. The depolarization ratio is a measure of the degree of symmetry of the vibrational center. Since the resonance Raman bands of heme compounds arise mainly from vibrations of atoms in the porphyrin macrocycle, a change in symmetry of porphyrin will cause changes in the polarization of the scattering, which can be detected from the depolarization ratio. In heme proteins, the symmetry is even further reduced because of the different fifth and sixth ligands to iron. The depolarization ratio can also be influenced by other environmental factors such as peripheral substitution (Pezolet et a!., 1973; Mendelsohn et al., 1975a; Verma et al., 1974). It also depends on whether Soret excitation or a, f3 excitation is used (Callahan and Babcock, 1981).
~!!
Polarization
347
A difference in polarization is reported for normal and sickle cell oxyhemoglobin; this is especially apparent in the N-Ca-band region. The Raman technique may prove to be useful for detecting abnormal hemoglobin by observing polarization (Barrett, 1979). 6.1.
INVERSE DEPOLARIZATION
One interesting finding is that some heme proteins have resonance Raman lines with inverse polarization. The depolarization ratio (p = 11-/Iii) becomes greater than or reaches an infinitely high value (00) (Spiro and Strekas, 1972; Felton et aI., 1976). The origin of inverse polarization is related to an antisymmetrical molecular-scattering tensor that is nonzero near resonance. The antisymmetry is associated with a particular class of vibrations of the tetragonal heme chromophores. This aspect was extensively studied by Pezolet et aI. (1973). They indicated that as the symmetry of the heme group is lowered by interaction with the protein, the changes are reflected in the three invariants of the Raman tensors. This may provide a method for probing changes in heme protein structure. Those interested in the theoretical aspects of inversely polarized modes of heme compounds are advised to read the articles by Zgierski and Pawlikowski (1978) and Shelnutt (1980a).
*
6.2.
HEME COMPOUNDS
Inverse polarization is quite common for heme compounds (Figure 12.13). Many prominent bands such as the 1585 and 1310 cm- 1 bands display inverse polarization, thus the depolarization ratio p = 11- /1 11 is infinity or a very large number when cytochrome c is excited at the 514.5-cm - I line. As the excitation wavelength is lowered to 457.9 and 363.8 nm, the depolarization ratios become lower (Nafie et aI., 1973). Similarly, Collins et aI. (1973, 1976) observed that the depolarization ratio depends strongly on laser frequency, and symmetry assignments of heme proteins based on the depolarization ratio should not be based on data at a single frequency. For the anomalously polarized peaks (p > the depolarization increases to a maximum when the laser frequency is midway between the a- and ,a-absorption bands. Hemin does not show any inversely polarized bands, whereas the imidazole complex shows several such bands using the a- and ,a-absorbance for excitation (Verma and Bernstein, 1974b). Cytochrome oxidase contains cytochromes a and a 3 • The Soret band for reduced cytochrome oxidase occurs at 440 nm. Raman spectra of cytochrome oxidase lack noticeable inverse polarization (Nafie et aI., 1973). This observation confirms the fact that when excited near the a-band, the depolarization ratio for heme a vibrations is much lower than for protoheme compounds (Babcock et aI., 1979a).
*),
348
Hemes and Porphyrins It'>
5145
III
'"
~~:ll:;:mg
.~
1-----
~
It'>
a
-
$
0
"
!!?
~I
'----.:.-.JL...
4579
It'>
tot1>
III
'"to-
It'>
a
It'>
t1>
III
~.
to-
;_N
I}
FIGURE 12.13. Examples of inverse polarization in cytochrome c. See the bands at l3l3 and 1585 cm- I when the compound is excited at 5145 A (top). The inverse polarization depends on the wavelength of laser excitation. See the 1313 and 1585 cm -1 bands at 4579-A excitation, the depolarization ratios become low (bottom). The figure was reproduced from Nafie et al. (1973).
6.3.
METALLOPORPHYRINS
The anomalously polarized Raman bands observed in heme compounds are also commonly found in other metalloporphyrins. The extent of anomalous polarization is unique for each compound, and it can be used to probe the structure of the chromophore in solution (Sunder et al., 1975; Asher and Sauer, 1976). Resonance Raman spectra of Ni(II)-, Co(II)-, and Cu(II)-mesoporphyrin IX dimethyl ester reveal that intensity (excitation-profile effect) and depolarization of Raman bands vary with the exciting wavelength. Anomalously polarized bands appear at 1305 and 1602 em-I for the Ni(II) compound, 1308 and 1597 em-I for the Co(II) derivative, and 1313 and 1581 cm- I for the Cu(II) compound (Verma et al., 1974). Ni, Cu, and Pd complexes of octamethylporphyrin and mesotetraphenylporphyrin also show several such anomalously polarized vibrations (Mendelsohn et aI., 1975). In spectra of copper porphin, such bands are found at 1322 and 1587 cm- I using an excitation line of 514.5 nm. Copper porphin has a unique structure with no peripheral substituents except hydrogen (Figure 12.14). It is interesting
Quarternary Structure 2
a
349
3
13
5 7
v
6
2
a
3
5 7
6
FIGURE 12.14. Structure of Cu·porphins (bottom) and porphin (top).
to observe that depolarization ratios for the two anomalously polarized bands remain constant at 45 and over all the exciting wavelengths in the a- and ,B-absorption regions, whereas some polarized bands show variation in their depolarization ratios (Verma and Bernstein, 1974c). 7.
QUATERNARY STRUCTURE
Biologically active hemoglobin is a tetramer. The mechanism of the cooperative oxygen binding of hemoglobin has been the subject of intensive study. The oxygen-binding curve of hemoglobin is sigmoidal; oxygenation becomes easier as the hemoglobin units in the tetramer become saturated. An important question arises concerning the mechanism of such an allosteric effect. Is the protein moiety responsible, or is there any change in heme structure? The primary structure of the four deoxyhemoglobin subunits is identical to that of the subunits of oxyhemoglobin, but they differ in quaternary structures, or in how the detailed protein three-dimensional structures are oriented in relation to each subunit. Two types of quaternary structure of methemoglobin have been proposed (Perutz, 1970). The T structure has low affinity for oxygen, whereas the R
350
Hemes and Porphyrins
structure has high affinity. There is a difference in the free energy of oxygen binding for the two structures. The difference in free energy is called the free energy of cooperativity and is about 3.6 kcal/mol. The T state is favored by deoxyhemoglobin and the R state is preferred by oxyhemoglobin. The cyrstal structures of ligated hemoglobins in the R form and deoxyhemoglobin in the T form do show the difference (Shelnutt, 1980b). In the R structure, the phenylalanine COl is in van der Waals contact (3.5 A) with the heme and very nearly parallel to it. The relative positions of phenylalanine residues of COl and G5 with respect to the heme for Rand T types of hemoglobins are shown in Figure 12.15. One question is where and how the free energy of cooperation is stored in the hemoglobin molecule. One speculation is that a particular bond is twisted or stretched, and this energy is released when the twisted bond is relaxed and extended. Frequently a metal-ligand bond is proposed to carry such an enhanced tension (Hoard and Scheidt, 1973). However, there is no experimental proof to indicate the presence of such a bond (Hopfield, 1973; Scholler et ai., 1976; Kincaid et ai., 1979a, b; Scholler and Hoffman, 1979a, b; Warshel and Weiss, 1981).
G5
~" G5
~ DEOXY
LIG NDED
COl
ALPHA CHAINS
"'"'~'"'"
~~EOY COl
LIGA OED
BETA CHAINS
FIGURE 12.15. Positions of phenylalanine side chains (G5 and CDI) with respect to the heme in the R (carbonmonoxyhemoglobin) and T structures (deoxyhemoglobin). The figure was reproduced from the abstract for Shelnutt (1980), who obtained it from Cyrus Chothia. Permission to reproduce the figure was granted by Dr. Chothia.
~
auarternary Structure
351
The addition of inositol hexaphosphate to nitrosylhemoglobin (HbNO) changes its visible and uv spectra significantly, whereas no effect is observed with HbOz. IHP is believed to stabilize the T structure. This is thought to be due to a strong interaction between the IHP and deoxy-HbNO that is not present with oxy-HbNO. This may stabilize deoxy-HbNO in the presence of IHP. It was found that there is a significant intensity change in the Raman band at 1643 cm -I and a decrease in the intensity of the 1633-cm- I band. Apparently these two bands are sensitive to changes in the quarternary structure of the protein (Szabo and Barron, 1975). The addition of IHP induces a change from the R to T quaternary structure of HbNO. Under this condition, the ,B-chains maintain the hexacoordinate structure of ImFeNO, whereas the a-chains are pentacoordinates with the FeNO (Maxwell and Caughey, 1976). The resonance Raman spectrum indicates that a new band at 592 cm- I (Fe-NO stretching) is produced in addition to the 553-cm- 1 Fe-NO band observed without IHP. This suggests that iron-imidazole bonds are broken in the a-chains in the T quaternary structure (Stong et aI., 1980). Quaternary structures of aquomethemoglobin and fluoromethemoglobin are also changed by the addition of IHP. The IHP changes the allosteric equilibrium of high-spin methemoglobin derivatives in favor of the T state (Perutz et aI., 1974). It is reported that the change in quaternary structure alters the equilibrium of high-spin and low-spin azidomethemoglobin from 10/90 to 45/55% (Scholler and Hoffman, 1979a, b). There have been several attempts to resolve quaternary-structure differences by resonance Raman spectroscopy. In order to see whether Raman spectroscopy can detect changes in quaternary structure induced by IHP, the protein structurally sensitive line at 1370 cm- 1 was examined (Rousseau et aI., 1980a). In all liganded methemoglobins known to have a change in quaternary structure, the frequency of the 1370-cm - I line decreases. In those in which no quaternary-structure change occurs, such as met HbA(Im-) and met HbA(Nj), the frequency difference is very small (Rousseau et aI., 1980b). When the sample was excited at 441.6 nm, which is close to the Soret band, only the T form of methemoglobin was partially photoreduced (Kitagawa and Nagai, 1979). Thus the two different quaternary-structure forms have different chemical reactivities. It is reported that the Fe(II)-N(histidine) band at 206 cm- 1 and a peripheral-group bending mode at 348 cm -I are also sensitive to the quaternary-structure change (Hori and Kitagawa, 1980; Nagai et aI., 1980a, b, c; Nagai and Kitagawa, 1980). When HbCO is photolyzed, the original 214-cm- 1 band shifts to 225 cm- I. Since the 214-cm- 1 band is close to the 206-cm- 1 band, it is speculated that a conformational change of hemoglobin may have occurred, for the early period of photodissociation, from 30 ps to 20 ns (Irwin and Atkinson, 1981). The resonance Raman spectral difference observed for deoxy Rand T structures is possibly due to the protein environment, but interaction between
352
Hemes and Porphyrins
the protein structure and the heme via the axial ligand is not ruled out (Shelnutt et al., 1979a, b).
8.
PROTEIN ENVIRONMENT
Resonance Raman spectra of heme proteins arise from heme vibrational modes. There are conflicting reports on whether the protein environment affects the resonance Raman spectra. If proteins do not affect the porphyrin ring, then they should show the same Raman spectra as long as the heme components are identical. On the other hand, if the apoprotein moiety influences the electronic-charge density on the porphyrin ring, then it will affect the spectra. Some investigators concluded that resonance Raman spectra do not reflect changes in the protein environment (Loehr and Loehr, 1973; Woodruff et al., 1975; Spiro et al., 1978; Spiro and Burke, 1976). Differences in Raman frequencies of heme proteins that contain the same heme group are indeed small. These small differences are attributed by some investigators to differences in the proteins' environment (Kitagawa and lizuka, 1974; Adar and Erecinska, 1974; Yamamoto et al., 1976). When cytochrome c from different species (horse, cow, dog, pigeon, spider monkey, turtle, human, tuna, baker's yeast, Candida krusei, Rhodospiri/lum rubrum) is examined, there are small (0-6 cm -1) frequency differences in the heme vibrational modes (Shelnutt et al., 1979a; 1981). Similarly, chemically modified (on an argInine residue) human deoxyhemoglobin has come up to 2.2-cm- I difference at 1567 cm- I compared with the unmodified compound (Shelnutt et al., 1979b). These differences are attributed to the effect of the protein moiety. In NMR, heme resonances vary only when there is a large change in the protein environment (Keller and Wuthrich, 1978). A dramatic change takes place in the vibrational properties of the FeN(histidine) bond when deoxyhemog10bin is frozen. This probably suggests that the protein interacts with water and affects the quaternary structure which in turn influences the Fe-N bond (Ondrias et al., 19&1).
9.
GEOMETRY OF THE HEME RING
There are a number of Raman active-modes that are sensitive to the geometry of the heme. A particularly important one is the anomalously polarized line at about 1590 cm- I . The type of ligand has a great influence on the position of iron with respect to the porphyrin plane. In the absence of a sixth ligand (pentacoordinated complex), the iron atom is out of plane. With a sixth ligand (hexacoordinated complex), the iron atom is pulled in the opposite direction by the sixth ligand (Figure 12.16). So whether or not the iron is in the plane of the porphyrin depends on the relative effects of the fifth and sixth ligands on the iron atom.
~
Geometry of the Heme Ring
/ L-Fe
\
L
I
L-Fe-L
I
353
FIGURE 12.16. Position of the central iron atom in pentacoordinated (left) and hexacoordinated (right) complexes.
There are two hypotheses concerning the geometry of the heme ring. The doming model was proposed by Spiro and his co-workers (Spiro and Strekas, 1974; Stein et al., 1975). According to this model, the central iron atom is situated out of plane to the heme and located above the porphyrin plane. However, there is no clear evidence that the heme is considerably "domed" in deoxyhemoglobin or other high-spin heme proteins (Yu, 1977). According to core expansion theory (Spaulding et al., 1975), the iron atom may stay in the center of the porphyrin ring, but the ring expands and shrinks. As the ring shrinks, the iron atom moves above the plane of the porphyrin ring. The 1590-cm- 1 (1580-1610 cm- I ) line is structurally sensitive in metalloporphyrin spectra, and there is an empirical correlation between its frequency and the distance from the center of the porphyrin core to the pyrro1e nitrogen atoms, d(C/-N) (Felton et al., 1974; Spaulding et al., 1975) (Figure 12.17). This structurally sensitive line is anomalously polarized (Spiro and Strekas, 1972). The line may arise from stretching and bending of the methine-bridge bonds. According to the core expansion model, an increase in the C/-N distance corresponds to a linear decrease in this frequency (1580-1610 cm -I). Using this plot, Lanir et al. (1979) came to the conclusion that the distance C/-N is 2.033 ± 0.01 A at acidic pH. X-ray diffraction study of diaquo( a, {3, 'I, c5-tetraphenylporphinato)Fe(III)-perchlorate, an iron porphyrin
FIGURE 12.17. Position of central iron atom in heme, where d(C,-N) is defined as the distance between the center of the porphyrin core and the pyrrole nitrogen. The figure was reproduced from Spaulding et al. (1975) with permission of the copyright owner, American Chemical Society.
354
Hemes and Porphyrins
with two weak-field axial ligands, showed that the average Fe-N-bond distance in this heme model is only 2.04 A (Kastner et aI., 1978). The short distance of the Fe-N bond in cytochrome c high-spin hemes with two weak axial ligands can only be explained if the iron(III) lies in plane with the heme. Myoglobin forms a stable complex with hydrogen peroxide in the pH range 8-9. In order to study the nature of the complex, Campbell et aI. (1980) examined Raman spectra of complexed and uncomplexed myoglobin. The distances from the iron to the pyrrol~ nitrogen (Fe-Np ) and the imidazole nitrogen (Fe-N im ) are 2.08 and 2.09 A, respectively, according to stretchingvibration bands at 345 and 375 cm -I and from the available X-ray data. The distance from the center of the porphyrin to the pyrrole nitrogen atoms Cr-N, as estimated from the 1560-1590-cm- 1 bands, is 2.01 A. There is X-ray evidence to indicate that core expansion takes place in some compounds. For instance, in (meso-tetraphenylporphinato)bis(tetrahydrofurna)-Fe(II), the iron atom is centered in the porphyrin plane, causing a greater amount of radial core expansion than observed in other iron porphyrin structures (Reed et aI., 1980). This is a rather interesting exception, because high-spin Fe(lI) porphyrins are usually pentacoordinated complexes, with large out-of-plane iron displacement. Deoxyhemoglobin has high-spin Fe(II) and is a pentacoordinated complex in which the iron atom lies substantially out of plane to the porphyrin. It is known that the transition of the R to the T quaternary stfllcture in carp azidemethemoglobin accompanies the change of spin state from low to high. The change in the high-frequency region of 1300-1700 cm- I reflects the change of spin states. The low-frequency band changes from 270 to 263 cm -I. This band is sensitive to the iron-heme plane distance (Desbois et aI., 1980). Instead of the anomalous polarized line near 1590 cm- 1 that appears in the spectra of compounds, a line appears at the unusually low frequency of 1550 cm - I in spectra of leghemoglobin, which is found in the nitrogen-fixing bacteria in legume root nodules. This indicates an expanded porphyrin core in leghemoglobin (Armstrong et aI., 1980). The iron of hemoglobin can be replaced with cobalt(lI) without loss in the hemoglobin's ability to bind oxygen. Deoxy-Co(lI)Hb is low spin. In hexacoordinated Co(lI)-porphyrin, the cobalt atom is small enough to stay in the center of the porphyrin-ring plane (Woodruff et aI., 1974). An important question is what factor makes the core expand. The conclusion of Spiro et aI. (1979) is that the core size is probably due to the effect of axial ligands.
10.
MITOCHONDRIA
Oxidation is an important process in metabolism. Most of the foodstuffs such as the carbohydrates and lipids that we eat are highly reduced compounds. Metabolism of these compounds involves mainly the removal of hydrogens or
Mitochondria
355
electrons using molecular oxygen as the final electron acceptor (or hydrogen acceptor). This involves a series of electron carriers, and during this process ATP is produced through oxidative phosphorylation in mitochondria. Substrate
10.1.
->
NAD
->
cyt c
->
->
flavin coenzyme
->
cyt (a + a3) cytochrome oxidase
coQ
->
->
cytb
->
cytc]
O2
INTACT MITOCHONDRIA
Resonance Raman spectra of intact reduced mitochondria were obtained by Adar and Erecinska (1978) (Figure 12.18). They were able to identify the contributions from the individual cytochromes and to monitor the physical and biochemical states of hemes in biologically active membranes. The intensity of Raman spectra depends on the excitation wavelength relative to the peak positions of the a-, {3-, and y-absorption bands of cytochromes. Isolated cytochrome b has marker bands at 1306, 1342, 1538, and 1564 cm- I . In this membrane system, these bands shifted to lower frequencies. Incidentally, the formation of the cytochrome b-c, complex shifts these marker bands to lower frequencies. 10.2.
CYTOCHROMES bAND c
When the cytochrome b-c] complex is excited at 531 nm, there are anomalous features in the Raman spectra that are explained as interactions between band c-type hemes in the membrane preparation. For instance, one of the marker bands (1297 cm - I) is low compared with the usual position at 1305-1340 cm- I in other proteins containing heme b. The other marker band .... to
~u' 441Gnm
~
5
"£
!2
...., ....
<5 cfl ~ ~
2
~
u
£
S
FIGURE 12.18. Resonance Raman spectrum of whole reduced mitochondria excited at 441.6 nm. The figure was reproduced from Adar and Erecinska (1978) with permission of copyright owner, American Chemical Society.
356
Hemes and Porphyrins
for heme b at 1340 cm- I is present even when only cytochrome b is oxidized or when only cytochrome C I is reduced. The persistence of these bands can only be explained by the quantum-mechanical mixing of vibration levels (Adar and Erecinska, 1977). 10.3.
CYTOCHROME OXIDASE
Spin and oxidation properties of cytochrome oxidase are discussed elsewhere (Sections 3 and 5). In this section, cytochrome oxidase is discussed again in the context of the electron acceptor and electron donor to molecular oxygen. Cytochrome c oxidase (cytochrome oxidase) is the terminal enzyme in the mitochondrial electron-transport chain. Cytochrome oxidase is composed of two different heme proteins, cytochrome a and cytochrome a 3 , and 2 mol of copper. Resonance Raman spectroscopy can be used to distinguish between these two hemes. By the proper selection of reducing agents, Salmeen et aI. (1978a, b) were able to reduce cytochrome a while a 3 remained oxidized. The Raman spectrum of the resulting mixture lacked several bands observed in the spectrum of the fully reduced enzyme. Thus by observing vibrational spectral differences they proved that the two hemes are structurally dissimilar. The magnetic dissimilarity of cytochromes a and a 3 is known from the study of electron paramagnetic resonance. Both the a and the a 3 species contribute to the Raman spectrum of cytochrome oxidase, depending on the excitation wavelength and the oxidation state (Babcock et al., 1980). It was determined that the Raman bands of fully reduced cytochrome oxidase at 215,364, 1230, and 1670 cm- I were due only to Fe(II)-cytochrome a 3' Moreover, the 1670-cm - I band originates from the C=O stretching vibration of the formyl group of Fe(II)-cytochrome a3' The formyl group is the side chain of the porphyrin macroring. It appears in the resonance Raman spectrum because the formyl group is in the plane of porphyrin, so it conjugates with the 'IT-electron system of porphyrin. Recently it was proposed by Seiter et al. (1978) that a carboxylate group of aspartic or glutamic acid residues serves as a bridge between the heme-a 3 iron and one of the copper atoms in the oxidized cytochrome oxidase. It was speculated that this bridging is disrupted by reduction of the metal centers, and that the molecular-oxygen-binding site is blocked by the metal-metal bridging ligand until reduction is completed.
11. 11.1.
OTHER STUDIES EFFECT OF REDOX POTENTIAL
An interesting study was done by Adar and Erecinska (1977) on Raman intensity at 1298 cm - I as a function of redox potential, using cytochromes bS66 ' bS6J ' and c\ excited at 530.9 nm. Although there is a general tendency for
....iIi
References
357
the Raman intensity to decrease as the redox potential increases, the proper explanation of this result is difficult. For instance, the laser itself causes photoreduction of the samples, and the a-band of ferrous cytochrome b566 disappears as oxidation takes place (Adar and Erecinska, 1977). The electrochemical method was coupled with resonance Raman spectroscopy to titrate and determine potentiometrically the formal reduction potential of cytochrome c. From this study it was proposed that the redox potential of heme can be found by quantitative measurement of resonance Raman spectra (Anderson and Kincaid, 1978). 11.2.
TEMPERATURE
There is little change in the Raman frequencies of cytochrome c when the temperature is lowered. However, at lower temperatures, the Raman line is more enhanced and hence is better resolved (Champion et aI., 1976a). Of course, there is a practical advantage to taking Raman spectra at lower temperatures, since this prevents the overheating of the samples by the laser. Fe(II)-cytochrome cm isolated from the thermophilic bacterium Thermus thermophilus is an interesting heme protein as it is stable to heat. Even at 87°C there is no change in its resonance Raman spectrum, which indicates that methionine is still bound to the heme iron, so it retains the low-spin state (Hon-Nami et aI., 1980). However, temperature-induced spin transition can be observed for methemoglobin derivatives. The Raman band is observed to shift from the high-spin marker band (1480 cm - I) to the low-spin marker band (1505 cm- I) by cooling from room temperature to 77 K (Cho et aI., 1981).
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358
Hemes and Porphyrins
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_ro
.
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Hemes and Porphyrins
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Tsubaki, M., and Yu, N.-T. (1982). Resonance Raman investigation of nitric oxide bonding in nitrosylhemoglobin A and -myoglobin: Detection of bound N-O stretching and Fe-NO stretching vibrations from the hexacoordinated NO-heme complex. Biochemistry 21, 1140. Tsubaki, M., Nagai, K, and Kitagawa, T. (1980). Resonance Raman spectra of myoglobins reconstituted with spirographis and isospirographis hemes and iron 2,4-diformylprotoporphyrin. IX. Effect of formyl substitution at the heme periphery. Biochemistry 19, 379. Tsubaki, M., Srivastava, R. B., and Yu, N.-T. (1981). Temperature dependence of resonance Raman spectra of metmyoglobin and methemoglobin azide vibrations and iron-azide stretch. Biochemistry 20, 946. Tsubaki, M., Srivastava, R. B., and Yu, N.-T. (1982). Resonance Raman investigation of carbon monoxide bonding in (carbon monoxy) hemoglobin and -myoglobin: Detection of Fe-CO stretching and Fe-C-O bending vibrations and influence of the quaternary structure change. Biochemistry 21, 1132. Van Steelandt-Frentrup, J., Salmeen, I., and Babcock, G. (1981). A ferrous, high-spin heme a model for cytochrome a3 in the dioxygen reducing site of cytochrome oxidase. J. Am. Chem. Soc. 103, 5981. Verma, A L. (1976). Effect of peripheral substituents on resonance Raman spectra of metalloporphyrins and heme proteins. In Proc. Tnt. Conf. Raman Spectrose., 5th, E. D. Schmid, J. Brandmueller, and W. Kiefer, Eds., Hans Ferdinand Schulz Verlag, FreiburgjBr., Germany, pp. 198-199. Verma, A L., and Bernstein H. 1. (1974a). Resonance Raman spectra of metal-free porphin and some porphyrins. Biochern. Biophys. Res. Commun. 57, 255. Verma, A L., and Bernstein, H. 1. (1974b). Resonance Raman spectra of protohemin and protohemin-imidazole complex. J. Raman Spectrose. 2, 163. Verma, A L., and Bernstein, H. J. (1974c). Resonance Raman spectra of copper-porphin. .I. Chem. Phys. 61, 2560. Verma, A L., Mendelsohn, R., and Bernstein, H. 1. (1974). Resonance Raman spectra of the nickel, cobalt and copper chelates of mesoporphyrin. IX. Dimethyl ester. J. Chern. Phys. 61, 383. Walters, M. A, Spiro, T. G., Suslick, K S., and Collman, J. P. (1980). Resonance Raman spectra of (dioxygen)(porphyrinato)(hindered imidazole) iron (II) complexes: Implications for hemoglobin cooperativity. J. Am. Chern. Soc. 102,6857. Warshel, A, and Weiss, R. M. (1981). Energetics of heme-protein interactions in hemoglobin. J. Am. Chem. Soc. 103,446. Woodruff, W. H., and Farquharson, S. (1978). Time-resolved resonance Raman spectroscopy of hemoglobin derivatives: Heme structure changes in 7 nanoseconds. Science 201, 831. Woodruff, W. H., Spiro, T. G., and Yonetani, T. (1974). Resonance Raman spectra of cobalt-substituted hemoglobin: Cooperativity and displacement of the cobalt atom upon oxygenation. Proc. Nat. Acad. Sci. 71, 1065. Woodruff, W. H., Adams, D. H., Spiro, T. G., and Yonetani, T. (1975). Resonance Raman spectra of cobalt myoglobins and cobalt porphyrins. Evaluation of protein effects on porphyrin structure, J. Am. Chem. Soc. 97, 1695. Woodruff. W. H., Dallinger, R. F., Antalis, T. M., and Palmer, G. (1981). Resonance Raman spectroscopy of cytochrome oxidase using Soret excitation: Selective enhancement, indicator bands, and structural significance for cytochromes a and a3' Biochemistry 20, 1332. Yamamoto, T., Palmer, G., Gill, D., Salmeen, I. T., and Rimai, L. (1973). The valence and spin state of iron in oxyhemoglobin as inferred from resonance Raman spectroscopy. J. Bio/. Chem. 248, 5211. Yamamoto, T., Palmer, G., and Crespi, H. (1974). Resonance Raman studies of a c type algal cytochrome. Biochim. Biophys. Acta 349, 232. -
368
Hemes and Porphyrins
Yu, N.-T. (1977). Raman spectroscopy: A conformational probe in biochemistry. CRC Cril. Rev. Biochem., 4, 229. Yu, N.-T., and Tsubaki, M. (1980). Resonance Raman spectra of manganese myoglobin and its azide complex. Assignment of a new charge-transfer band to azide ('IT) .... porphyrin ('IT') transition. Biochemistry 19, 4657. Zgierski, M. Z., and Pawlikowski, M. (1978). Theory of depolarization dispersion of inversely polarized modes in heme proteins. Chern. Phys. Lett. 57, 438.
CHAPTER
Copper and other Metals in Biological Systems
1.
COPPER IN BIOLOGICAL SYSTEMS
There are many varieties of oxygen carriers in the animal kingdom. The most common and well known example is hemoglobin, which is found in all vertebrates and many invertebrates. Another iron-containing oxygen carrier is hemerythrin, which is a nonheme protein. The third major type is represented by copper-containing proteins the hemocyanins. Despite their common functions as molecular oxygen carriers, their chemical structures and properties are quite different. Copper is also found in many other proteins; cytochrome oxidase is a well-known example (Chapter 12, Sections 5 and 10). There are different copper sites in copper proteins and usually they are classified into three types, namely type 1, type 2, and type 3, based on their spectral properties. Type 1 copper sites are characterized by high absorbance near 600
370
Copper and Other Metals in Biological Systems
nm giving a deep blue color. Type 2 copper has less blue color. Type 3 Copper is less studied but the copper of this type in Rhus laccase is an antiferromagnetic-coupled cupric dimer. Ascorbate oxidase, polyphenoloxidase, tyrosinase, plastocyanin, amine oxidase, dopamine-,B-hydroxylase, and ceruloplasmin are all copper-containing enzymes. Copper-containing enzymes are the subject of intensive investigation using a variety of physical techniques such as ESR, CD, and absorption spectroscopy. Because of the relatively recent use of Raman spectroscopy in biological systems, fewer studies have been made using Raman spectroscopy. Despite this, Raman spectroscopy is becoming important in this field, and some important problems such as oxygen structure in oxyhemocyanin and the type of ligand in Cu-proteins have been studied by this technique. In this chapter, only the studies using Raman spectroscopy are reviewed.
1.1.
HEMOCYANIN
The blood of some nonvertebrates, such as the arthropods and mollusks, contains the blue pigment hemocyanin as the oxygen carrier. Hemocyanin is a metalloprotein containing 2 mol of copper per mole of protein. The molecular weight of hemocyanin is very large and ranges from half a million to over 10 million. One mole of copper binds to 2 mol of oxygen in a nonlinear fashion. There are several questions still to be answered concerning the oxidation state of the O2 in oxyhemocyanin and the oxidation state and arrangement of the two copper atoms. 1.1.1.
Oxygen in Hemocyanin
The oxygenated hemocyanin has a strong absorption band in the near uv at 350 nm and a weak band in the visible region at 570-580 nm. Thus one can obtain a resonance-enhanced spectrum of the blue-colored oxyhemocyanin by excitation with visible light such as 531-, 514.5-, 488-, 476.5-, and 457.9-nm laser lines. Both blue oxyhemocyanin and colorless deoxyhemocyanin are diamagnetic. The 570-580-nm absorption band is assigned to a charge-transfer transition of O{- to Cu(II) (Freedman et al., 1976). The origin of the 350-nm absorption band may be a simultaneous pair excitation (Larrabee et al., 1977). Oxygenation of hemocyanin produces two resonance-enhanced peaks at 742 and 282 em-I. The 742-cm- 1 peak is from the diatomic 0-0 stretching vibration determined by the replacement of 16 0 by 18 0, which causes a shift to 704 em-I. There is a small frequency difference in the peroxide (O{-) band depending on the source of hemocyanin. For instance, the vibration of this band is found at 744 em-I for arthropod hemocyanin, 749 em-I for mollusk hemocyanin (Freedman et al., 1976), 752 em-I for snail hemocyanin (Chen et al., 1979), and 741 em-I for the functionally active fragment (Gielens et al., 1980).
.J
':t
Copper in Biological Systems
CuliU -
0'0 _ CuUII
371
FIGURE 13.1. Simplified diagram of oxygen-binding site in oxyhemocyanin. The Cu-Oz-Cu is nonplanar and has JL-dioxygen-bridged geometry.
It is known that the stretching vibration of 0-0 is at 878 em -I for hydrogen peroxide (H 20 2 ) and at 738 em-I for Na 20 2 (Evans, 1969). If the O2
/
a
were the M- a type, the stretching vibration of 0-0 would occur around 1100-1140 em -I (Caughey et al., 1975; Szymanski et al., 1979). It is, therefore, reasonable to conclude that the bound oxygen is of the peroxide-ion type 01(Figure 13.1). A more-detailed model was proposed by Lontie and Gielens (1979) and is shown in Figure 13.2. The 282-cm- 1 peak is insensitive to isotopic substitution and is probably associated with the Cu-02 -Cu vibration (Loehr et al., 1974). The same conclusion was reached by Chen et al. (1979). However, the 282-cm- 1 band was assigned to the Cu(II)-imidazole stretching mode based on the frequency of the Cu(II)-imidazole complex as described by Larrabee et al. (1977). Further clarification of the assignment of this band is required. The 0-0 vibration of molecular oxygen appears at 1555 em-I, that of superoxide (a-a-H) at 1101 em-I, and that of peroxide (H-O-OH) at 878 em -I. Therefore, it is concluded that oxygen is bound as a peroxide ion in oxyhemocyanin. Tryptic fragments of hemocyanin retain the active site, as can be seen from the 0-0 stretching vibration (Gielens et al., 1980). Oxygen binding is an oxidative addition process in which O2 is reduced and the two Cu(II) centers of colorless deoxyhemocyanin are converted to the blue Cu(II) state (Thamann et al., 1977). 1.1.2.
Ligands
The Cu-N (imidazole) ligand vibration has been determined to be at 282 cm- I by a number of investigators (Table 13.2). For the functionally active fragment of tryptic hydrolysis, the Cu-N band was observed at 270 cm- l (Gie1ens et al., 1980). An excitation line in the visible region was used for most of the hemocyanin studies. The resonance spectrum of hemocyanin can be obtained by excitation at 350 cm -I. The 226- and 267-cm -I bands were assigned to Cu-N (imidazole) vibration modes; however, no Cu-O stretching mode was observed (Larrabee and Spiro, 1980). Addition of ethyleneglycol converts the blue color of oxyhemocyanin to purple. This suggests that some change occurs at or near the active site of
L 1m, / ' " ",1m Im-cu CU,lm Im/ '0-0..... 1m
FIGURE 13.2. A proposed model of the oxygen-binding site of oxyhemocyanin based on evidence from a combination of studies. 1m refers to the imidazole of histidine residues, and L refers to a bridging ligand, probably from a tyrosine residue. The figure was obtained from Lontie and Gielens, 1979.
372
Copper and Other Metals In Biological Systems
hemocyanin with the addition of ethyleneglycol. This color change takes place only with oxyhemocyanin and not with deoxyhemocyanin. Raman spectra of both purple and blue hemocyanins show a peroxide-dianion band around 750 cm -1, indicating that the color change is not due to the presence or absence of the peroxide dianion, but is probably due to a modification of the protein moiety, which eventually affects the copper active site (Mori et al., 1980; Nakahara et al., 1980). 1.2.
OTHER BLUE COPPER PROTEINS
There is a variety of copper-containing proteins with deep blue color due to their intense absorption near 600 nm. The intensities of the blue color are much stronger than those of simple cupric complexes. Tyrosinase is a copper-containing protein that interacts with molecular oxygen to hydroxylate monophenols. Oxytyrosinase and oxyhemocyanin have similar absorption and Raman spectra. The numerous low-frequency bands are probably due to the vibrational modes involving Cu-N and N-Cu-N. The 755-cm- t band originates from the 0-0 peroxide stretching vibration, as in the isotopic '80z-oxytyrosinase, the 755-cm- 1 band shifts to 714 cm- I • The copper atoms in oxytyrosinase are probably in a divalent state (Eickman et al., 1978). Stellacyanin isolated from the Japanese lac tree is a conj\Jgated protein containing mucopolysaccharides. It consists of 108 amino acid residues, 20% carbohydrate, and I mol of copper atoms. Ceruloplasmin is a plasma copper oxidase and has a blue color. A person with Wilson's disease has a low ceruloplasmin content. Ceruloplasmin and laccase contain six and four copper atoms, respectively. The intense absorption at 600 nm is believed to originate from S(Cys) ~ Cu(I1) ligand-to-metal charge-transfer transition. Raman spectroscopic evidence suggests that Cu-N bonds must also be present (Herve et al., 1981). Raman bands near 350-400 cm- I are assigned to Cu-N bond vibration for stellacyanin, ceruloplasmin, and laccase (Siiman et al., 1974). The band at 1640-1660 cm- I is assigned to C=O vibration involving Cu attached to the N atom of a peptide bond such as O-Cu
II
0----- Cu
I
or
- C - N-
II
I
- C - N-
The possibility of
o II -C-O-Cu is excluded, as a Cu-Gly-Gly complex gives a band at 1590 cm- I . Low-frequency Raman bands of ceruloplasmin at 415, 402, 382, 360, and 340 cm- I are assigned to the Cu-ligand stretching mode (Table 13.1). After .,j
Copper in Biological Systems
373
addition of azide and SCN- , the Cu(II)-ligand bonds of one of the two copper sites are disrupted, and the bands at 415,382, and 340 cm- I disappear. This suggests that the two copper atoms are not equivalent (Tosi et aI., 1975). There is no question that low-frequency bands arise from metal-ligand bond vibrations, but their exact assignments are difficult. Using CU-D-penicillamine as a model compound, it was assumed that the 375- and 427-cm- 1 bands originate from Cu-S stretching vibrations, and the 480-cm- 1 band is due to Cu-N vibration (Siiman and Carey, 1980). In addition to nitrogen-atom ligands, participation of cysteine-SH as a ligand in some blue copper proteins (stellacyanin, laccase, plastocyanin, ascorbate oxidase, and ceruloplasmin) is a possibility. The band near 260 cm -I in the blue copper proteins was assigned to the Cu-S vibrational mode by Siiman et aI. (1976). This assignment is reasonable. The 274-cm- 1 band of a model compound, Cu(II)NiSR), where SR is p-nitrobenzenethiolate or O-ethylcysteinate, is assigned to the Cu-S bond (Thompson et aI., 1977). However, instead of the cysteinyl (SH) group, methionine sulfur is proposed as a copper-binding site in blue copper proteins, (Ferris et aI., 1978). In the case of stellacyanin, which contains no methionine, a Cu-S (S from the disulfide bond) bond is suggested. There are also many vibrational bands in the 300-400 cm- 1 region for blue copper proteins. Judging from model compounds, some of these bands are suspected to be due to Fe(III)-S(Cys) and Fe-O stretching vibrational modes (Siiman and Carey, 1980). The Cu-N bond vibrations give intense bands near 415 and 380 cm -I. Moreover, weakly enhanced vibrational modes due to the amide are observed. They concluded that the "blue" copper site has a distorted fourcoordinated structure arising from the binding of copper to one cysteine sulfur
TABLE 13.1. Resonance Raman Frequencies of Blue Copper Proteins and Cu(II)- Sulfur Complexes in the CuN and CuS (mercaptide) Stretching and Ligand Bending Regions (Cm-').
Frequency and Assignment a
Compounds Ceruloplasmin I a Ceruloplasmin I b Azurin Azurin Tree laccase Plastocyanin Plastocyanin Stellacyanin CU(D-Pen) Cuddtc2 Assignments
422m 425m 434s 426s 420sh 425m
415m 415s 412s 419 415w
360m 374m 372m 383s 382s 379s
407s 408s 407w 408sh
388s 393w
Mostly CuN stretch
as: strong; m: medium; w: weak; sh: shoulder. Source:
and Garnier (1979).
340w
380m 402s
360w
350m 371s 367s CuS stretch
340sh 331w
337w 325w 312w Bending
The table was reproduced from Tosi
TABLE 13.2.
Frequency of Metal- Ligand Vibrations of Different Metalloproteins
Biological Molecule
MetalLigand
Wave Number (em-I)
Adrenodoxin Ferredoxin Hemerythrin
Fe-S Fe-S Fe-S Fe-O
279,350,397 345, 360 444 510
Nitrogenase Protocatechuate 3,4-dioxygenase Rubredoxin Ascorbate oxidase
Fe-S Fe-O
368 371,423,465
Fe-S Cu-S Cu-N Cu-N Cu-S Cu-N Cu-O
311, 365 260 380,415 350-400 375,427 480 282
N-Cu-N bending
119
Ceruloplasmin Copper- D- Penicillamine Hemocyanin
Alcohol dehydrogenase
Cu-N Cu-S Cu-N Cu-S Cu-N Cu-S Cu-N Cu-N Cu-O Zn-O
170-180 217-226 262-271 282-288 282 308-315 332-337 226,267 260 380,415 260 380,415 260 350-400 350 470 368
Acid phosphatase Selenocystine
Mn-S Se-Se
370 286-288
Cu-N
Cu-N Cu-N
Laccase Plastocyanin Stellacyanin Azurin
374
Reference Tang et al. (1975) Tanget al. (1975) Freier et al. (1979) Dunn et al. (1973, 1975) Freier et al. (1980) Levchenko et aI. (1980) Bull et aI. (1979) Long and Loehr (1970) Siiman et al. (1976) Siiman et aI. (1974) Siiman and Carey (1980) Siiman and Carey (1980) Loehr et aI. (1974) Freedman et al. (1976) Chen et al. (1979) Eickman et aI. (1978) Eickman et al. (1978) Eickman et al. (1978) Eickman et al. (1978) Eickman et aI. (1978) Larrabee et aI. (1977) Eickman et al. (1978) Eickman et al. (1978) Larrabee and Spiro (1980) Siiman et aI. (1976) Siiman et al. (1976) . Siiman et al. (1976) Siiman et aI. (1976) Siiman et aI. (1976) Siiman et al. (1974) Miskowski et al. (1975) Miskowski et al. (1975) Jagodzinski and Peticolas (1981 ) Sugiura et al. (1981) Lopez et al. (1981)
Copper in Biological Systems
375
and three nitrogen atoms, at least one of which is an amide nitrogen (Siiman et al., 1976). Rhus vermicifera laccase contains all three types of copper sites (types I, 2, and 3). The laccase without type 2 was prepared and called T2D (a type 2 depleted). The type 3 copper site in T2D laccase is reduced but can be oxidized by the addition of excess hydrogen peroxide. The oxidized T2D laccase gives a Raman spectrum similar to that of native laccase but not the reduced T2D. This experiment shows that the type 3 copper in T2D is in a reduced state (LuBien et al., 1981). Transferrin is an iron-containing protein (see Chapter II). Transferrin binds to Cu(II) to form a stable complex that shows an intense visible absorption band. When Cu(II)-transferrin is examined, the high-frequency region (6001700 cm -I) of the Raman spectrum has the same features as that of Fe(III)transferrin. The low-frequency region, by contrast, is more like that in the spectra of ceruloplasmin, stellacyanin, and laccase (Siiman et al., 1974). As to the binding site, it is concluded that Cu(II) binds to the phenol oxygen of tyrosine just as Fe(lII) chelates to the tyrosine residue in Fe(III)-transferrin (Tomimatsu et al., 1976; Gaber et al., 1974). The frequencies of various metal-ligand bond vibrations have been identified by many investigators. The Fe-ligand vibrations in heme compounds are summarized in Table 12.2, and Cu-N and Cu-S vibrational frequencies in ceruloplasmin and related compounds are summarized in Table 13.1. Frequencies of metal-ligand bond vibrations of many other compounds are summarized in Table 13.2. Azurins are a group of blue copper proteins found in bacteria. Azurin from Pseudomonas aeruginosa has a molecular weight of about 16,000. Copper(II) can be replaced with Ni(II), and both show resonance Raman spectra. Analysis of these spectra indicates both copper and nickel ions possessing cysteine and histidine as ligands (Ferris et al., 1979). 1.3.
Cu-PEPTIDE COMPLEX
The resonance Raman technique is useful in determining the site of the copper-peptide complex. For instance, Cu(II) forms a complex with poly(L-Tyr, L-Lys)n with an absorption maximum at 400 nm. Thus the resonance Raman spectrum can be obtained using the 457.9-nm Ar-ion laser line, which is still within the contour of the absorption band. There are two types of Cu(II)poly(L-Tyr, L-Lys) complexes. The first one occurs at pH 7.8, involving two amino nitrogens and two peptide nitrogens as ligands. The second complex is formed at alkaline pH and is coordinated with four peptide nitrogens (Tosi and Garnier, 1978; Garnier and Tosi, 1979). Normally, Cu(II) does not coordinate with the tyrosine phenolate oxygen atom, although it does coordinate with the peptide nitrogen. However, Cu(II) chelates to the phenolate oxygen atom in poly(Lys-Tyr) (Pastor et al., 1979). Glycylglycine complexes with Cu(II), but the site of attachment depends on pH. Using DC_, 15N_, and 2H-substituted dipeptides, the site of complexing
376
Copper and Other Metals in Biological Systems
and the exact assignment of some vibrational bands have been made (Takahashi et al., 1978). 2.
OTHER METALS
Many metal ions complex with a great variety of compounds. Usually they attach to specific molecular sites. Vibrational spectroscopy frequently can detect such complexing sites, since metal ions influence the vibrational modes of the functional groups to which they are attached. On some occasions, metal-ligand bonds can be detected and usually appear at relatively low frequency, because metal ions are greater in mass. Cyclic peptides are of great interest from a conformational viewpoint. Because of their cyclic nature, the conformation is less flexible than that of linear oligopeptides. The cyclo(L-Pro-GIY)3-expressed as C(PG)3-has interesting specific-cation-binding properties. It binds Li(I) and Na(I) selectively over K(I) and Rb(I). It also forms complexes of different stoichiometry with Mg(II), which binds selectively over Ba(II) and Ca(II). Upon cation complexation, the prolylcarbonyl stretching bands sharpen and upshift to 1690-1700 cm - I. The glycyl carbonyl stretching band is unaffected by Na(I), upshifted 15 cm - I by K(I), and downshifted to 1619 cm - I by Ca(III) complexation (Asher et al., 1980). Some peptides such as nonactin, monactin, and dinactin bi,nd to Na(l), K(I), Rb(I), Cs(l), Tl(l), and ammonium ions. When the peptides complex with a cation, the characteristic Raman spectra change, especially the ester carbonyl stretching frequencies (Asher et al., 1974, 1977). Histidine residues often serve as ligands in metalloenzymes such as carboxypeptidase, thermolysin, and carbonic anhydrase. Cobalt atom is frequently used as a substitute for the zinc atom in zinc enzymes. All zinc enzymes are colorless, but the substituted cobalt enzyme possesses a color that can be studied spectrophotometrically. For this reason, it is worthwhile to investigate the properties of the cobalt-imidazole complex. Unfortunately, the intensity of the resonance Raman bands of the Co(II)-;-imidazole complex are very weak. This is due to the absence of much involvement of the ligand orbitals in the visible transitions, which are largely forbidden d-d bands, and to the high energy of the ligand-cobalt charge-transfer states. Co(II)-carbonic anhydrase and Co(II)-carboxypeptidase show a very weak resonance Raman effect (Derry, 1974). Therefore, cobalt-substituted proteins have relatively limited use for resonance Raman study (Yoshida et al., 1975). Histidine forms a complex with Ni(III), Cu(II) or Zn(II) in a 2-to-l ratio, that is, (L-Hish-metal. Raman spectroscopic studies indicate that the site of coordination is the N-3 position (Itabashi and Itoh, 1980a, b). Ruthenium red inhibits calcium transport across mitochondrial membranes. It binds to various proteins, phospholipids, chelating agents, and mitochondria. Ruthenium red is used as a cytological reagent and has the structure [(NH3)5Ru-0-Ru(NH3)4-0-Ru(NH3)5]6+ . It seems that ruthenium red interacts with these biological materials in a rather specific fashion. Resonance
£!.
References
377
Raman spectra of free ruthenium red and ruthenium red complexed with Ca(II)-binding agents are quite different. Addition of Ca(II) can reverse these spectral changes. Moreover, Raman spectra of ruthenium phospholipids are different, enabling us to distinguish two classes of molecules (Friedman et aI., 1979). Ruthenium red is photosensitive; therefore, caution should be used in its use. Actually, photodegradation products can be detected from its resonance Raman spectrum (Itabashi et aI., 1981). Some enzymes such as acid phosphatase, pyruvate carboxylase, superoxide dismutase, and diamine oxidase, contain manganese as an integral part of the molecule. The resonance Raman spectrum of acid phosphatase (1230, 1298, 1508, and 1620 cm - I) shows bands very similar to the Fe(III)-transferrin and protocatechuate 3,4-dioxygenase tyrosine vibrational bands. It is concluded that Mn(III) also attaches to the tyrosine and cysteine residues of acid phosphatase (Sugiura et aI., 1980, 1981). Thiomolybdate, MoSJ- , plays and important biological role. It inhibits copper metabolism by the formation of a CuMoScprotein complex. Tetrathiomolybdate ions were detected in the hydrolysis products of nitrogenase, which is a Fe-Mo-protein complex. The Mo-S bond vibration is detected by Raman spectroscopy at 148, 158, 201, 429, 440, and 493 em-I. Copper or silver coordination to thiomolybdate influences the MoS bands by affecting Mo-S bond length (Muller et aI., 1981). Selenium belongs to the oxygen family and, although not a metal, is discussed in this chapter for convenience. As atomic weight increases from oxygen to sulfur, selenium, tellurium, and polonium, the metalloid property increases. More strictly speaking, selenium is a metalloid that is between metals and nonmetals. Selenium is a trace element and is present in a minute quantity in biological systems. It is known to be a part of the enzyme glutathione peroxidase (glutathione-H 20 2 oxidoreductase). As a first step toward the understanding of the selenium enzyme, Raman spectra of selenomethionine and selenocystine were obtained (Lopez et aI., 1981). Because of the relatively high atomic weight of selenium, the vibrations involving selenium appear at a low frequency. These are summarized in Table 13.2. Naturally occurring transferrin (see Chapter 11) contains iron, and Fe(III) can readily be replaced by Co(II) and Mn(III). Because of the similarity in the resonance Raman spectra to the natural Fe(III)-transferrin, both Co(III) and Mn(III) must occupy the same binding site as Fe(III) (Tomimatsu et aI., 1976).
REFERENCES Asher,1. M., Phillies, G. D. J., and Stanley, H. E. (1974). Nonactin and its alkali complexes-A Raman spectroscopic study. Biochem. Biophys. Res. Commun. 61, 1356. Asher,1. M., Phillies, G. D. J., Kim, B. J., and Stanley, H. E. (1977). Ion complexation in nonactin, monactin, and dinactin: A Raman spectroscopic study. Biopo/ymers 16, 157. Asher, I. M., Phillies, G. D. J., Geller, R. B., and Stanley, H. E. (1980). Cyclo(L-prolylglycyl)3 and its sodium, potassium, and calcium ion complexes: A Raman spectroscopic study. Biochemistry 19, 1805.
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Copper and Other Metals in Biological Systems
c., Ballou, D. P., and Salmeen, I. (1979). Raman spectrum of protocatechuate dioxygenase from Pseudomonas putida; new low frequency bands. Biochem. Biophys. Res. Commrtn. 87, 836. Caughey, W. S., Barlow, C. H., Maxwell, J. c., Volpe, J. A, and Wallace, W. 1. (1975). Reactions of oxygen with hemoglobin, cytochrome c oxidase and other hemeproteins. I. Hemeproteins: Ligation phenomena. Ann. N. Y. Acad. Sci. 244, 1. Chen, J. T., Shen, S. T., Chung, C. S., Chang, H., Wang, S. M., and Li, N. C. (1979). Achatina lulica hemocyanin and its interactions with imidazole, potassium cyanide, and fluoride as studied by spectrophotometry and nuclear magnetic resonance and resonance Raman spectroscopy. Biochemistry 18, 3097. Derry, R. E. (1974). Characterization of zinc containing metalloproteins by resonance Raman spectroscopy. Master's thesis, Portland State University, Portland, Ore. Dunn, J. B. R., Shriver, D. F., and Klotz, I. M. (1973). Resonance Raman studies of the electronic state of oxygen in hemerythrin. Proc. Nat. Acad. Sci. 79,2582. Dunn, J. B. R., Shriver, D. F., and Klotz, I. M. (1975). Resonance Raman studies of hemerythrinligand complexes. Biochemistry 14, 2689. Bull,
Eickman, N. c., Solomon, E. I., Larrabee, 1. A, Spiro, T. G., and Lerch, K. (1978). Ultraviolet resonance Raman study of oxytyrosinase. Comparison with oxyhemocyanins. J. Am. Chem. Soc. 100, 6529. Evans, J. C. (1969). The peroxide-ion fundamental frequency. Chem. Commun. 682. Ferris, N. S., Woodruff, W. H., Rorabacher, D. B., Jones, T. E., and Ochrymowycz, I.. A (1978). Resonance Raman spectra of copper-sulfur complexes and the blue copper protein question. J. Am. Chem. Soc. 100, 5939. Ferris, N. S., Woodruff, W. H., Tennent, D. L., and McMillin, D. R. (1979). Native azurin and its nickel(lI) derivative: A resonance Raman study. Biochem. Biophys. Res. Commun. 88,288. Freedman, T. B., Loehr, 1. S., and Loehr, T. M. (1976). A resonance Raman study of the copper protein, hemocyanin. New evidence for the structure of the oxygen-binding site. J. Am. Chem. Soc. 98, 2809. Freier, S. M., Duff, I.. 1.., Van Duyne, R. P., and Klotz, 1. M. (1979). Resonance Raman studies and structure of a sulfide complex of methemerythrin. Biochemistry 24, 5372. Freier, S. M., Duff, I.. 1.., Shriver, D. F., and Klotz, I. M. (1980). Resonance Raman spectroscopy of iron-oxygen vibrations in hemerythrin. Arch. Biochem. Biophys. 205, 449. Friedman, 1. M., Rousseau, D. 1.., Navon, G., Rosenfeld, S., Glynn, P., and Lyons, K. B. (1979). Ruthenium red as a resonance Raman probe of Ca2+ binding sites in biological materials. Arch. Biochem. Biophys. 193, 14. Gaber, B. P., Miskowski, V., and Spiro, T. G. (1974). Resonance Raman scattering from iron(III)and copper(II)-transferrin and an iron(III) model compound. A spectroscopic interpretation of the transferrin binding site. J. Am. Chem. Soc. 96, 6868. Garnier, A, and Tosi, I.. (1979). Cupric complexes of poly(L-lysine, L-tyrosine): Spectroscopic determination of structure in aqueous solution. J. lnorg. Biochem. 10, 147. Gielens, c., Maes, G., Zeegers-Huyskens, T., and Lontie, R. (1980). Raman resonance studies of functional fragments of helix pomatia .Bc-haemocyanin. J. lnorg. Biochem. 13, 41. Herve, M., Garnier, A, Tosi, 1.., and Steinbuch, M. (1981). Spectroscopic and photoreduction studies of copper chromophores in ceruloplasmin. Eur. J. Biochem. 116, 177. ltabashi, M., and ltoh, K. (1980a). Raman scattering study on coordination structures of Cu(Il)-L-histidine( 1: 2) in aqueous solutions. Bull. Chem. Soc. Japan 53, 3131. ltabashi, M., and ltoh, K. (I 980b). Tautomerism of imidazole group and its coordination site structure in metal complexes: Raman scattering study. In Proc. Vllth Int. Conf. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 610-611. ltabashi, M., Shoji, K., and ltoh, K. (1981). Reinvestigation of resonance Raman spectrum of Ruthenium red and its photodegradation. Chem. Lett. 491.
References
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Jagodzinski, P. W., and Peticolas, W. L. (1981). Resonance enhanced Raman identification of the zinc-oxygen bond in a horse liver alcohol dehydrogenase-nicotinamide adenine dinucleotidcaldehyde transient chemical intermediate. J. Am. Chern. Soc. 103,234. Larrabee, 1. A, and Spiro, T. G. (1980). Structural studies of the hemocyanin active site. 2. Resonance Raman spectroscopy. J. Am. ChenL Soc. 102,4217. Larrabee, 1. A, Spiro, T. G., Ferris, N. S., Woodruff, W. H., Maltese, W. A, and Kerr, M. S. (1977). Resonance Raman stndy of mollusc and arthropod hemocyanins using ultraviolet excitation: Copper environment and subunit inhomogeneity. J. A m. Chern. Soc. 99, 1979. Levchenko, L. A, Poschupkina, O. S., Sadkov, A P., Marakushev, S. A, Mikhailov, G. M., and Borod'ko, Yu. G. (1980). Spectroscopic investigation of FeMo-COFACTOR. Coenzyme A as one of the probable components of an active site of nitrogenase. Biochem. Biophys. Res. Commun. 96, 1384. Loehr, J. S., Freedman, T. B., and Loehr, T. M. (1974). Oxygen binding to hemocyanin: A resonance Raman spectroscopic study. Biochem. Biophys. Res. Commllll. 56, 510. Lontie, R. and Gielens, C. (1979). MIII/uscan and Arthropodan Haemocyanins. Metal/oproteins, U. Weser, Ed., Thieme, Stuttgart. Long, II, T. V., and Loehr, T. M. (1970). The possible determination of iron coordination in nonheme iron proteins using laser-Raman spectroscopy. Rubredoxin. J. Am. Chern. Soc. 92, 6384. Lopez, L., Jao, T. C., and Rudzinski, W. E. (1981). Tbe Raman spectra of selenomethionine and selenocystine. J. Inorg. Biochem. 14, 177. LuBien, C. D., Winkler, M. E., Thamann, T. J., Scott, R. A, Co, M. S., Hodgson, K. 0., and Solomon, E. 1. (1981). Chemical and spectroscopic properties of the binuclear copper active site in Rhus laccase: direct confirmation of a reduced binuclear type 3 copper site in type 2 depleted laccase and intramolecular coupling of the type 3 to the type I and type 2 copper sites. J. Am. Chem. Soc. 103, 7014. Miskowski, V., Tang, S. P. W., Spiro, T. G., Shapiro, E., and Moss, T. H. (1975). Copper coordination group in blue copper proteins. Evidence from resonance Raman spectra. Biochemistry 14, 1244. Mori, W., Suzuki, S., Kimura, M., Sugiura, Y., and Nakahara, A (1980). Characterization of the purple hemocyanin of Septioteuthis lessoniana. J. lnorg. Biochem. 13, 89. Miiller, A, Filgueira, R. R., Jaegermann, W., and Che, S. (1981). Resonance Raman spectroscopic identification of coordinating Mosl- in systems of bioinorganic interest. Natumissenschalten 68,93. Nakahara, A, Mori, W., and Suzuki, S. (1980). The active site of Septoteuthis lessoniana hemocyanin. Adv. Chern. Ser. 191, 341. Pastor, 1. M., Gamier, A, and Tosi, L. (1979). Absorption, circular dichroism and resonance Raman spectra of Cu(lI)-poly(L-glutamic, L-tyrosine) complexes. Evidence of phenolate coordination. Inorg. Chim. Acta 37, L549. Suman, 0., and Carey, P. R. (1980). Resonance Raman spectra of some ferric and cupric thiolate complexes. J. Inorg. Chem. 12, 353. Siiman, 0., Young, N. M., and Carey, P. R. (1974). Resonance Raman studies of "blue" copper proteins. J. Am. Chern. Soc. 96, 5583. Siiman, 0., Young, N. M., and Carey, P. R. (1976). Resonance Raman spectra of "blue" copper proteins and the nature of their copper sites. J. Am. Chem. Soc. 98, 744. Sugiura, Y, Kawabe, H., Tanaka, H. (1980). New manganese(III)-containing acid phosphatase. Evidence for an intense charge-transfer band and tyrosine phenolate coordination. J. Am. Chern. Soc. 102,6581. Sugiura, Y., Kawabe, H., Tanaka, H., Fujimoto, S., and Obara, A (1981). Purification, enzymic properties, and active site environment of a novel manganese(lIl)-containing acid phosphatase. J. Bioi. Chern. 256, 10664.
380
Copper and Other Metals In Biological Systems
Szymanski, T., Cape, T. W., Van Duyne, R. P., and Basolo, F. (1979). Determination of the resolution 0-0 stretching frequency of a monomeric dioxygen cobalt complex by resonance Raman spectroscopy. J. Chem. Soc., Chern. Commun. 5. Takahashi, S., Miyazawa, T, and Tasumi, M. (1978). Vibrational spectra of 13C_, 15N_ and D (C)-enriched glycylglycines and their Cu(II) complexes in aqueous solutions. Indian J. Pure Appl. Phys. 16,412. Tang, S.-P. W., Spiro, T G., Antanaitis, C, Moss, T B., Holm, R. H., Herskovitz, T, and Mortensen, L. E. (1975). Resonance Raman spectroscopic evidence for structural variation among bacterial ferredoxin, HiPIP, and Fe4S4(SCH2Ph)~- . Biochem. Biophys. Res. Commun. 62, I. Thamann, T 1., Loehr, J. S., and Loehr, T M. (1977). Resonance Raman study of oxyhemocyanin with unsymmetrically labeled oxygen. J. Am. Chem. Soc. 99,4187. Thompson,1. S., Marks, T 1., and (bers,1. A (1977). Blue copper proteins: synthesis, spectra, and structures of Cu I N 3(SR) and Cu II N 3(SR) active site analogues. Proc. Nat. Acad. Sci. USA 74, 3114. Tomimatsu, Y, Kint, S., and Scherer, 1. R. (1976). Resonance Raman spectra of iron(III), copper(II), cobalt(III) and manganese(III)-transferrin and of bis(2, 4, 6-trichlorophenolato)diimidazolecopper(II) monohydrate, a possible model of copper(II) binding to transferrins. Biochemistry 15, 4918. Tosi, L., and Gamier, A (1978). The formation and structure of a Cu(II)-poly(L-lysine, L-tyrosine) complex. Absorption and resonance spectral evidence of phenolate coordination. Inorg. Chim. Acta 29, L261. Tosi, L., and Garnier, A (1979). Circular dichroism and resonance Raman spectra of the Cu(II)-Cu(I) complex of D-penicillamine. The Cu(Cys) stretching mode in blue copper proteins. Biochem. Biophys. Res. Commun. 91, 1273. Tosi, L., Garnier, A, Herve, M., and Steinbuch, M. (1975). Ceruloplasmin-anion interaction-a resonance Raman spectroscopic study. Biochem. Biophys. Res. Commun. 65, 100. Yoshida, eM., Freedman, T. B., and Loehr, T. M. (1975). Resonance Raman study of cobalt(II)-irnidazole complexes as models for metalloproteins. J. Am. Chem. Soc. 97, 1028.
..J
CHAPTER
Al
-----mJr
Photosynthetic Pigments and Vitamin 8 12 Photosynthesis is a direct conversion of light energy into chemical energy by the chlorophyll-mediated synthesis of ATP in green plants, and by certain algae and bacteria. Chlorophyll is not the only compound involved in the energy-transfer process. It is well known that accessory pigments such as carotenoids and phycobilins are somehow involved in the photoreaction. They also serve as receptors of light energy. In photosynthetic organisms, photosynthetic reactions take place at specialized sites called reaction centers. How chlorophylls, carotenoids, quinones, and nonheme iron atoms within the reaction center are arranged and function is still unclear and is a matter of considerable interest. Using various excitation wavelengths, resonance Raman spectra of the pigments involved in photosynthesis can be obtained. The arrangement within the cell of these compounds is not random, and the
382
Photosynthetic Pigments and Vitamin 8 12
relative spatial arrangement within the cell is probably very crucial for photoenergy transfer. There have been intensive investigations in this field using resonance Raman spectroscopy. Dr. M. Lutz and his associates are contributing significantly in this area.
1.
PHOTOSYNTHETIC PIGMENTS
Chlorophylls a and b are green pigments that are structurally very similar. The difference is that one of the -CH 3 groups is replaced by a -CHO group in chlorophyll b. Electrons are delocalized over a large part of the macroring (Figure 14.1). 1.1.
RAMAN-BAND ASSIGNMENT
Chlorophylls show complex Raman bands from 200 to 1800 cm- '. The bands above 800 cm- I arise mainly from tetrapyrrolic-macrocyde vibrations. Aggregation of the chlorophylls induces characteristic spectral changes in the lowfrequency regions (100-700 cm- I ), and these low-frequency bands involve Mg-pyrrole-ligand vibrations.
C2 HS
H
H
H 3C CH2 H
I
CH2
I
C02C20H39
FIGURE 14.1. Structure of chlorophyll b. The structure of chlorophyll a is the same, except that it has a methyl group at position 3, where chlorophyll b has a formyl group.
TABLE 14.1. Resonance Raman frequencies d (cm- 1) observed in the carbonyl-stretching region for Chi a and for Chi b in chlorophyll-protein complexes at 30 K
Chloroplasts (mean values) ChI a O ChI bb 441.6 c 465.8 c 1616
1618 1630 1640
Tobacco CPU 465.8
441.6
1615sh 1634m I642sh
1615sh 1631m 1639wsh
1653 1661
CPI 441.6
Spinach CPI 441.6
Phormidium luridum CPI 441.6
Anabaena cylindrica CPI 441.6
Euglena gracilis CPI 441.6
1614m
1616m
1615m
1615m
1659w
1616m I630vwsh 1642vw 1653vwsh 1664w
I654vwsh I664wsh
1654vwsh 1664w
I654vwsh I662wsh
1656vwsh 1664vwsh
1670w 1679w 1688w
1673w 1680w 1691vwsh
1674w 1681wsh I690wsh
1674w 1680w
~ 1673w 1681w I690vwsh
1673wsh 1683w
~
~
1661w 1670 1681 1689 1694
I694w
1702
1705vwsh 1700vwsh
1705wsh
1705vwsh 1702wsh
1704vwsh Source: Table reproduced from Lutz et al. (1979). aFrom seven species. bFrom four species (Lutz 1975, 1977 and Lutz et al., 1979). c441.6, 465.8: excitation wavelength. d m: medium relative intensity, w: weak, v: very, sh: shoulder. Ca-C m are methine bridges.
1703vwsh
Assignments Phorbin, JI(Ca:":"':C m ) ChI b, JI(3-C=O) ChI b, JI(3-C=O) ChI a, JI(9-C=O) ChI a, JI(9-C=O) ChI b, JI(3-C=O) ChI a, JI(9-C=O) ChI, a, JI(9-C=O) ChI a, JI(9-C=O) ChI b, JI(9-C=O) ChI a, JI(9-C=O) ChI b, JI(3-C=O)
384
Photosynthetic Pigments and Vitamin 8 '2
Chlorophyll a contains two ester carbonyls and one keto carbonyl group; chlorophyll b contains one additional aldehyde carbonyl group. Although the 9-keto carbonyl of chlorophylls a and b and the 3-formyl carbonyl of chlorophyll b are shown in Raman spectra, no Raman activity is seen for ester carbonyl groups, because they lack conjugation with the phorbin-ring 1T-electron system. Within living cells, chlorophylls coexist ~ith proteins as. complexes (CP). In chloroplasts, or CP complexes, the C=O stretching vibrations appear in the 1620-1750-cm- I region, and most are downshifted from the free C=O stretching frequencies, but the carbonyl vibration has multiple bands and their analysis is not a simple matter. Resolution of carbonyl bands increases when the samples are cooled to 35 K. The 1630- and 1640-cm- I bands of the 35 K spectra are assigned to the aldehyde-carbonyl stretching mode of ChI b. The 1695-cm- 1 band is assigned to the free 9-ketone carbonyls of Chi b. For Chi a, the C(9)=O stretching modes appear in the 1650-1705 cm - I region (Lutz, 1975). Different carbonyl vibration bands and their assignments are shown in Table 14.1. Multiplicity of carbonyl bands in both chlorophylls a and b suggests that the group is probably involved in complex binding with proteins under natural conditions. In addition to the C(9)=O keto carbonyl group of chlorophyll a, the 1560-, 1585-, and 1617-cm- 1 bands originate from C:'::':C stretching modes of the parent ring without any noticeable participation from nitrogen motion (Lutz et aI., 1975). The low-frequency bands at 308-318 cm- I arise from the Mg-N bond in chlorophyll a, and are sensitive to the number of ligands bound to the magnesium ion (Lutz, 1974, 1975; Lutz et aI., 1975). For chlorophyll band CP IIa, the 312-cm- 1 band is assigned to Mg:..:..:.:N vibrations of the pentacoordinated-type binding to a single external ligand. The 304-cm- I component is for the hexacoordinated type. However, out-of-parent ring-plane Mg-axial-1igand bonds are unlikely to give rise to resonance-enhanced Raman bands in chlorophyll spectra. One piece of evidence is that the monomeric ch10rophyllacetone complex does not give a 31 O-cm- I band. In the complex, the carbony1s of two acetone molecules coordinate with the magnesium from each side of the phorbin plane (Lutz, 1974, 1975). There are additional low-frequency bands (100-300 cm- I ) for chlorophylls a and b and chloroplasts, but these bands have not been assigned (Lutz, 1972). There is an extensive review on the assignment of bands for free chlorophylls a and b and the pigments in chloroplasts (Lutz, 1977).
1.2.
PLANT PHOTOSYNTHETIC SYSTEM
The chloroplast is the site of photosynthesis in plants and photosynthetic eucaryotic organisms and contains large quantities of chlorophylls a and b. It also contains carotenoids. For procaryotes such as blue-green algae and the purple and green bacteria, the photoenergy-trapping process takes place in
Photosynthetic Pigments
385
chromatophores. Absorption peaks of these three pigments appear at different wavelengths; therefore, selective resonance-enhanced Raman bands can be obtained for each compound in its native environment by varying the excitation frequency. Below 450 nm the chlorophyll a Raman spectrum can be obtained; 450-475 nm is used for chlorophyll b, and wavelengths above 475 nm are used for the carotenoids (Lutz and Breton, 1973). P-700 is a pigment that undergoes bleaching when the cell is illuminated. It consists of chlorophyll a and a specific type of protein. P-700 constitutes only a small fraction (1 : 400 ratio) of the total chlorophyll in chloroplasts, and it may be considered a special form of chlorophyll a. There are two major complexes of CPo CP I is the P-700 ChI a-protein complex, whereas CP II is the CW alb-protein complex. These complexes probably represent the native states of chlorophyll. Raman spectra of chlorophyll in chlorophyll-protein complexes, intact cells, and cWoroplasts are nearly identical. This indicates that the protein-chlorophyll binding sites are probably the same in the intact membranes as in the CP complexes (Lutz et al., I978b, 1979; Pashchenko et al., 1980). What parts of chlorophyll molecules are involved in intermolecular associations is a matter of great importance. Several forms of chlorophyll a (ChI 66z , Ch1 668 , Ch1 675 , Ch1 651 , Ch1 686 , Ch1 693 ) are formed as a result of pigment-protein interaction of the C(9) = 0 keto group of the cWorophyll a molecule. On the other hand, other forms of chlorophylls such as Chl 700 and Chl 7lO are formed by molecular aggregation (Gadzhiev et al., 1981).
1.3.
BACTERIAL PHOTOSYNTHETIC SYSTEM
There are differences in the schemes of green plants, algae, and bacteria. Bacteria tend to use HzS, Na Z SZ0 3, succinate, acetate, and other organic compounds as hydrogen donors instead of HzO. Thus bacterial photosynthesis does not produce Oz. Instead of being located in the well-defined chloroplasts found in higher plants, bacterial cWorophyll is located in particles attached to lamellar extensions of the plasma membrane. In photosynthesis, transfer of excitation energy, separation of charge, and transfer of electrons take place very rapidly. These processes occur through ordered arrangements of photosynthetic molecules in membranes. An isolated photochemical reaction center can be prepared from bacterial chromophores. The isolation of such a photochemical reaction center greatly facilitates the study of the photoreaction mechanism of bacterial photosynthesis. The reaction center of Rhodopseudomonas spheroides contains four molecules of bacteriochlorophyll and two molecules of bacteriopheophytin (a derivative of chlorophyll) bound in an undetermined manner to a protein. By excitation with 600- and 529-nm laser light, resonance Raman spectra of bacteriochlorophyll or bacteriopheophytin can be selectively obtained. When the spectrum of the reaction center of R. spheroides is compared with those of isolated components, many Raman bands differ markedly in both frequency
386
Photosynthetic Pigments and Vitamin 8 12
and relative intensity. This suggests that these molecules interact with each other in the reaction center (Lutz and Kleo, 1976). Chlorophyll molecules must interact with the surrounding molecules in a specially oriented manner in the membrane. Even the aggregated chlorophyll molecules have Raman spectra different from that of the monomer, suggesting that aggregation takes place through interactions of the acetyl or keto carbonyl group of one molecule with the magnesium atom of another (Cotton and Van Duyne, 1981). In the first step of the photochemical reaction, an excited chlorophyll donates an electron to an acceptor to form a chlorophyll radical, normally cluster-expressed as ChI + '. Therefore, it is important to determine the chemical nature of the chlorophyll cation. When the spectrum of the bacteriochlorophyll a cation is compared with that of the parent molecule, significant differences in both band frequencies and intensities are found (Cotton and Van Duyne, 1978; Cotton et a1., 1980). Many Raman bands in the region of 1300-1700 cm-\ are shifted to lower frequencies. These are due to the delocalized C"-"-C stretching and bending motions of the pyrrolic ring. From this it can be said that the positive charge of the bacterial Chl+ cation becomes conjugated with part of the dihydrophorbin ring, resulting in a weakening of C=-C bonds, including the methine Ca=-Cm bridges, but C=-N and Mg-N bonds remain unaffected (Kleo and Lutz, 1978; Lutz and KIeo, 1979). One technique used to examine the photosynthetic system is to control the temperature-dependent intensity of Stokes Raman scattering by chlorophyll b and carotenoids. As the temperature is increased from 4.2 K, the shape and intensity of the spectral bands of pigments in green monocellular algae (Chlorella pyrenoidosa) change at 230 and 261 K. Finally, the spectrum becomes the spectral curve of living cells. Spectra obtained in this way represent a collective structural change of all pigments and other compounds (Drissler and MacFarlane, 1978; Drissler, 1980). 1.4.
CAROTENOIDS IN PHOTOSYNTHESIS
Carotenoids are discussed in Chapter 9, and in this section only their relationship to photosynthesis is discussed. Why carotenoids are present in the photosynthetic apparatus is not clear. Possible reasons may be that they are used in energy transfer within the photosynthetic apparatus, they protect cWorophylls from oxidation, or they are essential for structural integrity. Carotenoids can be found in chloroplasts together with chlorophylls a and b; the Raman spectra of carotenoids in the natural environment within chloro~ plasts can be obtained. Analysis indicates that four different molecular species of carotenoids are contributing to the spectrum (Lutz and Breton, 1973). There is evidence that carotenoids are also present as an integral part of the complex in the photochemical reaction center in certain photosynthetic purple bacteria (A thiorhodaceae). Resonance Raman spectra of these carotenoids are quite different from those of membrane-bound carotenoids (Lutz and Agalidis, 1978). Carotenoids found in the photochemical reaction center of photosynthetic bacteria are spheroidenes, and most spheroidenes in nature possess an
Vitamin 8 12
387
all-trans conformation. Unusual Raman spectra of the spheroidene present in the preparation of reaction centers is interpreted to mean that this compound is present in a di-cis conformation. Cis-spheroidenes are unstable and isomerize to the all-trans forms upon extraction from the reaction center. If the extraction is done rapidly in subdued light, followed by immediate freezing, the Raman spectrum shows features of the native form. This demonstrates that spheroidenes are in a cis conformation in the native photochemical reaction centers (Lutz et al., 1978a, b; Agalidis et al., 1980). Actually the reaction center of bacterial photosynthesis is quite complicated and contains not only chlorophylls and carotenoids but also bacteriopheophytins, phycocyanin, quinones, and nonheme iron. Using different excitation wavelengths, respective resonance Raman spectra can be selectively produced (Kubota et al., 1981). Raman spectra of various reaction-center-bound chromophores can be obtained from preparations of R. spheroides and other Athiorhodaceae. For instance, resonance Raman spectra of carotenoids, bacteriopheophyrins, and bacteriochlorophylls can be obtained by using the excitation wavelengths of 441.6, 528-535 and 545-550, and 580-610 nm, respectively. How these compounds are arranged in the photochemical reaction center of bacteria is a matter of great importance. Unfortunately, we still don't know the details, except that these compounds are bound to polypeptides at specific sites (Lutz, 1980a, b). Phycocyanin is an auxiliary pigment of the blue-green algae and transfers the absorbed energy via allophycocyanin to the photosystem II reaction center. The resonance Raman band of the C=C stretching vibration (1525 cm -I) of carotenoids in intact Anacystis nidulans is very sensitive to 625-nm light. Carotenoids do not absorb at this wavelength; therefore, the change in the resonance Raman spectrum cannot be due to the carotenoids themselves. The change is explained as being due to a specific phycocyanin-carotenoid interaction in A. nidulans, and the interaction probably involves the 'IT-electron system of both compounds (Szalontai and Van de Ven, 1981). Excited triple states of chlorophylls and polyenes have been recognized in the photosynthetic system, although nothing is known about the role of these triple states in photosynthesis. For the first step toward understanding this problem, resonance Raman spectroscopy was used to study the excited triple states of chlorophyll a, J3-carotene, and canthaxanthin. The results suggest that the relaxed triple conformations of all-trans-J3-carotene and canthaxanthin are different (Jensen and Wilbrandt, 1980). The Raman spectra of all-trans-J3-carotene indicate that the C=C bond order is decreased, and that the molecule may be substantially twisted probably at the C(15)=C(15') bond in the triple state (Jensen, 1980; Jensen and Wilbrandt, 1980).
2.
VITAMIN 8 12
Vitamin B12 , also called cyanocobalamin, was first isolated in 1948 and is essential in the human diet (Figure 14.2). Vitamin B12 has strong absorption bands in the near uv (340-390 om) and the visible regions (440-540 nm),
388
Photosynthetic Pigments and Vitamin B 12
C=N
Vitamin B12
FIGURE 14.2.
Structure of vitamin B I2 (cyanocobalamin).
which originate from 'TT-'TT* electronic transitions. Resonance Raman spectra of cyanocobalamin can be obtained by laser excitation coincident in wavelength with the visible absorption maxima. Dicyanocobinamide, which ::8 considerably different chemically, gives essentially identical spectra (Mayer et al., 1973a). This is reasonable, since the resonance Raman spectra are due to the resonance-enhanced vibrations associated with the common corrin ring system (George and Mendelsohn, 1973). However, an alteration in the corrin-ring chromophor induces large changes in the Raman spectra (Mayer et al., 1973b). Another possible reason for different spectra by changing the corrin-ring structure is the nonplanar nature of the corrinoid nucleus. The corrinoid ring occurs in several different forms depending on the nature of the metal and substituents. Conformational differences are largely responsible for different resonance spectra of aquocobalamin (vitamin B1u ) and cobalamin-Co(l) (vitamin B I2s )' No bands can be assigned to the vibration of the sixth ligand such as Co-C or C :::::: N. Even the in-plane Co-N stretching mode is not observed in the cobalamin spectra (Wozniak and Spiro, 1973). When a ring-stretching vibration at the 1504 cm - 1 scattering line is used, the concentration of vitamin B12 can be measured at values as low as 10- 6 M, with results reproducible within 2%. The intensity of this line is pH dependent. The change at pH 3 is due to deprotonation of benzimidazole and formation of the cobalt-ligand bond. The inflection at pH 8 corresponds to deprotonation of the coordinated water molecule (Tsai and Morris, 1975). The relative intensity of Raman bands depends on the excitation wavelength. The 1500-cm - I band of cobalt corrinoids is enhanced by visible-light excitation, whereas the 1550-cm -1 band is enhanced by UV-light excitation. This is because the electronic-transition mechanisms differ for visible light and
References 3
5
389
7
8
10
12
18 17
15
13
FIGURE 14.3. The nucleus of the cobalt corrinoid is composed of a 'IT-electron system.
UV absorption (Mayer et aI., 1973c; Salama and Spiro, 1977). As can be seen in Figure 14.3, the corrin 'IT-system is not symmetrical. Vibration along one axis (X) and along another axis (Y) should give different types of Raman spectra; these different vibrational modes may eventually appear as different Raman band patterns when irradiated with laser light of different wavelengths.
REFERENCES Agalidis, 1., Lutz, M., and Reiss-Husson, F. (1980). Binding of carotenoids on reaction centers from Rhodopseudomonas sphaeroides R 26. Biochim. Biophys. Acta 589, 264. Cotton, T. M., and Van Duyne, R. P. (1978). Resonance Raman spectroelectrochemistry of bacteriochlorophyll and bacteriochlorophyll cation radical. Biochem. Biophys. Res. Commun. 82,424. Cotton, T. M., and Van Duyne, R. P. (1981). Characterization of bacteriochlorophyll interactions in vitro by resonance Raman spectroscopy. J. Am. Chem. Soc. 103, 6020. Cotton, T. M., Parks, K. D., and Van Duyne, R. P. (1980). Resonance Raman spectra of bacteriochlorophyll and its electrogenerated cation radical. Excitation of the Soret bands by use of stimulated Raman scattering from Hz and D z . J. Am. Chem. Soc. 102,6399. Drissler, F. (1980). Discovery of phase transitions in photosynthetic systems. Phys. Lett. 77A, 207. Drissler, F., and MacFarlane, R. M. (1978). Enhanced anti-Stokes Raman scattering from living cells of Chlorella pyrenoidosa. Phys. Lett. 69A, 65. Gadzhiev, Z. I., Godzhaev, N. M., Gorokhov, V. v., Churin, A. A., Pashchenko, V. Z., and Rubin, L. B. (1981). Study of certain features of intermolecular interactions of chlorophyll a in vivo by resonance Raman spectroscopy. Dokl. Akad. Nauk. SSSR 261, 497. George, W.O., and Mendelsohn, R. (1973). Resonance Raman spectrum of cyanocobalamin (vitamin BIZ)' Appl. Spectrosc. 27, 390. Jensen, N. H. (1980). Photosynthetic Pigments and Model Compounds Studied by Pulse Radiolysis, Risj
390
Photosynthetic Pigments and Vitamin B 12
Kubota, K., Kuroda, S., and Koyama, Y. (1981). Resonance inverse Raman spectra of p-carotene and carotenoid in photosynthetic bacteria. Biopolymers 20, 2701. Lutz, M. (1972). Spectroscopic Raman de resonance de pigments vegetaux en solution et indus dans des lamelles chloroplastiques. C. R. Acad. Sc. 275, B497. Lutz, M. (1974). Resonance Raman spectra of chlorophyll in solution. J. Ramall Spectrosc. 2, 497. Lutz, M. (1975). Resonance Raman spectroscopy of the chlorophylls in photosynthetic structures at low temperature. In Lasers ill Chemistry alld Biophysics, Elsevier Scientific, Amsterdam, pp. 451-463. Lutz, M. (1977). Antenna chlorophyll in photosynthetic membranes, a study by resonance Raman spectroscopy. Biochim. Biophys. Acta 460,408. Lutz, M. (1980a). Resonance Raman studies of the bacterial photosynthetic reaction center. In Proc. VIlth 1111. Conf. Raman Speclrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 520-23. Lutz, M. (1980b). Bonding interactions on pigments in bacterial antenna and reaction centers. In Proc. 5th Int. Congo PhotosYII., Athens, in press. Lutz, M., and Agalidis, I. (1978). Resonance Raman scattering of cis conformers of C-40 carotenoids. Evidence for cis-carotenoids in bacterial reaction centers. In Proc. Sixth Int. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, Philadelphia, Rheine, pp. 162-163. Lutz, M., and Breton, 1. (1973). Chlorophyll association in the chloroplast: Resonance Raman spectroscopy. Biochem. Biophys. Res. Commun. 53, 413. Lutz, M., and Kleo, 1. (1976). Resonance Raman scattering of bacteriochlorophyll, bacteriopheophytin and spheroidene in reaction centers of Rhodopseudomonas spheroides. Biochem. Biophys. Res. Commun. 69, 711. . Lutz, M., and Kleo, 1. (1979). Bacteriochlorophyll a cation radical in solution and in reaction centers of Rhodopseudomollas sphaeroides. Resonance Raman scattering. Biochim. Biophys. Acta 546, 365. Lutz, M., Kleo, 1., Gilet, R., Henry, M., Plus, R., and Leickman, J. P. (1975). Vibrational spectra of chlorophylls a and b labelled with 26Mg and 15N. In Proc. 2nd Int. Conf. Stable Isotopes, E. R. Klein and P. D. Klein, Eds. U.S. Dept. Commerce, Springfield, Va., pp. 462-469. Lutz, M., Agalidis, I., Hervo, G., Cogdell, R. J., and Reiss-Husson, F. (1978a). On the state of carotenoids bound to reaction centers of photosynthetic bacteria: A resonance Raman study. Biochim. Biophys. Acta 503, 287. Lutz, M., Brown, J. S., and Remy, R. (1978b). Resonance Raman scattering of chlorophyll-protein complexes. In Proc. Sixth Int. Conf. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter, Eds., Heyden, London, pp. 158-~9. Lutz, M., Brown, J. S., and Remy, R. (1979). Resonance Raman spectroscopy of chlorophyll-protein complexes. In Chlorophyll Organization and Energy Transfer in Photosynthesis, Ciba Foundation Symposium 61, Exerpta Medica. Elsevier North-Holland, pp. 105-125. Mayer, E., Gardiner, D. J., and Hester, R. E. (I 973a). Resonance Raman spectra of vitamin B I2 and dicyanocobalamin. Biochim. Biophys. Acta 297, 568. Mayer, E., Gardiner, D. J., and Hester, R. E. (1973b). Resonance Raman spectra of vitamin B I2 and some cobalt corrinoid derivatives. Chern. Soc. London, J. Faraday Trans. Il, 69, 1350. Mayer, E., Gardiner, D. J., and Hester, R. E. (1973c). The selective enhancement of band intensities in resonance Raman spectra of cobalt corrinoids. Mol. Phys. 26, 783. Pashchenko, V. Z., Gadzhiev, Z. I., Churin, A A, and Rubin, L. B. (1980). Investigation of the association of chlorophyll a by the spectroscopic method of combined resonance. Rep. USSR A cad. Sci. 251, 995. Rimai, L., Kilponen, R. G., and Gill, D. (1970). Resonance-enhanced Raman spectra of visual pigments in intact bovine retinas at low temperatures. Biochem. Biophys. Res. Commun. 41, 492.
References
391
Salama, S., and Spiro, T. G. (1977). Visible and near-ultraviolet resonance Raman spectra of photolabile vitamin B I2 derivatives with a rapid-flow technique. J. Raman Spectrosc. 6, 57. Szalontai, B., and Van de Ven, M. (1981). Raman spectroscopic evidence for phycocyanin-carotenoid interaction in Anacystis nidulans. FEBS Lett. 131, ISS. Tsai, c.-W., and Morris, M. D. (1975). Application of resonance Raman spectrometry to the determination of vitamin B 12 . Anal. Chim. Acta 76, 193. Wozniak, W. T., and Spiro, T. G. (1973). Resonance Raman spectra of vitamin B I2 derivatives. J. Am. Chem. Soc. 95, 3402.
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Eyes, Teeth, and Muscles
As the application of laser Raman spectroscopy to biological systems is relatively new, not many tissues have been examined yet. This does not mean that this technique is not applicable. On the contrary, it will probably become very commonly used in biology and medicine in the future as biologists learn more about Raman spectroscopy. In this chapter, four tissues (lens, cornea, teeth, and muscle) are discussed. Mammary tissues, both normal and carcinoma, are discussed in Chapter 18.
1.
OCULAR LENSES
The lens transmits and refracts visible light which eventually converges on the retina (Figure 15.1). The image is formed on the retina, where the photosensitive rods and cones convert the light energy into nerve impulses.
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396
Eyes, Teeth, and Muscles
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LENS FIGURE 15.1.
Diagrams showing an eye and a lens.
The lens is composed of a mixture of proteins. The lens does not contain blood vessels or nerves. Lens proteins are roughly grouped into water soluble proteins (a-, {3-, and y-crystallin) and insoluble albuminoid. A healthy lens, being transparent to visible light, possesses a unique structure of proteins and' water. If this structure is altered in some manner, opacities or cataracts can appear that eventually can impair vision and cause blindness. The transparency of the human lens and cataractogenesis is not well understood. Recently, laser Raman spectroscopy has been applied to studying lens proteins from both normal and cataractous lenses. Raman spectroscopy is a noninvasive, sensitive, and selective probe of lens structures at the molecular level.
Ocular Lenses
1.1.
397
PEPTIDE BACKBONE
Bovine and rabbit lens proteins have predominantly antiparallel f3-structure (Yu et al., 1974; Schachar and Solin, 1975; Thomas and Schepler, 1980a, b). On the other hand, a-crystallin is the major protein in bird and reptile lenses. Conformational analysis by Raman spectroscopy indicates that a-crystallin is chiefly in the a-helical conformation. Heat treatment of a-crystallin converts it to a coagulated opaque protein and the conformation changes from a- to f3-structure (Yu et al., 1977). Large-scale spectral changes occur in the lens proteins with TCA (trichloroacetic acid) induced opacity (McKenzie, 1979). Apparently some types of lens opacification can be correlated to conformational changes in lens proteins.
1.2.
SULFHYDRYL GROUP AND DISULFIDE BOND
Besides the conformational differences in lens proteins, bovine and bird lens proteins also differ in their sulfhydryl-group content. Bovine lens is rich in sulfhydryl groups, as can be seen from the peak at 2582 em- I, but such a peak is absent in chick lens (Kuck et al., 1976). The sulfhydryl groups are derived
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FIGURE 15.2. Variation of sulfhydryl-group content (monitored at 2582 em-I) along the visual axis of a' six month mouse lens. The intensity at 2731 em - I is used as an indicator of protein concentration. The figure was reproduced from Askren et al. (1979) by permission of the copyright owner, Academic Press.
398
Eyes, Teeth, and Muscles
from cysteine residues as well as from the reduced form of glutathione present in lens. There is a variation in sulfhydryl-group content (protein -SH plus glutathione -SH) along the visual axis of the rat and mouse lenses (Figure 15.2). There are two maxima in the cortex, one anterior and one posterior, and a central minimum. The sulfhydryl content, as measured by the 2582-cm- I band, decreases with age. These results are interpreted in terms of a 2SH ---> S -S conversion, which occurs as agin-g proceeds (Askren et al., 1979; East et al., 1978). A decrease in the -SH content of the lens accompanies an increase in the S-S content on aging, but the decrease in concentration of the former is not equivalent to the increase in the latter. This suggests that IX and f3 crystallins, which are low in -SH, have faster synthesis rates than y-crystallin, which is high in -SH. Continuous irradiation of mice in vivo with long-wave uv light produces decreases in both -SH and S-S concentrations in the lens nucleus and cortex (East et al., 1978). Yu and East (1975) showed that the sulfhydryl groups of the bovine lens proteins are unaffected by heat denaturation at 100°C. This suggests that the opacification of a lens does not necessarily involve the oxidation of sulfhydryl groups. The incidence of cataracts increases as aging proceeds. Yet studies of -SH and S-S content of lens with aging and opacification by heating give different results. Obviously, more investigation is needed in this area. 1.3.
OTHERS
Direct application of laser Raman spectroscopy to human eyes has a great potential for monitoring the development of cataracts. Since the retina is very sensitive to laser damage, conventional Raman techniques are not suitable. Recently a sensitive multichannel optical detector, such as a silicon intensified target coupled to an optical multichannel analyzer, has been used to monitor intact lens with very low laser power (Mathies and Yu, 1978). This new technique may eventually be used to detect the opacity of the lens directly or precataractous condition. Comparisons of the Raman spectra of cataractous and normal rabbit lens in 3280-3400 cm-\ region, which is the H 20 stretching vibration mode, have been made. The increase in the H 20 vibrational intensity is explained by the increase in water content in the cataractous area. This is probably due to thermal damage to the lens protein-water network caused by laser light and the increase in water content at the injury site. The original hydrogen bonding system is probably disrupted; this eventually causes cataract formation in the lens (Thomas and Schepler, 1980a, b). 2.
CORNEA
The cornea is the transparent tissue separating the forward interior of the eye from the exterior; it is composed almost entirely of collagen. Collagen consists
Teeth
399
of three fibrils (a-chains) twisted about a common axis, resulting in a triplestranded coil. Individual collagen fibrils are inefficient light transmitters, but when they are arranged in lamellar fashion, the whole cornea becomes transparent. Corneal collagen also absorbs water, causing the cornea to become opaque. Corneal collagen fibrils do not dissociate appreciably upon heating at 70°C, as there are no changes in Raman spectra before and after heat treatment (Goheen et aI., 1978). The swelling induced by heating and aging is apparently caused by water's being absorbed and remaining between collagen fibrils, rather than drastic changes in secondary structure.
3.
TEETH
Teeth are very rich in calcium, magnesium, fluoride, and phosphorus. The outermost layer of the teeth is enamel, the hardest substance that can be found in the human body (Figure 15.3). Underneath the enamel is dentin, which has a composition similar to that of bone. Raman spectra of teeth (enamel and dentin) show bands at 432, 525, 595, 924, 1006, 1045, and 1072 cm - I (Figure 15.4). Four of them arise from pol- vibration modes, as teeth contain hydroxylapatite, Ca lO(P04 MOHh. The bands at 432, 595, 924, and 1045 cm - I correspond to P2' P4' P P and P3 vibrational bands of hydroxylapatite (Figure 15.5). Raman spectra have even been obtained from 15,000-year-old
FIGURE 15.3. Cut face of modem human tooth. E, enamel; D, dentin; dp, in living tissue, this space contains dental pulp, which includes blood vessels, nerves, lymphatics, and others. The figure was kindly supplied by Dr. Masa-Oki Yamada.
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teeth which have been embedded in marble stone (Yamada et aI., 1979). The most prominent difference between ancient teeth and modern teeth is that the relative fluorescence is twice as high in ancient teeth. Shark teeth show different Raman spectra from those of human teeth (Nishigori et aI., 1979). It seems that Raman spectroscopy can be used in paleontological investigation as well as to analyze differences between teeth from different animals.
4.
MUSCLE
Muscle consists of many different compounds such as myosin, actin, tropomyosin, troponin, ATP, and so forth. Application of Raman spectroscopy to muscle is still in the infant stage, but it is worthwhile reviewing the use of this unique tool. It is useful to review the structure of muscle briefly before Raman spectroscopic studies of muscle and its components are discussed. The morphology of skeletal muscle is quite different from that of other types of cells. Skeletal muscle cells are extremely elongated, with several nuclei usually found at the periphery of the cell. Repeating patterns are found throughout muscle (Figure 15.6). A sarcomere extends from one Z line to another. The elongated b!Jndles of fibers are composed of myosin and actin filaments. The thicker filaments are made up of myosin with a diameter of about 10 nm, and the thinner filaments are composed of actin with a diameter of 6 nm. The difference in the size between myosin and actin can best be seen in a cross section of a muscle bundle (Figure 15.7 and 8). The length of the myosin
FIGURE 15.6. Ultrastructure of normal muscle-longitudinal view. Z, Z line; H, H band; M, mitochondria. The electron micrograph was made by the author.
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bundles corresponds to the length of the A band (Figure 15.7). When contraction takes place, the actin filaments slide toward the M band, and the light I band disappears. 4.1.
MYOSIN
Myosin is the major component of muscle and accounts for half of the myofibrillar protein. Myosin has a large size, with a molecular weight of
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FIGURE 15.8. Ultrastructure of normal muscle-cross sectional view. The large dot is myosin; the small dot, actin (electron micrograph by M. Stringer of the author's laboratory).
480,000. The amide I band appears at 1650 cm- I , which indicates the presence of a-helical conformation. The amide III band also shows that the protein has typical a-helical conformation, and appears at 1265 and 1304 cm -I. Myosin also contains nonhelical structure, as indicated by the band at 1244 cm- I . The 1244-cm - I band represents either ,a-structure, random coil, or a mixture of both (Carew et al., 1975). The conformation of myosin can be influenced by some cations, and the transition can be detected using amide I and III bands, and bands at 900 and 940 cm- I • For instance, CaCl 2 transforms myosin from a-to ,a-structure, and LiBr denatures the protein, thus increasing the random-coil structure (Barrett et al., 1978). 4.2.
MYOSIN SUBFRAGMENTS
Light meromyosin (LMM) and subfragment I, S-I, are characteristic of the helical-tail and globular-head portions of the myosin molecule. The LMM consists of two a-helical chains wound into a rodlike shape, somewhat similar to a supercoil. The S-I contains about 35% ,a-structure, as measured by optical rotary dispersion. Raman spectra of LMM and S-I seem to reflect these structural differences in the amide III region. The LMM lacks bands in the
404
Eyes, Teeth, and Muscles
l244-l265-cm - 1 region, whereas the S-l has a band in the l240-l280-cm - 1 region. The LMM also has a band near 1306 cm- I , whereas the S-l does not. The l306-cm -1 band probably represents the coiled helices of the fibrous tail portion, whereas the 1265-cm- I band is associated with a-helical portions of the globular heads (Asher et a1., 1976). 4.3.
TROPOMYOSIN AND TROPONIN
Both tropomyosin and troponin, components of skeletal muscle, regulate the interaction of actin and myosin. Tropomyosin has the amide I band at 1655 cm -1, the amide III band at 1254 cm -1, and the band at 940 cm - 1. Judging from these bands, tropomyosin has mainly a-helical structure. The a-content, however, decreases rapidly as the pH is raised above 9.5 (Frushour and Koenig, 1974). Nadeau et a1. (1982) showed that the amide I band is centered at 1646 cm -1. Troponin C is the Ca(II)-binding subunit of troponin. Addition of Ca(II) ion to troponin C produces perturbation in the amide III region, showing evidence for increased a-helical content (Carew et a1., 1980). 4.4.
WHOLE MUSCLE
As can be seen from Figure 15.9, the Raman spectrum of whole muscle is strikingly similar to that of myosin. This is reasonable, as myosin is the major component and accounts for approximately 50% of muscle proteins (Asher et a1., 1976). An examination of the Raman spectrum of intact single muscle fibers indicates that some proteins are bound to intracellular-membrane fi-carotene. Carotenoid pigments are known to give strong resonance Raman bands at 1530 and 1160 cm- I that originate from -C=C- and =C-C= stretching vibrations of the conjugated polyene chains. Indeed, strong bands at 1521 and 1156 em- 1 are clearly visible in the Raman spectrum' of a single muscle fiber (Pezolet et a1., 1978a, b). The amide I at 1648 em - I and a strong skeletal C-C stretching band at 939 em-I also indicate that muscle fiber contains large amounts of a-helix proteins. However, a-helix content decreases when the myofibrils are isolated. Apparently the state of myofibrils in situ is different from that of myofibrils outside intact muscle tissue (Pezolet et al., 1980b). When the O-H stretching vibration bands (3100-3700 em-I) of an intact single muscle fiber are examined, there is no appreciable difference between the shape and relative intensity of the bands due to the water molecules located inside the muscle fiber and the shape and intensity of the corresponding bands in the spectrum of pure water. This suggests that there is little specially structured intracellular water. Since ice and liquid water have different Raman spectra in the OH vibrational region, it is possible to examine frozen and unfrozen water in a frozen intact muscle cell. By this method it is found that about 20% of the water molecules remain in the supercooled state at - 5°C.
References
405
-f:j.v(cm) FIGURE 15.9. Raman spectra of myosin (a, b) and whole muscle (c). The figure was reproduced [rom Asher et al. (1976) by permission of copyright owner, Raven Press.
which corresponds to I g of water per gram of fiber dry weight. This amount of water is probably truly the "bound water" (Pezolet et al., I978a, b). This value is somewhat larger than that found with the calorimetric method (Aubin et al., 1980). These studies suggest that a constant amount of water is bound to the protein molecules, and that this amount does not vary with water content of the muscle. 4.5.
CONTRACTILE STATE AND PROTEIN STRUCTURE
Because good-quality Raman spectra can be obtained from a single intact muscle fiber, the Raman spectroscopic technique can be used to correlate the protein structure to the state of muscle contraction. This was beautifully done by Pezolet et al. (1980a). A predominantly a-helical structure of muscle proteins remained even when the muscle was contracted or relaxed in the presence of ATP and Ca(II). However, the contraction induced a decrease in the scattering intensity of some of the Raman bands that are due to the acidic and tryptophan residues, showing that these amino acids are involved in the generation of tension.
REFERENCES Asher, l. M., Carew, E. B., and Stanley, H. E. (1976). Laser Raman spectroscopy: A new probe of the molecular conformations of intact muscle and its components. In Physiology of Smooth Muscle, E. Bulbring and M. F. Shuba, Eds., Raven. New York. DO. 229-238.
406
Eyes, Teeth, and Muscles
Askren, C. c., Yu, N.-T., and Kuck, Jr., 1. F. R. (1979). Variation of the concentration of sulfhydryl along tbe visual axis of aging lenses by laser Raman optical dissection technique. Exp. Eye Res. 29, 647. Aubin, M., Prud'Homme, R. E., Pezolet, M., and Caille, J.-P. (1980). Calorimetric study of water in muscle tissue. Biochim. Biophys. Acta 631, 90. Barrett, T. W., Peticolas, W. L., and Robson, R. M. (1978). Laser Raman light-scattering observations of conformational changes in myosin induced by inorganic salts. Biophys. J. 23, 349. Carew, E. B., Asher, 1. M., and Stanley, H. E. (1975). Laser Raman spectroscopy-new probe of myosin substructure. Science 188, 933. Carew, E. B., Leavis, P. c., Stanley, H. E., and Gergely, J. (1980). A laser Raman spectroscopic study of Ca2+ binding to troponin c. Biophys. J. 30, 351. East, E. 1., Cbang, R. C. c., and Yu, N.-T. (1978). Raman spectroscopic measurement of total sulfhydryl in intact lens as affected by aging and ultraviolet irradiation. J. BioI. Chern. 253, 1436. Frushour, B. G., and Koenig, 1. L. (1974). Raman spectroscopic study of tropomyosin denaturation. Biopolymers 13, 1809. Goheen, S. c., Lis, L. J., and Kauffman, 1. W. (1978). Raman spectroscopy of intact feline corneal collagen. Biochim. Biophys. Acta 536, 197. Kuck, Jr., 1. F. R., East, E. 1., and Yu, N.-T. (1976). Prevalence of a-helical form in avian lens proteins. Exp. Eye Res. 23, 9. Mathies, R., and Yu, N.-T. (1978). Raman spectroscopy with intensified vidicon detectors: A study of intact bovine lens proteins. J. Raman Spectrosc. 7, 349. McKenzie, C. R. (1979). Raman and infrared studies of biologically relevant systems: The ocular lens and glutathione. Diss. Abstr. Int. B. 39, 4707. Nadeau, J., Pezolet, M., Williams, D. L., Jr., and Swenson, C. A. (1982). Personal communication. Nishigori, K, Yamada, M.-O., Fujimori, K, Chikamori, K, and Yamashita, S. (1979). Comparative studies on fish tooth tissues by micronuorometry and Raman-spectroscopy. In Acta Histochern. Cytochem., 20th meeting, JSHC, Kyoto, Part I, p. 599. Pezolet, M., Pigeon-Gosselin, M., and Caille, 1.-P. (1978a). Laser Raman investigation of intact single fibers protein conformations. Biochim. Biophys. Acta 533, 263. Pezolet, M., Pigeon-Gosselin, M., Savoie, R., and Caille, 1.-P. (1978b). Laser Raman investigation of intact single muscle fibers on the state of water in muscle tissue. Biochirn. Biophys. Acta 544,394. Pezolet, M., Pigeon-Gosselin, M., Nadeau, J., and Caille, 1.-P. (1980a). A molecular probe of tbe contractile state of intact single muscle fibers. Biophys. J. 31, I. • Pezolet, M., Pigeon-Gosselin, M., Nadeau, J., and Caille, 1.-P. (1980b). Laser Raman scattering as a probe of muscle structure. In Proc. Vllth Int. Conf. Raman Spectrosc., W. F. Murphy, Ed., North-Holland, Amsterdam and New York, pp. 600-603. Scbachar, R. A., and Solin, S. A. (1975). The microscopic protein structure of the lens with a theory for cataract formation as determined by Raman spectroscopy of intact bovine lens. Invest. Ophthalmol. Vis. Sci. 14, 380. Thomas, D. M., and Schepler, K. L. (1980a). Raman spectra of normal and ultraviolet-induced cataractous rabbit lens. Invest. Ophthalmol. Vis. Sci. 19, 904. Thomas, D. M., and Schepler, K. L. (1980b). Laser Raman spectroscopy of ultraviolet-induced cataracts in rabbits and monkeys. Gov. Rep. Announce. Index (U.S.) 80, 4924. Also U.S. Air Force Report SAM-TR79-40, p. 1. Yamada, M.-O., Horibe, H., Fujimori, K, Yamashita, S., and Yamashita, T. (1979). Paleobiophysical studies on tooth tissues by micronuorometry and Raman-spectrometry. Cel/. Molec. Bioi. 25, 167.
~
References
407
Yu, N.-T., and East, E. 1. (1975). Laser Raman spectroscopic studies of ocular lens and its isolated protein fractions. J. Bioi. Chem., 250, 2196. Yu, N.-T., Jo, B. H., Chang, R. C. c., and Huber, 1. D. (1974). Single-crystal Raman spectra of native insulin structures of insulin fibrils, glucagon fibrils, and intact calf lens. Arch. Biochem. Biophys. 160,614. Yu, N.-T., East, E. J., and Chang, R. C. C. (1977). Raman spectra of bird and reptile lens proteins. Exp. Eye Res. 24, 321.
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CHAPTER
Raman Microprobe, MOLE
The Raman microprobe is called by different names, such as the Raman microscope, the Delhaye molecular microprobe, an optical microscope using the Raman effect, the Raman laser microprobe, or simply MOLE (molecular optical laser examiner). It is a combination of a microscope and a Raman spectrometer. MOLE was developed mainly by the efforts of three laboratories: Black Engineering, Inc.; the U.S. National Bureau of Standards; and Centre National de la Recherche Scientifique at Lille, France (Hirschfeld, 1973; Rosasco et al., 1975; Delhaye and Dharmelincourt, 1975). MOLE gives a standard microscopic image as well as a two-dimensional microscopic image of a sample at a certain shift from the exciting frequency (Raman lines). The only part of the sample that appears in the second case is the one that exhibits the Raman effect at the particular wave number under study. In this way heterogeneous samples can be differentiated under the microscope. Also, the Raman spectrum of a sample from a narrowly focused area can be obtained. From the spectrum we can determine the type of compound present in that particular area. MOLE provides the composition of the sample at the molecular level
412
Raman Microprobe, MOLE
usually without damaging. Unlike an electron microscope, which must be operated under high vacuum, the MOLE gives an image using almost any condition, such as the presence of water, air, or at high temperatures. MOLE is used for the qualitative and quantitative localization on a microscopic scale of organic, inorganic, and biological samples. One advantage of MOLE is that it allows visualizing a very small area of the sample, down to 1 X 1 !-tm in size. This is useful to identify the nature of a compound in a tiny spot in a gem or crystal or to see and identify the nature of a pollutant. In a sample with a homogeneous surface, the Raman spectrum obtained from a small area represents the spectrum of the sample's overall area. For instance, the spectrum of a unit 1 !-tm in length of calf thymus DNA fiber obtained by MOLE is identical to the spectrum obtained by a conventional method from a much larger area of the sample (Adar, 1978). It is well known that a laser is a relatively high-energy beam. So it is important to determine whether heating effects on samples may cause problems. Investigation indicates that heating effects become less severe as the particle size decreases (Etz and Blaha, 1980). However, sample sensitivity to the tightly focused, hence very intense, laser beam sets a practical upper limit on incident laser power (about 100-150 mW). There have been some technical improvements in MOLE recently. For instance, the attachment of a multichannel optical detector shortens the time required for obtaining a Raman spectrum, and this also avoids excess heat absorption by samples (Etz, 1980; Etz et al., 1980). There are excellent review articles by Rosasco (1980) and by Adar (1981) on the basic theory as well as applications.
1.
THEORY
The basic principle of the Raman microprobe is somewhat similar to that of the electron microprobe (Figure 16.1). In the electron microprobe, an electron beam is focused in vacuum on the surface of a solid sample. By analyzing the scattered secondary electrons and emitted characteristic X rays, one can determine the position and the concentration of a given element on the surface. In the Raman microprobe, photons (light) are used, and from the scattered Raman light one can detect and identify the compounds and their structures (Delhaye and Dhamelincourt, 1975; Dhamelincourt et al., 1979). MOLE consists of two components; one is a microscope (imaging system) and the other is a Raman spectrometer (spectral mode) (Figure 16.2). From the Raman spectrometer component of MOLE we can get a Raman spectrum, from which we can identify the nature of a compound. The sample is illuminated by a laser (vo ) beam, and the sample emits Rayleigh-scattered (vo ) and Raman-scattered light (vo ± vI' Vo ± V 2 " ' " Vo ± vn ). The frequencies vI ... vn are characteristic of the molecular vibrations of a molecule. The total image of the sample can be obtained in the microscope when Vo is used. This image is the same as that seen with a conventional
...
Theory
413
Excitations
Electrons
Backscattered or secondary electrons Fluorescence X-rays
Electron Microprobe
hv o hv Rayleigh o + h{v l - vol Raman
Raman Microprobe
FIGURE 16.1. The basic principle of the Raman microprobe and an electron microprobe. There is an analogy between these two probes.
microscope. But with the microscope in MOLE, the light can be filtered to give "n" If the sample is observed through MOLE with the frequency of "0 - "n' one sees the image of the p~ion of the sample that gives the Raman-scattered light. However, the frequency one selects to reconstruct the image must be well above the background. All other parts of the sample are not seen under this condition (Figure 16.3). The overall appearance of MOLE is shown in Figure 16.4. "0 -
mlcr05COpe con~rol
optical
fil~er
cra-~recorder
'screen
Raman
spectrometer
FIGURE 16.2. A simplified diagram of a Raman microprobe. The figure was reproduced from Ballan-Dufranpis et aI. (1979) by permission of the copyright owner. Biologie Cellulaire.
414
Raman Microprobe, MOLE
>-.
Raman spectrum of substance A
~
OM ~
ctl
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CIl ~
~
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Raman spectrum of substance B Raman spectrum '~ubstance C
V
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/
Image obtained using a particular Raman line
FIGURE 16.3. A diagram showing why MOLE can show the localization of a particular constituent in a heterogeneous sample. The figure was reproduced from Dhamelincourt and Bisson (1977).
FIGURE 16.4. An overall view of MOLE. The photograph was kindly supplied by Dr. M. Delhaye and Dr. 1. Oswalt, Jobin-Yvon Instruments, France.
2.
CLINICAL AND HISTOLOGICAL APPLICATIONS
The Raman microprobe is useful for the in situ analysis of cellular inclusions. Several examples are shown in this section. The foreign material within a lymph node from a patient with a silicone elastomer finger-joint prosthesis was detected by MOLE (Abraham and Etz,
Clinical and Histological Applications
415
1979). The light micrograph of the lymph node shows the foreign material within multinucleated giant cells (Figure 16.5). The Raman spectra of the microscopic foreign inclusion in the lymph node and of the prosthesis are identical (Figure 16.6). The foreign material inside a histological section of fish liver was identified by MOLE as disordered graphite carbon (Delhaye et al., 1979). Purine compounds in tissue can be detected by MOLE. As a result of nucleic acid and protein degradation, certain purine compounds such as uric acid, guanine, and xanthine accumulate in cells and tissues. In situ study of such bioaccumulated compounds has been difficult because of the lack of specific histochemical reagents for purines. Moreover, their chemical structures are similar, so exact identification has been difficult. By the use of MOLE, uric acid, guanine, or xanthine have been identified in spider cuticle, fish skin, Blatella fat granule, and other animal tissues (Ballan-Dufran9ais et al., 1979) (Figures 16.7 and 16.8). The formation of stones (renal lithiasis) in different parts of the urinary system is a common disease for persons of all ages. How the stone is formed is not known, but it is widely believed that it is due to a metabolic abnormality. Kidney stones can be made of inorganic salts such as calcium phosphate or calcium carbonate, or they can be organic stones such as cysteine stone, oxalate stone, or uric acid stone. The chief constituent of a kidney stone is calcium. Identification of kidney-stone composition is important, so a physician can advise a patient what dietary course should prevent another kidney stone from forming. MOLE not only can locate the kidney stone in the nephron, but it can also determine its composition. For instance, for one type of stone, the Raman
FIGURE J6.5. A section of lymph node with foreign bodies of silicone rubber within multinucleated giant ceIls. The arrow indicates typical inclusions of silicone polymer. The star shows the cytoplasmic area analyzed to obtain the micro-Raman spectrum of the host tissue matrix. The figure was reproduced from Abraham and Etz (1979) by permission of the American Association for the Advancement of Science. The photograph was kindly supplied by Drs. Abraham and Etz.
416
Raman Microprobe, MOLE
)
(a)
I)
CYTOPLASM OF GIANT CELL
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I
I
I
2
u
>f-
FOREIGN BODY IN GIANT CELL
(a)
IL
3000
I,
~
I
I"lj I 0
~ f~
I
0
SILICONE ELASTOMER
2000
1000
o
RAMAN SHIFT (em-I)
FIGURE 16.6. Spectra recorded in the Raman probe microanalysis of a deparaffinized standard 5-JLtn section of lymph node. The figure was reproduced from Abraham and Etz (1979) by copyright permission of the American Association for the Advancement of Science. The photograph was kindly supplied by Drs. Abraham and Etz.
spectrum shows three distinct bands at 1075, 970, and 278 cm - I. The 1078-cm- 1 band corresponds to the carbonate C-O vibrational mode and the 970-cm- 1 band to the symmetrical P-O vibration, and the 278-cm- 1 band indicates the presence of calcite CaC03 . Therefore, the kidney stone is made up of calcium phosphate and calcium carbonate. Mineralizations in kidneys of rabbits intoxicated by HgC1 2 , periwinkle concretions, and conjunctive cells were also studied by MOLE (Truchet et al., 1980). There are two types of oxalate stones: calcium oxalate monohydrate and calcium oxalate dihydrate. These two types of crystals can be readily distinguished by MOLE. The monohydrate shows a Raman line at 1462 cm- I, and the dihydrate shows a line at 1476 cm- I . Likewise, cysteine stone can be readily detected by the characteristic vibrational band of S-S stretching at 499 cm- I. Other types of kidney stones such as uric, calcium phosphate, and calcium carbonate stones can also be detected by their characteristic Raman lines. Thus MOLE is a powerful tool in the analysis of calculi (Daudon et al., 1980). Myxomycetes are mushrooms that secrete mineralized grains whose chemical identity was previously unknown. Using MOLE, the grains were found to .J
FIGURE 16.7. (A) Histological section of spider (X680). C, cuticle; E, epithelium; G, guanine concretions; 0, approximate analyzed area. (B) Histological section of fish (XSOO). S, skin; G, needles of guanine; 0, approximate analyzed area. The photograph was reproduced from BallanDufran~ais et al. (1979) by permission of the copyright owner, Biologie Cellulaire.
418
Raman Microprobe, MOLE
a
b
c 500
1000
1500
.b.\lcm 1
FIGURE 16.8. Raman spectra obtained from MOLE. (A) Spider guanophore. (8) Standard guanine. (C) Fish guanophore. Note the striking similarities in the spectra. These results demonstrate that the concretions seen in the micrograph are pure guanine. The spectra were reproduced from Ballan-Dufran~s et aI. (1979) by permission of the copyright owner, Biologie Cellulaire.
be composed of calcite, since they show the characteristic Raman lines at 1090, 715, and 280 em-I (Locquin and laeschke-Boyer, 1980). Enamel formation is common in animal tissues. Enamel formation is characterized by the nucleation of a mineral deposit and its growth during the secretory stage of ameloblasts. Fully grown extracellular enamel eventually becomes 99% mineral in the tissue. The vibrational analysis of hydroxylapatite permits the detection of enamel in the rat incisor (Casciani and Etz, 1979). Examination of mollusk tissue shows a very distinct band at 474 em - I that is an indication of the S-S stretching vibration. This is due to the presence of CuS in intermediate compounds degraded from hemocyanin, which is a copper-sulfur compound (Ballan-Dufranljais et al., 1979). Pyrocystis lunula is a dinoflagellated algae that emits light. The substance responsible for light emission has not yet been identified. Using MOLE, the
Environmental Applications
419
FIGURE 16.9. The image was obtained using one of the resonance Raman lines of carotenoids at 1527 cm - I. Localization of carotenoids in carrot-root cell. The photograph was kindly supplied by Dr. M. Delhaye. The source of the photograph is P. Bisson, These de 3eme cycle, Universite de Lille, Juillet 1977.
location of bioluminescent materials within the cells have been identified. Judging from the Raman spectrum, the compound may be luciferin (Arrio et aI., 1980). Localization of carotenoids (see Chapter 14, Section 1.4) in a cell of a carrot root is shown in Figure 16.9. Carotenoids are polyene compounds that give resonance Raman spectra. Using one of the carotenoid resonance Raman lines, one can obtain an image of the carotenoid localization. The same kind of Raman image can easily be obtained from a variety of living cells, including both plants and algae (Cavagnat et aI., 1981).
3.
ENVIRONMENTAL APPLICATIONS
Wide use of pesticides causes considerable environmental problems. Many pesticides are absorbed by cattle, fish, and humans, and these pesticides usually become embedded in the tissues. Identification of the absorbed pesticides is an important, yet difficult, problem. Interferential contrast microscopy shows the pesticide location; only the Raman microprobe allows their identification without extraction (Delhaye et aI., 1979; Arrio et aI., 1980). Many ciliated protozoans ingest nutrients and inert, insoluble pollutants by endocytosis. The important question is whether ingested pesticides are metabolized or remain intact. Raman microprobe investigations indicate the pesticides 4,4'-dichlorodiphenyl and j3-endosulfan remain intact, with no chemical modification occurring in the cells (Dive et aI., 1980a, b). Some polynuclear aromatic hydrocarbons are potent carcinogens. Such compounds are often found in polluted urban air produced by photochemical reactions of miscellaneous hydrocarbons released to the air after combustion of
420
Raman Microprobe, MOLE
fossil fuels. Pyrene and phenanthrene are detected in airborne particles as small as 30 !tm (Etz, 1979).
4.
OTHER APPLICATIONS
MOLE is also applied to paleontology. Both calcite and argonite are calcium carbonate, but they have different crystal forms. Thus they have slightly different carbonate vibration bands. When MOLE was us~d, the calcite nature of protozoan Ammonia beccarii was found to be calcareous lamellae and that of Hoeglundina elegans, aragnonitic in nature (Venec-Peyre and Jaeschke-Boyer, 1978).
REFERENCES Abraham, J. L., and Etz, E. S. (1979). Molecular microanalysis of pathological specimens in situ with a laser-Raman microprobe. Science 206, 716. Adar, F. (1978). Use of the molecular microprobe to record Raman spectra of a single mitochondrion and a fiber of calf thymus DNA. Fron. BioI. Energ. 1, 592. Adar, F. (1981). Developments in Raman microanalysis. In Microbeam Analysis, H. Geiss, Ed., San Francisco Press, San Francisco, pp. 67-72. Arrio, B., Dupaix, A., Fresneau, C., Lecuyer, B., and Volfin, P. (1980). Etudes spectroscopiques in vivo de cellules vegeiales bioluminescentes: Pyrocystis lunula. L'Actualite Chimique 4, 18.
c., Truchet, M., and Dhamelincourt P. (1979). Interest of Raman laser microprobe (mole) for the identification of purinic concretions in histological sections. BioI. Cellulaire 36, 51.
Ballan-Dufran~s,
Casciani, F. S., and Etz, E. S. (1979). Raman microprobe study of biological mineralization in situ: Enamel of the rat incisor. In Microbeam Analysis, Dale E. Newbury, Ed., San Francisco Press, San Francisco, pp. 169-172. Cavagnat, R., Cruege, F., and Pham, V. H. (1981). Biological applications of resonance Raman spectroscopy and resonance micro-Raman spectroscopy. Biochimie 63, 927. Daudon, M., Jaeschke-Boyer, H., Protat, M. F., and Reveillaud, R. J. (1980). La microsonde Mole et l'analyse des calculs urinaires. Perspectives et realites. L'Actualite Chimique 4,25. Delhaye, M., and Dhamelincourt, P. (1975). Raman microprobe and microscope with laser excitation. J. Raman Spectrosc. 3, 33. Delhaye, M., Dhamelincourt, P., and Wallart, F. (1979). Analysis of particulates by Raman microprobe. Toxicol. Environm. Chem. Rev. 3, 73. Dhamelincourt, P., and Bisson, P. (1977). Principle and realization of an optical microscope using the Raman effect. Microsc. Acta 79, 267. Dhamelincourt, P., Wallart, F., Leclercq, M., N'Guyen, A. T., and Landon, D. O. (1979). Laser Raman molecular microprobe (MOLE). Anal. Chem. 51, 414A. Dive, D., Devynck, J. M., Leroy, G., Coustaut, D., and Moschetto, Y. (1980a). Principles and examples of applications of the Raman microprobe in the biological area. L'Actual. Chimique 4,24. Dive, D., Devynck, J. M., Leroy, G., Fourmaux, M. N., and Moschetto, Y. (1980b). Identification of intracellular particles of pesticides in ciliate protozoa by Raman microprobe. Experientia 36,832.
References
421
Etz, E. S. (1979). Raman microprobe analysis: principles and applications. In Scanning Electron Microscopy, SEM, AMF O'Hare, IIl., pp. 67-92. Etz, E. S. (1980). New Raman microprobe with multichannel optical detector. In National Bureau of Standards Dimensions 11, I. Etz, E. S., and Blaha, 1. J. (1980). Scope and limitations of single particle analysis by Raman microprobe spectroscopy. In Proc. Spec. Sess. on Particle Anal., 13th Ann. Conf. Microbeam Anal. Soc., Ann Arbor, Michigan, pp. 153-197. Etz, E. S., Adar, F., Landon, D.O., and Steinbach, W. R. (1980). A new Raman microprobe with multichannel optical detector-characteristics and applications. In Proc. 10th N. E. Regional ACS Mtg. Abstract, Potsdam, N.Y. Hirschfeld, T. (1973). Raman microprobe: vibrational spectroscopy in the Femtogram range. Opt. Soc. Am. 63, 476. Locquin, M., and Jaeschke-Boyer, H. (1980). Structure calcitique inframicroscopique de grains de secretion microniques chez les myxomycetes. L 'Actual. Chimique 4, 22. Rosasco, G. J. (1980). Raman microprobe spectroscopy. In Advances ill Infrared and Raman Spectroscopy, Vol. 7, R. J. H. Clark and R. H. Hester, Eds., Heyden, London, pp. 223-282. Rosasco, G. J., Etz, E. S., and Cassatt, W. A. (1975). The analysis of discrete fine particles by Raman spectroscopy. Appl. Spectrosc. 29, 396. Truchet, M., Martoja, M., Martoja, R., and Ballan-Dufran~s,C. (1980). Applications de la mole a l'mineraux sur coupes histologiques. L'Actual. Chimique, p. 15. Venec-Peyre, M.-T., and Jaeschke-Boyer, H. (1978). Micropaleontologie. Application de la microsonde moleculaire illaser Mole a l'etude du test du quelques Foraminiferes calcaires. C. R. Acad. Sc. 287, 607.
CHAPTER
Clinical and Environmental Applications
CLINICAL DIAGNOSIS: TISSUES AND CELLS
le application of Raman spectroscopy to clinical problems is still in its infant 1ge, but it has a great future potential. In recent years, the Raman spectra of blood have proved to be sensitive to rious pathological disturbances affecting human blood plasma, and this ~thod of diagnosis claimed to be superior to conventional clinical tests such the sedimentation rate of erythrocytes. In the Stokes-frequency range )00-4000 cm -I), the overall spectra and fluorescence backgrounds of blood lsma taken from patients suffering mammary carcinoma and pseudolcinous papillary cystodeno carcinoma were shown to differ considerably m the spectra of normal blood plasma (Larsson and Hellgren, 1974). In jition, the low-frequency Raman spectra (up to 200 cm - I) of human
Clinical Diagnosis: Tissues and Cells
423
mammary carcinoma and normal human mammary tissues appear to differ in that the shift lines, at given frequencies, seen with normal tissues tend to become much broader and resolved into two or three separate lines in the spectra of homologous malignant mammary tissues (Figure 17.1) (Webb et aI., 1977a). Similar alterations in the millimeter microwave spectra of tumor versus normal cells including mammary tissues also have been observed (Webb and Booth, 1971; Webb et aI., 1977b). Thus both Raman and microwave spectroscopy of blood sera and tissues may become diagnostic tools for the rapid detection of mammary carcinoma as well as other malignant diseases (Webb, 1976; Webb and Lee, 1977; Webb, 1980). One of the more interesting facets of the Raman spectroscopy of intact cells is found with microbial cells. The Raman spectra of bacterial cells when they are in the resting state have been found to be essentially devoid of shift lines, whereas the spectra of growing cells display many lines of a complex nature (Webb and Stoneham, 1976a, b, 1977; Webb et aI., 1977b; Webb, 1980). In addition, at normal physiological temperatures, the spectra of algal cells show only lines assigned to carotenoid pigments; the expected lines from the large chlorophyll molecules apparently are absent and may be seen only in the spectra of such cells when they are held at temperatures below 261 K (Drissler, 1980). Such observations have led to the suggestion (Webb, 1980) that many of the shift lines seen in the spectra of active cells arise from Raman-active energy states induced in vivo by metabolic activities. This suggestion was based not only on the lack of lines from resting cells but also on the finding that the "active cell" Raman spectra were altered when growing cells were exposed to
B
A
lcm-1 J FIGURE 17.1. Raman spectra of (A) normal human mammary tissue, (B) normal right-side and (C) left side 2-3 cm mammary carcinoma of human. The figure was reproduced from Webb et al. (l977a) by permission of the copyright owner, John Wiley.
424
Clinical and Environmental Applications
microwave fields (from 40-140 GHz) during the determinations of their Raman spectra. Microwaves of such frequencies have been shown to affect the growth and metabolic activities of both bacteria (Webb and Dodds, 1968; Webb and Booth, 1969; Berteaud et al., 1975) and yeast cells (Grundler et al., 1977). It appears, therefore, that normal in vivo metabolic activities may be followed by Raman spectroscopy, and this same technique may be used to assess the influence of imposed microwaves stresses on them. It is possible, therefore, that with refinements in techniques, Raman spectroscopy may prove to be a useful tool in the assessment of the mechanisms by which the functions of the nervous and cardiovascular systems become impaired when they are exposed to microwave fields of low intensities, as has been reported in numerous publications. The infection of animal and microbial cells by viruses is known to alter metabolic processes severely, as does the physical or chemical induction of prophages and latent viruses (such as the Herpes complex) normally carried by some bacterial and animal cells, respectively. It has been found that the microwave spectra of bacterial cells carrying prophages differ in small detail from those of the same cell strain not possessing the virus (Lee and Webb, 1977). The "active" Raman spectra of these two cell types also differ, especially when the prophage in the carrier strain is induced to multiply in vivo by agents such as X rays, uv light, and chemical carcinogens (Webb, 1981). The presence, in vivo, of a carried virus or viral infection from outside apparently may be detected, therefore, by Raman spectroscopy, and such a procedure could enhance greatly the rapid clinical diagnosis and detection of viral diseases. Differences were observed in the Raman lines of a bacteria cell (Escherichia coli) suspension when illuminated by laser light of 10-400 mW for only 1 s (O'Sullivan and Santo, 1981). Raman spectroscopy may become an analytical technique to detect the state of bacteria or tissues in a metabolically active condition.
2.
CLINICAL ANALYSIS
Catecholamines (adrenaline and noradrenaline, also called epinephrine and norepinephrine) are neurotransmitters that play an important role in biological systems. With hypertension, pheochromocytoma, and other disorders, catecholamine levels in urine are elevated. Clinical analysis of adrenaline (epinephrine) and noradrenaline (norepinephrine) concentrations is important, as in some nerve disorders the levels of these compounds vary from normal values. Analysis is also important because these compounds are used as drugs for the treatment of some disorders. The normal procedure for the analysis is a fluorometric technique that requires the conversion of catecholamines to trihydroxyindoles. A variety of quenching agents and other fluorescent materials can interfere with this analysis. Resonance Raman spectroscopy can also be used to analyze for these compounds. For this analysis, catecholamines are
Environmental Problems
425
first oxidized by ferricyanamide to the corresponding aminochromes. The C=N+ stretching vibration mode of aminochromes can be resonance enhanced. Aminochromes are determined at 1480 cm -1 for adrenaline, 1425 cm- I for noradrenaline, and 1415 cm- I for dopamine (Morris, 1975, 1976; Rahaman and Morris, 1976). In order to establish Raman spectroscopy as a tool for the analysis of sympathomimetic amines, Raman spectra of amphetamine, methamphetamine, benzamphetamine, diphenhydramine, phendimetrazine, phentermine, ephedrine, and related drugs were obtained. It was concluded that these drugs can be differentiated by Raman spectroscopic techniques (Bass, 1978). Phenothiazines are drugs widely used as antipsychotics in the treatment of psychosis and can be analyzed by Raman spectroscopy due to the presence of the C-S bond (Kure and Morris, 1976). Barbiturates are drugs widely used for neuropsychiatric therapy in cases of insomnia and as sedatives and anticonvulsants. Widespread use of barbiturates has made their detection and analysis an ever-increasing problem in clinical, forensic, and toxicological laboratories. All barbituric acids contain the pyrimidine-ring structure. The strong Raman bands at 629 ± 8 cm - I (free acid) and 642 ± 4 cm ~ I (sodium salts) are due to the breathing vibrations of the pyrimidine ring. Most barbiturates are readily distinguished from each other by analysis of specific Raman band frequencies (Willis et al., 1972). Sulfonamides are well-known drugs commonly used as antibiotics. By diazotization, colorless drugs are converted to colored compounds from which resonance Raman spectra can be obtained. The detection limit is as low as 2 X 10- 8 M (Sato et al., 1980). Many drugs interact with nucleic acids. These are discussed in Section 6.4., Chapter 5. Many drugs show polymorphism in the solid state; therefore, they exist in several crystalline forms having different physical properties and different absorption rates when administered as drugs. Griseofulvin is a drug used in the treatment of fungus infections. This compound exists in the solid state in the unsolvated form and can form solvates with different solvents. The manner of its solvation (solute-solvent interaction) can be studied by Raman spectroscopy. In the benzene solvate, only weak van der Waals interactions exist between solute and solvent. However, in solvates with chloroform and bromoform, weak hydrogen bonding exists between the proton of the solvent and the keto group of the benzofuran ring in griseofulvin. When desolvation occurs, the crystal does not go through any intermediate, and the lattice reverts to the structure of unsolvated griseofulvin (Bolton and Prasad, 1981).
3.
ENVIRONMENTAL PROBLEMS
One important aspect in environmental research is to analyze pollutants quantitatively and also to identify compounds. Most compounds we would need to find in air, water, soil, and biological tissue are usually present in very
426
Clinical and Environmental Applications
small amounts. In order to identify a compound and its quantity, analytical methods must be simple, rapid, and accurate. Raman spectroscopy can be an ideal analytical tool for certain compounds. 3.1.
PESTICIDES
The detection of residual pesticides in air, soils, water, and animal tissues is an important step in environmental control. There has been considerable effort to use Raman spectroscopy for such analysis. Raman spectra of parathion, Guthion, ethion, Furadan, Baygon, Endosulfan, dieldrin, lindant, Aroclor, and 0, P' DDT [1-( O-chlorophenyl)-l-( p-chlorophenyl)-2, 2, 2-trichloroethane] were obtained by Vickers et aI. (1973). The work so far is preliminary, but it does indicate the feasibility of this technique. Raman spectra of chlorinated, sulfur-containing, phosphorus-containing and many other pesticides such as DDT, TDE, Perthane, methoxychlor, Dicofol, Terradifon, heptachlor, chlordane, dieldrin, endrin and Endosulfan, Pepulate, CDEC (2-chloroallyldiethyldithiocarbamate), thiram, maneb, zineb, ferbam, malathion, ethion, Methylparathion, parathion, EPN, O,O-diethyl0-(2, 4-dichlorophenyl)phosphorothioate, Dichlorvos, Mevinphos, tributyl phosphorotrithioite, 2, 4-dichlorobenzyltributylphosphonium chloride, IPC (isopropyl-N-phenylcarbamate, propham), CIPC (chloropropham), EPTC (ethyl-N, N-dipropyl-3-chlorophenylcarbamate), and nicotine were obtained by Nicholas et aI. (1976a, b, c). These are common pesticides currently used in agriculture in the United States. By comparing the fingerprint bands, they concluded that Raman spectroscopy is potentially useful for analysis of these pesticides. An attempt was made to detect chlorinated-hydrocarbon pesticides on thin-layer-chromatography plates by Raman spectroscopy. It was found that Raman detection requires at least 200 p,g of sample compared with a few micrograms needed for Fourier-transform IR spectroscopy (Gomez-Taylor et aI., 1976). This indicates that this application of Raman spectroscopy is potentially useful; yet it needs further refinement to be' a good analytical tool for pesticide detection. Phenylamide pesticides do not exhibit resonance Raman spectra, but when converted to azo-dye derivatives, they show good resonance Raman spectra that can be used for analysis of the pesticides (Higuchi et aI., 1980). 3.2.
FOOD ADDITIVES
The use of coloring additives in food is common practice in the food industry. The conventional methods of analyzing these dyes are tedious and time consuming; usually such analysis involves chromatographic and colorimetric methods. Dyes are suitable for analysis by resonance Raman spectroscopic methods, as each dye has its own absorption characteristic. Raman spectra of
References
427
artificial dyes FD & C Red, Nos. 2, 4, and 40 were obtained. Each dye has a unique Raman fingerprint, permitting identification of concentrations as low as 5 ppm. This result is promising, as the dye can be analyzed in situ without going through time-consuming separation steps (Brown and Lynch, 1976). The concentration of nitrate in water can be readily analyzed by Raman spectroscopy using a nitrate vibration line at 1045 cm -1. The detection limit is about 2 ppm. For waste and treated water, analysis becomes harder, due to the strong luminescence. With potassium iodide as a quencher, the sensitivity becomes comparable to that of pure water (Furuya et al., 1979).
4.
OTHERS
Bacterial spores contain about 10% dipicolinate. It is thought that this factor is responsible for the high resistance in the presence of calcium ion of spores to heat and other environmental stresses. The analysis of the carboxyl-group vibrational mode of Raman spectra indicates that the carboxyl group is not a simple calcium-salt type. From this, it was concluded that dipicolinate in the spore protoplasts is not the simple calcium salt. Calcium ion and dipicolinate are probably involved in some type of interaction, the nature of which is still unknown (Woodruff et al., 1974). Carbon monoxide is known to alter the in vivo metabolic activity of bacteria. Exposure of E. coli to carbon monoxide changes its metabolic activity and its metabolic time clock. This change can be followed by observing the l273-cm -1 Raman band. As the bacteria and CO interact, the intensity of the l273-cm -1 band decreases (Stoneham and Webb, 1976). Some bacteria contain carotenoids; thus from their resonance Raman spectra, bacteria can be differentiated. Some bacteria exhibit pronounced carotenoid overtone and combination bands (Howard et al., 1980).
REFERENCES Bass, V. C. (1978). The identification of sympathomimetic amines by Raman spectroscopy. Forens. Sci. 11, 57. Berteaud, M. A J., Dardalhon, M., Rebeyrotle, N., and Averbeck, M. D. (1975). Action d'un rayonnement electromagnetique illonguerev d'onde millimetrique sur la croissance bacterienne. C. R. Acad. Sci. 281, 843. Bolton, B. A, and Prasad, P. N. (1981). Laser Raman investigation of pharmaceutical solids: Griseofulvin and its solvates. J. Pharm. Sci. 70, 789. Brown, C. W., and Lynch, P. F. (1976). Identification of FD & C dyes by resonance Raman spectroscopy. J. Food Sci. 41, 1231. Drissler, F. (1980). Discovery of phase transitions in photosynthetic systems. Phys. Lett. 77A, 207. Furuya, N., Matsuyuki, A, Higuchi, S., and Tanaka, S. (1979). Determination of nitrate ion in waste and treated waters by laser Raman spectroscopy. Water Res. 13,371.
428
Clinical and Environmental Applications
Gomez-Taylor, M. M., Kuehl, D., and Griffiths, P. R. (1976). Vibrational spectrometry of pesticides and related materials on thin layer chromatography adsorbents. Appl. Spectrosc. 30, 447. Grundler, W., and Keilmann, F. (1978). Nonthermal effects of millimeter microwaves on yeast growth. Z. Naturforsch. 33C, 15. Grundler, W., Keilmann, F., and Frohlich, H. (1977). Resonant growth rate response of yeast cells irradiated by weak microwaves. Phys. Lett. 62A, 463. Higuchi, S., Aiko, 0., and Tanaka, S. (1980). Determination of trace amounts of some phenylamide pesticides by resonance Raman spectroscopy. Anal. Chirn. Acta 116, 1. Howard, Jr., W. F., Nelson, W. H., and Sperry, 1. F. (1980). A resonance Raman method for the rapid detection and identification of bacteria in water. Appl. Spectrosc. 34, 72. Kure, B., and Morris, M. D. (1976). Raman spectra of phenothiazine and some pharmaceutical derivatives. Talanta 23, 398. Larsson, K., and Hellgren, L. (1974). Combined Raman and fluorescence scattering from human blood plasma. Experientia 30,481. Lee, R. A, and Webb, S. J. (1977). Possible detection of in vivo viruses by fine structure millimeter microwave spectroscopy between 68 and 76 GHz. IRCS Med. Sci. 5, 222. Morris, M. D. (1975). Resonance Raman spectra of the aminochromes of some biochemically important catecholamines. Anal. Chern. 47, 2453. Morris, M. D. (1976). Determination of catecholamines by resonance Raman spectroscopy of their aminochromes. Anal. Lett. 9, 469. Nicholas, M. L., Powell, D. L., Williams, T. R., and Bromund, R. H. (I 976a). Reference Raman spectra of DDT and five structurally related pesticides and of five pesticides containing the norbomene group. J. Assoc. Offic. Anal. Chern. 59, 197. Nicholas, M. L., Powell, D. L., Williams, T. R., and Huff, S. R. (1976b). Reference Raman spectra of ten phosphorus-containing pesticides. J. Assoc. Offic. A nal. Chern. 59, 1071. Nicholas, M. L., Powell, D. L., Williams, T. R., Thompson, R. Q., and Oliver, N. H. (I 976c). Reference Raman spectra of eleven miscellaneous pesticides. J. Assoc. Offic. Anal. Chern. 59, 1266. O'Sullivan, R. A, and Santo, L. (1981). Experimental aspects in Raman spectroscopy of microorganisms. Abstr., Spec. Issue Can. J. Spectrosc., June. Rahaman, M. S., and Morris, M. D. (1976). Determination of adrenaline and noradrenaline by resonance Raman spectrometry. Talanta 23, 65. Sato, S., Higuchi, S., and Tanaka, S. (1980). Determination of small amounts of some sulfonamide drugs by resonance Raman spectroscopy. Anal. Chirn. Acta 120, 200. Stoneham, M. E., and Webb, S. J. (1976). Action of carbon monoxide on bacteria as seen by laser-Raman spectroscopy. Int. Res. Cornrnun. Sys., Med. Sci. 4, 520. Vickers, R. S., Chan, P. W., and Johnsen, R. E. (1973). Laser excited Raman and fluorescence spectra of some important pesticides. Spectrosc. Lett. 6, 131. Webb, S. 1. (1976). Nutrition Time and Motion in Metabolism and Genetics. Thomas, Springfield, Ill. Webb, S. 1. (1980). Laser-Raman spectroscopy of living cells. Phys. Repts. 60, 201. Webb, S. J. (1981). Detection and inquction of in vivo viruses as seen by laser-Raman spectroscopy. Personal communication. Webb, S. J., and Booth, AD. (1969). Absorption of microwaves by microorganisms. Nature 222, 1199.
Webb, S. J., and Booth, A D. (1971). Microwave absorption by normal and tumor cells. Science
174,72. Webb, S. 1., and Dodds, D. E. (1968). Inhibition of bacterial cell growth by 136 gc microwaves. Nature 218, 374.
References
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Webb, S. J., and Lee, R. A. (1977). Microwave and laser-Raman spectra of normal and tumour human mammary tissue. IRCS Med. Sci. U.K. 5, 102. Webb, S. 1., and Stoneham, M. E. (1976a). The effect of microwaves on the metabolic time clocks of normal and tumour cells. A laser-Raman study. IRCS J. Med. Sci. UK 4, 10. Webb, S. 1., and Stoneham, M. E. (1976b). Action of CO on bacteria as seen by laser-Raman spectroscopy. IRCS J. Med. Sci. U.K. 4, 520. Webb, S. J., and Stoneham, M. E. (1977). Resonances between 10 II and 10 12 Hz in active bacterial cells as seen by laser Raman spectroscopy. Phys. Lett. 6OA, 267. Webb, S. J., Lee, R. A., and Stoneham, M. E. (I 977a). Possible viral involvement in human mammary carcinoma: A microwave and laser-Raman study. Int. 1. Quant. Chem.: Quallt. BioI. Symp. 4, 277. Webb, S. J., Stoneham, M. E., and Frolich, A. (1977b). Evidence for non-thermal excitation of energy levels in active biological systems. Phys. Lell. 63A, 407. Willis. Jr., J. N., Cook, R. B., and Jankow, R. (1972). Raman spectroscopy of some common barbiturates. Anal. Chern. 44, 1228. Woodruff, W. H., Spiro, T. G., Gilvarg, C. (1974). Raman spectroscopy in vivo: Evidence of the structure of dipicolinate in intact spores of Bacillus megaterium. Biochem. Biophys. Res.
Commull.58, 197.
IIII~I~I~~~EN~~~I~II
About Professor
C.V.Raman The essence of science is independent thinking, hard work and not equipment.
-G. V. Raman
I think it is appropriate to present briefly a personal history of Professor C. v. Raman, an eminent Indian physicist, whose keen mind and astute observations led to the discovery of the scattering effect that bears his name. C. V. Raman was born in Thiruvanaikkaval near Trichinopoly, southern India, on November 7, 1888, as one of seven children. He obtained his master's degree in physics from Madras University and came to Calcutta in August 1907 as an officer of the Accountant General of Bengal. His research on spectroscopy after office hours made him an honorary worker at the Indian Association for the Cultivation of Science. In July 1917, C. V. Raman was appointed Palit Professor and Head of the Department of Physics of the Calcutta University of Science (Chakravarti, 1978).
Appendix
431
During a voyage to England, Raman was struck with the idea that the deep blue color of the sea might be due to light scattering by water molecules, independent of the presence of any suspended matter. He considered that the color of the sea was fundamentally the same phenomenon as that observed in the sky, namely, the scattering of light by the molecules in the air (Raman, 1921; Ramanathan, 1978). Upon his return to India, Raman extended Lord Rayleigh's work on the scattering of light and extensively published papers on the light scattering and magnetism of liquids and solids. He published 57 papers before reporting on the effect now known as Raman scattering. The first line spectrum showing the Raman effect was made in the evening of February 28, 1928, in the laboratory of the Indian Association for the Cultivation of Science at 210 Bow Bazar Street, Calcutta. Professor Raman observed that highly purified organic liquids contained weak scattering components of frequencies not originally present in the incident light. A fascinating fact is that he observed the new radiation by eye. Raman suspected that this scattered frequency was analogous to the Compton effect. In 1923, the American physicist Arthur Holly Compton in Chicago found that the scattering of X rays by the electrons in a graphite target gave rise to a second X ray band with shifted wavelength. Raman therefore concluded that there were new types of scattered light that must be optical analogs of the Compton effect. The first announcement of the discovery of modified scattered radiation in scattering was made by Professor Raman in an inaugural lecture delivered under the title "A New Radiation," on Friday, March 16, 1928, at a meeting of the South Indian Association in Bangalore (Krishnan, 1978). The text was published in the Indian Journal of Physics, 2, 387-398 (1928). The text of the speech consisted of nine parts. They were (i) "Introduction," (ii) "A New Phenomenon," (iii) "Its Universality," (iv) "Line-Spectrum of New Radiation," (v) "Nature of the New Radiation," (vi) "Relation to Thermodynamics," (vii) "Coherent or Non-Coherent Radiation?" (viii) "Possible X-Ray Analogues," (ix) "Conclusion." Professor Raman was very confident about his finding and his first sentence was, "I propose this evening to speak to you on a new kind of radiation or light-emission from atoms and molecules." Then he explained the phenomena of fluorescence and light scattering. In "A New Phenomenon," he described the history of the investigation made in Calcutta. The first hint that he and his assistant, Mr. Seshagiri Rao, found, in December 1921, was that the depolarization of light transversely scattered by distilled water increased very markedly when a violet filter was placed in the incident-light path. Later, in 1922, using dust-free methyl and ethyl alcohol, he noticed that the colors of scattered light from the different liquids did not match. In the summer of 1923, Professor Raman and Dr. K. R. Ramanathan suspected that the difference in color was due to scattered light. Ramanathan described this as "a trace of fluorescence." Since 1923, the investigation of this "weak fluorescence" was the main effort of Professor Raman and his associates. Although the effect was called "feeble
432
Appendix
fluorescence" at the time, Professor Raman thought this was not fluorescence. In his speech he said, " ... the impression left on my mind at the time was that we had here an entirely new type of secondary radiation distinct from what is usually described as fluorescence." After Professor Compton's study of X-ray scattering, Professor Raman accelerated his light-scattering study. He used a powerful beam of sunlight from a heliostat concentrated by a 7-in. telescope objective combined with a short-focus lens. The light was passed through a blue-violet filter and then through the liquid under examination. The new type of radiation, Professor Raman observed, was not restricted to liquids. Mr. K. S. Krishnan and Professor Raman observed the same phenomenon in the gases CO2 and N 20. They noticed that the scattering power of gases was much weaker than that of liquids. It was further determined that this new type of radiation could be observed in crystals such as ice. Thus the phenomenon was observed with the three phases of matter. In his inaugural lecture in 1928, Professor Raman explained the first Raman spectra made on photographic plates. He was not satisfied by the discovery alone, but proposed to explain the theory of the new phenomenon. He clearly stated, "As a tentative explanation, we may adopt the language of the quantum theory, and say that the incident quantum of radiation is partially absorbed by the molecule, and that the unabsorbed part is scattered." He had already predicted the usefulness of Raman scattering in the study of molecular structure by saying, "The measurement of the frequencies of the new spectral lines thus opens a rleW pathway of research into molecular spectra, particularly those in the infra-red region." The prediction was emphasized again in the final part of his speech, "We are obviously only at the fringe of a fascinating new region of experimental research which promises to throw light on diverse problems relating to radiation and wave theory, X-ray optics, atomic and molecular spectra, fluorescence and scattering, thermodynamics and chemistry. It all remains to be worked out." All of these points he made in 1928. Raman recognized the importance of the scattering phenomenon and stated in his letter to the Spex Speaker that from September 1921 to February 1922, he thought" the molecular scattering of light in a transparent medium is a universal phenomenon exhibited in various degrees by all such materials, viz., crystalline solids, glasses, liquid and gases" (Raman, 1966). During the period September 1927 to February 1928, his thoughts became more specific about light scattering, and he said, "It began with my attempts to reconcile the apparent conflict between the ideas of wave-optics and Einstein's idea that light consists of discrete quanta of energy. That a reconciliation was possible between these two concepts of the nature of light was demonstrated by my success in deducing on the basis of the classical wave-principles, the existence, as well as the characteristics, of the two types of X-ray scattering, respectively with and without a change of frequency" (Raman, 1966). C. V. Raman and K. S. Krishnan submitted a paper to Nature on February 16, 1928, under the title "A New Type of Secondary Radiation." It was published in Nature, 121, 501-502 (1928). Again, on March 6, 1928, C. V.
Appendix
433
Raman submitted a one-page paper, "A Change of Wave-Length in Light Scattering," to Nature, and the article was published in volume 121, page 619, in 1928. With these announcements and publications, "a new radiation," presently known as the Raman effect, was revealed to the world. The equipment originally used by Raman consisted of a quartz mercury lamp, a container for the sample, a condensing lens, and a pocket spectroscope. Later, more-powerful mercury arcs, large-aperture spectrographs, and photographic recordings were used. In order for the photographic plate to have a good exposure, an exposure of hours and even days was given. Raman was not aware of the theory that predicted the scattering effect he observed. In 1923 an Austrian physicist, A. Smekal, published a paper predicting the frequency shift of inelastic light scattering (Andrews, 1978). Actually, there are other people who, independently, predicted the Raman effect. Rocard concluded from his research that scattered light must have a different spectral structure than incident light (Rocard, 1928). Brillouin delivered to the French Academy of Science a report that Rocard had published a note on Raman scattering in Comptes Rendues on March 23, 1928. One week later, Fabry reported to the French Academy on a note by Cabannes (1928), who predicted the Raman effect as early as 1924. Landsberg and Mandelstam at the Institute for Theoretical Physics, without knowing of Raman's publication, submitted a paper to the German journal Naturwissenschaften on the same effect Raman observed (Landsberg and Mandelstam, 1928). So the effect was predicted by several investigators before Professor Raman actually observed the effect. However, Professor Raman was the first person to publish an actual spectrum of scattered light containing shifted frequencies. Readers who are interested in the historical aspects of Raman spectroscopy are advised to read the article by BrandmUller and Kiefer (1978). The news of Raman's discovery did not evoke much interest in London at that time, because very few physicists were actively working on the problem of light scattering. Dr. R. W. Wood, an American physicist working on light scattering, immediately checked the Raman effect after reading the letter by Raman (Lord, 1978). Wood rechecked his own spectroscopic plates taken earlier and found evidence of the same effect Raman observed. He cabled his results and published them in Nature (Wood, 1928). In the beginning of his cable he stated, "PROF. RAMAN's brilliant and surprising discovery that transparent substances illuminated by very intense monochromatic light scatter radiations of modified wavelength, and that frequency of difference between emitted radiations and one exciting medium is identical with frequency of infrared absorption bands opens up wholly new fields in the study of molecular structure." At the end of the cable, he praised Professor Raman's discovery by saying, "It appears to me that this very beautiful discovery, which resulted from Raman's long and patient study of phenomena of light scattering, is one of most convincing proofs, of quantum theory of light which we have at present time."
434
Appendix
Wood introduced the new terminology of anti-Stokes scattering to the higher frequency of Raman lines (Brandmiiller and Kiefer, 1978). At the end of 1928, Wood, Rasetti, and McLennan showed that the Raman displacements actually represented the vibrational or rotational frequencies of the diatomic molecules, liquid nitrogen and oxygen (Asundi, 1978). Wood observed the Raman effect of HCI gas and obtained the Raman line at 2886.0 cm -I that coincided with 2885.9 cm -I, a value well established by its IR spectrum for this frequency and left no doubt as to its origin. The energy associated with the 2886 cm - I band actually corresponds to an energy transition from the ground state to the first excited vibrational level. In 1931, the book entitled Der Smekal-Raman Effect by K. W. F. Kohlrausch was published in Germany, and this upset Professor Raman very much. In 1938 the second edition of this book appeared, again under the same title. By this time the entire scientific community was unanimously calling this effect just the Raman effect. When the 1928 Nobel Prize in physics was awarded to Richardson, it was expected by several scientists that the 1929 prize would be awarded to Raman, but instead it was given to de Broglie, who discovered the matter wave. But in 1930 Professor Raman was rightly awarded the prize. His Nobel lecture was delivered on December 11, 1930, and consisted of nine parts (Raman, 1931). The first one was "The Color of the Sea." This shows how interested he was in the deep blue color of the sea, probably recalling his first voyage to England through the Red Sea and the Mediterranean, where he first thought of the light-scattering phenomenon. The titles of the other sections of the Nobel lecture were (ii) "The Theory of Fluctuation," (iii) "The Anisotropy of Molecules," (iv) "A New Phenomenon," (v) "The Optical Analogue of the Compton Effect," (vi) "Its Spectroscopic Character," (vii) "Interpretation of the Effect," (viii) "The Significance of the Effect," and (ix) "Some Concluding Remarks." In the concluding remarks he praised the works of Dr. McLennan on liquified gases and Dr. Wood and Dr. Rasetti on their light-scattering work. Professor Raman not only contributed to the fundamental knowledge of light scattering but also to the application of the Raman effect to practical problems. One example is the identification of diamond from its Raman spectrum (Bhagavantam, 1978). Professor Raman's last scholarly work was published in Nature in 1945, when he was 57 years old. The title was "Scattering of light in crystals," andit was his 94th paper (Raman, 1945). His first paper was entitled "The Doppler effect in the molecular scattering of radiation" and also was published in Nature (Raman, 1919). Throughout his life of 82 years, he devoted himself to the study of the phenomena of light scattering. It seems he was always absorbed in something to do with color. A student, Dr. S. Ramaseshan, wrote, "A few months before he died, I remember, while walking with him one evening amongst the eucalyptus groves that he loved, he stopped me in his characteristic manner and pointing towards the sky said: 'Have you seen
References
435
anything so beautiful?' " Till his end he was fascinated by the color of the sky, the effect of light scattering. Readers who are interested in original papers of Professor C. V. Raman are directed to the book published by the Indian Academy of Science (1978). Since the discovery of the Raman effect in 1928 and its subsequent recognition by world scientists, there has been rapid progress in this field, as can be witnessed by the 1800 scientific papers and Raman spectra on over 2500 compounds published by 1939 (Long, 1978). The total number of publications on Raman spectroscopy exceeds 23,600 up to the end of 1978 (Krishnan and Shankar, 1980). I had an occasion, in 1979, to give a seminar at the Biophysics Unit of the Indian Institute of Science in Banga10re, and was impressed by the quality of scientists at the institute. Professor Raman was the director of the Indian Institute of Science in 1933. Even based on the standards of the United States and Europe, the Indian Institute of Science is a first-class research institute. Here I observed the influence of Professor Raman, where he is still remembered and respected by scientists of the younger generation. As a scientist who works on Raman spectroscopy, the visit to Banga10re was one of the most memorable and moving events in my life. I wish he could have lived to see the tremendous development in the biological application of Raman spectroscopy. He died in 1970, about the time the biological application of Raman spectroscopy began. REFERENCES Andrews, D. (1978). The discovery of the Raman effect. New Sci. 16, March 1978,722. Asundi, R. K. (1978). Reminiscences relating to the discovery of the Raman effect. Curro Sci., 47, 192. Bhagavantam, S. (1978). The discovery of the Raman effect, reminiscences of Sir C. V. Raman. In Proc. Sixth Int. Can/. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter (Eds.). Heyden, London, Philadelphia, Rheine, pp. 3-12. Brandmiiller, J. and Kiefer, W. (1978). Physicists' view, fifty years of Raman spectroscopy. Spex Speaker 23, 310. Chakravarti, R. N. (1978). Fifty years of Raman effect: 1928-1978. J. Inst. Chem. (India), SO, 187. Krishnan, R. S. (1978). Fifty years of Raman effect-some recent developments. Curro Sci. 47, 196. Krishnan, R. S. and Shankar, R. K. (1980). Progress of research on the Raman effect: A statistical analysis. J. Indian Inst. Sci., 62, 53. Landsberg, G. and Mandelstam, L. (1928). Eine neue Erscheinung bei der Lichtzerstreuung in Kristallen. Naturwissenschaften, 16, 772. Long, D. A. (1978). Raman spectroscopy in Europe for the past fifty years. In Proc. Sixth Int. Can/. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter (Eds.). Heyden, London, Philadelphia, Rheine, pp. 13-27. Lord, R. C. (1978). The early days of Raman spectroscopy in the United States. In Proc. 6th Int. Can/. Raman Spectrosc., E. D. Schmid, R. S. Krishnan, W. Kiefer, and H. W. Schrotter (Eds.). Heyden, London, Philadelphia, Rheine, pp. 29-37.
436
Appendix
Raman, C. V. (1919). The Doppler effect in the molecular scattering of radiation. Nature, 103, 165. Raman, C. V. (1921). The colour of the sea. Nature, 108,367. Raman, C. V. (1928). A new radiation. Indian 1. Phys., 2, 387. Raman, C. V. (1931). The molecular scattering of light, Nobel lecture. Les Prix Nobel, en 1930. Stockholm, Imprimerie Royale, P. A. Norstedt. Raman, C. V. (1945). Scattering of light in crystals. Nature ISS, 196. Raman, C. V. (1966). Laser Raman spectroscopy. The Spex Speaker, XI, I. Ramanathan, K. R. (1978). Some reminiscences of my association with Professor Raman. Curl'. Sci. 47, 179. Rocard, Y. (1928). Les nouvelles radiations diffusees. C. R. Ac. Sci., 186, 1107. Wood, R. W. (1928). Wave-length shifts in scattered light. Nature 122, 349.
Index
A,165 Absorption band, 18.317 Absorption of light, 3-6 Absorption spectra, heme compounds, 317 2-Acetamido-2-deoxY-D--glucose, 240, 246 Acetyl group, carbohydrates, 245 N-Acetyl group, 246 N-Acetylg!ucosamine, 235-239, 245, 246 N-Acetylhistidine methylamide, 82 N-Acetylprolinamide, 80 N-Acetylserine methylamide, 82 N-Acetyltyrosine methylamide, 82 Acid anhydride, 36 Acid phosphatase, 83, 377 Acoustic accordionlike motion, 191 Acridine orange, 57 Actin, 401 Actinomycin, 163 Active Raman spectra of cells, 424 Adenine, 59. 148,150,154,155,160,184 Adenosine, 159 ADP, 162, 163,283 Adrenaline, 424 Adrenaredoxin, 307 Adrenodoxin, 309, 374 Adriamycin, 165 Airborne particles, 420 (L-Alah,82 L-Ala-L-Ala,82 Ala-Gly, 82 Albumin, 209 Alcohol dehydrogenase, 124, 127, 128,374 Aldehyde, 36, 137 Alkylation, 156, 157
Alkylmercury, 160 Allosteric effect, 349 AII-rrans-retinal, 272, 273, 275-277 Alpha-cystalline, 397 Alpha (a)-helix, 70, 72-75, 79, 80-83, 85, 98, 99,107,108,176,179,180,184,403,405 Amide, 79, 80, 137 Amide I band, 68-70, 73, 74, 76-83, 85, 97, 98, 103,108,177, 182,200,214,225,403 Amide II band, 68, 71 Amide III band, 40, 68, 70, 71, 73, 74, 76-83, 85, 86, 97, 98, 103, 108, 182, 225, 403 Amide IV band, 71, 72 Amide V band, 71, 72 Amide VI band, 71, 72 Amide VII band, 71, 72 Amide A band, 67 Amide B band, 67 4-Amidino-4'-dimethylaminoazobenzene, 127 Amine, 311 Amine oxidase, 370 D--Amino acid, 81, 82 Aminochromes, 425 2-Amino-2-deoxY-D--glucose, 240 4-Amino-3-nitro-rrans-cinnamic acid, 121 AMP, 143, 152, 159, 160, 166, 167 Amphetamine, 425 Amphipathic molecule, 188 Amphotericin B, 217, 218 Amplitude, definition, 4, 13 Amylose, 250, 251 Anisotropy, 34, 434 Antibiotic-lipid interaction, 213-218 Antibiotics, 69, 165,425 "'27
438
Index
Antibonding, 327 Antiferromagnetism, 370 Antifreeze glycoprotein, 108 Antiparallel-{3-sheet (ant iparallel-pleated sheet), 73, 75, 82,107,129 Anti-Stokes, 6-14, 55, 56, 434 Anti-Stokes Raman scattering: definition, 6, 7 origin, 8-12 Antisymmetrical molecular-scattering tensor, 347 ApA, 167 Apamin,83 ApG,136 ApU, 150 Aquocobalamin, 388 Arginine, 209 Argininylvasopressin, 93 Aroclor, 426 Aromatic hydrocarbons, 419 Ascorbate oxidase, 370, 373, 374 Aspartic acid, 300 Assignment of vibrations. rule, 27 Astaxanthin, 261 Asymmetrical ring vibrational mode, 235 Asymmetrical stretching vi brat ion, 21, 25, 27, 29,31,32,198,199,210,245,247,320 ATP, 161-163,222,223,270,283,284.40 1,405 ATPase, 223 Axial-ligand, 332-339, 341-343 Azo group, 38, 121-123, 126, 127, 130 Azurin, 373-375 Bacteria, 424, 427 Bacterial photosynthetic system, 385-386 Bacteriophage, 180-182 Bacteriophosphyrin, 387 Bacteriorhodopsin, 283-293 Barbiturates, 425 Barium (Ba), 159, 376 Base stacking, 142, 150, 155 Base vibrations, 135-138 Bathorhodopsin (prelumirhodopsin), 272, 274-277, 280 Baygon,426 Bonding vibrations, 20, 26, 27, 30, 32, 204, 274-276 Benzamphetamine, 425 Benzene, 23, 27, 33, 107, 202 S-Benzyl-Cys- Pro- Leu-Gly NH" 77 Beta (f3)-sheet, 70, 72-76, 78-83, 85, 86, 98, 99, 107, 176, 179, 180, 182,212,214, 222, 403. See also Antiparallel-{3-sheet Beta (f3)-turn (3 10 helix), 72, 76-79, 85, 176 Bilayer, 188, 189,202,208,211,213,214,217 Binuclear-cluster-type Fe-S linkage, 306 Biological membranes, 187-225 Bioluminescent bacterium, 57 Bleomycin, 164
Blood, 108, 259, 422 Blood coagulation, 108 Bonding, 327 Bound water, 405 Breathing vibration. 32, 33, 90, 100, 101 cyclic compound, 32 Breathing vibration, proteins, 100. 101 Bromide, 213, 332 B-type cytochrome, 318 Buried tyrosine, 87-89 Cadmium (Cd), 126, 159 Calcium (Ca), 158, 159, 161,219,223,376,377, 399,404,405 Calcium carbonate stones, 416 Calcium oxalate, 416 Calcium phosphate stones, 416 Calculi, 416 Carbohydrate, 129,234-253 Carbohydrate-lipid interaction, 251, 252 Carbon dioxide, 8, 9, 21, 23-26 Carbonic anhydrase, 85, 107, 125-127 Carbon monoxide ligand, 334, 335 Carbonmonoxyhemoglobin (HbCo), 57, 335, 350 Carbon tetrachloride (CCI,). 39, 40 Carbonyl (C=O) stretching vibration, 36, 38, 57,68,70,79,96,97,136,137,151-153, 159,160,175,17"5,206,208,215,246, 247,277,324,326,356,384 Carbonyl stretching vibration, carbohydrate, 246-248 Carbonyl group: carbohydrates, 245, 246, 248 stretching vibration, nucleic acids, 136, 139 vibration, 36, 68,137,245,246,311,312, 198, 199 Carboxyhemoglobin, 334 Carboxypeptidase A, 121, 122 Cardiolipin, 210 {3-Carotene, 219, 223, 256-260, 387 Carotenoiq, 223, 224, 256-262, 381, 386, 387, 419,423 in carrot-root cell, 419 in membranes, 223, 224 in photosynthesis, 386, 387 Carrot, 258, 419 CARS (coherent anti-Stokes Raman spectroscopy), 41, 54-58, 257, 265, 334 flavins, 265 Cataracts, 396-398 Catecholamines, 424 C-C stretching vibration lipids, 192-196, 201-204,207,210,213,216, 219, 221,22~ CDEC (2-chloroallydithiocarbamate), 426 Cerebronic acid, 205 Cerebroside, 205, 206 Ceruloplasmin, 370, 372-375 Cesium (Cs), 376
Index C-H vibration, 27, 31, 32, 36, 234, 244, 245, 248, 249, 274, 285, carbohydrates, 244, 245, 248, 249 lipids, 194-199,201,203-207,210,213,214, 218,219,221,224,225 methane, 34 methylene group, 27, 31 methyl group, 31, 32, 274-276, 285, 286 protein, 65, 81, 96 Channel protein, 189 Chemical modification, proteins, 106, 107 Chitin, 246, 251 Chlordane, 426 Chloride, 213, 332, 334 Chlorinated-hydrocarbon pesticides, 426 Chloroperoxidase, 332 [I-O-Chlorophenyl-)-I-(P-chlorophenyl)2,2,2-trichloroethane],426 Chlorophyll, 382-385, 423 Chlorophyll a, 382-384 Chlorophyll a-protein complex, 385 Chlorophyll b, 382-384 Cholesterol, 208, 215, 217, 218, 221 Chondroitinsulfatc, 240, 241, 247, 251 Chromatin, 135, 182 Chromium (Cr), 158,302, 330 Chromosome, 182-185 Chymotrypsin, 70, 86, 100, 10 I, 103, 120, 125 Chymotrypsinogen, 71, 75, 76 CIPC (chloropham), 426 Circular dichroism (CD), 65,66,75,81,84,86, 300 Cis amide, 79, 80 Cis isomer, 199,200 Classical mechanics of Raman scattering, 12-14 Clinical analysis, 424, 425 Clinical application, 422 Clinical diagnosis, 422 CMP, 142, 166 Cobalt (Co), 158, 159, 161,302,309,321,323, 327, 348, 354, 376, 377, 389, 427 Cobalt corrinoid, 389 Coherent light. 46, 47 Collagen, 398, 399 Combination band, 39, 67 Compton effect, 434 Computer, 50-53 Computer subtraction, 290 Conformation: carbohydrates, 249-251 nucleic acid, 145-153 protein, 73-78 Contractile state of muscle, 405 Co(II)-carbonic anhydrase, 376 Co(II)-carboxypeptidase, 376 Co(II)- imidaZOle, 376 Co(II)-mesoporphyrin IX, 348 Co(II)-octaethylporphyrins, 321
439
Co(JI)-porphyrin, 354 Copper, 158, 159,161,162,213,209,302, 321-323" 325, 326, 348, 369-376, 418 in biological systems, 369-376 Copper (Cu)-N bond vibration, 371-375 Copper (Cu)-S bond vibration, 372-375 Copper-D-penicillamine, 374 Copper protein, 372 Core expansion theory, 353, 354 Cornea, 398, 399 Corrinoid, 389 CoO stretching vibration, 334, 335 CPG, 150 Cryoglobulin, 130 CoS stretching vibration, 38, 93-96, 104, 322, 425 cytochrome c, 322 C-type cytochrome, 318 Cubane-type Fe-S linkage, 307 Cu-mesotetraphenylporphyrin, 348 Cu-octamethylporphyrin, 348 Cu-peptide complex, 375, 376 CuS, 373, 418 Cu(II)-mesoporphyrin IX, 348 Cu(II)-octaethylporphyrin, 321 Cu(II)-sulfur complexes, 373 Cu(II)-transferrin, 375 Cu(JI)-uroporphyrin I, 322 Cyanide (CN), 326, 334, 388 Cyanocobalamin, 388 Cyclic compounds, 32, 33,240, 249, 250, 252, 376 Cyclic oligoamylose, 252 Cycloheptaamylose, 252 Cyclohexaamylose, 240, 249, 250 Cyclo(L-Pro-Gly)" 376 Cysteinato ligand, 311 Cysteine, 96, 97, 372 Cystine, 94-96, 309 Cytidine, 143, 159, 160 Cytochrome a, 318, 324, 331, 347 Cytochrome a oxidase, 356 Cytochrome aJ, 318, 324, 326, 342, 343, 347, 356 Cytochrome aJ oxidase, 356 Cytochrome b, 317, 344, 355 Cytochrome b" 319, 344, 346 Cytochrome b ,6l , 356 Cytochrome b"2, 333 Cytochrome b"" 356, 357 Cytochrome c, 53, 54, 57, 59, 317, 319, 322, 323,331,333,337,340,342-345,348, 352, 355, 357 Cytochrome c', 332, 333, 343 Cytochrome C1, 344, 355, 356 Cytochrome CJ, 333, 344 Cytochrome em, 333 Cytochrome Cm, 323 Cytochrome Cm, 323
440
Index
Cytochrome cd, 59 Cytochrome c oxidase, 210, 345, 346, 356 Cytochrome c peroxidase, 331, 344, 346 Cytochrome!,333 Cytocrhome oxidase, 318, 344, 345, 347, 356 Cytochrome P-450, 330, 331, 340, 342, 344 Cytosine, 142, 147, 148, 154, 155, 175 Dark-adapted bacteriorhodopsin, 58, 283, 292" 293 DDT,426 O,P' DDT, 426 Degenerate, 24, 205 Degenerate bending, 24 Degree of antisymmetry, 35 Degree of freedom, 12, 25, 27 Delhaye molecular microprobe, 411 Denaturation, proteins, 103-106 Deoxyhemoglobin, 334, 339 Depolarization, 346, 349 Depolarization ratio, definition, 33-36 Dermatan sulfate. 247 Deuterium exchange, 38, 70, 71, 79, 80, 85, 97,101,107,135,136,140,141,152, 197,198,202,236-240,246,252,277, 278, 289 Deuteroheme, 323 Dextransulfate, 247 Diamine oxidase, 377 Diatomic molecule, 21, 22 2,4- Dichl oro benzylt ri butyl Ph os p honi u m chloride, 426 cis-Dichlorodiamine platinum, /60 Dichlorvos, 426 Dicofol, 426 Dieldrin, 426 0- 0- Diethy1- 0-(2,4-d ic hloro phe nyl)phosphorothioate, 426 Dila uroylphosphatidylethanola mine (D LPE), 203-205 4- Di met hyla mi no-3-n it ro( cr-benza mid 0 cinnamate), 118, 119 Dimyristoylphosphatidylcholine (D M PC), 198,202,203,210,212,213,218,224 Dinactin, 69, 215 Dioxygenase, 303, 305, 374 Dipalmitoylethanolamine, 202 Dipalmitoylphosphatidic acid, 203 DipalmitoylphosphatidyJcholine (DPPC), 194, 198,201,203,205,206,208,210, 211,212,218,220 Di pal mi toylph os p ha tid ylet ha nola mi ne (DPPE),219 Diphenhydramine, 425 Dipiocolinate, 427 Dipole moment, 14, 15, 20, 22, 23-26, 28-30 Distamycin A, 164, /65 Disteraoylphosphatidylcholine (DS PC), 202 Disulfide (S-S) stretching vibration, 38, 66, 83, 91-94,96,104-106,176,179, 180,310, 373,397,398,418
DNA, 57,145-149,155,157,160,163-166, 180-185,412 Dopamine-{3-hydroxylase, 370 Dopier effect, 434 Diamagnetism, 330 Drugs, 128, 129, 163-165, 252, 425 interaction with nucleic acids, 163-165 Dual-beam method, 279 Dye laser, 48 Egg white protein, 93 Elbow-bending, 102 Electric dipole transition moments, 17 Electric field, 4, 12 Electric vector, 3, 4 Electromagnetic molecular electronic resonance, 103 Electromagnetic wave, 3-5 Electron cloud, 8, 9, 21-30 Electronic time gate, 41 Emulsion, 188, 189 Enamel,418 Endoplasmic reticulum, 204, 205 Endosulfan, 419, 426 Endrin, 426 Enhl'drina schis/osa toxin, 71 Environmental applications, 419, 420, 422, 425-427 Enzyme-drug interactio"l' 128, 129 Enzyme-inhibitor complexes, 125-128, 305 Enzyme, 117-129 Ephedrine, 425 EPIC (ethyl-N-dipropyl-3chlorophenylcarbamate),426 Epinephrine, 424 EPR, 303, 331 Equatorial C-H vibration, 235, 239, 240, 241 Erythrocyte, 36, 210, 212, 219, 220, 222, 223 Erythrocyte ghost, 36, 220, 222, 223 ESR, 207, 209, 222 Ester, 36, 68,137, 190 Ethidium ~romide, 165 Ethion,426 Ethylene, 23, 27 Etiporphyrins, 325, 326, 337, 341 Excitation profile, 47,262, 322 carotenoids, 262 Excitation wavelength (frequency), 18,40,41 Exposed tyrosine, 87-89 Eye, 270-272, 281, 395, 398 Fatty acids, 188, 190-/92, 198, 199,207,208 Fe-N bond, 334, 337-340, 343, 352 Fe-O stretching, 299, 30 I, 304, 312, 341, 335-339 Ferbam, 426 Fermi resonance, 87, 196 Ferredoxin, 306-308, 374 Fe-S, 306-310 Fibrin, 108 Fibrinogen, 108,209
Index Fifth ligand, 333, 342, 352, 353 Fish guanophore, 418 Flavin, 57, 262-266 Flavinadeninedinucleotide (F A D), 57, 262-265 Flavinmononucleotide (FMN), 57, 262-265 Flavocytochrome c, 266 F1avocytochrome Cm, 265, 266 Flavodoxin, 264 Flavoenzyme, 262-266 Florescence, 40, 57. 65, 103,203, 303 Fluorescence spectroscopy, 65 Fluoride (F-), 334, 399 Food additives, 426-427 Foreign bodies of silicone rubber within multinucleated giant cells, 415 Formyl group, 324-326 Free energy of cooperation, 350 Frequency, definition, 4 Frequency difference, 10 Frequency doubler, 18 Fundamental vibrations, 20, 24, 25, 27, 67, 196,200 Furadan.426 {3-(2-Furyl)acryloylphosphate, 124 Galactosamine, 245 Galactose, 206, 212 Gamma (I')-turn, 72, 76, 78 Gauche conformation, 91, 92, 93, 95,192-196, 205,206,208,209,211-213,219,220 Gauche form, methionine, 95 Gauche-gauche-gauche conformation, 91-93 Gauche-gauche-trans conformation, 91-93 Gel, 75,152,153,192,193,197,202,205,208, 211,215,218 Gcl phase, 191-193,201,202,205,207,215 Geometry of the heme ring, 352-354 Glucagon, 83, 210 Glucosamine, 235-239, 245 Glucose, 235-240 Glucose oxidase, 264 Glucose sulfate, 247 Glucuronic acid, 235-239, 246 Glutamic acid, 79, 300 Glutathione peroxidase, 377 Glyceraldehyde-3-phosphate dehydrogenase, 124 Glycerol, 188, 202, 203 Glycophorin,212 Glycoprotein, 108,212 Glycosidic linkage, 235, 241, 249 Glycosphingolipid, 212 Glycylglycine complexes with Cu(II), 375 Gly-Gly,82 Gly-Tyr,87 GMP, 135, 143, 153, 156, 159, 160, 166 GpA,136 OpC, 150 OpG,153 Oramicidin A, 214, 215
441
Oramidicin D, 216 Gramidicin S, 77, 216 Griseofulvin, 425 Group frequency, 36-38 Guanine(G), 142, 148, 151, 153-155, 159, 160, 175,415,418 Guthion, 426 Hapten, 129, 130 Hard metal, 158 Harmonic oscillation, 19 Heat treatment, nucleic acids, 141 Hematoporphyrin, 321 Heme, 266, 316-357 structure of, 324 Heme 0, 344, 345, 347 Heme 03,345 Heme IX, 344 Hemerythrin, 299-301, 374 Hemeundecapeptide, 317 Hemocyanin, 370, 371, 374 oxygen in, 370, 371 Hemoglobin, 57, 330, 334-342, 344, 346, 349-352, 354 Heparin, 247, 25 I Heparin sulfate, 247 Heptachlor, 426 Hexacoordinated complex, 353 Hexacoordination, 342 Hexopyranoside, 241 Hinge-bending, 102 Histidine, 91,107,209,300,311,333 Histone, 70, 182-185 Hooke's law, 19 Horseradish peroxidase, 344, 346 Hyaluronic acid, 240,241,242,246 Hydralation, 84, 101 Hydrocarbon backbone, 201-203 Hydrochloric acid (HCI), 21,434 Hydrogen bond, 38, 79, 87,153,212,214,219, 242, 252, 273 nucleic acids, 150, 151, 153, 154 protein, 79, 81, 87, 106 tyrosine hydroxyl group, 87 Hydroxo ligand, 311 1- Hydro xy(2,4-d ini trop henylazo)-2,5naphthalenedisulfonic acid, 130 Hydroxy, (-OH) group, 36, 96 Hydroxylapatite, 399, 400 IgA,130 IgO, 86,101,129,130 IgM,130 Imidazole, 91 Immunoglobulins, 102, 129, 130 IMP, 142 Inaugural lecture, Raman, 431 Indole-ring vibration, 89, 90 Induced-dipole moment, 8, 12, 14, 15,21 Inosine, 142 Inositol hexaohosohate, 351
442
Index
In-phase stretching vibration, 33 In-plane vibration, CO" 24-26 methyl group, 31 Insulin, 73, 83, 104, 108,210 Intensity of Raman lines, 10-12 Intermolecular vibrations, 101, 102 Internal vibrations, 100, 101 Inverse polarization, 347 Iodide 252, 332 Iodine, 213, 252 Iodine-starch complex, 252 lon-lipid interaction, 213 IPC (isopropyl-N-phenylcarbamate, prophamJ, 426 I R (infrared), 4, 5, 15, 16,21-28,33,36,42,43, 45,50,58,71,78,81,82,97,99,159, 185, 190, 222, 234, 242, 255, 334, 426 Iron (Fe), 158,298-312,327-332,338,339, 341-343, 346, 352, 377 Iron (Fe) ligand band, 337-343, 352 Iron-sulfur proteins, 306-310 Isoalloxazine, 263 Isoleucine, 89 Isomers, 326 Isorhodopsin, 274-277 Isotopes, 38-40 Isotopic substitution, 38, 40 Isozymes, 128, 129
(n
Jet-stream technique, 280 Kerasin, 205 Keratin, 79 Keratin sulfate, 247 Keto carbonyl group, 386 Ketone, 36, 137 Kidney stones, 415 Laccase, 373-375 Lactoferrin, 302, 304 ,a-Lactoglobulin, 74 Lactose, 240 Lamellar liquid, 207 Lapemis hardwickii toxin, 70, 71 Laser, 9, 18, 40, 44-49, 97, 334 Laser tubes, 18,47-49,51 Lateral interaction, 207 Lateral packing, 197, 199,208 Laricauda semifasciara neurotoxin, 103 Lattice vibrations, 102, 191,252 Lauric acid, 191, 192 Lead (Pb), 159 Leghemoglobin, 336, 343, 354 Lens, 396, 398 Leucine, 89 Ligand, 332, 371, 372 Ligand-sensitive Raman bands, 333 Light meromyosin, 403 Lignoceric acid, 205
Lindant, 426 Linkage in bacteriorhodopsin, 288-293 Linkage in rhodopsin, 277-279 Lipid, 187-225 Lipid-antibiotic interaction, 213-218 Lipid-carbohydrate interaction, 251, 252 Lipid-ion interaction, 213 Lipid-lipid interaction, 207, 208 Lipid-protein interaction, 209-213, 221 Liposomes, 188, 189,211-214 Lippert's equation, 85 Liquid crystalline, 192, 193, 197,201,202,205, 207, 208,211,215,218 Lobster shell, 261 Longitudinal acoustic vibrations, 101, 191 Low-frequency vibration, 99-102,191,192, 194, 235, 340, 341 lipids, 190- J 92, 194, 320 proteins, 99-102 Lumazine protein, 57 Luciferin, 419 Lumazine protein, 264, 265 Luminavin, 266 Lumirhodopsin,272 Lutein, 260 Lycopene, 257, 258 Lymph node, 414-416 Lymphocytes, 225, 226 Lyophilization, 83, 84 Lysine, 130, 209, 288, 333 Lysozyme, 52, 78, 81, 83, 86, 93, 101, 103, 104,212 Magnesium (Mg), 144, 156, 158, 159, 161-163, 213, 224, 341, 376, 377, 382, 384, 399 Magnetic field, 4 Magnetic vector, 3,4 Malathion, 426 Maltose, 240, 250 Maltotriose, 250 Mammary carcinoma, 422 Mammary tissus:s, 423 Maneb,426 Manganese (Mn), 158, 159,302,321,377 Maser, 45 Maximum number of vibrations, 21, 27 Melittin, 210, 211 Melting temperature (T m ), 154,191,192,197, 201-203, 205, 206 Melting, tRNA, 154 Membrane, 187-225 Membrane potential, 260, 262 Membrane proteins, 222-224 Mercury (Hg), 158-161,416 Mercury lamp, 45 Mesoheme, 323 Mesotetra-(a,a,a,a-pivaloylamidophenyl) porphyrin, 337, 342 Mesotetraoxyhemoglobin, 337
Index
Meso-tet ra phenylporphinato -bis(tetrahydrofurna)-Fe(II),354 Messenger RNA (mRNA), 155 Metabolic time process, 427 Metal ion, interaction with nucleic acids, 158-163 Metarhodopsin, 272, 281 Methamphetamine, 425 Methemerythrin, 300 Methemoglobin, 343, 351 Methionine, 66, 94-96, 333, 373 Methotrexate, 128 Methoxychlor, 426 Methylene vibration, 86, 204, 210 Methyl group (CH,) vibrations, 31,32, 101, 199,204, 205, 214, 222, 274, 276, 382 Methylene group, vibrations, 27, 31, 199 7- Methylguanosine, 156, 157 Methylelaidate, 200 Methyllaurate, 200 Methyllinoelaidate, 200 Methyllinoleate, 200 Methyllinolenate, 200 Methyloleate, 200 Methylparathion, 426 Meting, nucleic acid, 147-150, 154 Metmyoglobin, 343 Mevinphos, 426 Micelle, 188, 189 Microbial cells, 423, 424 Microprobe, 413 Microwave, 45, 424 Midtransition temperature (Tm), 191, 192 Mineralization in kidneys, 416 Mitochondria, 209, 354-356 Mixed vibration, 70 Mojave toxin, 83 MOLE (molecular optical laser examiner), 411,414,418,420 clinical and histological applications, 414-419 Molybdenum (Mo), 312, 377 Monactin,69 Monochromator, 48, 50 Monolayer, 188, 189 Monooxygenase, 303 Monostearin, 251, 252 Multichannel analyzer, 53, 398, 412 Multilayer, 188, 189,207 Multiple wavelength detectors, 53 Mu (Il)-oxobridged ligand, 311 MUSCle, 80, 395, 401-405 whole, 405 Mu (Il)-sulfide bridge, 300 Mutual exclusion, principle of, 23 Myelin sheath, 212 Myoglobin, 321, 330, 336, 338-341, 344, 354 Myoglobin-carbon monoxide complex (MbCO), 334, 335
443
Myosin, 401, 402, 405 Myosin subfragments, 403, 404 Myotoxin a, 70 NAD,137 NADPH, 128, 167 Neoprontosil, 126 Nerves, 224, 225 Netropsin, 164 Neurotoxins, 103-105 New yellow enzyme, 265 N-H stretching vibration, 38, 67, 68, 70, 245 N-H vibration, carbohydrates, 245 Nickel (Ni), 159, 319-321, 348 Nicotine, 426 Ni-mesotetraphenylporphyrin, 348 Ni-octaethylporphyrin, 320 Ni-octamethylporphyrin, 348 Nitrate analysis, 427 Nitrobenzene group, 118 Nitrogenase, 312, 374 Nitro group vibration, 97 Nitrogen molecular (N2), 21 Nitrosylhemoglobin (HbNO), 351 Ni(Il)-octaethylporphyrin, 321 Ni(1I)-mesoporphyrin IX, 348 NMR, 202, 222, 352 Nobel lecture, 434 Nobel Prize, 434 Nonactin, 215, 216 Noncoherent light, 48 Nonheme iron compounds, 298-312 Non-histone proteins, 183, 184 Non-totally symmetrical vibrations, 35, 36 Noradrenaline, 424 Norepinephrine, 424 Normal coordinate, 20 Nucleic acid, 59, 103, 134-167 Nucleoprotein, 174-185 Nucleosome, 182-185 Nystatin, 218 Occular lenses, 395 Octaethylporphyrin, 319, 330, 338, 339 Octahedral, 342 O-H vibration, 20, 242-244, 248 carbohydrates, 242-244, 248 definition, 20 Old yellow enzyme, 265 Omega-helix, 78 O,P' DDT, 426 Opsin, 272, 282, 283 Optical maser, 45 Optical microscope using Raman effect, 411 Order parameter, 207 Out-of-plane vibration: CO 2 , 24, 27 methyl group, 31 Ovalbumin, 76, 107
444
Index
Overtone, 67, 196 Ovorubin, 262 Ovotransferrin, 302, 304 Oxalate stones, 416 Oxidation state, 343-346 Oxidation-state marker band, 344 Oxidative phosphorylation uncouplers, 222 Oxygenase, 303-306 Oxygenation, 335-337 Oxygen (0,), bound to hemerythrin, 299, 300 Oxygen (0-0) stretching vibration, 299, 300, 321, 335-33~ 339, 370-372 Oxyhemocyanin, 300 Oxyhemoglobin (HbO,), 300, 335, 336, 339, 351 Oxymyoglobin, 321 Oxytocin, 70, 71, 77 Oxytocin agonist, 83 Oxytocin antagonist, 83 Oxytyrosinase, 372 Paleontology, 420 Papain, 117-120 Paradium (Pd), 348 Paramagnetism, 327 Parathion, 426 Pd-mesotetraphenylporphyrin, 348 Pd-octamethylporphyrin, 348 Pelamis platurus neurotoxin, 105 [I -Penicillamine,2-leucine]-oxytocin, 92 [I-Penicillamine]-oxytocin, 78, 92 Pentacoordinated complex, 352, 353 Pentacoordination,342 Peptide backbone, 66, 70, 103, 104,397 Peptide bond vibration, 66-72, 75 Pepulate, 426 Perchlorate (ClO~), 213 Permanent electric dipole moment, 21 Peroxidase, 123, 322, 331, 338, 339, 343 Peroxide, 300 Peroxo ligand, 311 Perthane, 426 Pesticides, 419, 426 Phage, 180, 181,211 Phage fd, 181,211 Phage MS2, 180 Phage, P22, 180 Phage PfI, 181 Phage RI?, 180 Phendimetrazine, 425 Phenolate, 311 Phentermine, 425 Phenylalanine, 90, 350 Phenylamide, 426 2-Phenylethane-boronic acid, 125 Phosphate, 128, 139 Phosphatidylcholine (PC), 204, 205, 208, 215 Phosphatidylethanolamine (PE), 197, 204, 205
Phosphodiester-bond stretching vibrations, 141-143,145,147,151,154,155, 157-159,161,176,178 Phosphoionic bond vibrations, 144, 145, 155, 158,162,178,195,196,212,213,219, 220 Phospholipid, 187-225 Phosphorus, 399 Phosphorus-containing pesticides, 426 Photochemical cycle, 272, 284 Photomultiplier, 41,48, 50 Photoreduction, 345, 351 Photosynthetic pigments, 382-387 Phrenosine, 205, 206 pH treatment, nucleic acids, 141 Phycobilins, 381 Phycocyanin, 26 J, 387 Pi (7T)-helix, 72 Planck's constant, 17, 19 Plant photosynthetic system, 384-385 Plant viruses, 175-180 Plastocyanin, 370, 373, 374 Platelet, 260 Platinum (Pt), 159, 160,321 Polarizability, 8, 12, 13, 14, 21-30, 34 isotropic, 34 Polarizability derivative
(~Q)' 21-26, 28-30
Polarizability ellipsoid, 21-3') Polarizability tensor, 17 Polarization, 346-349 Polarizer, 33-35 Poly(A), 135, 138-140, 142, 143, 150, 152, 15~ 158, 167 Poly(A,G) • poly(U), lSI Poly(A) • poly(C), 145 Poly(A). poly(U), 147, 150 Poly(A-U). poly(A-U), 150 Poly(,B-benzyl-L-Asp), 69, 78 Poly(C). 138-140, 142,143,151,152,158 Poly d (A-T), 157 Poly(DL-Ala), III Poly(G), 138, 139, 144 Poly(G) • poly(C), 144 Poly(Y-benzyl-L-Glu), 69, 73, 82 Poly(Gly), 69, 73, 74, 99 Poly(I), 138, 139, 142, 151, 152 Polyiodide, 252 Poly(L-Ala), 69, 73-75, 80, 81, 100 Poly( L-Ala-Gly), 74, 75 PolY(L-Arg), 158 Poly(LGlu), 69, 73, 80, 81, 97, 98 PolY(L-His),69 POlY(L-Leu), 69, 80 PolY(L-Lys), 69, 73, 74, 80, 86, 97, 98, 158, 211 PolY(L-Met), 80 POlY(L-Pro),68 Poly(L-Tyr, L-Lys), 375
Index Poly(L-Val), 74, 82, 83, 99 Polypeptides, interaction with nucleic acids, 157, 158 Polyphenoloxidase, 370 Poly(U), 136, 138-140, 142, 144, 152, 167 Population inversion, 45-47 Porphin, 348, 349 Porphyrins, 316 Potassium (K), 216, 376 Praseodymium (Pr), 159 Pre melting, 202 Preresonance Raman spectroscopy, 16,97,98, 165-167 Principle of mutual exclusion, 23 Probe laser, 55 Proflavin, 266 Proinsulin, 108 Pro-Leu-GlyNH 2 ,77-80 Protein, 65-108 effect on water, 84 Protein environment, hemeproteins, 352 Protein-lipid interaction, 209-213, 221 Protocatechuate, 374 Protocatechuate 3,4-dioxygenase, 128, 303-305 Protocatechuate 3,4-dioxygenase tyrosine, 377 Protoheme, 323 Protonated Schiff base, 277-281, 288-293 Pseudo-Raman, 103 Pulsed laser, 41, 53 Pumping, 45, 46 Pump laser, 55 Purine compounds, 415 Purine-ring vibrations, 136 Putidaredoxin, 307 Pyrazine, 23, 27 Pyrimidine dimer, 147 Pyrimidine-portion vibrations, 136, 137 Pyrocatechase, 304-306 Pyroglutamyl residue, 79, 80 Pyrole, 23, 27 Pyruvate carboxylase, 377 Qualitative estimation, phospholipid conformation, 206, 207 Quantitative estimation, protein structures, 84-86 Quaternary structure, 332, 349, 351, 352 Ramachandran angles, 66, 70 Raman, C. V., 430 first paper, 434 last pa per, 434 Raman band assignment, 382-384 Raman circular intensity differential (Raman ClD), 58, 59 Raman difference spectroscopy (ROS), 53-55 Raman dispersion, 47, 48, 318
445
Raman hyperchromicity (Raman hyperchromism), 15, 16, 152, 175 Raman hypochromism, 15, 16 Raman microprobe, 411, 413 Raman microscope, 41 I Raman optical activity (ROA), 58, 59 Raman-scattering tensor elements, 17 Raman spectra: background, 103 bacteriorhodopsin, 58, 291 benzene, 16 blood plasma, 259 ca rbon tetrachloride, 39 carrot (live root and canned juice), tomato fruit, {3-caroteine and Iycopene, 258 copper-etioporphyrin isomers, 325 cyanomethyl-2-aceta mide-2-deoxy-l-thio {3-D-glucopyranoside, 248 cytochrome c (ROS spectrum), 55 cytochrome c (resonance Raman), 345, 348 dipalmitoylphosphatidylcholine (0 M PC), 194 ON A (type A and type B), 146 erythrocyte ghost, 220, 221 fatty acid (low frequency region), 192 glucose, 243 hydroxylapatite, 400 lens, 397 lysozyme, 52, 104 mammary tissue (normal and cancerous), 423 a-methyl-D-glucoside and {3-anomer, 244 mitochondria (whole), 355 mole spectra: foreign body in lymph node, 416 crystals in tissues, 418 myosine and whole muscle, 405 oxyhemoglobin, 336, 341 papain-substrate complex for resonance Raman, 119 all-trans-retinal-n-butyla mine hydrochloride, 278 rhodopsin, 279 sea snake neurotoxin, 105 of solid and aqueous samples, proteins, 83, 84 sulfhydryl group of sea snake neurotoxin, 96 sulfhydryl group of TMV proteins, 179 teeth,400 tobacco mosaic virus, 178 its coat protein, 178 its RNA, 179 tyrosine bands, 87 {3-uridine-5'-phosphoric acid, 166 Raman spectra background, 103 Raman spectrometer, 48, 50-53 Random coil, 73, 74, 78-81, 85, 103, 179, 180 Ratio of anti-Stokes to Stokes lines, 10-12 Rayleigh scattering, 6, 7, 9, 13, 191
446
Index
Reaction center, 260, 385, 386 Redox potential, 356 Reduced mass, 38 Relative-intensity parameter, 211, 215 Renal lithiasis, 415 Resonance Raman spectroscopy: bacterial cells, 422, 423 bacterial rhodopsin, 283-293 carotenoids, 256-262, 419 CARS, 54-58 chlorophylls, 381-387 copper and other metal compounds, 369-377 definition, 16-19 drug analysis, 424, 425 enzymes, 117-128 flavins, 262-266 food additives, 425-426 hapten, 129, 130 hemes and porphyrins, 316-357 laser for, 47 membranes, 219 nonheme iron compounds, 298-312 nucleic acids, 165-167 rhodopsin, 270-282 sulfhydryl group, 97 theory, 16-19 visual pigments, 270-282 vitamin B 12 , 387-389 Retina, 270, 271 Retinal, 272, 275-277 II-cis-Retinal, 272, 273, 275, 277 13-cis-Retinal, 274-277 all-trans-Retinal,272-276 R form, heme proteins, 343, 350 Rhodopsin, 270-283 Ribonuclease, 75, 76, 81, 83, 86, 87, 90, 93,157 Ribose, 138, 139 Ribose-phosphate vibrations, 139 Ribose-ring vibrations, 136 Ribosome, 153, 154, 155, 156 RNA, 144, 145,147,153,155,156,165,166, 175-180 Rocking vibration, 31, 204, 275, 285, 286 Rotational quantum level, 5 R state, heme proteins, 335, 349, 351, 354 Rubidium (Rb), 376 Rubredoxin, 308, 309, 374 Ruthenium (Ru), 376 Ruthenium red, 376, 377 Sarcoplasmic reticulum membranes, 223, 224 Scaling constant, 85 Scattering: definition, 6-14 elastic, 6, 7 inelastic, 6, 7 Schiff base, 277-279, 288-293 Scissoring vibration, 204, 205 Sea snake neurotoxin, 85, 87, 89, 90, 96, 97, 103-106 Secondary IinkaJ!e, bacteriorhodopsin, 288
Selenium (Se), 374, 377 Selenocystine, 374 Serum transferrin, 304 Side chain, 86-97, 104-106,322-325 proteins, 86-97 Signal averaging, 41, 53 Signal-to-noise ratio, 41, 53 Silver (Ag), 158, 159, 264 Simian virus 40 (SV40), 225 Sixth ligand, 333, 342, 352, 353 Skew-skew conformation, 20 I Smoothing, 52 Snake neurotoxin, 83, 85, 87, 89,93,96,97, 103-106 Sodium (Na), 158 Soft metal, 158 Sonication, 203 Soret band, 317, 346 Spheroidene, 260 Sphingomyelin, 203-205 Sphingosine, 206 Spider, 417 Spider cuticle, 415 Spin, 309, 327, 330, 331, 334, 346, 357 Spin labelling, 207, 218 Spin-sensitive Raman bands, 326, 331 Spin state, 326-332 Square pyramidal, 342 Starch, 249-252 Starch-iodine complex, 252 Steady-state concentration, 285 Stearic acid, 192, 205 Stellacyanin, 372-374 Stimulated-emission process, 45, 46 Stokes effect (Stokes Raman scattering), 6-14, 56 Streptococcus faecalis membrane, 224 Streptomyces subtilisin inhibitor, 93 Subunit interactions, 101, 102 Succinate-cytochrome c reductase, 318 Sulfates, 213, 245, 247 Sulfhydryl groufJ (-SH), 38, 66, 96, 97, 106, 176, 177, 179,234,263,397,398 Sulfonamide, 425 Sulfonamide inhibitor, 125, 126 4-Sulfonamido-4'-aminoazobenzene, 126 4-Sulfonamido-4'-dimethylaminoazobenzene, 126 Sulfur-containing pesticides, 426 Sulfur dioxide (SO,), 27-30 Superoxide, 300 Superoxide dis mutase, 377 Superoxo ligand, 311 Surface-enhanced Raman spectroscopy (SERS),59 Symmetrical ring vibration, 241 Symmetrical stretching vibration, 21-24, 27, 28,31,32,39,91, 197, 199,203-205, 245, 247,299,312, 320 Symmetry: center of, 23
Index Stokes and anti-Stokes lines, 9, 10 of vibrations, hemes, 319-321 Synthetic polypeptide, 69, 73, 82, 83 Tautomer, histidine. 91 Tautomerism, 135 TDE,426 Teeth, 395, 399, 400 Temperature effect: cryoglobulin, 130 heme, 357 lipids, 192, 197 nucleic acids, 147-149 on Raman intensity, 10-12 Terminal CH" 197, 203 Terradifon, 426 Tetraglycine,( GLY)" 82 Tetrahedron-type Fe-S linkage, 307 Tetraphenylporphyrin, 346 Theory: Raman microprobe, 412-414 Raman spectroscopy, 3-43 Thermotropic transition (T m), 191, 192 Time-resolved resonance Raman spectrocopy, 257 Thiocyanate (CNS-), 213 Thiohexopyranoside, 241 Thiomolybdate, 377 Thiram, 426 Thymidylate synthetase, 123, 124 Thymine, 136, 148, 160, 175 Thymocytes, 225 Thyroid releasing factor, 79, 80 Tobacco mosaic virus (TMV), 97,175-177 Tolbutamide, 252 Toluene, 15, 16 Tomato, 258 Torsional angles (Ramachandran angles), 66, 70 Torsional vibrations, 101 Totally symmetrical vibrations, 35. 36 Total number of vibrations, 20 Trans amide, 79 Trans conformation, 192-195, 199-20 I, 205, 206, 208, 209 Trans~rrin, 302, 303, 375, 377 Transfer RNA (tRNA), 150, 153-156, 167 Trans-gauche-gauche conformation, 92, 93 Trans-gauche·trans conformation, 91-93, 104 Trans isomer, 199, 200 Transition enthalpy, 206 Transition temperature (Tm ), 191, 192, 197, 211,215 Transmembrane potential, 224 Triatomic molecules: linear, 21 nonlinear, 25, 27 Tributyl phosphorotrithioite, 426 Trichinopoly, 430 Triclinic paraffin, 191 Triclinic polyethylene, 191
447
Trielaidin, 200 Trilinoelaidin, 200 Trilinolein, 200 Trilinolenin, 200 Triolein, 200 Tripalmitin, 204 Tropomyosin. 80, 401, 404 Traponin, 401, 404 Trypanocidal drug, 165 Trypsin, 127 Trypsin inhibitor, 86 Tryptophan. 66, 89, 90,97, 106, 107 T state, heme compounds, 335, 349, 350, 351, 354 Tumor, 423 Tunable laser, 18,48 Turnip yellow mosaic virus, 177, 179, 180 Twisting vibration, 31, 200, 204 Type 1 copper, 369, 370 Type 2 copper, 369, 370, 375 Type 3 copper, 369, 370, 375 Tyrosinase, 370 Tyrosine, 66, 83, 87-89, 90, 107, 122, 177, 283, 300, 304-306, 383 UMP, 160 Unsaturated fatly acids, 199·201 Unstacking of bases, 154 UpA,I50 Uracil (U), 142, 151, 154, 155, 175 Uric acid, 415 Uric stones, 416 Uridine, 143, 159 I3-Uaridine-5'-phosphoric acid, 165, 166 Uteroferrin, 303 UTP, 166 Valine, 89, 217 Valinomycin, 69, 198, 217 Vibration, origins. heme, 319-322 Vibrational assignment: amide I, 74, 76, 77, 81, 98 amide III, 73, 74, 76, 77, 81, 86, 98 bacteriorhodopsin, 285-287 carbohydrates, 236-240, 246. 247 chlorophylls, 383 CuN and CuS, 373 FeS, 309, 310 Fe tyrosine, 304 heme oxidation sensitive bands, 344 isomers of etioporphyrins, 325 ligand of hemes, 338, 339, 340 lipids, 198, 199,204,205 low frequency bands of heme, 341 metal ligand, 311 metal ligand bonds of different metalloproteins, 374 nucleic acids, 138-140, 142, 143, 154, 158, 161 0-0 stretching, 300 rhodopsin, 274-277
448
Index
side chain of hemes, 323 tyrosine, 87 Vibrational circular dichroism (V eo), 58, 59 Vibrational modes, hemes, 317 Vibrational quantum level, 5, 8, 17 Vibronic coupling, 319 Vinyl groups, 322-324 Virtual state, 8, 16, 17 Viruses, 174-182 Visual axis, 397, 398 Visual pigment, 270-283 Vitamin A, 272 Vitamin BJl, 57, 387 Wagging vibration, 31, 204. 274
Wavelength, definition, 4 Wave number, definition, 4
Xanthine, 415 Xanthophyll,257 X-ray diffraction data. 46, 70, 73. 75, ' 81-83,85,86,93,108,145,147, 215, 250, 323, 354
Zinc (Zn), 73,122,124,125,127,159 Zincon, 127 Zineb,426 Zn(II)-octaethylporphyrin, 321
